Rotaviruses: Methods & Protocols (Methods in Molecular Medicine)

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M E T H O D S I N M O L E C U L A R M E D I C I N E TM

Rotaviruses Methods and Protocols Edited by

James Gray Ulrich Desselberger

Humana Press

Rotaviruses

METHODS IN MOLECULAR MEDICINE

TM

John M. Walker, SERIES EDITOR 46. Angiogenesis: Reviews and Protocols, edited by J. Clifford Murray, 2000 45. Hepatocellular Carcinoma Methods and Protocols, edited by Nagy A. Habib, 2000 44. Asthma: Mechanisms and Protocols, edited by K. Fan Chung and Ian Adcock, 2000 43. Muscular Dystrophy: Methods and Protocols, edited by Katherine B. Bushby and Louise Anderson, 2000 42. Vaccine Adjuvants: Preparation Methods and Research Protocols, edited by Derek T. O’Hagan, 2000 41. Celiac Disease: Methods and Protocols, edited by Michael N. Marsh, 2000 40. Diagnostic and Therapeutic Antibodies, edited by Andrew J. T. George and Catherine E. Urch, 2000 39. Ovarian Cancer: Methods and Protocols, edited by John M. S. Bartlett, 2000 38. Aging Methods and Protocols, edited by Yvonne A. Barnett and Christopher P. Barnett, 2000 37. Electrically Mediated Delivery of Molecules to Cells, edited by Mark J. Jaroszeski, Richard Heller, and Richard Gilbert, 2000 36. Septic Shock Methods and Protocols, edited by Thomas J. Evans, 2000 35. Gene Therapy of Cancer: Methods and Protocols, edited by Wolfgang Walther and Ulrike Stein, 2000

34. Rotaviruses: Methods and Protocols, edited by James Gray and Ulrich Desselberger, 2000 33. Cytomegalovirus Protocols, edited by John Sinclair, 2000 32. Alzheimer’s Disease: Methods and Protocols, edited by Nigel M. Hooper, 1999 31. Hemostasis and Thrombosis Protocols: Methods in Molecular Medicine, edited by David J. Perry and K. John Pasi, 1999 30. Vascular Disease: Molecular Biology and Gene Therapy Protocols, edited by Andrew H. Baker, 1999 29. DNA Vaccines: Methods and Protocols, edited by Douglas B. Lowrie and Robert Whalen, 1999 28. Cytotoxic Drug Resistance Mechanisms, edited by Robert Brown and Uta Böger-Brown, 1999 27. Clinical Applications of Capillary Electrophoresis, edited by Stephen M. Palfrey, 1999 26. Quantitative PCR Protocols, edited by Bernd Kochanowski and Udo Reischl, 1999 25. Drug Targeting, edited by G. E. Francis and Cristina Delgado, 1999 24. Antiviral Methods and Protocols, edited by Derek Kinchington and Raymond F. Schinazi, 2000 23. Peptidomimetics Protocols, edited by Wieslaw M. Kazmierski, 1999 22. Neurodegeneration Methods and Protocols, edited by Jean Harry and Hugh A. Tilson, 1999

M E T H O D S I N M O L E C U L A R M ED I C I N E

TM

Rotaviruses Methods and Protocols Edited by

James Gray Ulrich Desselberger Clinical Microbiology and Public Health Laboratory Addenbrooke's Hospital Cambridge, UK

Humana Press

Totowa, New Jersey

© 2000 Humana Press Inc. 999 Riverview Drive, Suite 208 Totowa, New Jersey 07512 All rights reserved. No part of this book may be reproduced, stored in a retrieval system, or transmitted in any form or by any means, electronic, mechanical, photocopying, microfilming, recording, or otherwise without written permission from the Publisher. Methods in Molecular Medicine™ is a trademark of The Humana Press Inc. The content and opinions expressed in this book are the sole work of the authors and editors, who have warranted due diligence in the creation and issuance of their work. The publisher, editors, and authors are not responsible for errors or omissions or for any consequences arising from the information or opinions presented in this book and make no warranty, express or implied, with respect to its contents. This publication is printed on acid-free paper. ∞ ANSI Z39.48-1984 (American Standards Institute) Permanence of Paper for Printed Library Materials. Cover design by Patricia F. Cleary. For additional copies, pricing for bulk purchases, and/or information about other Humana titles, contact Humana at the above address or at any of the following numbers: Tel: 973-256-1699; Fax: 973-256-8341; E-mail: [email protected], or visit our Website at www.humanapress.com Photocopy Authorization Policy: Authorization to photocopy items for internal or personal use, or the internal or personal use of specific clients, is granted by Humana Press Inc., provided that the base fee of US $10.00 per copy, plus US $00.25 per page, is paid directly to the Copyright Clearance Center at 222 Rosewood Drive, Danvers, MA 01923. For those organizations that have been granted a photocopy license from the CCC, a separate system of payment has been arranged and is acceptable to Humana Press Inc. The fee code for users of the Transactional Reporting Service is: [0-89603-736-3/00 $10.00 + $00.25]. Printed in the United States of America. 10 9 8 7 6 5 4 3 2 1

Library of Congress Cataloging-in-Publication Data Rotaviruses: methods and protocols / edited by James Gray, Ulrich Desselberger. p. cm. -- (Methods in molecular medicine ; 34) Includes bibliographical references and index. ISBN 0-89603-736-3 (alk. paper) 1. Rotavirus infections Laboratory manuals. 2. Rotaviruses laboratory manuals. I. Gray, James (James J.) II. Desselberger, U. III. Series. [DNLM: 1. Rotavirus Laboratory Manuals. 2. Rotavirus Infections -- Virology Laboratory Manuals. QW 25 R842 2000] QR201.R67R67 2000 616'.0194--DC21 DNLM/DLC for Library of Congress 99-23881 CIP

Preface This is among several volumes in the series Methods in Molecular Medicine that concentrate on a relatively specialized topic: Rotaviruses, one genus within the Reoviridae family. Rotaviruses are the most frequent cause of infantile gastroenteritis worldwide and a significant cause of death, following severe diarrhea and dehydration, in infants and young children of developing countries. Recently, a live attenuated tetravalent rotavirus vaccine has been licensed in the United States, and any widespread use of a rotavirus vaccine will be a further milestone in viral vaccinology. Structure, replication, and various functions of rotaviruses have been thoroughly investigated, and their medical importance clearly justifies and attracts interest to a detailed presentation of the modern methods and approaches used. In organizing this collection we considered it important to strike a balance of presentation among molecular and other modern techniques applied in rotavirus research, accompanied by the relevant background information and review material needed to render this collection attractive to the widest audience. A short introductory chapter (U. Desselberger) sets the scene. The enormous progress made in elucidating the detailed structure of rotaviruses using cryoelectron microscopy and complex computer imaging techniques is presented in the chapter by B. V. Prasad and M. K. Estes. Owing to easy propagation of some rotaviruses in tissue culture and to the application of molecular labeling, blotting, and specialized electrophoretic techniques, details of rotavirus replication, some in common with other Reoviridae and others specific to themselves, have been unraveled and are described in the chapter by J. T. Patton, V. Chizhikov, Z. Taraporelawa, and D. Chen. J. M. Gilbert and H. B. Greenberg contribute some recently developed methods to study the still evolving mechanisms of interaction of viral receptor(s) with the host cell and of viral penetration as the initial steps of viral replication. Because of the segmented nature of their genomes, rotaviruses (like other segmented RNA viruses: reo-, influenza -, bunyaviruses, etc.) have, from the very beginning of their identification, elicited the interest of viral geneticists, and R. F. Ramig’s chapter describes some of the methods used in this context. Rotaviruses have a very wide animal reservoir, and animal models (gnotobiotic piglets, calves, rabbits, mice) have significantly contributed to our understanding of pathogenesis, the immune response, and the study of the

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Preface

most relevant correlates of protection. Three chapters are devoted to these important issues: L. S. Saif and L. A. Ward review pathogenesis models; K. K. Macartney and P. A Offit, the application of immunological techniques; and M. A. Franco and H. B. Greenberg, the application of mouse genetics to the study and recognition of the significance of different branches of the immune response for protection. Properly controlled animal models have also been crucial for studying and dissecting the immune responses to rotavirus vaccine candidates of various kinds, and M. Ciarlet and M. E. Conner provide a comprehensive overview of the methods applied in this context in smaller animal models. We considered inclusion of a chapter on methods of human rotavirus vaccinology, but abstained since it would lead readers too far away from the framework of this book series. In both humans and various mammals, rotaviruses exhibit a high degree of diversity of cocirculating strains, and reliable methods for detection, typing (by serological and, increasingly, molecular techniques), and phylogenetic grouping (based on genomic nucleotide sequence information) are prerequisite to any understanding of the detailed epidemiology and also to carrying out implementation studies on widely used vaccines. M. Iturriza Gómara, J. Green, and J. J. Gray review and describe the techniques applied for this purpose. Some of the epidemiological tools used in rotavirus surveillance are discussed by D. Brown and M. Ramsay. In a final chapter U. Desselberger and M. Estes have attempted to identify topics of future research and have come up with a number of relevant items, hoping that readers may be stimulated. The editors have made an effort to produce a standard layout in all chapters, to convey better the application of the methods. Introductory remarks are followed by sections on Materials needed and the Methods proper; added Notes often reflect personal experience of the authors with the methods conveyed and are worth reading and considering; and Reference lists were intended to be up-to-date. The editors wish to convey their sincere thanks to all contributors for providing their chapters in time and in smooth interaction. The Publishers have been understanding and very helpful, and we wish to thank Tom Lanigan, Craig Adams, Fran Lipton, and John Morgan. We are confident that the contributions speak for themselves and hope that readers will have some gain from and enjoy reading them.

J. J. Gray U. Desselberger

Contents Preface ............................................................................................................ v Contributors .................................................................................................... ix 1 Rotaviruses: Basic Facts ......................................................................... 1 Ulrich Desselberger 2 Electron Cryomicroscopy and Computer Image Processing Techniques: Use in Structure–Function Studies of Rotavirus ........... 9 B. V. Venkataram Prasad and Mary K. Estes 3 Virus Replication .................................................................................... 33 John T. Patton, Vladimir Chizhikov, Zenobia Taraporewala, and Dayue Chen 4 Rotavirus Entry into Tissue Culture Cells ............................................. 67 Joanna M. Gilbert and Harry B. Greenberg 5 Mixed Infections with Rotaviruses: Protocols for Reassortment, Complementation, and Other Assays ............................................... 79 Robert F. Ramig 6 Pathogenesis and Animal Models ....................................................... 101 Linda J. Saif and Lucy A. Ward 7 Immunologic Methods and Correlates of Protection .......................... 119 Kristine K. Macartney and Paul A. Offit 8 In Vivo Study of Immunity to Rotaviruses: Selected Methods in Mice ............................................................... 133 Manuel A. Franco and Harry B. Greenberg 9 Evaluation of Rotavirus Vaccines in Small Animal Models ................ Max Ciarlet and Margaret E. Conner 10 Methods of Rotavirus Detection, Sero- and Genotyping, Sequencing, and Phylogenetic Analysis ........................................ Miren Iturriza Gómara, Jon Green, and Jim Gray 11 Epidemiology of Group A Rotaviruses: Surveillance and Burden of Disease Studies ................................. Mary Ramsay and David Brown 12 Future Rotavirus Research ................................................................. Ulrich Desselberger and Mary K. Estes Index ............................................................................................................

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147

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217 239 259

CONTRIBUTORS

DAVID BROWN • Enteric and Respiratory Virus Laboratory, Central Public Health Laboratory, Public Health Laboratory Service, London, UK DAYUE CHEN • Laboratory of Infectious Diseases, National Institutes of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD VLADIMIR CHIZHIKOV • Laboratory of Infectious Diseases, National Institutes of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD MAX CIARLET • Division of Molecular Virology, Baylor College of Medicine, Houston, TX MARGARET E. CONNER • Division of Molecular Virology, Baylor College of Medicine; Veterans Affairs Medical Center, Houston, TX ULRICH DESSELBERGER • Clinical Microbiology and Public Health Laboratory, Addenbrooke’s Hospital, Cambridge, UK MARY K. ESTES • Division of Molecular Virology, Baylor College of Medicine, Houston, TX MANUEL A. FRANCO • Stanford University School of Medicine, Stanford, CA JOANNA M. GILBERT • Stanford University School of Medicine, Stanford CA JIM GRAY • Clinical Microbiology and Public Health Laboratory, Addenbrooke’s Hospital, Cambridge, UK JON GREEN • Enteric and Respiratory Virus Laboratory, Central Public Health Laboratory, Public Health Laboratory Service, London, UK HARRY B. GREENBERG • Stanford University School of Medicine, Stanford CA MIREN ITURRIZA GÓMARA • Clinical Microbiology and Public Health Laboratory, Addenbrooke’s Hospital, Cambridge, UK KRISTINE K. MACARTNEY • Pediatric Infectious Diseases, Childrens’ Hospital of Philadelphia, Philadelphia, PA PAUL A. OFFIT • Pediatric Infectious Diseases, Childrens’ Hospital of Philadelphia, Philadelphia, PA JOHN T. PATTON • Laboratory of Infectious Diseases, National Institutes of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD

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Contributors

B. V. VENKATARAM PRASAD • Verna and Marrs Department of Biochemistry and W. M. Keck Center for Computational Biology, Baylor College of Medicine, Houston, TX ROBERT F. RAMIG • Division of Molecular Virology, Baylor College of Medicine, Houston TX MARY RAMSAY • Public Health Laboratory Service, Communicable Disease Surveillance Centre, London, UK LINDA J. SAIF • Food and Animal Health Research Program, Department of Veterinary Preventive Medicine, Ohio Agricultural Research and Development Center, The Ohio State University, Wooster, OH ZENOBIA TARAPOREWALA • Laboratory of Infectious Diseases, National Institutes of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD LUCY A. WARD • Department of Veterinary Preventive Medicine, Ohio Agricultural Research and Development Center, The Ohio State University, Wooster, OH

Rotaviruses: Basic Facts

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1 Rotaviruses: Basic Facts Ulrich Desselberger 1. Introduction Rotaviruses (RVs) are the chief etiologic agent of viral gastroenteritis in infants and young children, and in the young of a large variety of animal species. Since the discovery of RVs in man 25 yr ago, much has been learned about their genome and protein composition; their three-dimensional structure; their replication, pathogenesis and clinical pattern; the host’s immune response; and the epidemiology. Measures of individual treatment have recently been complemented by the licensure in the United States of a tetravalent (TV), live attenuated rhesus rotaviruses (RRV)-based, human reassortant vaccine which may to be universally applied. The brief introductory description mostly follows recent reviews (1–4) in which more special references can be found. 2. The RV Genome The genome of RVs consists of 11 segments of double-stranded RNA (dsRNA) with conserved 5' and 3' ends, ranging from 667 bp (segment 11) to 3302 bp (segment 1) in size (SA11 simian RV strain), and totaling 6120 kDa, or 18,555 bp. 3. Gene-Protein Assignment This is complete for several RV strains, and is shown for the SA11 strain in Table 1. With the exception of two genes (RNA 9 and 11), all genes are monocistronic, and the untranslated 5' and 3' regions are very small. 4. RV Proteins There are six structural viral proteins (VPs: termed VP1, VP2, VP3, VP4, VP6, and VP7) and five nonstructural proteins (termed NSP1–NSP5). The From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ

1

2

Size (bp)

1 2 3 4

3302 2690 2591 2362

VP1 VP2 VP3 VP4

5

1611

6 7a

1356 1104

8a

1059

9a

1062

10

751

11

667

Posttranslational modification

125.0 102.7 88.0 86.7

– Myristylation – Proteolytic cleavage (VP5* + VP8*)

VP5 (NS53) VP6 VP8 (NS35) VP9 (NS34) VP7 (1) VP7 (2)

58.6

–

44.8 36.7

Myristylation –

34.6

–

VP10 (NS29) VP11 (NS26)

20.3

Cleavage of signal sequence; glycosylation Glycosylation

21.7

Phosphorylation

37.4 33.9

Location and function Inner core protein; RNA polymerase Inner core protein; RNA binding; leucine zipper Inner core protein; guanylyl transferase; methalyse Surface protein (dimer) Hemagglutinin Neutralization antigen (serotype specific) Fusogenic protein Virulence Pathogenicity Nonstructural (?) Zinc fingers; assembly Inner capsid protein (trimer); group and subgroup antigen Nonstructural; RNA replication? Nonstructural; RNA binding Surface glycoprotein Neutralization antigen (serotype specific); Ca2+ binding site? Nonstructural; intracellular receptor; morphogenesis; enterotoxin Nonstructural

aGene protein coding assignment for SA11 RV strain; assignment different in other strains. Adapted with permission from ref. 4.

Desselberger

RNA segment

Protein product Deduced mol Designation wt (kDa)

2

Table 1 Gene-Protein Assignments, Protein Location, and Function of Group A RVs (SA11 RV/strain)

Rotaviruses: Basic Facts

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functions of all proteins are summarized in Table 1 and reviewed below (as far as known): 1. 2. 3. 4.

VP2 provides a scaffolding function of the inner core (see Subheading 5.). VP6 carries group- and subgroup-specifying determinants. VP7 and VP4 both carry neutralizable, and thus type-specifying, epitopes. VP7 is a glycoprotein of 326 amino acids (aa) (a second in-frame initiation codon lies 30 codons downstream), with nine variable regions contributing to type specificity at varying degrees (see Subheading 6.). 5. VP4 is a nonglycosylated protein of 776 aa, and has a large number of functions: It is the viral hemagglutinin; it is posttranslationally cleaved (in aa positions 241 and 247, the latter being the preferred cleavage site) into the larger VP5* and the smaller VP8* subunits, and cleavage of VP4 enhances infectivity; it is the determinant for protease-enhanced plaque formation and growth restriction; it interacts with the cellular receptor; it has a fusion domain (of still unclear function); it is a virulence determinant. 6. The NSPs have various functions in replication (see Subheading 7.). NSP4 has been found to act as a viral enterotoxin.

5. RV Particle Structure The particle is of icosahedral symmetry, measures 75 nm in diameter, and consists of three layers: 1. The core layer, formed by VP2, and containing the viral genome and the proteins VP1 (the RNA-dependent RNA polymerase) and VP3 (a guanylyltransferase and methylase) (these proteins may have other enzyme functions); 2. The inner capsid (intermediate layer), consisting of 260 VP6 trimers, which are interrupted by 132 aqueous channels of three different kinds in relation to the capsid’s symmetry; 3. The outer capsid (third layer), consisting of 260 VP7 trimers and 60 spike-like VP4 dimers. VP4 interacts with VP7 and VP6.

6. Virus Classification According to VP6 reactivities, there are at least seven different groups (groups A–E are confirmed by complete crossreactivities; groups F and G are likely to be new groups). Within group A, subgroups I, II, I + II, and non-I, non-II are distinguished (according to reactivities of VP6 with two monoclonal antibodies). Because there are two neutralizable outer capsid proteins (VP4 and VP7), a dual classification system has emerged (1), similar to the dual classification established for influenzaviruses (distinguishing different hemagglutinins and neuraminidases): 1. There are at least 14 different VP7-specific types, termed G-types (derived from glycoprotein);

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Desselberger 2. There are at least 19 different VP4-specific types, termed P-types (derived from protease-sensitive protein).

All G- and P-types can be unambiguously distinguished by sequencing of the relevant genes (genotypes). All G-genotypes have been characterized as serotypes; however, this is not the case for all P-genotypes. Therefore, the following nomenclature has been agreed upon: Each virus has a P-type, indicated by an order number for the serotype and by an order number in square brackets for the genotype, and a G-serotype, indicated by an order number (coinciding with genotype number). Thus, the human Wa strain is defined as P1A[8]G1 (P-serotype 1A, P-genotype 8; G-sero/genotype 1); the equine RV L338 is P[18]G13 (P-genotype 18, P-serotype not determined; G-sero/genotype 13); and so on. Because VP4 and VP7 are coded for by different RNA segments (RNA4, and RNA7–9, respectively), various combinations of G- and P-types can be observed in vivo and in vitro, both in man and in animals (4). 7. RV Replication The major features of RV replication are shown in Fig. 1, and can be described as a sequence of the following steps (all replication takes place in the cytoplasm): 1. Adsorption to cellular receptor(s) and receptor-mediated endocytosis, or direct penetration. 2. Messenger RNA (mRNA) production in the cytoplasm from single-shelled (bilayered) subviral particles. 3. mRNA translation to synthesize six structural and five nonstructural proteins (see Subheading 4.). 4. Assembly of single-shelled particles containing VP2, VP1, VP3, and VP6, and a full complement of 11 single-stranded RNAs (ssRNAs); involvement of NSP2 and NSP5; formation of dsRNA (replication) within particle precursors (no free dsRNA or negative ssRNA in cytoplasm); formation of aggregates of bilayered particles (pseudocrystals termed “viroplasm”); 5. Particle maturation to double-shelled (triple-layered) particles: a. Glycosylation of VP7 in rough endoplasmic reticulum (RER), NSP4 acting as an intracelluar receptor for bilayered particles. b. Transiently enveloped particles in RER containing VP4 and VP7. c. Envelope removal. 6. Liberation of double-shelled (triple-layered) infectious particles (virions) by cell lysis.

For further details, see ref. 1; for questions to be investigated further, see Chapter 12. Double infection of cells with two different RV strains leads to simultaneous replication of the genes and synthesis of proteins of both viruses, and, at

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Fig. 1. Major features of the RV replication cycle. Adapted with permission from ref. 1. the stage of assembly, the formation of reassortants of various gene segment combinations; those also include the emergence of different G/P reassortants (see Subheading 6.). 8. Pathogenesis and Animal Models RVs infect the apical cells of the villi of the small intestine, causing cell death and desquamation. At the zenith of the disease, up to 1011 virus particles/mL stool have been counted, concomitant with infection of all susceptible cells in a very short period. The necrosis of the apical villi reduces digestion, causing diarrhea because of primary maladsorption, and leads to villous atrophy (Fig. 2). This is compensated for by a reactive crypt cell hyperplasia accompanied by increased secretion, which also contributes to the diarrhea. Recovery is by replacement of villous epithelium by enteroblasts ascending from crypts. The disease process takes 5–7 d (5). The viral factors determining pathogenicity of RVs have been investigated in several animal models. The product of RNA segment 4, VP4 (Table 1), is likely to be a major determinant, but products of other structural genes (RNA3 coding for VP3; RNAs 8 or 9 coding VP7) and of some of the nonstructural genes (RNA5 coding for NSP1; RNA 8 coding for NSP2, and RNA 10 coding for NSP4) have also

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Fig. 2. Mid-small intestine of gnotobiotic calves. (A) Healthy control animal. (B) animal inoculated experimentally with bovine RV. The normally extended finger-like villi (A) have become stunted and misshapen (atrophy), and the enterocytes of the upper part of the villi are disarranged and swollen. Adapted with permission from ref. 5. been associated with pathogenicity (for review, see ref. 6). Host factors, such as age and host restriction of viral replication, are involved as well in determining pathogenicity, but are less clearly defined. 9. Clinical Symptoms and Treatment After a short incubation period of 24–48 h, the onset of illness is sudden, with watery diarrhea, vomiting, and rapid dehydration. Untreated RV infection is a major cause of infantile death in developing countries. It should be noted, though, that clinical symptoms after RV infection vary widely, and asymptomatic infections in neonates with so-called “nursery” strains have been described. Treatment is by oral or parenteral rehydration with oral rehydration solution (ORS) formulae, which have been approved by the World Health Organization for worldwide treatment in developing countries, and by some drugs (7). 10. Diagnosis Because of the high number of virus particles in feces during the acute illness, diagnosis is easy, using electron microscopy, passive particle agglutination tests, or enzyme-linked immunosorbent assays. 11. Immune Response and Correlates of Protection Acute RV infection is followed by a virus-specific, humoral immune response comprising immunoglobulin (Ig)M, IgG, and IgA antibodies, and by

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a cell-mediated immune response of RV-specific cytotoxic T-cells in the lamina propria of gut tissue. After prolonged research and discussions, it has become increasingly clear that RV-specific, local secretory IgA antibodies (copro-IgA) represent the best correlate of protection (8–10). 12. Epidemiology RVs are the main etiologic agents of serious diarrheal disease in infants and young children under 2 yr of age throughout the world. For developing countries, approx 125 million cases of RV infection occur annually in children under 5 yr of age, of which 18 million are moderately severe to severe; almost 900,000 children die annually from RV infections in these countries. For the USA it is estimated that RV infections cause an estimated 1 million cases of severe diarrhea and approx 150 deaths per annum. RVs are transmitted mostly by the fecal–oral route. A high degree of resistance to physical inactivation, the large number of virus particles shed, and the very low diarrhea dose 50% ensure that infection is also easily taken up from environmental sources, as demonstrated by tenacious nosocomial infections once a clinical ward has been contaminated. Animals infected by various RV types may act as a reservoir for human RV infections. The epidemiology of RVs in complex. Group A RVs are the major cause of human infections. Outbreaks with a strict seasonal winter pattern occur in temperate climates, in tropical regions infections are spread more evenly throughout the year. At any one time and site, there is cocirculation of RVs of different G- and P-types. Viruses of multiple different G/P-type combinations have been isolated. However, G1–G4 viruses represent over 95% of the human strains co-circulating worldwide, and G1 viruses approx 50%. The P/G combinations found are P1A[8]G1, P1B[4]G2, P1A[8]G3, and P1A[8]G4. Within one country, the relative incidence figures for the different types show regional differences, as well as changes over time, and changes in relative incidences of different types are unpredictable. 13. Prevention and Control Since August 1998, a TV, RRV based human reassortant vaccine has been licensed in the United States for universal use, and a decision of licensure for Europe is pending. The vaccine does not prevent infection to a significant degree (i.e., does not produce sterilizing immunity), but has been shown to prevent severe disease with an efficacy of 80%. The vaccine carries G1–G4 epitopes of human RV strains (G1, G2, and G4 on RRV monoreassortants, G3 on RRV). It remains to be seen to what extent the vaccine will produce heterotypic immunity, and whether, upon extensive use of the vaccine, new RV types emerge in man (for review, see refs. 11,12).

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References 1. Estes, M. K. (1996) Rotaviruses and their replication, in Fields Virology, 3rd ed., (Fields, B. N., Knipe, D. M., Howley, P. M., et al., eds.), Lippincott-Raven, Philadelphia, pp. 1625–1655. 2. Kapikian, A. Z. and Chanock, R. M. (1996) Rotaviruses, in Fields Virology, 3rd ed., (Fields, B. N., Knipe, D. M., Howley, P. M., et al., eds.), Lippincott-Raven, Philadelphia, pp. 1657–1708. 3. Iqbal, N. and Shaw, R. D. (1997) Rotaviruses. in Clinical Virology, (Richman, D. D., Whitley, R. J., and Hayden, R. G. eds.), Churchill Livingstone, New York-Edinburgh-London, pp. 765–785. 4. Desselberger, U. (1998) Reoviruses, in Topley and Wilson’s Microbiology and Microbial Infections. 9th ed. vol. 1: Virology, (Mahy, B. W. J. and Collier, L., eds.), E. Arnold, London-Sydney-Auckland, pp. 537–550. 5. Greenberg, H. B., Clark, H. F., and Offit, P. A. (1994) Rotavirus pathology and pathophysiology in Rotaviruses (Ramig, R. F., ed), Springer Verlag, Berlin-Heidelberg, pp. 256–283. 6. Burke, B. and Desselberger, U. (1996) Rotavirus pathogenicity. Virology 218, 299–305. 7. Desselberger, U. (1999) Rotavirus infection: guidelines for treatment and prevention. Drugs 58, 447–452. 8. Offit, P. A. (1994). Rotaviruses: immunological determinants of protection against infection and disease. Adv. Virus. Res. 44, 161–202. 9. Yuan, L. J., Ward, L. A., Rosen, B. I., To, T. L., and Saif, L. J. (1996) Systemic and intestinal antibody secreting cell responses and correlates of protective immunity to human rotaviruses in a gnotobiotic pig model of disease. J. Virol. 70, 3075–3083. 10. Moser, C. A., Cookinham, S., Coffin, S. E., Clark, H. F., and Offit, P. A. (1998) Relative importance of rotavirus-specific effector and memory B cells in protection against challenge. J. Virol. 72, 1108–1114. 11. Vesikari, T. (1997) Rotavirus vaccines against diarrhoeal disease. Lancet 350, 1538–1541. 12. Desselberger, U. (1998) Towards rotavirus vaccines. Rev. Med. Virol. 8, 43–52.

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2 Electron Cryomicroscopy and Computer Image Processing Techniques Use in Structure–Function Studies of Rotavirus B. V. Venkataram Prasad and Mary K. Estes 1. Introduction Rotavirus (RV), a double-stranded (ds)RNA virus in the family Reoviridae, is a complex, relatively large (diameter, including spikes = 1000 Å), nonenveloped icosahedral virus. Once RV was recognized as a major human pathogen, it was extensively studied using modern molecular genetic and biological techniques, as discussed elsewhere in this book. These studies provided basic information about gene-coding assignments, protein processing, genome expression and replication, viral morphogenesis, and pathogenesis (1). In addition, molecular epidemiological studies, coupled with the characterization of neutralizing monoclonal antibodies (MAbs) and sequencing of the genes that encode the neutralizing antigens, provided an understanding at the molecular level of the antigenic and genetic variability of the RVs. Medical relevance, intriguing structural complexity, and several unique strategies in the morphogenesis of RVs have provoked extensive structural studies on these viruses in recent years (2–10). A detailed architectural description of these complex viruses, including the topographical locations of the various structural proteins and their stoichiometric proportions, was obtained as the resolution of these techniques improved. Together with molecular biological studies, structural studies are permitting a dissection of the molecular mechanisms that underlie biological processes of the virus, such as cell entry, neutralization, transcription, gene expression, and virus assembly (8). This chapter reviews current knowledge of RV structure and the methods used in structural analysis. From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ

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2. Computer Image Processing of Electron Cryomicrographs Over the past four decades, X-ray crystallography has been the technique of choice to study three-dimensional (3-D) structures of biological macromolecules and macromolecular assemblies at atomic resolution. Structures of several animal viruses have been studied using this technique (11–12; see ref. 13 for a brief review of the technique). Until now, the largest structure that has been studied using this technique is simian vacuolating virus 40, ~500 Å in diameter (14). However, recently, the structure of bluetongue virus (BTV) cores, ~680 Å in diameter, a virus in another genus of the Reoviridae family, has been determined to near-atomic resolution, using X-ray crystallographic techniques (15). X-ray crystallographic techniques have also been used on individual viral proteins (VPs), such as the hemagglutinin and neuramindase proteins of influenza virus (12), several proteins of HIV (16,17) and, most recently, VP6 of RV (18). Attempts are being made to determine the atomic resolution structure of the entire RV and subassemblies of RV using X-ray crystallographic techniques. Prerequisite to successful structural analysis by X-ray crystallography is the ability to obtain crystals, of the specimen of interest, which diffract X-rays to atomic resolution. In the past decade, advances in electron cryomicroscopy (cryo-EM) and computer image processing techniques have provided another powerful tool for studying the 3-D structures of biological macromolecules and assemblies. This technique does not require the specimen to be in a crystalline state. Another advantage of this technique that it allows structural studies of the specimen not only in its native state, but also under various physiological conditions. For example, this technique has been used to study complexes of antibody-bound virus (3,19–22) and receptor-bound virus (23), transcribing virus (9), and specimens under varied chemical states, such as different pH and ionic strengths (24). All information obtained on structure–function relationships in RVs has come from using cryo-EM and computer image processing techniques. The following sections briefly discuss these techniques, and then describe how they have been useful in dissecting structure–function correlations in RVs.

2.1. Computer Image Processing Because of the large depth of focus of conventional electron microscopes, transmission electron micrographs, in effect, represent two-dimensional (2-D) projections of the specimen. Inference of the detailed 3-D structure by direct examination of electron micrographs is therefore often a difficult task. Over the past two decades, computerized procedures have been developed to reconstruct the 3-D structure of a specimen from such projections. These procedures offer an objective way of extracting 3-D structural information from electron

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micrographs. Similar procedures of reconstruction from projections are used in other contexts, such as diagnostic tomography. To determine the 3-D structure of any object, it is necessary to combine the information content from different views of that object. These different views are sometimes provided by the specimen lying in different orientations, as in the case of icosahedral viruses, or they may be obtained by tilting the specimen in the microscope. 3-D reconstruction from electron micrographs is based on what is known as the projection theorem, which states that the 2-D Fourier transform of a projection of a 3-D object is a central section, normal to the direction of the view, of the 3-D Fourier transform of the object. If all the different orientations of the specimen can be identified with respect to a common frame of reference, the 3-D Fourier transform of the specimen can be built from the 2-D Fourier transforms of different views. Fourier inversion of the 3-D transform thus obtained gives the 3-D structure of the specimen. For icosahedral viruses, Crowther et al. (25–27) have developed image-processing procedures to determine their 3-D structure from micrographs. The first step in the computer processing of electron micrographs is converting the image into digitized data by microdensitometry. Regions in the micrographs with a sufficient number of particles are digitized using a computer-controlled microdensitometer. The digitized regions are put into the computer, where each particle is windowed and centered inside a box. The orientation of each particle is determined in a computer from its Fourier transform using the so-called common lines procedure (26). Orientations of enough particles are determined so that they evenly represent the asymmetric unit of an icosahedron. The number of particles required for a reconstruction depends on the size of the specimen and the resolution sought. Typically, to attain a resolution of 20 Å for RV, 200 particles with unique orientations are used. For resolution closer to 10 Å, the number of particles with unique orientations have to be in the thousands. Particle images with well-determined orientation parameters are then combined in Fourier space using cylindrical expansion methods, to obtain a 3-D Fourier transform (26). An inverse Fourier transform then gives the 3-D structure. Orientation parameters of the particles are refined, either with respect to one another, using the cross-common lines, or with respect to the projections obtained from the initial 3-D map, in an iterative fashion. The visualization and interpretation of the 3-D reconstruction are carried out using computer graphics software. These procedures have been modified and refined in the past few years by several laboratories (27–31). A simplified outline of the image processing scheme is shown in Fig. 1. A detailed description of every aspect of the processing protocol is beyond the scope of this chapter but can be found in ref. 29. Computations of difference maps between closely related

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Fig. 1. A brief schematic representation of the major steps involved in the 3-D reconstruction of the RV structure.

strains (e.g., wild-type and mutant [4]), or viruses in different states (6), allow additional structural information to be obtained.

2.2. Electron Cryomicroscopy In conventional transmission EM one normally uses metal-shadowing or negative-staining techniques to enhance image contrast. These preparative techniques may potentially alter the structure of the specimen, and sometimes may destroy fragile structural features because of chemical modification, dehydration, and desiccation. As a result, these preparative techniques are not suitable, particularly when the goal is to obtain 3-D structural information. However, for diagnostic purposes, these conventional techniques are adequate, and have proven very useful. In 1975, Taylor and Glaeser (32) introduced a method of embedding the specimen in a thin layer of ice, and imaging at low temperatures using a low electron dose. Since then, several laboratories have been involved in the improvement of this technique (for review, see ref. 33). Cryo-preparative techniques not only provide good contrast but also preserve the structural integrity of the specimen. One further advantage of structural analysis using electron images of ice-embedded specimens is the ability to retrieve the details of the internal structure of particles. This contrasts with

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structure determination from images of negatively stained specimens, in which it is possible to obtain only surface features. The image contrast in the absence of stains or metal decoration is caused by the scattering density difference between protein and ice. Cryo-EM involves three principal steps: quick-freezing of the specimen, transfer of the specimen to the microscope, and examination of the specimen in the microscope at low temperature. The first step is carried out using a mechanical guillotine-type plunging device (34). The specimen grid is held by a tweezer in the quick-freezing device. After the excess liquid on the grid is blotted using filter paper, the specimen grid is rapidly plunged into liquid ethane (cryogen) at its melting point. This rapid freezing produces a thin layer of vitreous or amorphous ice in which the specimen is embedded. In the second step, the frozen specimen is initially transferred to a cryospecimen holder maintained at liquid nitrogen temperature (–173°C) in a workstation. Then the cryo-specimen holder is quickly transferred to the electron microscope. The specimen is examined in the microscope at a temperature below –160°C, using low electron doses (typically 5–10 e – /Å 2 on the specimen). Several technical problems associated with this technique have been addressed during the past few years, which have made this a successful technique (for details, see refs. 33,35).

2.3. Recent Developments in Cryo-EM In recent years, there have been exciting developments in the high-resolution electron imaging of biological specimens: for example, the use of computer-controlled spot-scan imaging with medium-high-voltage electrons to reduce beam-induced motion images and to increase the efficiency of recording high resolution (7,36–39); the use of a field emission gun with intermediate-high-voltage electrons to increase the high-resolution image contrast by improving temporal and spatial coherence (40–42), and the use of energy-filtered electrons to remove the background intensity caused by inelastic scattering, and to increase the scattering signal from the specimen (43). Advances in computer image processing schemes have also been made, taking advantage of high-speed, high-performance computers and multiprocessor computers (44). The future looks promising for attaining 10 Å resolution and higher by cryo-EM, for icosahedral particles. The structure of the hepatitis B virus has already been determined to a resolution of ~8 Å, using cryo-EM techniques (41,42).

2.4. Combined Use of X-ray Crystallography and Cryo-EM Techniques An exciting trend in recent years is the marriage of X-ray crystallography and cryo-EM techniques (45). Several studies have shown that these two

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techniques are not mutually exclusive, but can effectively complement one another, particularly in studying large macromolecular assemblies. If the structures of individual components of an assembly are available at atomic resolution, this information can be fitted into the structure of the whole assembly determined by cryo-EM techniques, to enhance understanding of the molecular interactions. The complementarity of these two techniques offers a significant step forward in structural biology, as is shown in structural studies on such viruses as adenovirus (46), flock-house virus (47), rhinovirus (48), and Sindbis virus (49). Recent structural analysis of the BTV core is an excellent example of the combined use of cryo-EM and X-ray crystallography techniques to determine the structure of a complex macromolecular assembly at atomic resolution (50). Similarly, the cryo-EM structure of Norwalk virus has been very useful for determining the structure at atomic resolution of this virus by X-ray crystallographic data (51). Such a combinatorial approach is being used not only on icosahedral viruses, but also in the structural studies of ribosomes (52), which lack any symmetry, and other nonicosahedral complex structures, such as bacteriophage φ29 (53). 3. 3-D Structure of RV 3-D structural studies using cryo-EM techniques have been carried out on two RV strains: simian RV SA11 (2–4) and rhesus RV (5,6). The structural features seen in these two strains of RV are very similar, because both have three concentric layers of protein. A surface representation of a 3-D reconstruction of triple-layered (mature) RV along the icosahedral threefold axes is shown in Fig. 2A. Some of the structural features inside the RV structure are shown in a cross-sectional slice taken from the 3-D reconstruction of RV in Fig. 2B. The structure, which is based on a left-handed T = 13 icosahedral lattice, exhibits distinct structural features, including aqueous channels and surface spikes. The overall diameter of the particle, including the spikes, is 1000 Å.

3.1. Outer Layer Biochemical experiments have shown that the outer shell of the RV contains two proteins, VP7 and VP4 (54,55). The major component is VP7, a glycoprotein. A close examination of surface features (Fig. 2A) indicates that VP7 molecules cluster into triangular-shaped trimers surrounding the aqueous channels, on the T = 13 icosahedral lattice. The VP7 trimers have a small depression at the center, and they are connected to one another on the sides. Whether VP7 molecules aggregate into trimers prior to the assembly of the shell is not known, nor whether they cluster into trimers on their assembly into the icosahedral shell. The T = 13 icosahedral symmetry of the VP7 layer dictates that each virion has 780 molecules (or 260 trimers) of VP7. The overall thickness of the VP7 layer is ~35 Å.

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Fig. 2. (A) 3-D structure of the triple-layered particle (TLP) at a resolution of 23 Å. A set of icosahedral symmetry axes and the locations of the three types of channels are indicated. The surface is a left-handed T = 13 icosahedral lattice. In such a lattice, a fivefold axis is reached from its neighboring fivefold axis by stepping over three six-coordinated positions and taking a left turn. (B) Central cross section extracted from the 3-D map of the virus. The mass density breaks up into three distinct shells between the radii of 210 and 500 Å. These three shells are between the radii 500 and 340 Å, 340 and 270 Å, and 270 and 210 Å. The VP4 spike density is also shown; notice the lower domain of VP4 inside one of the channels. (C) Surface representation of the 3-D structure with anti(VP4) Fab bound. (D) Interaction of VP4 spikes with the VP6 layer. Scale bar = 20 Å.

3.2. Aqueous Channels A distinctive feature of the RV structure is the presence of aqueous channels at all of the five- and six-coordinated positions on the T = 13 icosahedral lattice (see Fig. 2A). These channels penetrate into the outer two layers of the structure and are ~140 Å deep. They have been classified into three types based on their location with respect to the icosahedral symmetry axes. Type I channels run down the icosahedral fivefold axes, type II channels are those on the

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six-coordinated positions surrounding 2the fivefold axes, and type III channels are those on the six-coordinated positions neighboring the icosahedral threefold axes. In each virion, there are 132 channels: 12 type I, 60 type II, and 60 type III channels. Type II and III channels are about 55-Å wide at the outer surface of the virus. The type I channels, in contrast, have a narrower and more circular opening of ~40 Å in diameter. All the channels constrict before widening in the interior, and have their maximum width at the position close to the surface of the inner shell proteins, as seen in the side view of the type III channels in Fig. 2B.

3.3. Surface Spikes Another distinctive feature of the RV structure is surface spikes. From the surface of the particles, 60 spikes extend to a length of 120 Å (Fig. 2A). The RV spikes are located at the outer edge of the type II channels that surround the icosahedral fivefold axes. Each spike has a distinct bilobed structure at the distal end. Each lobe has a diameter of ~25 Å. These lobes are individually connected to rod-shaped densities that are separated from each other by a hole that is ~20 Å in diameter. These rod-shaped densities, with a left-handed helical twist, merge together as they approach the surface of the outer layer, making two points of contact with the vertices of the triangular-shaped VP7 trimers. When the spikes on RV were first discovered, Prasad et al. (2) predicted that these surface projections were made up of VP4, which was confirmed by immunolabeling studies using MAbs against VP4 (3). Two antigen-binding fragments (Fab) molecules bound to the sides of the distal bilobes of each spike (Fig. 2C). The volume of the spike indicates that each spike can accommodate two molecules of VP4. Thus, each virion contains 120 copies of VP4. The observed structural features of individual spikes, as described above, and in vivo radiolabeling studies, further support the idea that the spikes are dimers of VP4 (4). Gel filtration analyses of expressed VP4 also have provided biochemical evidence that VP4 is an oligomer, probably a dimer. Interactions between these oligomers are apparently maintained by hydrophobic interactions, because these are readily disrupted by detergents (56), which explains why dimers of VP4 have not been detected by simple analysis of VP4 in virus disrupted by sodium dodecylsulfate (SDS) and separated by SDS-polyacrylamide gel electrophoresis.

3.4. Internal Domain of VP4 From the 3-D structure of the native virus alone, it is not possible to determine whether there is an inward extension of the VP4 spikes. That is, do the spikes terminate at the surface of the virion, or do they penetrate into the VP7 layer? Based on the volume of VP4 calculated from the 3-D reconstructions of

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the native and the Fab-bound virions, Prasad et al. (3) estimated that about 30 kDa of VP4 is buried inside the virion surface. By computing a difference map between the structures of the native strain and a reassortant strain of RV that lacks spikes, the existence of a large internal domain of VP4 beneath the VP7 surface was confirmed (4). This domain of VP4 is centered inside the type II channel, in close association with the walls of the channel made of trimers of VP6, as shown in Fig. 2D (the structure of the VP6 shell is described in more detail in Subheading 3.5.). Similar results were obtained by difference imaging of the virus structure at normal pH and alkaline pH (6). At alkaline pH, the RV spikes fall off without affecting the integrity of the VP7 layer (6,57). The finding that VP4 interacts extensively with the VP6 layer may have implications in the morphogenesis of the RV particles. Shaw et al. (4) have suggested that VP4 may play an important role, along with the nonstructural protein (NSP4), in facilitating the budding of progeny double-layered particles (DLPs) through the endoplasmic reticulum (ER) membrane, and that assembly of VP4 onto newly made particles occurs prior to VP7 assembly. An interesting aspect of RV infection is trypsin-enhanced infectivity (54,58). Trypsin, present at the natural site of infection, cleaves VP4 at a conserved arginine residue to produce VP8* (28 kDa, aa 1–247) and VP5* (~60 kDa, aa 248–776). Trypsin cleavage of VP4 is accompanied by a significant increase in RV infectivity (54,58), which is associated with enhanced cell entry (59,60). Although the overall structure of the VP4 spike is visualized in the cryo-EM reconstructions, the topographical locations of the proteolytic fragments VP5* and VP8* are not known. The MAb that was used in the structural studies by Prasad et al. (3), 2G4, is a VP5*-specific antibody, which binds to the sides of the distal lobes of the VP4-spike, suggesting that these distal globular domains of VP4 contain some portions of VP5*. The MAb 2G4 is a neutralizing antibody, and has been shown to block virus penetration, but not cell attachment. The exposed region of the distal tip of the spike is probably involved in initial attachment to cells, and the region of the spike binding to 2G4 molecules may be involved in cell penetration.

3.5. The Intermediate Layer Treatment of intact triple-layered virions with chelating agents (e.g., ethylenediamine tetra-acetic acid) removes the outer shell, reduces infectivity by several log steps, and exposes the inner shell proteins. The resulting DLPs are indistinguishable from those produced in infected cells (61). Electron micrographs of DLPs embedded in vitreous ice show that these particles are 705 Å in diameter, with a bristly surface. The 3-D structure of the DLPs (Fig. 3A) has been determined to ~19 Å, using cryo-EM techniques (7). The protein mass is mostly concentrated into 260 morphological units positioned at

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Fig. 3. (A) Surface representation of the 3-D structure of the double-layered particle (DLP) at a resolution of ~19 Å. A set of icosahedral symmetry axes and the locations of the three types of channels are indicated. (B) VP6 computationally isolated from the 3-D map of the DLP. (C) Surface representation of the VP2 layer extracted from the structure of DLP; the threshold used represents 120 molecules of VP2. (D) As viewed at a higher threshold, the boundaries of two molecules of VP2 that make the icosahedral asymmetric unit are denoted as A and B. Scale bar = 200 Å.

all the local and strict threefold axes of the T = 13 icosahedral lattice. The location and shape of the capsomeres strongly suggest a trimeric clustering of the inner capsid protein, VP6. This has been confirmed by separating VPs on nondenaturing conditions (62,63). These 260 capsomeric units are arranged in such a way that there are channels at all five- and six-coordinated centers. These channels are in register with the channels in the VP7 layer. The structure of the VP6 molecule (Fig. 3B) appears to have two domains: the globular upper domain, and a slender lower domain. The upper globular domains of individual monomers interact with one another in stabilizing the trimer. The sides of the lower domain seem to be involved in intercapsomeric interactions, and the bottom part of the lower domain interacts

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with the VP2 layer. VP6 molecules interact with VP7 through their upper domain. Flat upper domains of the VP6 trimers protrude into the VP7 layer, so that the close contact between the two shells occurs primarily around the local and strict threefold axes. The triangular-shaped VP7 trimers spread across the VP6 trimers, and portions of the VP6 trimers are visible through the outer-shell channels (Fig. 2A). It is possible that small molecules or small enzymes may be able to interact with portions of the VP6 layer, even in the presence of the outer VP7 layer. Consistent with this idea, at least one MAb to VP6 has been shown to bind to both triple-layered particles and DLPs (64). Recently, the structure of VP6 has been determined by X-ray crystallographic analysis to 2.8 Å (18). The tertiary structure of VP6 bears several similarities to the structure of VP7 of BTV (65), which is the counterpart of VP6 in RV. Both these proteins consist of two distinct domains: a distal eight-stranded β-barrel domain and a lower α-helical domain. The distal β-barrel domain interacts with VP7; the α-helical domain interacts with VP2. Fitting of the X-ray structure of VP6 into the cryo-EM structure of RV is in progress, to delineate the regions of VP6 that interact with VP7, VP4, and VP2.

3.6. Inner Layer Although early protein and EM analyses defined the composition of the outer two layers of the virion with certainty, the existence and composition of the third shell initially was based on conjecture. The total mass density in the outer shell accounts for 780 and 120 molecules of VP7 and VP4, respectively. The volume of the protein in the intermediate shell accounts for 780 molecules of VP6. On the basis of the radial density profile, and the fact that VP2 is the most abundant of the remaining three structural proteins, it was proposed that the density between the radii of 230 and 270 Å is caused by the shell formed by VP2 molecules (66). The existence of an inner shell was confirmed when single-layered (core-like) particles were produced by expression of VP2 alone (67). 3-D structural analysis of recombinant particles containing VP2 alone (2-virus-like particles [VLPs]) has indicated that the VP2 layer extends from a radius of about 230 Å to 270 Å, in agreement with interpretation based on the radial density profile computed from the 3-D structure of the virion. The structure determination of 2-VLPs to a resolution closer to ~20 Å has been hampered by extensive aggregation of 2-VLPs. However, knowing the radial extension of the VP2 layer, the structural features of the VP2 layer has been deduced from the reconstruction of native RV particles or recombinant particles (68) containing VP2 and VP6, referred to as 2/6-VLPs (10). In contrast to the VP7 and VP6 layers, the VP2 layer is a rather featureless, continuous bed of density (Fig. 3C). Examination of the structure of this layer at a slightly higher

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threshold of protein mass reveals the arrangement of 120 VP2 molecules on the T = 1 icosahedral lattice (Fig. 3D). Each asymmetric unit of the icosahedron consists of two molecules, referred to as type A and type B monomers. The type A monomer lies close to the fivefold axis, the type B monomer originates slightly away from the fivefold axis, and extends toward the threefold axis. The T = 1 icosahedral organization, with 120 molecules, poses an interesting structural question about how this layer, consisting of 120 molecules, interacts with the VP6 layer, consisting of 780 molecules. Such a T = 1 icosahedral structure, with 120 molecules, is unique, because it renders the two molecules in the asymmetric unit quasiequivalent, and has been observed in other dsRNA viruses besides RV. In the Reoviridae family, both aquareovirus (69) and BTV (15) have inner layers composed of 120 subunits on a T = 1 icosahedral lattice. A T = 1 symmetry with dimers is also observed in other dsRNA viruses outside Reoviridae: The fungal viruses L-A of Saccaromyces cerevisiae and P4 of Ustilago maydis exhibit similar organization (70). This unique T = 1 organization may be of fundamental significance in the endogenous transcription of the genome of dsRNA viruses.

3.7. Internal Organization In addition to VP7, VP4, VP6, and VP2 reviewed so far, the RV structure should account for VP1 and VP3, the remaining structural proteins, and the genomic RNA. In the 3-D structure of RV, several internal features are seen. To understand the internal organization of RV and to interpret the internal structural features in terms of minor proteins and the genomic RNA, a twofold strategy was used by Prasad et al. (7). First, difference imaging between native DLPs and various recombinant VLPs was used to identify the internal features. Second, a higher-resolution (~19 Å) structure of the DLP was carried out, to delineate the internal features more clearly.

3.7.1. Locations of VP1 and VP3 Although VP1 and VP3 are present in small amounts, they play an important role in the endogenous transcription process of RV. Several biochemical studies have indicated that VP1 is the RNA-dependent RNA polymerase (71) and VP3 is the guanylyltransferase (72). The structure of 1/2/3/6-VLPs shows flower-shaped features attached to the inside tip of the VP2 at all the fivefold positions (Fig. 4B), which is absent in 2/6-VLPs (Fig. 4A); it is also absent in 1/2/6-VLPs and 3/2/6-VLPs. These studies suggest that the flower-shaped feature becomes discernible only when VP1 and VP3 are both present, and that this structural feature represents a complex of VP1 and VP3. Although these studies strongly suggest the location of VP1 and VP3, the observed shape of the proposed VP1–VP3 is uncertain, because of the reconstruction procedures that

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Fig. 4. (A) Identical cross-section of 2/6-VLP structure. Notice the absence of the flower-shaped structural feature seen in B. (B) Cross-section (85 Å thick) of 1/2/3/6-VLP structure. Flower-shaped structures are seen attached to the inside surface of VP2 at all the icosahedral fivefold axes. (C) Cutaway from the 19-Å structure of the DLP, exposing the mass density caused by the genomic RNA. (D) Dodecahedral shell of the ordered RNA, extracted from the 19-Å structure. Scale bar = 200 Å.

implicitly use icosahedral symmetry. The shape that has been observed can only be real if each virion has 60 molecules of VP1–VP3, or this flower-shaped complex has an internal fivefold symmetry. The volume occupied by each flower-shaped structure, assuming a protein density of 1.30 g/cm3, accounts for about 25% of the expected mass of a complex of VP1 (125 kDa) and VP3 (98 kDa). Biochemical data indicate that there are 12 molecules of VP1 and 12 VP3 molecules per virion (72). The remaining portions of VP1 and VP3 may extend further inside the radius of 160 Å, and are transparent to structural analysis, either because they are disordered or because they move away from the fivefold axes, and lack any semblance of icosahedral symmetry. Thus, the flower-shaped structure may represent the structurally discernible portion of the VP1–VP3 enzyme complex.

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3.7.2. Location of Genomic RNA Comparison of the radial density profiles computed from the structures of DLPs and VLPs clearly indicate that mass density caused by the genomic RNA lies inside the radius of 230 Å (7). At this radius, the DLP structure shows strong mass density surrounding the fivefold axes, and between the neighboring threefold axes (Fig. 4C). This mass density is completely absent in the 2/6-VLP and the 1/3/2/6-VLP structures. The density attributed to ordered RNA in the 23-Å DLP is clearly resolved into parallel strands of tube-like mass density that form a dodecahedral shell in the 19 Å structure of the DLPs, determined using micrographs taken by using a 400 keV electron microscope. These tubes of mass density have an average diameter of 20 Å, typical of a dsRNA double helix (Fig. 4D). These strands encircle the VP1–VP3 complex located at the fivefold axis. Icosahedral packing of RNA does not imply 60-fold repetition in the gene sequence, but indicates that portions of the dsRNA occupy an icosahedrally equivalent volume in the structure. Icosahedrally ordered nucleic acid has been previously observed in small virus structures by X-ray crystallography (73–77). The maximum number of base pairs previously visualized in a virus structure is about 300 (76). In the RV structure, ordered RNA accounts for ~4500 of a total 18,555 bp. The 19 Å-DLP structure shows that there are several points of contact between the inwardly protruding portions of VP2 and the RNA surrounding each fivefold axis (7). These observations are consistent with biochemical results that show VP2 has dsRNA binding activity (78). Thus, VP2, which is icosahedrally assembled,

Fig. 5. (A) 3-D structure of transcribing DLP. (B) Proposed pathway for the nascent mRNA molecule exiting through a type I channel.

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appears to be responsible for inducing icosahedral ordering on closely interacting portions of RNA. Such extensive ordering of dsRNA is also seen in the recently determined X-ray structure of the BTV core (15), which is the equivalent of the RV DLP. The extensive ordering of the genome, described above, is perhaps critical for endogenous transcriptase activity. It may facilitate an orchestrated movement of the genome through the enzyme complex. This motion could be driven by the continuous exit of newly synthesized mRNA. The pores surrounding the fivefold axes, in the VP2 layer, are large enough to permit mRNA to exit, although it is also possible that VP2 may undergo conformational changes during transcription, to facilitate the exit of mRNA. Understanding of this process will be made clear by observing changes in the RNA and the VPs during active transcription, and also under conditions in which the DLP has been rendered transcriptionally incompetent. Though it has not been possible, from any of the current structural studies, to address the question of where the different segments of dsRNA are located inside the structure, it is tempting to speculate that a substantial portion of each of the 11 segments is ordered around a fivefold axis, and that each segment interacts with a specific VP1–VP3 complex.

3.8. Structure of Actively Transcribing DLPs From earlier studies, the authors had postulated that channels in the capsid layers would be used for the import of the precursors necessary for transcription and the exit of nascent mRNA molecules (2). As described in Subheading 3.5., there are 132 channels in the VP6 layer. Which of these channels are used for export of the nascent transcripts? To determine how the mRNA transcripts are translocated through the intact DLPs during transcription, the 3-D structure of actively transcribing DLPs has been determined (9). There are two chief inferences from these studies: First, the DLPs remain structurally intact during transcription; second, the nascent transcripts exit through the type I channels in the VP6 layer (Fig. 5A). The observation that the DLPs remain structurally intact is especially important, considering that the particles are capable of unlimited transcription, provided that the necessary RNA precursors are continuously supplied (79). The structural integrity of the particles is suggested not only from the images and the reconstruction, but also from the observation that the comparison of particle images with projections of the reconstruction by cross-common lines yields phase residual values that are very similar to those observed for nontranscribing DLPs. These results strongly imply that icosahedral symmetry is maintained to the same degree in both structures, at least to the resolution that these structures have been analyzed. Why is the structural integrity of DLP necessary for transcription? Biochemical studies show that no structural protein by itself is capable of carrying

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out transcription, which occurs within the confines of the intact DLPs. Structural and biochemical studies seem to indicate that the VP2 protein may play a scaffolding role in the architecture of RV. The inner surface of the VP2 layer not only provides a structural support for the RNA, but also helps to properly position the transcription complex consisting of VP1 and VP3. Recent biochemical studies have shown that the amino terminus of the VP2 molecule possesses nonspecific single-stranded RNA and dsRNA binding activity, and also that it is essential for the incorporation of VP1 and VP3 (80). Structural analysis of 2/6-VLPs, with an intact and a truncated aminoterminus of VP2, have shown that the amino-terminus is located at the inside surface of the VP2 layer, and close to the fivefold axis (10). The unique arrangement of VP2 molecules on a T = 1 icosahedral lattice, discussed previously (Fig. 3D), may be particularly useful in carrying out the dual roles of providing structural support to the RNA and transcription complex. It is likely that type A monomers of VP2, as they interact in a head-to-head manner at the fivefold axis, are exclusively involved in the interactions with VP1 and VP3, which are anchored to the inner surface of the VP2 capsid layer, along the fivefold axis; the N-terminal portions of type B monomers may provide the necessary interactions with RNA. The outer surface of the VP2 layer provides a structural platform for the assembly of VP6 trimers, preventing aggregation of the core particles (VP2, VP1, VP3, and the genome), which are known to be highly hydrophobic (81). The assembly of VP6 provides well-defined conduits for the exiting RNA molecules. From structural and biochemical studies, it can be hypothesized that the structural integrity of the DLP is necessary for the observed transcriptional efficiency and continuous reinitiation, because of the need to hold the components of the transcription machinery in their proper arrangements throughout repeated cycles of initiation–elongation, as well as to enable the efficient and continuous release of the mRNA transcripts.

3.9. Model for Exit Pathway of mRNA Although structural studies on the actively transcribing particle strongly suggest that the type I channels are used for exporting the nascent mRNA molecules, the precise pathway from the site of synthesis to the exterior of the particle remains to be elucidated. Based on the observed internal organization (7), and the structure of the transcribing particle, a model for the exit pathway in RV has been proposed (9). The RNA synthesis probably occurs within the core of the virion, very near to the fivefold axes, because this is the location of the transcriptional complexes composed of VP1 and VP3. Newly transcribed mRNA then exits the core through the channels in the VP2 layer, which are immediately adjacent to the fivefold axis, and probably in clos-

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est proximity to the site of synthesis. Once through the VP2 layer, the exiting strands of mRNA migrate out of the particle through the type I channels (at the fivefold axes) in the VP6 layer (Fig. 5B). In the electron images of the actively transcribing particles, 3–4 strands of mRNA, associated with the particles, have been observed (9). From these observations, it appears that multiple mRNA transcripts can be released simultaneously from an actively transcribing particle. Each genome segment may be transcribed by a specific polymerase complex, and the resulting transcript may exit the particle through the channel system at the fivefold axis adjacent to its site of synthesis. The mechanism of transcription and mRNA release proposed in these studies might also explain why no dsRNA virus with more than 12 genome segments has ever been found. 3-D structural studies using cryo-EM, similar to those described by Lawton et al. (9) on transcribing RV particles, have been carried out on orthoreoviruses (M. Yeager, personal communication). These studies have confirmed the finding of earlier studies using classical Kleinschmidt techniques, that the mRNA release occurs through the fivefold vertices in these viruses (82). In orthoreoviruses, the transcription complex is also suggested to be anchored to the inner surface of the innermost capsid layer at the fivefold axes (83). The release of mRNA through the fivefold vertices is probably a common characteristic of segmented dsRNA viruses, including BTV (15,84), aquareovirus (69), and the bacteriophage φ6 (85). 4. Conclusions and Future Challenges Structural studies on RVs have helped to provide a foundation for understanding the molecular mechanisms underlying some of the functions of these viruses, and are just the beginning of obtaining a detailed understanding of the structure–function relationships in this complex and large virus. Present and future studies are aimed at answering several questions. Are there specific receptors for RVs and how do these viruses initiate the cell-entry process? How does the transcription take place inside intact DLPs? How does the virus encapsidate a correct set of genome segments? What are the roles of the nonstructural proteins in virus replication and self-assembly, and what molecular interactions regulate their functions? What is the molecular mechanism of budding of the progeny DLPs into the ER membrane? The authors anticipate that a more complete picture of how these viruses replicate will emerge as structural analyses improve in resolution, either by cryo-EM or X-ray crystallography, or combinations thereof, in conjunction with advances in biochemical and molecular biological studies. Obtaining detailed molecular and structural information should also allow the development of more effective strategies to combat, prevent, or treat the clinical outcome of infections with these viruses. Increasing knowledge of the structure and

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function of the RV genes and proteins, and the ability to produce VLPs of various components, are already being exploited to develop interesting vaccine candidates (see Chapters 3 and 9). Acknowledgments The authors’ work is supported in part by grants from the NIH (AI 36040 and DK 31044), the W. M. Keck Foundation and the National Center for Research Resources (RR 02250). We thank J. Lawton and B. Pesavento for useful discussions and their help in making the figures. References 1. Estes, M. K. (1996) Rotaviruses and their replication, in Virology (Fields, B. N., Knipe, D. M., and Howley, P. M., eds.), Lippencott Raven, Philadelphia, pp. 1625–1655. 2. Prasad, B. V. V., Wang, G. J., Clerx, J. P., and Chiu, W. (1988) Three-dimensional structure of rotavirus. J. Mol. Biol. 199, 269–275. 3. Prasad, B. V. V., Burns, J. W., Marietta, E., Estes, M. K., and Chiu, W. (1990) Localization of VP4 neutralization sites in rotavirus by three-dimensional cryo-electron microscopy. Nature 343, 476–479. 4. Shaw, A. L., Rothnagel, R., Chen, D., Ramig, R. F., Chiu, W., and Prasad, B. V. V. (1993) Three-dimensional visualization of the rotavirus hemagglutinin structure. Cell 74, 693–701. 5. Yeager, M., Dryden, K. A., Olson, N. H., Greenberg, H. B., and Baker, T. S. (1990) Three-dimensional structure of rhesus rotavirus by cryoelectron microscopy and image reconstruction. J. Cell Biol. 110, 2133–2144. 6. Yeager, M., Berriman, J. A., Baker, T. S., and Bellamy, A. R. (1994) Three-dimensional structure of the rotavirus haemagglutinin VP4 by cryo- electron microscopy and difference map analysis. EMBO J. 13, 1011–1018. 7. Prasad, B. V. V., Rothnagel, R., Zeng, C. Q., Jakana, J., Lawton, J. A., Chiu, W., and Estes, M. K. (1996) Visualization of ordered genomic RNA and localization of transcriptional complexes in rotavirus. Nature 382, 471–473. 8. Prasad, B. V. V. and Estes, M. K. (1997) Molecular basis of rotavirus replication: structure-function correlations, in Structural Biology of Viruses (Chiu, W., Burnett, R., and Garcia, R., eds.), Oxford University Press, New York and Oxford, pp. 239–268. 9. Lawton, J. A., Estes, M. K., and Prasad, B. V. V. (1997) Three-dimensional visualization of mRNA release from actively transcribing rotavirus particles Nature Struct. Biol. 4, 118–121. 10. Lawton, J. A., Zeng, C. Q., Mukherjee, S. K., Cohen, J., Estes, M. K., and Prasad, B. V. V. (1997) Three-dimensional structural analysis of recombinant rotavirus-like particles with intact and amino-terminal-deleted VP2: implications for the architecture of the VP2 capsid layer. J. Virol. 71, 7353–7360. 11. Rossmann, M. G. and Johnson, J. E. (1989) Icosahedral RNA virus structure. Annu. Rev. Biochem. 58, 533–573.

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12. Harrison, S., Skehel, J. J., and Wiley, D. (1996) Virus structure, in Fields Virology (Fields, B. N., Knipe, D. M. and Howley, P. M., eds.), Lippincott Raven, Philadelphia, pp. 59–99. 13. Baker, T. S. and Johnson, J. E. (1997) Principles of virus structure determination, in Structural Biology of Viruses (Chiu, W., Burnett, R., and Garcia, R., eds.), Oxford University Press, New York and Oxford, pp. 38–79. 14. Liddington, R. C., Yan, Y., Moulai, J., Sahli, R., Benjamin, T. L., and Harrison, S. C. (1991) Structure of simian virus 40 at 3.8-Å resolution. Nature 354, 278–284. 15. Grimes, J. M., Burroughs, J. N., Gouet, P., Diprose, J. M., Malby, R., Zientara, S., Mertens, P. C. P., and Stuart, D. I. (1998) The atomic structure of the bluetongue virus core. Nature 395, 470–478. 16. Hodge, C. N., Straatsma, T. P., McCammon, J. A., and Wlodawer, A. (1997) Rational design of HIV protease inhibitors, in Structural Biology of Viruses (Chiu, W., Burnett, R., and Garcia, R, ed.), Oxford University Press, New York and Oxford, pp. 451–473. 17. Kwong, P. D., Wyatt, R., Robinson, J., Sweet, R. W., Sodroski, J., and Hendrickson, W. A. (1998) Structure of an HIV gp120 envelope glycoprotein in complex with the CD4 receptor and a neutralizing human antibody. Nature 393, 648–659. 18. Petitpas, I., Lepault, J., Vachette, P., Charpilienne, A., Mathieu, M., Kohli, E., Pothier, P., Cohen, J., and Rey, F. A. (1998) Crystallization an preliminary x-ray analysis of rotavirus protein VP6. J. Virol. 72, 7615–7619. 19. Wang, G. J., Porta, C., Chen, Z. G., Baker, T. S., and Johnson, J. E. (1992) Identification of a Fab interaction footprint site on an icosahedral virus by cryoelectron microscopy and X-ray crystallography. Nature 355, 275–278. 20 Smith, T. J., Olson, N. H., Cheng, R. H., Chase, E. S., and Baker, T. S. (1993) Structure of a human rhinovirus-bivalently bound antibody complex: implications for viral neutralization and antibody flexibility. Proc. Natl. Acad. Sci. USA 90, 7015–7018. 21. Smith, T. J., Cheng, R. H., Olson, N. H., Peterson, P., Chase, E., Kuhn, R. J., and Baker, T. S. (1995) Putative receptor binding sites on alphaviruses as visualized by cryoelectron microscopy. Proc. Natl. Acad. Sci. USA 92, 10,648–10,652. 22. Trus, B. L., Newcomb, W. W., Booy, F. P., Brown, J. C., and Steven, A. C. (1992) Distinct monoclonal antibodies separately label the hexons or the pentons of herpes simplex virus capsid. Proc. Natl. Acad. Sci. USA 89, 11,508–11,512. 23. Olson, N. H., Kolatkar, P. R., Oliveira, M. A., Cheng, R. H., Greve, J. M., McClelland, A., Baker, T. S., and Rossmann, M. G. (1993) Structure of a human rhinovirus complexed with its receptor molecule. Proc. Natl. Acad. Sci. USA 90, 507–511. 24. Wang, G. J., Hewlett, M., and Chiu, W. (1991) Structural variation of La Crosse virions under different chemical and physical conditions. Virology 184, 455–459. 25. Crowther, R. A., DeRosier, D. J., and Klug, A. (1970) The reconstruction of a threedimensional structure from projections and its applications to electron microscopy. Proc. Roy. Soc. Lond. A 317, 319–340.

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43. Frank, J., Zhu, J., Penczek, P., Li, Y., Srivastava, S., Verschoor, A., et al. (1995) Model of protein synthesis based on cryo-electron microscopy of the E. coli ribosome. Nature 376, 441–444. 44. Zhou, Z. H., Chiu, W., Haskell, K., Spears, H., Jr., Jakana, J., Rixon, F. J., and Scott, L. R. (1998) Refinement of herpesvirus B-capsid structure on parallel supercomputers. Biophys. J. 74, 576–588. 45. Baker, T. S. and Johnson, J. E. (1996) Low resolution meets high: towards a resolution continuum from cells to atoms. Curr. Opin. Struct. Biol. 6, 585–594. 46. Stewart, P. L., Fuller, S. D., and Burnett, R. M. (1993) Difference imaging of adenovirus: bridging the resolution gap between X-ray crystallography and electron microscopy. EMBO J. 12, 2589–2599. 47. Cheng, R. H., Reddy, V. S., Olson, N. H., Fisher, A. J., Baker, T. S., and Johnson, J. E. (1994) Functional implications of quasi-equivalence in a T = 3 icosahedral animal virus established by cryo-electron microscopy and X-ray crystallography. Structure 2, 271–282. 48. Smith, T. J., Chase, E. S., Schmidt, T. J., Olson, N. H., and Baker, T. S. (1996) Neutralizing antibody to human rhinovirus 14 penetrates the receptor-binding canyon. Nature 383, 350–354. 49. Cheng, R. H., Kuhn, R. J., Olson, N. H., Rossmann, M. G., Choi, H. K., Smith, T. J., and Baker, T. S. (1995) Nucleocapsid and glycoprotein organization in an enveloped virus. Cell 80, 621–630. 50. Grimes, J. M., Jakana, J., Ghosh, M., Basak, A. K., Roy, P., Chiu, W., Stuart, D. I., and Prasad, B. V. (1997) An atomic model of the outer layer of the bluetongue virus core derived from X-ray crystallography and electron cryomicroscopy. Structure 5, 885–893. 51. Prasad, B. V. V., Hardy, M. E., Dokland, Bella, J., M., Rossmann, M. G., and Estes, M. K. (1999) X-ray crystallographic structure of the Norwalk virus capsid. Science, in press. 52. Ban, N., Freeborn, B., Nissen, P., Penczek, P., Grassucci, R. A., Sweet, R., et al. (1998) A 9 Å resolution X-ray crystallographic map of the large ribosomal subunit. Cell 93, 1105–1115. 53. Tao, Y., Olson, N. M., Xu, W., Anderson, D. L., Rossmann, M. G., and Baker, T. S. (1998) Assembly of tailed bacterial virus and its genome release studied in three-dimensions. Cell 95, 431–437. 54. Estes, M. K., Graham, D. Y., and Mason, B. B. (1981) Proteolytic enhancement of rotavirus infectivity: molecular mechanisms. J. Virol. 39, 879–888. 55. Arias, C. F., Lopez, S., and Espejo, R. T. (1982) Gene protein products of SA11 simian rotavirus genome. J. Virol. 41, 42–50. 56. Zhou, Z., Crawford, S., and Estes, M. K. Personal communication. 57. Anthony, I. D., Bullivant, S., Dayal, S., Bellamy, A. R., and Berriman, J. A. (1991) Rotavirus spike structure and polypeptide composition. J. Virol. 65, 4334–4340. 58. Arias, C. F., Romero, P., Alvarez, V., and Lopez, S. (1996) Trypsin activation pathway of rotavirus infectivity. J. Virol. 70, 5832–5839.

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59. Kaljot, K. T., Shaw, R. D., Rubin, D. H., and Greenberg, H. B. (1988) Infectious rotavirus enters cells by direct cell membrane penetration, not by endocytosis. J. Virol. 62, 1136–1144. 60. Keljo, D. J., Kuhn, M., and Smith, A. (1988) Acidification of endosomes is not important for the entry of rotavirus into the cell. J. Pediatr. Gastroenterol. Nutr. 7, 257–263. 61. Cohen, J. and Dobos, P. (1979) Cell free transcription and translation of rotavirus RNA. Biochem. Biophys. Res. Commun. 88, 791–796. 62. Sabara, M., Ready, K. F. M., Frenchick, P. J., and Babiuk, L. A. (1987) Biochemical evidence for the oligomeric arrangement of bovine rotavirus nucleocapsid protein and its possible significance in the immunogenicity of this protein. J. Gen. Virol. 68, 123–133. 63. Shen, S., Burke, B., and Desselberger, U. (1994) Rearrangement of the VP6 gene of a group A rotavirus in combination with a point mutation affecting trimer stability. J. Virol. 68, 1682–1688. 64. Tosser, G., Labbé, M., Bremont, M., and Cohen, J. (1992) Expression of the major capsid protein VP6 of group C rotavirus and synthesis of chimeric single-shelled particles by using recombinant baculoviruses. J. Virol. 66, 5825–5831. 65. Grimes, J., Basak, A. K., Roy, P., and Stuart, D. (1995) The crystal structure of bluetongue virus VP7. Nature 373, 167–170. 66. Prasad, B. V. V. and Chiu, W. (1994) Structure of rotavirus. Curr. Top. Microbiol. Immunol. 185, 9–29. 67. Labbé, M., Charpilienne, A., Crawford, S. E., Estes, M. K., and Cohen, J. (1991) Expression of rotavirus VP2 produces empty corelike particles. J. Virol. 65, 2946–2952. 68. Crawford, S. E., Labbé, M., Cohen, J., Burroughs, M. H., Zhou, Y. J., and Estes, M. K. (1994) Characterization of virus-like particles produced by the expression of rotavirus capsid proteins in insect cells. J. Virol. 68, 5945–5952. 69. Shaw, A. L., Samal, S. K., Subramanian, K., and Prasad, B. V. (1996) Structure of aquareovirus shows how the different geometries of the two layers of the capsid are reconciled to provide symmetrical interactions and stabilization. Structure 4, 957–967. 70. Cheng, R. H., Caston, J. R., Wang, G. J., Gu, F., Smith, T. J., Baker, T. S., et al. (1994) Fungal virus capsids, cytoplasmic compartments for the replication of double-stranded RNA, formed as icosahedral shells of asymmetric Gag dimers. J. Mol. Biol. 244, 255–258. 71. Valenzuela, S., Pizarro, J., Sandino, A. M., Vasquez, M., Fernandez, J., Hernandez, O., Patton, J., and Spencer, E. (1991) Photoaffinity labeling of rotavirus VP1 with 8-azido-ATP: identification of the viral RNA polymerase. J. Virol. 65, 3964–3967. 72. Liu, M., Mattion, N. M., and Estes, M. K. (1992) Rotavirus VP3 expressed in insect cells possesses guanylyltransferase activity. Virology 188, 77–84. 73. Chen, Z. G., Stauffacher, C., Li, Y., Schmidt, T., Bomu, W., Kamer, G., Shanks, M., Lomonossoff, G., and Johnson, J. E. (1989) Protein-RNA interactions in an icosahedral virus at 3.0 A resolution. Science 245, 154–159. 74. Tsao, J., Chapman, M. S., Agbandje, M., Keller, W., Smith, K., Wu, H., Luo, M., Rossmann, M. G., and Compans, R. W. (1991) The three-dimensional structure of canine parvovirus and its functional implications. Science 251, 1456–1464.

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75. McKenna, R., Xia, D., Willingmann, P., Ilag, L. L., Krishnaswamy, S., Rossmann, M. G., Olson, N. H., Baker, T. S., and Incardona, N. L. (1992) Atomic structure of single-stranded DNA bacteriophage phi X174 and its functional implications. Nature 355, 137–143. 76. Fisher, A. J. and Johnson, J. E. (1993) Ordered duplex RNA controls capsid architecture in an icosahedral animal virus. Nature 361, 176–179. 77. Larson, S. B., Koszelak, S., Day, J., Greenwood, A., Dodds, J. A., and McPherson, A. (1993) Double-helical RNA in satellite tobacco mosaic virus. Nature 361, 179–182. 78. Labbé, M., Baudoux, P., Charpilienne, A., Poncet, D., and Cohen, J. (1994) Identification of the nucleic acid binding domain of the rotavirus VP2 protein. J. Gen. Virol. 75, 3423–3430. 79. Cohen, J. (1977) Ribonucleic acid polymerase activity associated with purified calf rotavirus. J. Gen. Virol. 36, 395–402. 80. Zeng, C. Q., Estes, M. K., Charpilienne, A., and Cohen, J. (1998) The N terminus of rotavirus VP2 is necessary for encapsidation of VP1 and VP3. J. Virol. 72, 201–208. 81. Zeng, C. Q., Labbé, M., Cohen, J., Prasad, B. V. V., Chen, D., Ramig, R. F., and Estes, M. K. (1994) Characterization of rotavirus VP2 particles. Virology 201, 55–65. 82. Bartlett, N. M., Gillies, S. C., Bullivant, S., and Bellamy, A. R. (1974) Electron microscopy study of reovirus reaction cores. J. Virol. 14, 315–326. 83. Dryden, K. A., Farsetta, D. L., Wang, G., Keegan, J. M., Fields, B. N., Baker, T. S., and Nibert, M. L. (1998) Internal structures containing transcriptase-related proteins in top component particles of mammalian orthoreovirus. Virology 245, 33–46. 84. Prasad, B. V. V., Yamaguchi, S., and Roy, P. (1992) Three-dimensional structure of single-shelled bluetongue virus. J. Virol. 66, 2135–2142. 85. Gottlieb, P., Strassman, J., Qiao, X. Y., Frucht, A., and Mindich, L. (1990) In vitro replication, packaging, and transcription of the segmented double-stranded RNA genome of bacteriophage phi 6: studies with procapsids assembled from plasmid-encoded proteins. J. Bacteriol. 172, 5774–5782.

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3 Virus Replication John T. Patton, Vladimir Chizhikov, Zenobia Taraporewala, and Dayue Chen 1. Introduction The effort to understand the molecular biology of rotaviruses (RVs) has led to the development of procedures that can be used to study the replication and transcription of the RV genome, the assembly and structure of the rotavirion, and the structure and function of RV proteins. Because it is not possible to provide a detailed description of all the techniques developed, this chapter stresses only those that have broad application, or which represent important new technical advances. In particular, this chapter emphasizes procedures used to prepare large amounts of purified triple- (TLP), double- (DLP), and single-layered (core) RV particles; to synthesize viral RNAs in vitro, through the transcriptase and replicase activities associated with RV particles; to evaluate the RNA-binding activity of RV proteins; and to assemble core-like and virus-like particles (CLPs and VLPs, respectively) via the expression of RV recombinant proteins.

1.1. Growth In Vitro With the discovery two decades ago that trypsin-like proteases enhance the infectivity of RVs (1), cultivation of these viruses to higher titers became possible, and the large amounts of virions necessary for the study of their structural and enzymatic properties could be purified. This Subheading describes protocols routinely used in the authors’ laboratory to propagate and titer RVs and to generate cores and DLPs from triple-layered rotavirions.

1.1.1. Virus Propagation Although many cell lines will support the growth of group A RVs, MA104 cells (fetal monkey kidney cells) are the most commonly used for their From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ

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propagation. MA104 cells can be grown to confluency in either minimum essential medium or Medium 199 containing 5% fetal bovine serum (FBS). However, in this laboratory, higher virus yields are obtained with the latter.

1.1.2. Preparation of Virions Containing Uncleaved Virion Protein VP4 The presence of trypsin in the medium during RV propagation will result in the cleavage of the virion protein VP4 into VP5* (60kDa) and VP8* (27kDa) (2) and, thus, virus purified from trypsin-containing media will contain little if any intact VP4. To obtain purified virus that contains only intact VP4, cells are infected at a high multiplicity of infection (MOI) (3–10 plaque-forming units [PFU]/cell) with trysin-activated virus, washed several times with serumfree medium following absorption, and maintained in trypsin-free medium containing protease inhibitors (see Subheading 3.2.2.).

1.1.3. Purification of Triple-Layered and Double-Layered Virions To generate the large quantities of RV particles that are often necessary for structural and biochemical analyses, the authors’ usually purify virus from 50 150-cm2 flasks of infected cell lysate. The virions are recovered from the cell lysate after treatment with trichlorotrifluoroethane, followed by high-speed centrifugation in cesium chloride (CsCl) to separate triple- and double-layered virions (see Subheading 3.2.3.). Triple-layered virions can be easily converted to DLPs through the removal of the VP4–VP7 outer shell by chelating agents such as ethylenediaminetetraacetic acid (EDTA) or ethylene glycoltetraacetic acid (EGTA) (3,4; see Subheading 3.2.4.).

1.1.4. Preparation of Cores DLPs can be converted to core particles by chaotropic agents such as calcium chloride (CaCl2) or sodium thiocyanate (5,6; see Subheading 3.2.5.).

1.2. Cell-Free Synthesis of Viral RNA In the infected cell, the RV RNA-dependent RNA polymerase, VP1, serves dual functions: as the viral transcriptase with which it catalyzes the synthesis of capped, but nonpolyadenylated messenger (m)RNAs (7); and as the viral replicase with which it catalyzes the synthesis of double- stranded (ds)RNA (8). During transcription, the RNA polymerase uses genomic dsRNA as the template to synthesize mRNA; during RNA replication, the RNA polymerase uses mRNA as the template to synthesize minus-strand RNA, resulting in the formation of dsRNA. The RNA polymerase activity, associated with DLPs and open core particles, allows the in vitro synthesis of viral mRNA and dsRNA, respectively (9,10).

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1.2.1. Transcription by DLPs In the presence of ribonucleotides, the transcriptase of DLPs can use the endogenous dsRNA genome as the template for synthesis of capped and methylated mRNA (9,11). The ability of DLPs to efficiently transcribe the genome is dependent on the presence of the VP6 shell, because removal of this protein from such particles yields core particles that are transcriptionally inactive (5,12). Several lines of evidence suggest that VP3 may be the viral guanylyltransferase, including the observation that, when incubated with guanosine 5'-triphosphate (GTP), the protein will form a covalently linked VP3–guanosine 5'-monophosphate (GMP) complex (13,14). Less is known about the identity of the methyltransferase of the DLP. Because a cell-free system has not been developed that supports the synthesis of mRNA from exogenous genomic dsRNA, many questions regarding the molecular details of the transcription process remain unanswered. The preparation of RV mRNA is described in Subheading 3.3.1.

1.2.2. Preparation of Plus-Sense Template RNA from cDNAs Full-length complementary (c)DNAs, corresponding to all the genome segments of SA11 and bovine RV and to some segments of many other RVs have been synthesized. Sequencing of the cDNAs has shown that the mRNAs of the group A RVs generally begin with the sequence 5'-GGC-3' and end with the sequence 5'-ACC-3' (15). To prepare viral mRNAs that have authentic 5' and 3' ends, the full-length cDNAs can be placed into a vector immediately downstream of the promoter for T7 RNA polymerase and upstream of a SacII site. The sequence of the joined T7 promoter (lower case) and the 5'-end of the viral cDNA (upper case) should be 5'-taatacgactcactataGGC-3' and the sequence of the joined 3'-end of the cDNA (upper case) and SacII site (underlined) should be 5'-ACCgcgg-3'. As illustrated below, digestion of the vector with SacII leaves a 3'-overhang that can be removed by the 3' to 5' exonuclease activity of T4 DNA polymerase. Run-off transcription of the treated vector (see Subheading 3.2.2.) will produce viral mRNAs that have authentic 5'- and 3'- termini. 5'-ACCgcgg-3' 3'-TGGcgcc-5'

SacII

→

5'-ACCgc 3'-TGG

gg-3' cgcc-5'

T4 DNA pol

→

5'-ACC

gg-3'

3'-TGG

cc-5'

Polymerase chain reaction (PCR) amplification is an alternate approach for preparing DNA templates for the synthesis of RV mRNAs with authentic 5' and 3'-termini (16). In this case, the plus-sense primer will contain the T7 promoter sequence, followed directly by the sequence formed by the first 18 nucleotides of one of the viral mRNAs, and the minus-strand primer will be the inverse complement of the sequence formed by the last 18 nucleotides of the

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homologous mRNA. The template used for PCR amplification can be either the cDNA product of a reverse transcription reaction of viral mRNA or a cDNA that has been cloned into a DNA plasmid. If Taq DNA polymerase is used to generate the template for transcription, then the amplified product should be treated with T4 DNA polymerase, to remove the nontemplated A residues that may have been added to the 3'-ends of the cDNAs. If pfu or vent DNA polymerase is used instead, then the nontemplated addition of an A residue will not occur, and therefore treatment of the amplified DNA with T4 DNA polymerase is not required. The cDNA templates are transcribed with T7 RNA polymerase under the condition provided by the supplier of the enzyme. With a T7 Megascript Kit (Ambion, Austin, TX), transcription of the cDNA templates will yield 50–100 µg viral mRNA. The quantity of the RNA is determined by spectrophotometry, and the quality of the RNA is assessed by electrophoresis on a 5% polyacrylamide gel containing 7 M urea (see Subheading 3.7.2.). RNA in the gel can be detected with ethidium bromide or by silver staining.

1.2.3. RNA Replication by Open Cores The replicase activity associated with open cores can use viral transcripts made from DLPs or from cDNAs as templates for the sythesis of dsRNA (10). Coupled with baculovirus (BV)-expressed core proteins (VP1, VP2, and VP3), the open core system has proved to be an important tool for understanding the mechanism of RV RNA replication. Recent studies have shown that a conserved stretch of seven nucleotides at the 3'-end of the viral mRNAs forms an essential cis-acting signal for minus-strand synthesis (16,17); the 3'-end of the viral mRNA must be single stranded for efficient minus-strand synthesis (18); and both VP1 and VP2 are required for RV replicase activity (19,20,21). A step-by-step protocol for the in vitro synthesis of minus-strand RNA by open cores is described in Subheadings 3.4.1. and 3.4.2.

1.2.4. Capping-Related Activities of Open Cores VP3 and other guanylyltransferases function in mRNA capping by transferring GMP to the phosphate end of the nascent RNAs (22). During this process, the guanylyltransferase first interacts with GTP and generates a guanylyltransferase–GMP intermediate complex, or, for RVs, a reversible and covalently-linked VP3–GMP intermediate complex (13,14). The guanylyltransferase activity then transfers GMP from the intermediate complex to the pyrophosphate end of the nascent RNA, producing a GpppN cap structure in which N is a purine residue. Protocols for generating the VP3–GMP intermediate complex, and for capping the nascent RNA by the guanylyltransferase activity associated with open cores, are shown in Subheadings 3.4.3. and 3.4.4.

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1.3. Detecting RNA-Binding Activity of RV Proteins Of the 12 proteins encoded by the RV genome, at least one-half possess RNA-binding activity (23). Although the role of many of these proteins in RV replication is unclear, their function may be related to RNA synthesis and packaging, mRNA transport to the site of genome replication, and mRNA translation and the regulation of gene expression. Many methods have been used to analyze the RNA-binding activity of RV proteins although the most important has been the gel-shift assay (for VP1, VP3, nonstructural proteins NSP1, NSP2, NSP3, see refs. 19,21,24,25,26, respectively). Other methods that have also been useful for analysis of binding proteins include Northwestern blot assays (VP2) (26,27), RNA-capture assays with immunoadsorbed protein (NSP1, NSP2) (28,29) and ultraviolet (UV)-crosslinking assays (VP2, NSP2, NSP3) (20,25,26,30). RV proteins that have been used in RNA-binding assays have been obtained from infected cell lysates (25,27) and purified virions (19,21) and have been expressed in vitro with rabbit reticulocyte lysates (25) and in vivo with BV (18,26) and vaccinia expression vectors (28).

1.3.1. Gel-Shift Assays The electrophoretic mobility shift assay, or gel-shift assay, is the simplest and most rapid of the RNA-binding assays. Its usefulness for the study of RVs is perhaps best illustrated by the fact that, of the six known viral RNA-binding proteins, five have been characterized by the gel-shift assay. 1.3.1.1. PREPARATION OF PROBES

Choosing a suitable probe to use in the gel-shift assay often requires making assumptions about the location of the recognition signals in an RNA. In detecting and analyzing the RNA-binding activities of the RV proteins, the authors have assumed that the specific recognition signals for these proteins are more likely to reside in the 5' and 3'-untranslated regions of the viral mRNAs than in the open reading frame (ORF). Indeed, this idea is consistent with data showing that cis-acting signals in viral mRNAs, which function in RNA replication and in gene expression, reside at the ends of the mRNA (16,17). In designing probes, one must consider that many RNA-binding proteins may recognize defined structural motifs in an RNA, e.g., stem-loops, bulges, panhandles, rather than, or in addition to, the primary sequence of the RNA. Therefore, it may be worthwhile to determine whether the probe to be used in the gel-shift assay assumes a fold pattern that is similar to its predicted secondary structure in the full-length RNA from which it is derived. The secondary structures of RNAs are easily computed with the mfold program made available on the home page of Michael Zuker of Washington University (http://www.ibc.wustl.edu/~zuker).

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Methods for preparing 32P-labeled RNA probes and performing the gel-shift assay are described in Subheadings 3.5.1. and 3.5.2. 1.3.1.2. ASSESSING STABILITY OF PROBE–PROTEIN INTERACTIONS

The strength of the interaction between an RNA and a protein can be evaluated by monitoring the effect of including different concentrations of salt in the gel-shift assay. 1.3.1.3. DISTINGUISHING BETWEEN SPECIFIC AND NONSPECIFIC RNA-BINDING ACTIVITIES

Because RNA-binding proteins generally display some degree of nonspecific affinity for single-stranded RNA, competitive gel-shift assays should be performed to determine whether the RNA-binding activity observed for any protein is specific or nonspecific (21). In the competitive gel assay, cold competitor RNA and 32P-labeled probe are usually included in the reaction mixture at molar ratios of 1:1, 10:1, and 100:1. Particularly for those proteins that appear to have specific affinity for the probe, the competitor RNA should be of the same approximate size as the probe, so that the relative concentration of the probe to competitor RNA used in the assay will be about the same at the molar and mass level. By using dsRNA as the competitor RNA, the comparative gel-shift assay can also be used to evaluate the ability of RV proteins to bind dsRNA. The effect of the competitor RNA on the formation of protein–probe complexes is evaluated by electrophoresis of the reaction mixture on a nondenaturing 8% polyacrylamide gel, and by quantitation of the intensity of bands formed by the shifted probe with a phosphorimager. If the cold competitor RNA causes a loss of the complex formed between the 32P-labeled probe and the viral protein, this indicates that the complex represents a nonspecific interaction between the probe and protein. On the other hand, if the presence of cold competitor RNA does not affect the formation of the probe–protein complex, or causes a decrease in the amount of probe–protein complex formed, which is less than the molar ratio of competitor RNA:probe used in the assay, then the interaction between the viral protein and the probe probably includes a specific component. 1.3.1.4. IDENTIFYING PROTEINS WITH RNA-BINDING ACTIVITY

Often, an extract prepared from infected cells or virus particles is found to contain an RNA-binding activity, but the identity of the protein in the extract responsible for the activity is unknown. Four alternative procedures that can be used to identify the protein component of a probe–protein complex detected by the gel-shift assay are described in Subheading 3.5.3.1.

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1.3.2. Drawbacks of Gel-Shift Assay There are three significant limitations associated with using the gel-shift assay to study RNA-binding proteins. First, the assay does not work well for proteins that are poorly soluble, or that form multimeric structures of large or varying size. For example, analysis of the major core protein VP2 is difficult, if not impossible, by the gel-shift assay, because this protein self-assembles into poorly soluble structures that cannot migrate into nondenaturing gels (10). Second, the assay is not suitable for the analysis of proteins with weak or short-lived affinity for RNA, or for resolving RNA–protein complexes that are not stable in a Tris-glycine (pH 8.8) electrophoresis buffer system. Third, the gel-shift assay restricts the size of the RNA probe that can be used as the bait for detecting and analyzing the RNA-binding activity of a protein. This is particularly a problem when the location of a recognition signal on the RNAs is unknown. In such cases, a longer stretch of RNA, possibly a full-length mRNA in the case of RV, might serve as a more effective probe, compared to short RNA probes that represent fragments of the RNA.

1.4. RNA-Capture Assay with Immunoadsorbed Protein Most of the limitations associated with the gel-shift assay can be avoided by using the RNA-capture assay, in which a protein is immunoadsorbed onto an affinity matrix, such as protein-A Sepharose beads, and the ability of the immobilized protein to bind RNA is then evaluated by incubation with radiolabeled probes. This technique has been described in detail elsewhere (29), and therefore is only summarized here. 1. An extract containing the protein of interest is prepared under nondenaturing conditions from RV-infected cells or cells programmed to produce the protein with a recombinant expression vector. The lysate is then incubated with a monospecific monoclonal or polyclonal antiserum that recognizes the protein. Because some antibodies (Abs) might block or inactivate the RNA-binding activity of a protein by altering its conformation, it is advisable to test as many different monospecific antisera as possible for the ability to bind to the protein without affecting its activity. 2. The Ab-protein complex is incubated with protein-A or -G Sepharose beads in a buffer that is anticipated to allow the protein to retain its physiological conformation. Often, this precludes the presence of strong detergents, e.g., sodium dodecyl sulfate (SDS). The beads are then thoroughly washed with the same buffer to remove unbound protein. To verify that the protein of interest has been immunoadsorbed onto the beads, and to determine which, if any, other proteins are bound to the beads, a portion of the material is analyzed by SDS-12% polyacrylamide gel electrophoresis (PAGE). (Instead of using an Ab and protein-A or -G to immobilize the protein onto an affinity matrix, it is also

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possible to use iminodiacetic acid, coupled to Sepharose, to immobilize his-tagged RV recombinant proteins, and then to use this protein-loaded matrix in the RNA-binding assay). 3. The RNA-binding activity of the immunoadsorbed protein is assessed by incubating with radiolabeled RNA probes in the appropriate RNA-binding buffer for 1 h on a rotating wheel. Following several washes to remove the unbound probe, the bound probe is eluted under high-salt conditions, typically in the range of 0.8–1 M NaCl. Electrophoresis of a fraction of the eluted radiolabeled probe on a 5 or 8% polyacrylamide gel containing urea (see Subheading 3.7.2.) allows visualization and quantitation of the bound RNA by phosphorimaging. Alternatively, the RNA-binding activity of the protein can be analyzed by quantitating the radioactivity that is eluted from the beads with a scintillation counter.

For detecting and defining interactions between RNAs and proteins, the RNA-capture assay offers some technical advantages over other systems: 1. The RNA-capture assay provides a mechanism to study the RNA-binding properties of proteins that normally exist as oligomers, be they homo- or hetero-oligomeric; this is particularly important, because the strength and specificity of the RNA-binding activity of such a complex may differ significantly from that of any individual component of the complex. 2. The RNA-capture assay can also be used to evaluate the RNA-binding activity of a protein, without resorting to the use of strong denaturants and without a priori purification of the protein. The former is an important advantage over the Northwestern blot assay, in which the proteins must be treated with SDS, and electophoretically resolved before assaying their RNA-binding activity. 3. Compared to the gel-shift assay, the RNA-capture assay allows the use of probes that range in size from small to large, and that may even be full-length. Perhaps better than any other approach, because of the flexibility in probe size, this assay system can be used in a systematic manner, to map the precise site of a recognition signal for an RNA-binding protein, beginning with a large RNA. 4. Finally, a strong advantage of the RNA-capture assay is that the system can be used easily and rapidly to define the optimal conditions that promote the interaction between a protein and an RNA. For example, metal ions (zinc), nucleotides (adenosine triphosphate [ATP] or GTP), or Na or K salts, can be added to the RNA-binding buffer to determine whether they enhance the RNA-binding activity of a protein. Similarly, the effect of temperature, pH, buffering system, stabilizing agents (glycerol and bovine serum albumin [BSA]), and the presence of protease and RNase inhibitors on the formation of RNA–protein complexes can also be rapidly evaluated with this system.

1.5. Assembly of CLPs and VLPs from Recombinant Protein The expression of RV proteins with recombinant baculoviruses (rBVs) has been useful not only for analyzing the properties of the RV proteins, but,

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perhaps even more so, for defining the structure of the rotavirion. In particular, expression of VP2 alone, with VP6, or with VP6, VP7, and VP4, in Spodoptera frugiperda (Sf9) insect cells, infected with appropriate rBVs, has been used to generate single-layered CLPs, double-layered VLPs, and triple-layered VLPs, respectively (31–33). Analysis of the recombinant particles by cryoelectron microscopy and image reconstruction (see Chapter 2) has helped to establish the number, location, and arrangement (e.g., triangulation number) of structural proteins in the virion (34). More recently, the co-expression of recombinant rVP1 and/or rVP3 with VP2 and VP6 has also provided insight into the structure and location of the viral RNA polymerase and guanylyltransferase in the core of the virion (35; see Chapter 2). From an applied point of view, the fact that purified recombinant TLPs, when administered orally to animals, can induce a neutralizing Ab response, raises the possibility that rVLPs may be used to generate RV vaccines (see Chapter 9). Although BV vectors are noted for expressing high concentrations of recombinant protein, the production of protein by this system is a relatively slow process, because it requires several steps, including the cloning of a cDNA into a specialized transfer vector, the generation of a rBV with the transfer vector, and the selection and plaque purification of the rBV. Therefore, other expression systems may be more useful in situations in which many different recombinant proteins need to be generated (e.g., in mutagenesis studies), and when the recovery of large amounts of recombinant protein is not critical. As an alternative to the BV system, recombinant proteins can be produced by transfecting T7 transcription vectors containing cDNAs of RV genes into CV1 or MA104 cells infected with vTF7-3, a recombinant vaccinia virus that constitutively expresses T7 RNA polymerase (36). With respect to the study of RV replication, the vTF7-3 expression system has several benefits: The expression of RV proteins by the vTF7-3 system can reach levels that are as high as those found in RV-infected cells; the vTF7-3 system can express RV proteins in cells which are permissive for RV replication, and therefore is likely to support authentic modification of recombinant proteins; and structural proteins expressed by the vTF7-3 system can also assemble into CLPs and VLPs.

1.5.1. Expression and Purification of VLPs Made by BV Vectors The co-expression of various combinations of RV structural proteins with rBV vectors in insect cells has been use to produce large amounts of highly purified single-layered CLPs, and double- and triple-layered VLPs. Detailed protocols for producing and purifying the recombinant particles are given elsewhere in the literature (8,31–33,35,37), and therefore will not be repeated here (see Chapter 9). However, there are several features of these protocols that are worth noting.

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1. Double- and triple-layered VLPs are released into the media of infected Sf9 cells; CLPs will remain cell-associated (31,32). 2. The detergent sodium deoxycholate (DOC) is commonly used in the purification of CLPs, because it enhances their solubility and aids in their release from cellular material (33). Sonication has also been use to dissociate CLPs from host debris, and to break up CLP aggregates. 3. CLPs and double- and triple-layered VLPs have been successfully purified by centrifugation on isopycnic CsCl gradients (8,31,32). However, because CLPs can be somewhat unstable in CsCl, procedures have also been developed for the purification of CLPs by sedimentation on sucrose gradients (33,35,37). 4. Double- and triple-layered VLPs are stable upon extraction in the lipid solvent trichlorotrifluoroethane (32). 5. Concentration of CLPs may result in their conversion into sheet-like, helical, or elongated bristly structures (37). 6. VP2 expressed by BV vectors is extremely susceptible to proteolytic cleavage (37), and therefore protease inhibitors (e.g., leupeptin and aprotinin) must be added to all solutions used to produce and purify the protein.

1.5.2. Expression of CLPs with the Vaccinia-Virus vTF7-3 System Infection of mammalian cells with the recombinant vaccinia virus vTF7-3 leads to the production of T7 RNA polymerase, which in turn can direct the synthesis of mRNAs from T7 transcription vectors contained in the same cell. Because of the production of virus-encoded capping enzymes in vTF7-3-infected cells, a significant percentage of the T7 transcripts will gain 5'-cap structures that will enhance their ability to direct protein synthesis. By modification of transcription vectors, so that they produce T7 transcripts that initiate with sequences specifying an independent ribosome entry site, translation of the transcripts becomes cap-independent, which often will result in even higher levels of recombinant protein expression in the cell. The vTF7-3 expression system has been used successfully to express nearly all RV proteins, including those that are structural components of the virus and those that have RNA-binding activity. Subheadings 3.6.1. and 3.6.2. describe methods used to express and purify VP2 CLPs from vTF7-3-infected cells. Derivative protocols may be used to study the assembly of double- and triple-layered VLPs, and to identify, by mutagenesis, domains in the structural proteins that are important for virion assembly.

1.6. SDS-12% PAGE for Separation of dsRNAs and Proteins Viral dsRNAs and proteins can be electrophoretically resolved on discontinuous gels (100 × 100 × 1.5 mm) containing a resolving portion of 12% polyacrylamide and a stacking portion of 4% polyacrylamide (see Subheading 3.7.).

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2. Materials 2.1. Virus Propagation 1. 2. 3. 4. 5. 6. 7. 8. 9.

MA104 cell cultures. Medium 199. Phosphate buffered saline (PBS), pH 7.2. Trypsin (cryst., 5 µg/mL, 1 µg/mL), or 5 µg/mL pancreatin. EDTA. FBS. Cell culture flasks (150 cm2). Six-well tissue culture plates. 1% Nutrient agar: prepared by mixing an equal volume of a 2% sterile solution of Bacto agar (Difco, all locations) or cell culture grade agarose and 2X Medium 199 containing 25 µg/mL of diethylaminoethyl dextran, 200 U/mL penicillin, and 200 µg/mL streptomycin. 10. 1% Neutral red agar: prepared by mixing an equal volume of a 2% sterile solution of cell-culture grade agarose and 2X Medium 199 containing 50 µg/mL neutral red. 11. Centrifuge. 12. Incubator.

2.2. Virus Preparation 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.

12. 13. 14. 15. 16. 17.

Methionine-free cell culture medium. 35S-labeled amino acids. Medium 199. Aprotinin. Leupeptin. Trichlorotrifluoroethane. Sorvall (DuPont Co., Wilmington, DE) Omni-mixer. Centrifuge. Ultracentrifuge. Ultracentrifuge tubes. Tris-buffered saline (TBS): 8 g NaCl, 0.38 g KCl, 0.1 g Na2HPO4, 1 g dextrose, 3 g Tris-base, 0.1 g MgCl2, 0.1 g CaCl 2. Dissolve in distilled H2O, adjust to pH 7.4 with HCl, and make up to 1 L. CsCl. 21-gage needle. Sodium azide (NaN3). 0.5 M EDTA, pH 8.0. Spectrophotometer. SDS-12% PAGE (see Subheading 2.7.).

2.3. RV RNA Preparation 1. Transcription cocktail: 100 mM Tris-HCl, pH 7.8, 50 mM sodium acetate, 10 mM magnesium acetate, 1.0 mM dithiothreitol, 2.5 mM each nucleoside

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2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

triphosphate, 8.0 mM phospho(enol)pyruvate, 50 µg/mL pyruvate kinase, 1.0 mM S-adenosyl-L-methionine, 160 U/mL RNasin (Promega, Madison, WI). 10% SDS. Phenol:chloroform (1:1). Chloroform:isoamyl alcohol (24:1). 3 M sodium acetate. Ethanol. Centrifuge. RNase-free distilled water. Spectrophotometer. SacII restriction enzyme. T4 DNA polymerase. 10X T4 DNA polymerase buffer: 0.5 M NaCl, 0.1 M Tris-HCl, 0.1 M MgCl2, 10 mM dithiothreitol, pH 7.9 (New England BioLabs, Beverly, MA). 10 mg/mL BSA.

13.

2.4. Preparation and Use of Open Cores 1. Low-salt buffer (LSB): 2 mL 1 M Tris-HCl, pH 7.5, 1 mL 0.5 M Na 2EDTA, pH 7.5, 77 mg dithiothreitol. Make up to 1 L with distilled H 2O. 2. Digestion buffer: 340 U micrococcal nuclease, 10 mM Tris-HCl, pH 8.0, 10 mM NaCl, 1 mM CaCl2. 3. 1% agarose. 4. 50X TAE buffer: 242 g Tris-base, 136.1 g sodium acetate.3H2O, 19g Na2EDTA.2H2O Dissolve in distilled H2O, adjust to pH 7.2, and make up to 1 L. 5. EGTA. 6. 1 M Tris-HCl, pH 7.1. 7. 0.2 M magnesium acetate. 8. RNasin (40 U/µL). 9. 30% polyethylene glycol. 10. 10 mM nucleoside triphosphates. 11. Dithiothreitol. 12. [α-32P]-uridine triphosphate (UTP) (10 mCi/mL, 800 Ci/mmol). 13. RNase-free distilled water. 14. SDS-PAGE sample buffer (see Subheading 2.7.). 15. [α-32P]-GTP (10 mCi/mL, 800 Ci/mmol). 16. 5 mM MgCl2. 17. NaCl. 18. Phenol:chloroform (1:1). 19. Chloroform. 20. Sephadex G50. 21. SDS-12% polyacrylamide gel (see Subheading 2.7.). 22. 6 M urea-8% polyacrylamide gel (see Subheading 2.7.). 23. X-ray film. 24. Autoradiography cassette.

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2.5. Gel-Shift Assay 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18.

Ambion T7 MEGAshort transcription system. [α-32P]-UTP (10 mCi/mL, 800 Ci/mmol). 1.9 mM cold UTP. RNase-free DNase. Phenol:chloroform (1:1). Chloroform. 4 M NaCl. Ethanol. 6 M urea–8% polyacrylamide gel (see Subheading 2.7.). LSB. 8% polyacrylamide gel in Tris-glycine buffer (see Subheading 2.7.). Whatman 3M paper. X-ray film. Autoradiography cassette. 8% polyacrylamide gel containing 50 mM Tris-HCl, pH 8.8 (see Subheading 2.7.). SDS-PAGE sample buffer (see Subheading 2.7.). SDS-PAGE 4% stacking and 12% resolving gel (see Subheading 2.7.). Fluorographic enhancer (Amplify [Amersham, Piscataway, NJ] or Enhance [New Life, Boston, MA]). 19. 254 nm 25 W germicidal UV lamp. 20. RNase A.

2.6. Expression and Purification of VP2 CLPs from vTF7-3-Infected Cells 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15.

MA104 cell cultures. Medium 199. FBS. Penicillin (100 U/mL). Streptomycin (100 µg/mL). CO2 incubator. Cytosine β-D-arabinofuranoside (AraC). Vaccinia virus recombinant - vTF7-3. Qiagen Maxi-prep DNA purification kit. LipofectAMINE reagent (Gibco). Aprotinin. Leupeptin. PBS, pH 7.2. Centrifuge. Lysis buffer: 10 mM Tris-HCl, pH 7.4, 0.1 M EDTA, 1% DOC, 1 µg/mL aprotinin, 1 µg/mL leupeptin. 16. 20% sucrose (w/v) in 10 mM Tris-HCl, pH 7.4, 0.1 M EDTA, 0.2% deoxycholate (DOC). 17. Ultracentrifuge. 18. TE buffer: 10 mM Tris-HCl, pH 8.0, 0.1 mM EDTA.

46 19. 20. 21. 22.

Patton et al. Amicon Centricon microconcentrator. 3% phosphotungstic acid-NaOH, pH 7.2. Formvar-carbon coated grid. Transmission electron microscope.

2.7. Electrophoresis Systems 1. Electrophoresis power supply. 2. Electrophoresis tanks, vertical and horizontal. 3. Electrophoresis buffers: a. 10X SDS-PAGE running buffer: 105 g Tris-base, 504 g glycine, 35 g SDS, 3 L distilled H2O. Adjust to pH 9.1 with NaOH, and bring to a final volume of 3.5 L. b. SDS-PAGE sample buffer: 10 mL 50% glycerol (v/v), 1.25 mL 1 M Tris-HCl, pH 6.8, 5.0 mL 10% SDS (w/v), 8.0 mL distilled H2O, 150 µL 0.2% bromophenol blue in ethanol, 38.5 mg dithiothreitol. Bring to a final volume of 25 mL with distilled H2O. c. 10X TBE running buffer: 121.1 g Tris-base, 55 g boric acid, 7.5 g Na2EDTA. Bring to a final volume of 1 L with distilled H2O; pH should be 8.3. d. 2X Formamide sample buffer: 200 µL 0.5 M EDTA, pH 8.0, 200 µL 0.2% bromophenol blue in ethanol, 9.6 mL deionized formamide. 4. Electrophoresis gels: a. 12% Resolving gel: 15.5 mL 30% acrylamide (w/v), 6.4 mL 2% bis-acrylamide (w/v), 10.0 mL 1.5 M Tris-HCl, pH 8.8, 0.4 mL 10% SDS, 7.45 mL distilled H 2O, 20 µL TEMED, 0.2 mL 10% ammonium persulfate. b. 4% Stacking gel: 2.6 mL 30% acrylamide, 1.6 mL 2% bis-acrylamide, 2.5 mL 1 M Tris-HCl, pH 6.8, 0.2 mL 10% SDS, 13.7 mL distilled H2 O, 20 µL TEMED, 0.1 mL 10% ammonium persulfate. c. 6 M Urea-8% polyacrylamide gel: Dissolved 18.75 g of urea in 11.25 mL distilled H2O, then add 3.75 mL 10X TBE, 7.5 mL 40% acrylamide solution (38 g, acrylamide + 2 g bis-acrylamide in 100 mL distilled H 2 O). Add 7.5 µL TEMED and 0.38 mL 10% ammonium persulfate. d. 7 M Urea-5% polyacrylamide gel: Dissolve 16.8 g of urea in 17.5 mL distilled H 2O, then add 4 mL 10X TBE; 4 mL 50% acrylamide solution (50 g acrylamide + 1 g bis-acrylamide in 100 mL distilled H2O). Add 0.1 mL TEMED and 0.4 mL 10% ammonium persulfate.

3. Methods 3.1. Virus Propagation

3.1.1. Subculture of MA104 Cells 1. Disrupt a confluent cell monolayer of MA104 cells, by incubating in a solution of PBS containing 0.005% trypsin and 0.1% EDTA. 2. Resuspend the detached cells in serum-containing medium, (see Note 1). 3. Dilute the MA104 cell suspension from 1 in 5 to 1 in 10 in medium, and seed new cell culture flasks. The cells will produce a confluent monolayer in 4–7 d.

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3.1.2. Virus Propagation 1. Wash confluent MA104 cells maintained in 150-cm2 flasks, 3× with serum-free medium, or incubate several hours to overnight in one change of serum-free medium. 2. Prepare the viral inoculum by incubating virus stocks of known titer with 5 µg/mL trypsin for 30 min at 37°C, and diluting the activated virus stock to a volume of 5 mL/flask with serum-free medium, (see Note 2). 3. Remove the serum-free medium from the cell monolayer, and add 5 mL inoculum to each flask. 4. Incubate the flasks for 1 h at 37°C, with gentle swirling of the inoculum over the monolayer every 15 min. 5. Remove the inoculum and add 20 mL serum-free medium containing 1 µg/mL trypsin to the flasks, maintain the cells at 37°C until the cytopathic effect (CPE) is complete (see Note 3). 6. Freeze and thaw the infected cells 3× and remove large debris from the lysate by low-speed centrifugation (see Note 4). 7. The lysate is titered by plaque assay (see Subheading 3.1.3.), stored frozen at –20°C, and used as stock for subsequent propagation of the virus.

3.1.3. Titering Virus by Plaque Assay 1. Seed MA104 cells into six-well plates at a density that will allow the monolayers to reach confluency in 2–3 d (see Note 5). 2. Incubate the confluent monolayers in serum-free medium for 16–18 h, and inoculate with serial 10-fold dilutions of trypsin-activated virus (0.1 mL/well). 3. Incubate the plates at 37°C for 1 h, with occasional rocking to allow for virus adsorption. 4. Gently overlay the monolayers with 3.0 mL/well 1% nutrient agar or agarose containing pancreatin (3–5 µg/mL) or trypsin (1 µg/mL) cooled to 45°C. 5. Immediately after adding the overlay, swirl slowly, allowing the inoculum and overlay to mix. 6. Once solidified, incubate the plates for 3–4 d at 37°C. 7. Add 1 mL of a second overlay, consisting of 1% neutral red agar, cooled to 45°C, to each well, to aid plaque counting. 8. Count the plaques 3–5 d following the addition of the second overlay. Typically, the titer of rhesus rotavirus (RRV), simian rotavirus (SA11)-4F, and prototypic RVs will reach 1 × 108 PFU/mL.

3.2. Virus Preparation 3.2.1. Preparation of 35S-Labeled Virions When it is necessary to obtain 35S-labeled virions, monolayers are infected with virus and maintained in 85% methionine-free medium containing 5–10 µCi of 35S-labeled amino acids (Specific radioactivity: 1175 Ci/mmol) per mL. The infection is allowed to go until CPE is complete. Afterwards, the virus is purified from the lysate, as described in Subheading 3.2.3.

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3.2.2. Preparation of Virions Containing Uncleaved VP4 1. Infect MA104 cells at a high MOI (3–10 PFU/cell) with trypsin-activated virus. 2. Wash the cell sheet several times with serum-free medium following adsorption. 3. Maintain the cells in trypsin-free medium containing 1 µg/mL aprotinin or 0.5 µg/mL leupeptin (see Note 6). 4. Harvest the cells once the CPE has reached 70–80% (~24 h post infection).

3.2.3. Purification of Triple-Layered Virions 1. Combine the lysate with an equal volume of trichlorotrifluoroethane, and homogenize the mixture with a Sorvall Omni-mixer for three 1 min cycles on ice. 2. Centrifuge the homogenates at low speed to separate the organic and aqueous phases (4100g for 10 min in a Sorvall GSA rotor at 4°C). 3. Pellet the virus particles from the aqueous phase by centrifugation at 90,000g in a Beckman (Palo Alto, CA) SW28 rotor or at 45,000g in a Beckman Type 19 rotor for 2 h at 4°C. 4. Resuspend the pellet in TBS, and add CsCl to the virus suspension to achieve a density of 1.370 g/cm3, as determined by refractometry (see Note 7). 5. Centrifuge the mixture at 110,000g in a Beckman SW50.1 rotor for 20 h at 4°C. 6. Inspect the gradients in a darkened room with an inverted light source, to reveal two bands (see Note 8). 7. Recover the virus from the gradient by puncturing the bottom of the centrifuge tube with a 21-gage needle, and collect drops containing TLPs and DLPs, respectively, into separate tubes. 8. Remove the CsCl by dialyzing the virus extensively against TBS at 4°C. 9. Store the dialyzed samples at 4°C in the presence of 0.01% NaN3 (see Note 9).

3.2.4. Preparation of DLP Virus 1. Add 0.5 M EDTA, pH 8.0, to purified virus TLPs in TBS buffer, to give a final concentration of 10 mM EDTA. 2. Incubate for 1 h at 37°C with gentle periodic mixing. 3. Centrifuge the virus sample at 200,000g in a Beckman SW50.1 rotor for 1 h at 4°C, to pellet the DLPs. 4. Resuspend the pellet in TBS containing 10 mM EDTA. 5. Purify the DLPs by banding on a CsCl gradient (see Subheading 3.2.3 and Note 10).

3.2.5. Preparation of Cores 1. 2. 3. 4. 5. 6.

Dilute DLPs with TBS to an optical density OD 260 of 2 or less. Mix the sample with an equal volume of 2.0 M CaCl2 (see Note 11). Incubate for 1 h or more at 37°C, with frequent agitation (see Note 12). Centrifuge the sample at 16,000g for 2 min in an Eppendorf centrifuge at 4°C. Wash the pellet once with 0.5 mL 1 M CaCl2. Resuspend the cores in 2.5 µL TBS/mL of starting infected cell lysate.

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Fig. 1. Protein composition of purified TLPs (lane 1), DLPs (lane 2), and single-layered (core) particles (lane 3), as evaluated by SDS-12% PAGE. Virus was propagated in the presence of trypsin, causing the loss of VP4 and the appearance of VP5* and VP8* in the TLPs. Proteins in the gel were detected by staining with Coomassie blue R-250. 7. Dialyze extensively against TBS, to remove the residual CaCl2. 8. Purify the cores by centrifugation for 60 min at 200,000g in a Beckman SW50.1 rotor at 4°C through a 0.5-mL cushion of 40% sucrose in TBS (w/v). 9. Resuspend the pellets in TBS, and briefly sonicate to disrupt the core aggregates. 10. Adjust the suspension to a density of 1.45 g/cm3 with CsCl (by adding 3 g CsCl for every 4 mL core suspension). 11. Centrifuge the sample at 150,000g in a Beckman SW50.1 rotor for 20 h at 4°C. 12. Collect the cores by bottom puncture (see Note 13). 13. Dialize extensively against TBS to remove CsCl. 14. Determine the purity and quality of cores, DLPs and TLPs by SDS-12% PAGE (see Subheading 3.7., Fig. 1 and Note 14).

3.3. RV RNA Preparation 3.3.1. Preparation of mRNA from RV Particles 1. Pellet at least 100 µg CsCl gradient-purified DLPs in TBS by centrifugation at 200,000g in a Beckman SW50.1, or at 150,000g in a Beckman TLS55 rotor for 1 h at 4°C. 2. Resuspend in 1 mL transcription cocktail (see Note 15).

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3. Incubate for 2 h at 40°C (see Note 16). 4. Add 50 µL 10% SDS. 5. Extract the sample twice with an equal volume of phenol:chloroform (1:1, pH 4.5), and once with an equal volume of chloroform:isoamyl alcohol (24:1). 6. Add a 1/10 vol of 3 M sodium acetate and 3 vol of ethanol, mix, and incubate for 2 h at –70°C to precipitate the RNA transcripts. 7. Collect the RNA precipitate by centrifugation at 12,000g in a Sorvall SS34 rotor for 20 min at 4°C. 8. Resuspend the RNA pellet in 0.5 mL RNase-free water, and determine the concentration by spectrophotometry. The yield should be approx 200–300 µg RNA (see Note 17).

3.3.2. Production of Viral mRNA from Transcription Vector 1. Incubate 10 µg purified plasmid DNA with 30 U SacII restriction enzyme in a final volume of 50 µL for 1 h at 37°C. 2. Add 10 U T4 DNA polymerase, 10 µL 10X T4 DNA polymerase buffer, 1 µL 10 mg/mL BSA, 1 µL each of 2.5 mM dATP, dCTP, dGTP, and dTTP, and water to a final volume of 100 µL. 3. Incubate the reaction mixture for 20 min at room temperature. 4. Recover the plasmid by phenol:chloroform extraction and ethanol precipitation (see Subheading 3.3.1.).

3.4. Preparation and Use of RV Open Cores 3.4.1. Preparation of Open Cores 1. Prepare open cores by extensive dialysis of purified core particles (see Subheading 3.2.5.) against LSB at 4°C (see Note 18). 2. Incubate 100 µg open cores (see Note 19) in 1 mL digestion buffer, containing 340 U micrococcal nuclease, 10 mM Tris-HCl, pH 8.0, 10 mM NaCl, and 1 mM CaCl2 for 60 min at 30°C, to remove the dsRNA genome from open cores. 3. Confirm digestion of the dsRNA by electrophoresis of a 15-µL aliquot of the digest on a 1% agarose gel in TAE buffer. 4. Inactivate the micrococcal nuclease activity by adding EGTA to a final concentration of 3 mM to the digest.

3.4.2. Replicase Assay 1. Mix the following components: 1 µL 1 M Tris-HCl, pH 7.1; 1 µL 0.2 M magnesium acetate; 1 µL RNasin (40 U/µL); 1 µL 30% polyethylene glycol; 2.5 µL 10 mM mixture of nucleoside triphosphates (2.5 mM of each); 0.4 µL dithiothreitol; 1.5 µL [α-32P]-UTP (10 mCi/ml, 800 Ci/mmol); 2.5 µL open cores (100–300 µg/mL); 0.1–1.0 µg viral mRNA; RNase-free distilled water, to bring the reaction mix to a total volume 20 µL. 2. Incubate the mixture at 32°C for 2–3 h.

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Fig. 2. Replicase activity associated with untreated and micrococcal nuclease-treated open cores. The exogenous RNA added to some reaction mixtures (lanes 2 and 4) was produced in vitro by T7 transcription of a linearized plasmid containing a cDNA of the SA11 gene encoding NSP2 (gene 8). 32P-labeled dsRNA products recovered from the reaction mixtures were detected by SDS-12% PAGE and autoradiography. Note the appearance of radiolabeled background bands corresponding to the 11 genome segments made in the reaction mixture containing untreated open cores and no exogenous mRNA (lane 1). Micrococcal nuclease-treated open cores without exogenous RNA act as a negative control (lane 3). 3. Combine the reaction mixture with 50 µL SDS-PAGE sample buffer, and incubate for 15 min at 37°C. 4. Analyze the samples by SDS-12% PAGE (see Figs. 2 and 3, and Notes 20–23).

3.4.3. Formation and Detection of VP3–GMP Intermediate Complexes 1. Incubate 1–10 µg open cores with 100 µCi [α-32P]-GTP (800 Ci/mmol), 5 mM MgCl 2, and 50 mM NaCl in a final volume of 30 µL for 30 min at 37°C. (see Fig. 3 and Note 24). 2. Detect the 32P-labeled VP3 by SDS-12% PAGE and autoradiography.

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Fig. 3. The various biological activities of open cores and the influence of NaCl. (Top panel) 32P-labeled VP3–GMP intermediate complexes were formed by incubating 32P-GTP with open cores, and were detected by SDS-12% PAGE and autoradiography. Not only were 32P-labeled VP3–GMP complexes generated in reaction mixtures lacking NaCl, but, through an unknown mechanism, 32P-labeled VP1 was generated as well. In the absence of MgCl2, neither 32P-labeled VP3–GMP nor 32P-labeled VP1 were produced. The addition of NaCl to reaction mixtures prevented the formation of 32P-labeled VP1, but had little (<2-fold) effect on the formation of the 32 P-labeled VP3–GMP complex. (Second panel) RNA capping assays contained 32P-GTP, open cores, cold SP72-v3'40 RNA, and either no MgCl or MgCl and the 2 2 indicated level of NaCl. The products were analyzed by electrophoresis of reaction mixtures on a 6 M urea–8% polyacrylamide gel, and detected by autoradiography. As the results show, capping was dependent on MgCl2, and was nearly blocked at high

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3.4.4. Assay for 5'-Capping of RNA 1. Incubate 20 µg open cores in LSB, 5 mM MgCl2, 400 µCi [α-32P]-GTP (800 Ci/mmol), and 2.5 µg cold RNA in 160 µL for 3 h at 37°C, to produce capped and 32 P-GMP-labeled RNA (see Fig. 3 and Note 25). 2. Deproteinize the capped RNA by extraction with phenol:chloroform and chloroform; if necessary, unincorporated 32 P-GTP can be removed by passing the sample through a Sephadex G50 column. 3. Depending on the length of the RNA, the capped RNA can be detected by electrophoresis on either an SDS-12% polyacrylamide gel or a 6 M urea-8% polyacrylamide gel, followed by autoradiography or phosphorimaging.

3.5. Gel-Shift Assay 3.5.1. Preparation of 32P-Labeled RNA Probes 1. Prepare the 32P-labeled RNA probes for use in the gel-shift assay by run-off transcription with an Ambion T7 MEGAshort kit transcription (Austin, TX). The reaction conditions are as described by the manufacturer, except that 2.5 mCi/mL [32P]-UTP (800 Ci/mmol) and 1.9 mM cold UTP are included in the reaction mixture. 2. The templates used for transcription are generated either by PCR amplification of a viral cDNA or of a nonviral DNA using a plus-sense primer that also contains the promoter sequence for T7 RNA polymerase, or by linearizing a plasmid containing a promoter for T7 RNA polymerase at an appropriate site with a restriction enzyme. If necessary, T4 DNA polymerase can be used to produce blunt-ends the template before transcription (see Subheading 3.3.2.). 3. Incubate the transcription mixture for 8–18 h at 37°C. 4. Treat with RNase-free DNase to remove the DNA template. 5. Deproteinize by extracting sequentially with phenol:chloroform and chloroform. 6. Precipitate the RNA from the sample by adding 1/20 vol of 4 M NaCl and 2.5 vol of ethanol. Fig. 3. (continued) concentrations of NaCl (100–200 mM). (Third panel) Replicase assays contained 32P-UTP, open cores, no mRNA, or gene 8 mRNA and the indicated level of NaCl. The synthesis of 32P-labeled dsRNA in the reaction mixtures was quantitated by SDS-12% PAGE, followed by phosphorimaging. As the concentration of NaCl increased in the reaction mixtures, the level of dsRNA synthesis decreased. (Bottom panel) Gel-shift assays were performed to evaluate the RNA-binding activity of the core proteins. Reaction mixtures containing 32P-labeled SP72-v3'40 probe, open cores, no MgCl2, and the indicated amount of NaCl. The probe–protein complexes were detected by electrophoresis of the reaction mixtures on a nondenaturing 8% polyacrylamide gel. The presence of 50–150 mM NaCl had either no effect or an enhancing effect on the formation of VP1– and VP3–probe complexes. At 200 mM NaCl, the level of VP1–probe complexes formed in the reaction mixture was the same as that formed in reaction mixtures containing no added NaCl, but the level of VP3–probe complexes was reduced.

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7. Dissolve the probe in 20 µL RNase-free water, and gel-purify by electrophoresis on an 8% polyacrylamide gel containing 6 M urea, according to the protocol described in Subheading 3.7.

3.5.2. Assay Conditions (see Note 26) 1. Combine 1 pmol gel-purified 32P-labeled RNA probe with 2.5–5 µg open cores. 2. Adjust the mixture to a final volume of 10 µL with LSB. Include a control reaction mixture that contains the labeled probe, but lacks open cores. 3. Incubate the mixture for 60 min at room temperature (see Note 27). 4. Add 10 µL gel-shift sample buffer to the reaction mixture, and electrophorese the samples at 175 V on a nondenaturing 8% polyacrylamide gel in Tris-glycine buffer for 3–6 h (see Subheading 3.7. and Note 28). 5. Dry the gel onto Whatman 3M paper, and determine the location of the RNA probe in the gel by autoradiography (see Fig. 4 and Note 29).

Fig. 4 shows an example of the gel-shift pattern obtained with open cores and 32P-SP72-v3'40, an RNA probe that is 72 nucleotides long. The last 40 nucleotides of SP72-v3'40 are identical in sequence to the last 40 nucleotides of the SA11 gene 8 mRNA. The positions of the VP1–probe and VP3–probe complexes in the nondenaturing gel are indicated (lane 2, lower and upper bands, respectively). Despite containing a smaller protein, the VP3–probe complex migrates much slower in the gel than the VP1–probe complex. One explanation for this observation is that the VP3–probe complex may represent the interaction of a VP3 multimer with the probe. However, it is possible that the charge or Stokes radius of the protein is causing the slow rate of migration of the VP3–probe complex relative to the VP1–probe complex. Fig. 4 also exemplifies the usefulness of cold competitor RNAs as a tool to analyze the specificity of the interaction between viral proteins and probe in the gel-shift assay. The presence of the unlabeled luciferase RNA, Xenopus elongation factor-1 RNA, and brome mosaic virus RNA prevented the formation of a complex with VP3 and 32P-labeled SP72-v3'40, but not with VP1 and the same probe (21). Because the probe contains sequences present at the 3'-end of the RV gene 8 mRNA, the results of the gel-shift experiment suggest that VP3 does not specifically recognize the 3'-end of the mRNA, but VP1 does. Therefore, this experiment provides evidence that the 3'-terminus of the gene 8 mRNA may contain a specific recognition signal for VP1. As shown in Fig. 3, the RNA-binding activity of VP1 and VP3 was not inhibited by the presence of NaCl, except at 200 mM, at which the binding activity of VP3 was reduced by approx 30%. Indeed, in the presence of low concentrations of NaCl (50 mM), the RNA-binding activity of VP1 and VP3 was enhanced. Given that the physiological concentration of salt in the cell is

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Fig. 4. RNA-binding activities of open cores, as evaluated by the competitive gelshift assay. Open cores and 1 pmol 32P-labeled probe, SP72-v3'40, were incubated alone or with 5 pmol luciferase RNA, 1 pmol xenopus elongation factor-1 RNA, or 2 pmol brome mosaic virus RNA. Probe–protein complexes in the reaction mixtures were resolved by electrophoresis on a nondenaturing 8% polyacrylamide gel, and were detected by autoradiography. When open cores were not added to the reaction mixture, the upper and lower complexes were not detected (lane 1). Protein analysis has indicated that VP3 and VP1 are responsible for the formation of the upper and lower bands, respectively (lane 2). Note that in the presence of cold competitor RNA, the upper band is lost (lanes 3–5), indicating that the interaction between VP3 and the probe is nonspecific.

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taken as 100–150 mM, these data would indicate that VP1 and VP3 can form stable complexes with RNA in the infected cell. 3.5.3. Identifying Proteins with RNA-Binding Activity 3.5.3.1. DETECTION OF 35S-LABELED PROTEIN IN PROBE-PROTEIN COMPLEXES

This procedure requires the use of 35S-labeled viral protein in the gel-shift assay, and was used to establish that VP1, VP3, and NSP2 have RNA-binding activity (19,21,24). 1. Combine 35S-labeled open cores, or an extract prepared from 35S-labeled infected cells, with an appropriate 32P-labeled RNA probe. 2. Incubate for 30–60 min at room temperature (see Subheading 3.5.2.). 3. Resolve the probe–protein complexes in the mixture by electrophoresis on a nondenaturing 8% polyacrylamide gel containing 50 mM Tris-HCl, pH 8.8 (see Subheading 3.7.). 4. Dry the gel onto Whatman 3M paper and detect the positions of the probe–protein complexes by autoradiography. 5. With the autoradiograph as a guide, cut out portions of the gel containing probe-protein complexes. 6. Rehydrate the pieces for 5 min in 200 µL SDS-PAGE sample buffer at room temperature. 7. Separate the gel pieces from the Whatman paper with tweezers. 8. Incubate the gel pieces for 30 min more at 37°C in 200 µL of fresh sample buffer. 9. Insert the rehydrated pieces into the well of an SDS-PAGE 4% stacking gel layered over an SDS-PAGE 12% resolving gel (see Subheading 3.7.). Parallel wells are loaded, with appropriate viral protein markers and mol wt standards. 10. Following electrophoresis, soak the gel in a fluorographic enhancer agent such as Amplify or Enhance. 11. Detect the 35S-labeled protein in the gel by autoradiography.

3.5.3.2. CROSSLINKING OF RNA–BINDING PROTEIN TO 32P-LABELED PROBE

The basis for this procedure is that exposure of RNA–protein complexes to UV light can induce the formation of covalent linkages between the RNA and protein. 1. Incubate unlabeled open cores or extracts of infected cells with a high specific activity 32P-labeled probe, under the same conditions used for the gel-shift assay (see Subheading 3.5.2.). 2. Expose the reaction mixtures to UV light for various lengths of time (e.g., 0, 5, 10, 15, 30, 60 min) using a 254-nm 25-W germicidal lamp placed 3–4 cm above the samples. 3. Treat the samples with 10 µg/mL RNase A at 37°C, to remove all but the few nucleotides of the RNA probe that are covalently linked to the protein via exposure to UV light.

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4. Identify the 32P-labeled protein by co-electrophoresis on an SDS-12% polyacrylamide gel, with appropriate viral protein markers and mol wt standards (see Note 30).

3.5.3.3. RNA-BINDING ACTIVITY OF RECOMBINANT PROTEIN

Large amounts of nearly all the RV proteins have been produced with both recombinant eukaryotic (BV and vaccinia virus) and prokaryotic (bacterial) expression vectors. Some of the recombinant proteins have also been purified to homogeneity, or nearly so, and used in gel-shift assays to assess the potential RNA-binding activity of the protein. The obvious advantage of using purified recombinant proteins in the gel-shift assay is that detection of any RNA–protein complexes in the gel leads to the conclusion that the protein has RNA-binding activity. For example, experiments that showed that BV-expressed VP1 interacted with the 3'-specific viral probe SP72-v3'40 were useful for confirming results of earlier gel-shift assays, which suggested that VP1 has RNA-binding activity (18). Similarly, NSP1 produced in bacteria and NSP2 produced in CV1 cells, with the vaccinia virus vTF7-3 expression system, have also contributed to verifying and defining the RNA-binding activities of these proteins (28,29). The limitation of using recombinant protein to evaluate the RNA-binding activity is that the recombinant protein may behave differently than the naturally occurring protein. As an example, VP3 contained in open core preparations interacts readily with probes to form RNA–protein complexes that can be detected by the gel-shift assay. In contrast, recombinant VP3 produced with a BV expression vector is largely insoluble, and, as yet, has not been shown to have RNA-binding activity. 3.5.3.4. SUPERSHIFTING PROBE–RNA COMPLEXES WITH SPECIFIC ABS

Monospecific polyclonal and monoclonal antisera can be used together with the gel-shift assay as another approach for identifying proteins that interact with RNA probes to form complexes. The basis for this approach is that the specific interaction of an Ab molecule with the protein component of the probe–protein complex will dramatically increase the overall mass of the complex, which in turn will decrease its rate of migration during nondenaturing electrophoresis. Regarding application, binding of the Ab will cause the probe–protein complex to produce a band that is shifted upwards (supershifted) in the gel, relative to the same probe–protein complex that is Ab-free. Occasionally, binding of the Ab will generate a complex sufficiently large that it will not migrate into an 8% polyacrylamide gel (see Subheading 3.7.). Because monospecific and monoclonal antisera have been generated to most RV proteins, supershift assays are a reasonable approach for studying RNA–protein complexes formed with RV proteins. The quantity of Ab used in the assay must be empirically determined, because it will depend on the concentration of

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both the Ab and the protein, and on the strength of the affinity of the Ab for the protein. It is also critical that the antiserum be free of RNase activity, and that the binding activity of the Ab does not destabilize the probe–protein complex. Finally, it is important to realize that, although supershift assays can be used to identify a protein component of a probe–protein complex, it does not rule out the possibility that the complex may be heteromultimeric, and therefore may also contain other protein components.

3.6. Expression and Purification of VP2 CLPs from vTF7-3-Infected Cells 3.6.1. Infection and Transfection of Cells 1. The day prior to transfection, the appropriate number of 75-cm2 cell culture flasks are each seeded with 2 × 106 MA104 cells in 20 mL Medium 199 supplemented with 7% FBS, 100 U/mL penicillin, and 100 µg/mL streptomycin. The cells are incubated at 37°C in an atmosphere of 5% CO2-air, and should reach 80% confluency the following day. 2. On the day of transfection, the cell monolayers are washed with serum-free and antibiotic-free Medium 199. Afterwards, 20 mL Medium 199 containing 40 µg/mL AraC (Medium 199-AraC) is placed in each flask, and the cells are incubated for 1 h at 37°C in a CO2 incubator. AraC is an inhibitor of vaccinia virus DNA replication (38) and, in the authors’ experience, enhances the expression of recombinant proteins in vTF7-3-infected cells. 3. To prepare the virus inoculum, a vTF7-3 stock of known titer is vortexed vigorously, to disperse any clumps of material contained in the preparation (see Note 31). If this treatment is not adequate, then the stock should be sonicated for 15–30 s on ice. For each flask to be infected, 3 mL inoculum is prepared by diluting the vTF7-3 stock in Medium 199-AraC, to yield a MOI of 10 PFU/cell. Following removal of the medium from each flask, the inoculum is added, and the cells maintained for 1 h at 37°C in a CO2 incubator. During this period, the flasks are gently rocked every 10–15 min to allow the inoculum to swirl over the monolayer. 4. The production of VP2 CLPs requires a plasmid that contains a cDNA of the RV gene 2 ORF, positioned downstream from the promoter for T7 RNA polymerase. For the authors, plasmids that give the highest level of expression when transfected into cells are either purified by banding on CsCl-ethidium bromide gradients or with a Qiagen Maxi-prep DNA purification kit. About 30 min before the end of the vTF7-3 infection period, two solutions, A and B, are prepared, following the protocol provided by Life Technologies (Gaithersburg, MD) for transfecting cells with LipofectAMINE reagent. Solution A is made by adding 20 µg VP2 transcription vector to 700 µL Medium 199 containing 40 µg/mL AraC. Solution B is made by adding 100 µL LipofectAMINE reagent (Gibco) to 700 µL Medium 199, also containing AraC. Solutions A and B are combined,

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mixed gently, and then incubated for 30 min at room temperature. Afterwards, the combined solutions A and B are increased to a final volume of 6 mL by the addition of Medium 199-AraC. 5. Immediately before transfection, the vTF7-3 inoculum is removed from the flasks, and the cells are washed once with 10 mL Medium 199-AraC. The plasmid–LipofectAMINE mixture is gently layered over the cell monolayers and the flasks are incubated for 5 h at 37°C in a CO2 incubator. 6. Five h posttransfection, 6.0 mL Medium 199-AraC, containing 20% FBS, 100 U/mL penicillin, and 100 µg/mL streptomycin, are added to each flask. To inhibit the degradation of the expressed VP2, 1 µg/mL each of aprotinin and leupeptin are also added to the medium. The cells are then maintained at 37°C in a CO2 incubator. Overt cytopathic effects caused by infection of the cells with vTF7-3 virus will become apparent as early as 3–4 h postinfection. Although the infected cells will round up, the monolayer mostly will remain attached to the surface of the flask until time of harvest.

3.6.2. Purification of VP2 CLPs 1. At 24–36 h postinfection, the medium is removed from the flasks, and the cell monolayer is carefully washed with cold PBS. A rubber policeman or scraper is then used to scrape the cells into a small volume of cold PBS. 2. The cells are pelleted from the suspension by centrifugation at 1000g for 5 min at 4°C. The supernatant is removed, and the cell pellet is resuspended in 0.5 mL lysis buffer (10 mM Tris-HCl, pH 7.4, 0.1 mM EDTA, 1% DOC, 1 µg/mL each of aprotinin and leupeptin) per 75-cm2 cell culture flask harvested. The cell lysate is then incubated for 5 min at 15°C. 3. The lysate is transferred to a 1.5-mL microcentrifuge tube, and then clarified by centrifugation at 16,000g for 10 min at 4°C in a microcentrifuge. 4. The clarified lysate is layered onto 10-mL linear gradients of 5–20% sucrose (w/v) in 10 mM Tris-HCl, pH 7.4, 0.1 mM EDTA, and 0.2% DOC, and the gradients are centrifuged at 110,000g for 2 h in a Beckman SW 40.1 rotor at 4°C. 5. One-mL fractions are collected from the gradient, and the distribution of VP2 in the gradient is determined by SDS-PAGE of the fractions. Usually, VP2 forms a peak in the middle of the sucrose gradient. 6. Fractions containing VP2 are adjusted to 1 µg/mL each of aprotinin and leupeptin to inhibit protease activity, and are dialyzed against 10 mM Tris-HCl, pH 7.4, containing 0.1 mM EDTA, to remove sucrose. If necessary, VP2 in samples can be concentrated by centrifugation in an Amicon Centricon 30 microconcentrator. 7. To evaluate VP2 samples for the presence of CLPs by EM, a drop of the sample is placed on a Formvar-carbon-coated grid (EM Science, Gibbstown, NJ) and left for 1 min. The sample fluid is drawn off the grid with a piece of filter paper, and then a drop of 3% phosphotungstic acid-NaOH, pH 7.2, is added to the grid. After 1 min, the stain is removed with a piece of filter paper, and the grid is inspected for CLPs at a magnification of 50,000× (Fig. 5).

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Fig. 5. Production of VP2 CLPs and VP2-VP6 VLPs with the vTF7-3 expression system. MA104 cells were infected with vaccinia virus vTF7-3 and transfected with a VP2 transcription vector (A) alone, or in combination with a VP6 transcription vector (B and C). Particles were purified from the cell lysates, stained with phosphotungstic acid, and examined by electron microscopy. The bar is equivalent to 100 nm.

3.7. Electrophoresis Systems 3.7.1. SDS-12% Polyacrylamide Gel Electrophoresis 1. Prior to electrophoresis, protein samples are mixed with sample buffer, and are usually heated to 95°C for 3 min. (see Notes 32 and 33). 2. Electrophoresis is performed in 1X running buffer for 15 h at 30 mA for dsRNAs and 3–4 h at 60 mA for proteins. 3. Gels can be stained with ethidium bromide or silver, to detect dsRNAs, and with Coomassie blue R250 or silver, to detect proteins.

3.7.2. Urea–Polyacrylamide Gels for Purification of RNA Probes and Resolving Viral mRNAs 1. Immediately prior to electrophoresis, 32P-labeled probes in a volume of 20 µL distilled H2O are mixed with an equal volume of 2X formamide sample buffer, and are heated to 75°C for 10 min (see Note 34).

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2. The probes are electrophoresed on an 8% polyacrylamide gel containing 6 M urea (100 × 100 × 1.5 mm) in 1X TBE at 250 V, until the bromophenol dye front migrates to the bottom of the gel (2–3 h). 3. Positions of probes in wet gels are identified by autoradiography, and portions of the gel containing probes are cut out and soaked overnight at 37°C in 0.5 mL distilled H2O. 4. The eluted probes are extracted once with phenol:chloroform, and recovered by ethanol precipitation.

3.7.3. Nondenaturing Polyacrylamide Gels Used in the Gel-Shift Assays RNA–protein complexes are resolved by electrophoresis on a nondenaturing 8% polyacrylamide gel (100 × 100 × 1.5 mm) for 3–4 h at 175 V. Gels are dried and analyzed by autoradiography.

3.7.4. RNA Strand Separation Gel The plus-sense and minus-sense RNAs of RV dsRNAs can be resolved by electrophoresis on 1.75% agarose gels containing 6 M urea and 25 mM citrate buffer, pH 3.0. The details for this procedure have been published elsewhere (39). 4. Notes 1. Antibiotics (100 U/mL penicillin and 100 µg/mL streptomycin) can be included in the medium to reduce the possibility of bacterial contamination. 2. The concentration of virus in the inoculum should be sufficient to produce a MOI of 0.01–0.5 PFU per cell, depending on the growth characteristics of the virus. In general, slow-growing RVs require a higher MOI than fast-growing viruses, to obtain similar virus yields. 3. Under the conditions described in Subheading 3.1.2., CPEs are usually complete within 24–48 h of infection, especially for the well cell culture-adapted, fast-growing strains of RVs such as RRV and SA11. 4. Low-speed centrifugation is not necessarily required, if the lysate will be used for preparing CsCl-purified virus. 5. This can be accomplished by resuspending a confluent monolayer of MA104 cells (1–1.5 × 107 cells) from a 150-cm2 flask into 120 mL serum-containing medium and placing 3 mL cell suspension in each well of a six-well plate. 6. The protease inhibitors are also included in all buffers used in the purification of the virus. 7. A density of approx 1.37 g/mL can be produced by adding 2 g CsCl to 4 mL virus. 8. The upper band consists of triple-layered virions, and will have a density of 1.36 g/cm3; the lower band consists of DLPs and will have a density of 1.38 g/cm3. 9. The samples can be stored for at least 6 mo without noticeable degradation.

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10. If recovery of purified TLPs is not required, DLPs can be obtained directly from infected cell lysates as follows: Add 0.5 M EDTA (pH 8.0) to the lysates, to give a final concentration of 50 mM; extract the cell lysates with trichlorotrifluoroethane, and pellet the virus particles from the lysates, as described in Subheading 3.2.4. The pellets are resuspended in TBS, and the DLPs are purified by centrifugation in a CsCl gradient. To ensure high purity of the DLPs, an additional round of CsCl centrifugation is strongly recommended. 11. The efficiency of disruption of the VP6 layer of protein by CaCl2 decreases as the concentration of DLPs increases, and may differ, depending on the strain of virus used. Even in preparations in which CaCl2-treatment fails to remove as much as one-half of the VP6 layer of protein, the open core preparation will have a high level of replicase activity, and will lack transcriptase activity. 12. The treatment will cause the sample to become cloudy, and will eventually result in aggregation of the core particles. 13. The cores may appear as a flocculent band in the gradient, and will have a density of approx 1.45–1.50 g/cm3, depending on the amount of residual VP6 associated with the cores. 14. As shown in Fig. 1, purified DLPs should not contain detectable VP7 and VP4, or the trypsin-cleavage products of VP4, VP5*, and VP8*, and core particles should contain virtually no VP6. Although purified cores are not as stable as DLPs and TLPs, all three types of particles can be stored for several months at 4°C without loss of structural integrity. Frequently, as cores age, a specific degradation of VP2 generates a set of fragments that are slightly smaller than the wild-type protein, and which, upon SDS-PAGE, can obscure detection of VP3. Degradation of VP2 can be slowed by maintaining cores in the presence of 1 µg/mL leupeptin and aprotinin. 15. S-adenosyl-L-methionine is added to the reaction mixture, because it provides the methyl group for methylation of the cap structures added to the 5'-end of the mRNAs by guanylyltransferase. 32P-UTP can also be included in the reaction mixture, to label mRNA products. 16. If it is necessary to remove the endogenous dsRNA, the reaction mixture is centrifuged at 200,000g in a Beckman SW50.1 or TLS55 rotor for 1 h at 4°C. If necessary, the tubes can be filled by overlaying the reaction mixture with mineral oil. 17. Because of disruption of some DLPs during the transcription reaction, it is common to find some residual dsRNA contamination of the mRNA preparation. The residual dsRNA can be removed by lithium chloride precipitation (40). The quality of the mRNA preparation is assessed by electrophoresis of 1–2 µg on a 5% polyacrylamide gel containing 7 M urea (see Subheading 3.7.). A high quality preparation should contain all 11 transcripts, although the larger transcripts (of RNA segments 1–4) usually will be less abundant than the smaller transcripts. 18. Dialysis should continue until the sample has clarified (>24 h). The open cores can be used directly in replicase assays, or can be treated with micrococcal nuclease (see Subheading 3.4.1.2.) to remove endogenous dsRNA before being used in replicase assays.

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19. The concentration of protein can be determined with a Bio Rad (Hercules, CA) Protein Assay kit, or by co-electrophoresis with known amounts of BSA. The concentration of VP2 in the open core preparations is usually 100–300 µg/mL. 20. The 32P-labeled dsRNA products are detected by autoradiography, and can be quantified with a phosphorimager. Shown in Fig. 2 are the 32P-labeled dsRNA products made in reaction mixtures that contained gene 8 mRNA and untreated open cores (lane 2) or micrococcal nuclease-treated open cores (lane 4). 21. The product of assays performed with open cores that have not been treated with micrococcal nuclease, and that lack exogenous ssRNA will include low levels of 32P-labeled dsRNA (see Fig. 2, lane 1). 22. Synthesis of dsRNA will occur in the replicase assay if the open cores are replaced with purified rVP1 and rVP2 or CLPs containing rVP1 and rVP2. Synthesis of dsRNA with recombinant proteins is optimal when the molar ratio of rVP1 and rVP2 in the assay is approx 1:10 (8). The synthesis of dsRNA by rVP1 and rVP2 is much less efficient than the synthesis of dsRNA by open cores. 23. Synthesis of dsRNA by the replicase assay decreases as the level of salt in the reaction mixture increases (Fig. 3). At 200 mM NaCl, the amount of dsRNA produced in the assay will be at least 10-fold less than the amount of dsRNA produced in the control reaction mixture. 24. Formation of the VP3–GMP adduct is not affected by NaCl concentrations of 1–200 mM, but the presence of NaCl in the reaction mixture prevents the radiolabeling of VP1 with 32P-GTP (Fig. 3). 25. The 5'-terminal base of the nascent RNA can be either A or G. Like the formation of VP3–GMP intermediate complexes, the capping of RNA by open cores is a MgCl2-dependent process (Fig. 3). However, unlike the formation of VP3-GMP intermediate complexes, the capping of RNA by open cores is inhibited by the addition of NaCl to the assay. 26. Subheading 3.5.2. can be used to detect the RNA-binding activity of the RNA polymerase, VP1, and the guanylyltransferase, VP3, in open core preparations. Because both VP1 and VP3 have nonspecific affinity for RNA, any probe of 50–100 nucleotides can be used in the gel-shift assay. However, because there is evidence that VP1 also binds specifically to the 3'-end of viral mRNA, the authors generally use a probe that contains the 3'-terminal sequence of one of the viral mRNAs (19). 27. The temperature and length of incubation of the mixtures are not critical for the formation of RNA–protein complexes containing VP1 and VP3, but the authors normally incubate the mixtures for 60 min at room temperature. 28. The time of electrophoresis will vary from 3–6 h, depending on the length of the probe, and should be performed until free (unbound) probe, migrates near, but not off, the bottom of the gel. 29. Bands of probe that migrate in the gel more slowly than free probe indicate the presence of protein–probe complexes, and suggest that a protein with RNA-binding activity is contained in the reaction mixture. To verify that novel bands detected by the gel-shift assay do indeed represent complexes containing RNA and

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31.

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33.

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Patton et al. protein, reaction mixtures can be treated with 10 µg/mL RNase A and proteinase K, respectively, for 30 min at room temperature prior to electrophoresis. Crosslinking of radiolabeled nucleotides of the probe to the protein generates a 32P-labeled protein. The 32P-labeled protein may migrate slower in the gel, because the cross-linked nucleotides will add to the total mass of the protein (usually <1 kDa). The identity of the 32P-labeled protein can be verified by Western blot or immunoprecipitation assay using monospecific antiserum. UV-crosslinking has been used successfully to show that VP1, NSP2, and NSP3 have RNA-binding activity (24,26,41). BL2 experimentation: CAUTION: All procedures with vaccinia virus must be carried out using NIH-Biosafety Level 2 Guidelines. Smallpox vaccination is recommended for personnel who potentially may be exposed to vaccinia virus. VP4 and VP7 contain intramolecular disulfide bonds, which cause these proteins to migrate more rapidly when reducing agents are not added to the SDS-PAGE sample buffer (42). There is also some evidence that VP6 may also contain intrachain disulfide bonds (42). The intermediate shell of the RV triple-layered virion is formed by 260 trimers of VP6. The trimers are quite stable and are also formed by VP6 that is expressed by BV vectors containing the gene 6 cDNA (43), and by translation of the gene 6 mRNA in rabbit reticulocyte lysates (44). To detect trimers, VP6-containing preparations are incubated in SDS-PAGE sample buffer at 37°C for 5 min, and are then electrophoresed on an SDS-12% polyacrylamide gel. The trimer form of VP6 will migrate with a mol wt of 135–150 kDa. If, instead, the sample is heated to 100°C for 5 min, the VP6 trimers will be disrupted, and the VP6 will then migrate as a 45 kDa monomer upon SDS-12% PAGE. Viral mRNAs are denatured by heating in 2X formamide sample buffer. The samples are then electrophoresed on 5% polyacrylamide gels containing 7 M urea (100 × 100 × 0.75 mm) in 1X TBE running buffer at 200 V for 5 h. The mRNAs can be detected in the gel by staining with ethidium bromide or by silver staining.

References 1. Graham, D. Y. and Estes, M. K. (1980) Proteolytic enhancement of rotavirus infectivity: biological mechanisms. Virology 101, 432–439. 2. Espejo, R. T., Lopez, S., and Arias, C. (1981) Structural polypeptides of simian rotavirus SA11 and the effect of trypsin, J. Virol. 37, 156–160. 3. Cohen, J., Laporte, J., Charpilienne, A., and Scherrer, R. (1979) Activation of rotavirus RNA polymerase by calcium chelation. Arch. Virol. 60, 177–186. 4. Shirley, J. A., Beards, G. M., Thouless, M. E., and Flewett, T. H. (1981) The influence of divalent cations on the stability of human rotavirus. Arch. Virol. 67, 1–9. 5. Bican, P., Cohen, J., Charpilienne A., and Scherrer, R. (1982) Purification and characterization of bovine rotavirus cores. J. Virol. 43, 1113–1117. 6. Almeida, J. D., Bradburne, A. F., and Wreghitt, T. G. (1979) The effect of sodium thiocyanate on virus structure. J. Med. Virol. 4, 269–277. 7. Valenzuela, S., Pizarro, J., Sandino, A. M., Vasquez, M., Fernandez, J., Hernandez, O., Patton, J., and Spencer, E. (1991) Photoaffinity labeling of

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rotavirus VP1 with 8-azido-ATP: Identification of the viral RNA polymerase. J. Virol. 65, 3964–3967. Patton, J. T., Jones, M. T., Kalbach, A. N., He, Y.-W., and Xiaobo, J. (1997) Rotavirus RNA polymerase requires the core shell protein to synthesise the double-stranded RNA genome. J. Virol. 71, 9618–9626. Cohen, J. (1977) Ribonucleic acid polymerase activity associated with purified calf rotavirus. J. Gen. Virol. 36, 395–402. Chen, D. Y., Zeng, C. Q.-Y., Wentz, M. J., Gorziglia, M., Estes, M. K., and Ramig, R. F. (1994) Template dependent, in vitro replication of rotavirus RNA. J. Virol. 68, 7030–7039. Mason, B. B., Graham, D. Y., and Estes, M. K. (1980) In vitro transcription and translation of rotavirus SA11 gene products. J. Virol. 33, 1111–1121. Sandini, A. M., Jashes, M., Faundez, G., and Spencer, E. (1986) Role of the inner protein capsid on in vitro human rotavirus transcription. J. Virol. 60, 797–802. Pizarro, J. L., Sandino, A. M., Pizarro, J. M., Fernandez, J., and Spencer, E. (1991) Characterization of rotavirus guanylyltransferase activity associated with polypeptide VP3. J. Gen. Virol. 72, 325–332. Liu, M., Mattion, N. M., and Estes, M. K. (1992) Rotavirus VP3 expressed in insect cells possesses guanylyltransferase activity. Virology 188, 77–84. Desselberger, U. and McCrae, M. A. (1994) The rotavirus genome. Curr. Top. Microbiol. Immunol. 185, 31–66. Patton, J. T., Wentz, M., Xiaobo, J., and Ramig, R. F. (1996) Cis-acting signals that promote genome replication in rotavirus mRNAs. J. Virol. 70, 3961–3971. Wentz, M. J., Patton, J. T., and Ramig, R. F. (1996) The 3'-terminal consensus sequence of rotavirus mRNA is the minimal promoter of negative-strand RNA synthesis. J. Virol. 70, 7833–7841. Chen, D. and Patton, J. T. (1998) Rotavirus RNA replication requires a single-stranded 3'-terminus for efficient minus strand synthesis. J. Virol. 72, 7387–7396. Patton, J. T. (1996) Rotavirus VP1 alone specifically binds to the 3'-end of viral mRNA but the interaction is not sufficient to initiate minus-strand synthesis. J. Virol. 70, 7940–7947. Labbé, M., Baudoux, P., Charpilienne, A., Poncet, D., and Cohen, J. (1994) Identification of the nucleic acid binding domain of the rotavirus VP2 protein. J. Gen. Virol. 75, 3423–3430. Patton, J. T. and Chen, D. (1999) RNA binding and capping activities of proteins in rotavirus open cores. J. Virol. 73, 1382–1391. Shuman, S. (1995) Capping enzyme in eukariotic mRNA synthesis. Prog. Nucleic Acids Res. Mol. Biol. 50, 101–129. Patton, J. T. (1996) Structure and function of the rotavirus RNA-binding proteins. J. Gen. Virol. 76, 2633–2644. Brottier, P., Nandi, P., Bremont, M., and Cohen, J. (1992) Bovine rotavirus segment 5 protein expressed in the baculovirus system interacts with zinc and RNA. J. Gen. Virol. 73, 1931–1938. Kattoura, M., Clapp, L. L., and Patton, J. T. (1992) The rotavirus nonstructural protein, NS35, is a nonspecific RNA-binding protein. Virology 191, 698–708. Poncet, D., Aponte, C., and Cohen, J. (1994) Four nucleotides are the minimal requirement for RNA recognition by rotavirus non-structural protein NSP3. EMBO J. 13, 4165–4173. Boyle, J. F. and Holmes, K. V. (1986) RNA-binding proteins of bovine rotavirus. J. Virol. 58, 561–568.

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28. Hua, J., Chen, X., and Patton, J. T. (1994) Deletion mapping of the rotavirus metalloprotein NS53 (NSP1): The conserved cysteine-rich region is essential for virus-specific RNA binding. J. Virol. 68, 3990–4000. 29. Patton, J. T. and Hua, J. (1995) Using the RNA-capture assay to assess the RNA-binding activity of viral proteins, in Methods in Molecular Genetics, vol. 7: Molecular Virology Techniques (Adolph, A. W., ed.), Academic Press, New York, p. 373–387. 30. Poncet, D., Aponte, C., and Cohen, J. (1996) Structure and function of rotavirus nonstructural protein NSP3. Arch. Virol. 12, Suppl. S29–S35. 31. Zeng, Q.-Y., Wentz, M. J., Estes, M. K., and Ramig, R. F. (1996) Characterization and replicase activity of double-layered and single-layered rotavirus-like particles expressed from baculovirus recombinants. J. Virol. 70, 2736–2742. 32. Crawford, S. E., Labbé, M., Cohen, J., Burroughs, M. H., Zhou, Y. J., and Estes, M. K. (1994) Characterisation of virus-like particles produced by the expression of rotavirus capsid proteins in insect cells. J. Virol. 68, 5945–5952. 33. Labbé, M., Charpilienne, A., Crawford, S. E., Estes, M. K., and Cohen, J. (1991) Expression of rotavirus VP2 produces empty corelike particles. J. Virol. 65, 2946–2952. 34. Lawton, J. A., Zeng, C. Q., Mukherjee, S. K., Cohen, J., Estes, M. K., and Prasad, B. V. (1997) Three-dimentional structural analysis of recombinant rotavirus-like particles with intact and amino-terminal-deleted VP2: implications for the architecture of the VP2 capsid layer. J. Virol. 71, 7353–7360. 35. Zeng, C. Q.-Y., Estes, M. K., Charpilienne, A., and Cohen, J. (1998) The N terminus of rotavirus VP2 is necessary for encapsidation of VP1 and VP3. J. Virol. 72, 201–208. 36. Fuerst, T. R., Niles, E. G., Studier, F. W., and Moss, B. (1986) Eukaryotic transient-expression system based on recombinant vaccinia virus that synthesizes bacteriophage T7 RNA polymerase. Proc. Natl. Acad. Sci. USA 83, 8122–8126. 37. Zeng, C.Q.-Y., Labbé, M., Cohen, J., Prasad, B. V. V., Chen, D., Ramig, R. F., and Estes, M. K. (1994) Characterization of rotavirus VP2 particles. Virology 201, 55–65. 38. Cochran, M. A., Puckett, C., and Moss, B. (1985) In vitro mutagenesis of the promoter region for a vaccinia virus gene: evidence for tandem early and later regulatory signals. J. Virol. 54, 30–37. 39. Patton, J. T. and Stacey-Phipps, S. (1986) Electrophoretic separation of the plus and minus-strands of rotavirus SA11 double-stranded RNAs, J. Virol. Methods 13, 185–190. 40. Chen, D., Gombold, J. L., and Ramig, R. F. (1990) Intracellular RNA synthesis directed by temperature-sensitive mutants of simian rotavirus SA11. Virology 178, 143–151. 41. Poncet, D., Aponte, C., and Cohen, J. (1993) Rotavirus protein NSP3 (NS34) is bound to the 3’ end consensus sequence of viral mRNAs in infected cells. J. Virol. 67, 3159-3165. 42. Patton, J. T., Hua, J., and Mansell, E. A. (1993) Location of interchain disulfide bonds in VP5* and VP8* trypsin cleavage fragments of the rhesus rotavirus spike protein VP4. J. Virol. 67, 4848–4855. 43. Estes, M. K., Crawford, S. E., Penaranda, M. E., Petrie, B. L., Burns, J. W., Chan, W.-K., et al. (1987) Synthesis and immunogenicity of the rotavirus major capsid antigen using a baculovirus expression system. J. Virol. 61, 1488–1494. 44. Clapp, L. L. and Patton, J. T. (1991) Rotavirus morphogenesis: domains in the major inner capsid protein essential for binding to single-shelled particles and for trimerization. Virology 180, 697–708.

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4 Rotavirus Entry into Tissue Culture Cells Joanna M. Gilbert and Harry B. Greenberg 1. Introduction Rotavirus (RV) is a triple-protein-layered icosahedral virus, for which studies have established that the two outer-layer proteins, viral protein 4 (VP4) and viral protein 7 (VP7), are required for viral infectivity (1,2). VP7, a glycoprotein, is the major component of the outer-layer, but its role in viral entry is unclear. VP4 forms dimers extending out from the VP7-coated viral surface (3,4) and have been shown to be a determinant of host range and virulence, and is directly involved in cell attachment and RV entry into cells (5–8). Proteolytic cleavage of VP4 into two noncovalently associated subunits, VP8* and VP5* (2,9,10), significantly enhances viral infectivity (11–13). The mechanism of cell penetration during infection is not fully understood, but it probably consists of multiple steps, including binding, protease cleavage, and entry, in which binding is independent of the other two steps, and can occur without infection. VP4 is intimately involved in each of these steps of RV entry. Experiments have demonstrated that RV enters cells in a pH-independent fashion, and most, but not all, of the evidence indicates that RV penetrates cells directly through the plasma membrane rather than through the early endosomal membrane after receptor-mediated endocytosis (6,14–16). This chapter describes two different assays that measure the entry of RV into tissue culture cells: a cell-cell fusion (syncytia) assay (Fig. 1;17) and a toxin co-entry assay (Fig. 2;18–20). It is hypothesized that the ability of RV to induce both cell-to-cell fusion and toxin co-entry is related to its ability to penetrate the plasma membrane during an infection. The former assay, examining RV-induced cell fusion-from-without, allows direct visualization of the results of virus–cell interaction: RV-induced fusion of cellular plasma membranes. The toxin co-entry assay examines viral entry by determining the extent of α-sarcin From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ

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Fig. 1. Syncytia assay.

ingress into cells, concomitant with RV penetration. This toxin disrupts protein translation, and therefore allows quantitation of viral entry by measuring the decrease of protein synthesis (21,22). Both assays require cleavage of VP4 by the protease trypsin, and also depend on the presence of VP4 and VP7; particles depleted of both outer layer proteins are nonfunctioning in these assays (15–17,19,20,23). The assays can differentiate between cells that are permissive for RV infection from cells that are not permissive (19,20,23). Monoclonal antibodies, which neutralize RV infectivity, can be demonstrated to block both assays (17,19,20). Therefore, these assays provide us with methods to identify factors involved in RV entry into cells.

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Fig. 2. α-Sarcin coentry assay.

One major difficulty impeding the understanding of RV entry into cells is that a method to alter a specific RV gene product, and recover it in infectious virus, is not yet available. For molecular analysis, the authors have therefore employed VLPs (24) as an alternative to intact RV particles. VLPs are formed by co-infecting Spodoptera frugiperda 9 (Sf9) cells with four different recombinant baculoviruses (rBVs), each expressing one of the four main structural proteins of RV (VP2, VP4, VP6, or VP7). The co-expressed RV proteins assemble into particles that are similar to intact RV particles by electron microscopic, biochemical, and immunological criteria (24). The VLPs have also been shown to bind specifically to MA104 cells (24) and to function identically to intact RV particles in these entry assays (19,20,23).

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Ultimately, there are several major questions concerning the mechanism of RV entry into cells. Using recombinant VLPs, containing either VP4 or VP7 proteins engineered with specific mutations, allows the examination of the role of these proteins in RV entry at a molecular level (23). For example, recent work (25) has demonstrated that arginine residue 247 is the critical site for the proteolytic activation of VP4 required for penetration. VP4 is the primary protein involved in entry and contains two provocative sequence motifs that are hypothesized to be involved in virus penetration of host cells (26). One of these motifs is a region that has sequence similarity to the putative internal fusion peptides of Sindbis virus and Semliki Forest virus. This domain is proposed to form an amphipathic α-helix, with the hydrophobic face interacting directly to disrupt the cell plasma membrane. Using site-directed mutagenesis to reduce the hydrophobicity of this helix would allow one to examine the role of this region in RV entry. Fusion peptides have only been demonstrated to function in enveloped viruses. The second motif is a putative α-helical, coiled-coil domain that is conserved among viruses as diverse as retroviruses, alphaviruses, and paramyxoviruses (27). Using specifically mutated VLPs, whether this region is required for VP4 dimerization, or has a role in viral entry, can now be determined. Finally, it appears that there is a functional interaction between VP4 and VP7 (23,28), and they may cooperate in order for RV to penetrate host cells. The contributions of these individual proteins could be assessed using site-directed mutagenesis and one of the above entry assays. 2. Materials 1. The fetal African green monkey kidney cell line, MA104 (BioWhittaker, Walkersville, MD). 2. Dulbecco’s modified Eagle’s medium (DMEM; BioWhittaker), with 4.5 g/L glucose containing 100 IU/mL penicillin, 100 IU/mL streptomycin, 0.29 mg/mL L-glutamine (Irvine Scientific, Santa Ana, CA), 0.25 mg/mL amphotericin B (Gibco-BRL, Gaithersburg, MD), and 15, 10, 7%, or no heat-inactivated fetal bovine serum (hiFBS; Hyclone, Logan, UT), as indicated. 3. DMEM without cysteine or methionine (DMEM cys– met–; Gibco-BRL), with 4.5 g/L glucose containing 100 IU/mL penicillin, 100 IU/mL streptomycin, and 0.29 mg/mL L-glutamine. 4. Dulbecco’s phosphate buffered saline (DPBS) and DPBS calcium- and magnesiumfree (CMF; Gibco-BRL). 5. 0.5% trypsin in Versene (sodium ethylenediamine tetraacetic acid [NaEDTA], Gibco-BRL). 6. Trypsin (10 mg/mL in media without serum; Sigma, St. Louis, MO.; Type XIII, N-tosyl-L-phenylalanine chloromethyl ketone [TPCK]-treated). 7. N-α-p-tosyl-L-lysine chloromethyl ketone: 10 mM in media without serum (TLCK; Sigma). 8. 10 mM cholesterol (Sigma), freshly made in 100% ethanol.

RV Entry into Tissue Culture Cells 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25.

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α-sarcin (1 mg/mL in DMEM cys– met–, stored at –20°C; Sigma). Translabel (35S cysteine and methionine; ICN, Irvine, CA). 10% trichloroacetic acid (TCA). Absolute ethanol. 0.5% sodium dodecylsulfate (SDS) in 0.1 M sodium hydroxide (NaOH). 1 M hydrochloric acid (HCl). Aqueous scintillation fluor. The insect cell line derived from Sf9 (Invitrogen; San Diego, CA). Sf-900 II media (Gibco-BRL) supplemented with 100 IU/mL penicillin, 100 IU/mL streptomycin, 0.29 mg/mL L-glutamine, and containing either 2.5% or no hiFBS. 2X Graces media (Gibco-BRL). 2% low-melt agarose in double-distilled H2O (Gibco-BRL). The protease inhibitors pepstatin, aprotinin, and leupeptin (0.5 µg/mL; Sigma). Genentron (trichlorotrifluoroethane) (Dupont, Wilmington, DE. 10 mM Tris-HCL (Tris [hydroxymethyl] aminomethane), pH 7.4, (Dupont, Wilmington, DE) 100 mM NaCl, 5 mM CaCl 2 (TNC). 35% sucrose (w/v) in TNC. Cesium chloride (CsCl) (1.36 g/mL) in TNC. The BAC to BAC system rBV system (Gibco-BRL) containing the pFASTBAC subcloning and transposition vector and the DH10BAC Escherichia coli bacteria containing the BV genome.

3. Methods 3.1. Syncytia Assay 1. Plate 5 × 10 5 MA104 cells in 25-cm 2 flasks in 5 mL DMEM with 10% hiFBS, in an atmosphere of 5% CO 2-air at 37°C. 2. After 24 h, replace medium with 5 mL DMEM with 7% hiFBS containing 100 µM cholesterol, from the 10 mM cholesterol in ethanol stock, and incubate overnight (14–16 h) at 37°C in a CO 2 incubator (see Note 1). 3. Treat the RV or RV VLPs (see Subheading 3.3.) with 10 mg/mL trypsin at a final concentration of 20 µg/mL for 30 min at 37°C. Inactivate trypsin by adding TLCK to a final concentration of 100 µM. 4. Remove the cholesterol-supplemented cells from the flasks, first by washing with DPBS CMF, and then by treating with 1% trypsin in versene. 5. Centrifuge the cells at 1000g at room temperature. 6. Resuspend the cells in DMEM at 1 × 10 6 cells/mL and add 100 µL, containing 1 × 10 5 cells, to each 1.5-mL Eppendorf tube on ice. 7. Centrifuge samples for 30 s at 400g at 4°C, then carefully aspirate the supernatant without disturbing the cell pellet (see Note 2). 8. Resuspend the cell pellet in 100 µL virus in DMEM, on ice, at a dilution of 7.5 × 107 plaque forming units (PFU)/mL to 1 × 108 PFU/mL (approx 75–100 particles/cell), or the protein equivalent of VLPs (e.g., for a CsCl purified, titered stock of wild-type rhesus RV with a determined protein concentration

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Gilbert and Greenberg of 2.0 × 10 9 PFU/mg protein, 3.75 µg VLP protein is equivalent to approx 75 PFU for 1 × 10 5 cells). Incubate the samples on ice for 15 min, to allow the virus to bind to the cells. Centrifuge the tubes at 400g for 15 s, and then incubate at 37°C for 15 min. Gently resuspend the cell and virus pellet after addition of 400 µL DMEM. Plate the cells into six-well tissue culture dishes containing 1 mL/well DMEM with 15% hiFBS (10% hiFBS final). Incubate the plates for 2 h at 37°C in an atmosphere of 5% CO2 -air (see Note 3). Examine the wells for fusion, with an inverted phase-contrast microscope. Cells containing two or more nuclei are considered to have formed syncytia: Cells with two nuclei have undergone one fusion event; cells with three nuclei have undergone two fusion events, and so on. (see Note 4). Samples are tested in duplicate, and a minimum of 500 nuclei are counted per sample (five sets of approx 100 nuclei each) and the number of nuclei in polykaryons noted. Data are averaged and presented as number of nuclei in syncytia per 100 total nuclei counted.

3.2. α -Sarcin Assay 1. Plate 5 × 10 3 MA104 cells/well into 96-well dishes in DMEM with 10% hiFBS, and incubate at 37°C in an atmosphere of 5% CO2-air for 24–48 h (to confluency at approx 2.5 × 10 4 cells/well). 2. Aspirate the media and add DMEM cys – met –, and incubate at 37°C in an atmosphere of 5% CO2 for 1 h. 3. Treat the RVs or RV VLPs (see Subheading 3.3.) with 10 mg/mL trypsin at a final concentration of 10 µg/mL for 30 min at 37°C. Inactivate the trypsin by adding TLCK to a final concentration of 100 µM. 4. Aspirate the media, and incubate the cells with trypsin-activated virus or VLPs (at the amount or protein equivalent between 50 and 100 PFU/cell; J. Angel, personal communication) in DMEM cys – met –, in a volume of 50 µL, and add 50 µL α-sarcin in DMEM cys– met– at a final concentration of 100 µg/mL. 5. Incubate the cells at 37°C for 1 h in an atmosphere of 5% CO 2-air. 6. Remove the virus and α-sarcin, and wash the plates twice with DMEM cys – met – . 7. Overlay the cells with 100 µL DMEM cys – met – containing 0.1 µCi 35S Translabel/well, and incubate at 37°C for 1 h in an atmosphere of 5% CO 2. 8. Remove the media, and wash the plates twice with DMEM cys– met –. 9. Add 100 µL/well 10% TCA, and incubate at 4°C for 1 h (see Note 5). 10. Remove the TCA, and wash the wells twice with ethanol. Let the plates air-dry. 11. Lyse the cell monolayers with 50 µL/well 0.5% SDS in 0.1 M NaOH, and incubate at room temperature for 1 h (see Note 6). 12. Neutralize the NaOH with 5 µL 1 M HCl/well. Add the lysates to the scintillation fluor, and count samples. Data are expressed as a percent inhibition of protein synthesis, compared with control, untreated cells (see Notes 7 and 8).

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3.3. VLP Production 3.3.1. Cell Growth Sf9 cells are grown in monolayer culture in Sf-900 II media with 2.5% hiFBS in a humidified incubator at 28°C. Cells are subcultured by passaging at a 1 in 4 dilution every 3–4 d.

3.3.2. Virus Titration 1. BV stocks are grown in Sf9 cells in Sf-900 media without serum, to titers between 1 × 10 8 and 1 × 10 9 PFU/mL. To titer the virus, plate Sf9 cells into 6-well dishes at 2 × 10 6 cells/well for 1 h at 28°C in Sf-900 II media without serum (see Note 9). 2. Dilute the virus to be titered in 1.5 mL media without serum from 1 in 10–1 to 1 in 10–9 in 12 mm × 75 mm polystyrene fluorescence-activated cell sorter tubes. 3. Infect the Sf9 cells with 600 µL diluted virus mixture for 90 min at 28°C, rocking gently. 4. Aspirate media, and overlay with agar mix (1:1 2% low-melt agarose: 2X Graces’ media with 10% hiFBS), and allow the agar to set at room temperature for 20 min. Incubate at 28°C for 7–10 d, or until plaques are clearly visible (see Note 10).

3.3.3. Plaque Purification 1. To plaque-purify the virus, pick well-spaced plaques, and add them to microcentrifuge tubes containing 300 µL Sf-900 II media without serum. 2. Gently vortex the microcentrifuge tubes, and infect 0.5 × 106 Sf9 cells/well grown in 12-well dishes. Incubate the dishes at 28°C for 90 min, and then overlay with 1 mL Sf-900 II media without serum. 3. Incubate the virus and the cells until infection is complete (7–10 d). Harvest the supernatants containing the purified BV aseptically, and analyze the remaining cells for expression of the desired protein.

3.3.4. Virus Growth 1. To grow up virus, add 100 µL positive viral supernatant to a 25-cm2 flask containing 2.5 × 106 Sf9 cells in 750 µL Sf-900 II media without serum. Incubate the flask for 90 min at 28°C, rocking occasionally. 2. After incubation, overlay the cells with a final volume of 6 mL Sf-900 II media without serum. 3. Harvest the supernatant after all the cells are infected, approx 6–10 d at 28°C. Spin down the cell debris at 2000g, and store the supernatant at 4°C in the dark. For long-term storage, add 2% hiFBS, and store at –70°C. Virus stocks can be amplified from this initial stock.

3.3.5. VLP Preparation 1. Infect monolayers of Sf9 cells with the desired combination of recombinant rBVs (for the triple-layered particles required for entry studies, VP2, VP6,

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4. 5.

6.

7.

8. 9. 10.

Gilbert and Greenberg VP4, and VP7), at a multiplicity of infection (MOI) of 6, in Sf-900 II media in the absence of serum. Incubate the cells for 24–36 h at 28°C, and then remove the inoculum by aspiration. Overlay the cells with Sf-900 II media containing 0.5% hiFBS and the protease inhibitors pepstatin, aprotinin, and leupeptin (0.5 µg/mL final concentration of each). Incubate the cells at 28°C, with the addition of protease inhibitors daily until the cells are harvested at 5–6 d postinfection. Remove the cells from the flasks by shaking, and rinse the flasks with DPBS. Pellet the cells by centrifugation at 3300g for 15 min at 4°C, and save the supernatant on ice. Extract the cell pellet with Genentron, and add the aqueous layer to the supernatant on ice. Centrifuge the samples, first at 12,000g, in a JA-14 rotor (Beckman, Fullerton, CA) for 30 min at 4°C, and then at 200,000g in a Ti 45 rotor (Beckman) for 2 h at 4°C. Resuspend the pellet in TNC containing (0.5 µg/mL) aprotinin, (0.5 µg/mL) leupeptin, and (0.5 µg/mL) pepstatin, and centrifuge through a 35% (w/v) sucrose cushion in TNC at 300,000g for 90 min at 4°C. Resuspend the VLP pellet in TNC containing CsCl (1.36 g/mL), and centrifuge at 250,000g for 24–36 h at 4°C in an SW-40 rotor (Beckman). The VLP-containing bands are collected, and recentrifuged through CsCl. Collect the VLP band. The particles can be dialyzed extensively against TNC, and stored at 4°C for use in the short term. For longer-term storage, the particles should be stored in aliquots at –70°C. The VLPs are stable in CsCl at 4°C.

3.4.6. rBV Production rBVs can be easily prepared using the BAC to BAC system available from Gibco-BRL. cDNAs for VP4 and/or VP7, containing the desired changes, can be subcloned into the pFASTBAC vector. The mutant cDNAs can be transposed into the BV genome, resulting in the desired recombinant viruses. The mutant viruses can then be used in place of either wild-type VP4 or wild-type VP7 to create mutant VLPs, in order to examine the function of these mutants in RV entry. 4. Notes 1. To solubilize cholesterol in ethanol, warm the tube in a 37°C water bath, vortexing occasionally. When the cholesterol is added to the DMEM with 7% hiFBS, the media will become slightly cloudy. 2. If cells have been treated with too much cholesterol, they will not pellet at this g force, nor will they plate well after syncytia formation, making quantitation difficult. 3. If cells are left too long after plating (longer than 3.5 h), they will begin to die. 4. It is vital to include, as controls, cells alone, and cells incubated with untrypsinized virus, to establish the background levels of syncytia formation. Cells alone will usually have an average of three nuclei in syncytia/100 nuclei

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8.

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counted. An average of 4.5 nuclei in syncytia/100 nuclei counted can be expected with cells treated with untrypsinized virus. For trypsin-treated virus at an MOI of 75, the level of syncytia seen is approx 18 nuclei in syncytia/100 nuclei counted. Samples in TCA can be left overnight at 4°C. Samples in 0.5% SDS/0.1 M NaOH can be left overnight at 4°C. It is vital to include as controls, cells alone, cells incubated with α-sarcin alone, cells with trypsinized virus without α-sarcin, and cells incubated with untrypsinized virus with α-sarcin, to establish the background levels. Translation is expected to be diminished by 90% using virus at an MOI between 75 and 100. An alternative procedure is to examine the loss of translation using SDS polyacrylamide gel electrophoresis (PAGE). The assay should be scaled up to 24-well dishes. After the cells are labeled and washed, they are lysed in buffer containing 130 mM NaCl, 20 mM Tris (pH 7.4), 1 mM EDTA, 1.0 % Nonidet P-40, and 1 mM phenylmethyl sulfonylfluoride, and cleared of cellular debris by centrifugation at 13,000g for 15 min. The lysates are boiled in an equal volume of sample buffer (0.25 M Tris-HCl, pH 7.4, 30% sucrose, 8% SDS, and 50 mM dithiothreitol) for 5 min. These samples are run on SDS-PAGE gels, and examined by autoradiaography (F. Liprandi, J. Angel, and N. Shulman, personal communications). It is vital that cells are at the appropriate density, or they will overgrow and make detection of plaques impossible. It is crucial that the final concentration of agarose is not greater than 1%. The temperature of the agarose overlay should also not be greater than 28°C, because Sf9 cells are temperature-sensitive.

References 1. Bridger, J. C. and Woode, G. N. (1976) Characterization of two particle types of calf rotavirus. J. Gen. Virol. 31, 245–250. 2. Estes, M. K., Graham, D. Y., Smith, E. M., and Gerba, C. P. (1979) Rotavirus stability and inactivation. J. Gen. Virol. 43, 403–409. 3. Prasad, B. V. V., Burns, J. W., Marietta, E. Estes, M. K., and Chiu, W. (1990) Localization of VP4 neutralization sites in rotavirus by three-dimensional cryo-electron microscopy. Nature 343, 476–479. 4. Shaw, A. L., Rothnagel, R., Chen, D., Ramig, R. F., Chiu, W., and Prasad, B. V. V. (1993) Three-dimensional visualization of the rotavirus hemagglutinin structure. Cell 74, 693–701. 5. Greenberg, H. B., Flores, J., Kalica, A. R., Wyatt, R. G., and Jones, R. (1983) Gene coding assignments for growth restriction, neutralization and subgroup specificities of the W and DS-1 strains of human rotavirus. J. Gen. Virol. 64, 313–320. 6. Kaljot, K. T., Shaw, R. D., Rubin, D. H., and Greenberg, H. B. (1988) Infectious rotavirus enters cells by direct cell membrane penetration, not by endocytosis. J. Virol. 62, 1136–1144. 7. Offit, P. A., Blavat, G., Greenberg, H. B., and Clark, H. F. (1986) Molecular basis of rotavirus virulence: role of gene segment 4. J. Virol. 57, 46–49.

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8. Ruggeri, F. M. and Greenberg, H. B. (1991) Antibodies to the trypsin cleavage peptide VP8 neutralize rotavirus by inhibiting binding of virions to target cells in culture. J. Virol. 65, 2211–2219. 9. Espejo, R. T., Lopez, S., and Arias, C. (1981) Structural polypeptides of simian rotavirus SA11 and the effect of trypsin. J. Virol. 37, 156–160. 10. Lopez, S., Arias, C. F., Bell, J. R., Strauss, J. H., and Espejo, R. T. (1985) Primary structure of the cleavage site associated with trypsin enhancement of rotavirus SA11 infectivity. Virology 144, 11–19. 11. Babiuk, L. A., Mohammed, K., Spence, L., Fauvel, M., and Petro, R. (1977) Rotavirus isolation and cultivation in the presence of trypsin. J. Clin. Microbiol. 6, 610–617. 12. Barnett, B. B., Spendlove, R. S., and Clark, M. L. (1979) Effect of enzymes on rotavirus infectivity. J. Clin. Microbiol. 10, 111–113. 13. Clark, S. M., Roth, J. R., Clark, M. L., Barnett, B. B., and Spendlove, R. S. (1981) Trypsin enhancement of rotavirus infectivity: mechanism of enhancement. J. Virol. 39, 816–822. 14. Fukuhara, N., Yoshie, O., Kitaoka, S., and Konno, T. (1988) Role of VP3 in human rotavirus internalization after target cell attachment via VP7. J. Virol. 62, 2209–2218. 15. Nandi, P., Charpilienne, A., and Cohen, J. (1992) Interaction of rotavirus particles with liposomes. J. Virol. 66, 3363–3367. 16. Ruiz, M. C., Alonso, T. S., Charpilienne, A., Vasseur, M., Michelangeli, F., Cohen, J., and Alvarado, F. (1994) Rotavirus interaction with isolated membrane vesicles. J. Virol. 68, 4009–4016. 17. Falconer, M. M., Gilbert, J. M., Roper, A. M., Greenberg, H. B., and Gavora, J. S. (1995) Rotavirus-induced fusion-from-without in tissue culture cells. J. Virol. 69, 5582–5591. 18. Carrasco, L. (1994) Entry of animal viruses and macromolecules into cells. FEBS Lett. 350, 151–154. 19. Cuadras, M. A., Arias, C. F., and Lopez, S. (1997) Rotaviruses induce an early membrane permabilization of MA104 cells and do not require a low intracellular Ca2+ concentration to initiate their replication cycle. J. Virol. 71, 9065–9074. 20. Liprandi, F., Moros, Z., Gerder, M., Ludert, J. E., Pujol, F. H., Ruiz, M. C., Michelangeli, F., Charpilienne, A., and Cohen, J. (1997) Productive penetration of rotavirus in cultured cells induces coentry of the translation inhibitor α-sarcin. Virology 237, 430–438. 21. Brigotti, M., Rambelli, F., Zamboni, M., Montanaro, L., and Sperti, L. (1989) Effect of α-sarcin and ribosome-inactivating proteins on the interaction of elongation factos with ribosomes. Biochem. J. 257, 723–727. 22. Endo, Y. and Wool, I. G. (1982) The site of action of α-sarcin on eukaryotic ribosome. J. Biol. Chem. 257, 9054–9060. 23. Gilbert, J. M. and Greenberg, H. B. (1997) Virus-like particle induced fusion-from-without in tissue culture cells; role of outer-layer proteins VP4 and VP7. J. Virol. 71, 4555–4563.

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24. Crawford, S. E., Labbé, M., Cohen, J., Burroughs, M. H., Zhou, Y. J., and Estes, M. K. (1994) Characterization of virus-like particles produced by the expression of rotavirus capsid proteins in insect cells. J. Virol. 68, 5915–5922. 25. Gilbert, J. M. and Greenberg, H. B. (1998) Cleavage of rhesus rotavirus VP4 after arginine 247 is essential for rotavirus-like particle-induced fusion-from-without. J. Virol. 72, 5323–5327. 26. Mackow, E. R., Shaw, R. D., Matsui, S. M., Vo, P. T., Dang, M.-N., and Greenberg, H. B. (1988) The rhesus rotavirus gene encoding protein VP3: Location of amino acids involved in homologous and heterologous rotavirus neutralization and identification of a putative fusion region. Proc. Natl. Acad. Sci. USA 85, 645–649. 27. Buckland, R. and Wild, F. (1989) Leucine zipper motif extends. Nature 338, 547–548. 28. Mendez, E., Arias, C. F., and Lopez, S. (1996) Interactions between the two surface proteins of rotavirus may alter the receptor-binding specificity of the virus. J. Virol. 70, 1218–1222.

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5 Mixed Infections with Rotaviruses Protocols for Reassortment, Complementation, and Other Assays Robert F. Ramig 1. Introduction Mixed infection of tissue culture cells is the primary means of studying genetic and nongenetic interactions between viral mutants. The purpose of the mixed infection is to place two different viral genomes into the same cell, where interactions between the genomes and their encoded gene products can take place. In some cases, interactions are observed in the context of the mixed infected cell, but generally the yield of progeny virus from the mixed infected cells is assayed to reveal the result of the interactions. Here, methods for mixed infections between conditional-lethal mutants of the temperature-sensitive (ts) phenotype will be presented and considered. (Mutants with ts phenotype are designated according to the following nomenclature: tsA [778], in which ts indicates the phenotype, A indicates the mutant belongs to recombination group A, and [778] indicates the number of the specific mutant in group A.) Temperature-sensitive mutants are particularly useful for genetic studies, because they can be propagated at the permissive temperature (PT; 31°C in the case of rotavirus [RV] ts mutants), but the mutant phenotype is expressed at the nonpermissive temperature (NPT; 39°C in the case of RV ts mutants). The conditional-lethality of the ts mutations allows the use of selective conditions in the analysis of the progeny of the mixed infections, so that the results of genetic and nongenetic interactions can be easily revealed.

1.1. Mixed Infections for Recombination (Reassortment) Tests Along with other members of the Reoviridae, RVs have a segmented genome (1), which results in a high frequency of recombination by the mechanism of From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ

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reassortment of genome segments. Thus, mutations that reside on noncognate segments are expected to reassort at a high frequency; mutations that reside on cognate segments are expected to be unable to reassort. The all-or-none nature of recombination in the RVs confirms reassortment as the recombination mechanism (2,3). In recombination tests, one determines if two different mutant genomes can interact to form viable genomes with new combinations of genetic material. To allow the genomes of mutants tsA and tsB to interact, the mixed infection is performed at PT, so that the mutants can replicate and increase the size of the intracellular genome pools which, in turn, increases the probability of recombinational interaction. The formation of wild-type (WT, also called ts+; RV WT grows at both 31 and 39°C) recombinant genomes from the two ts mutant genomes is generally assayed, because this progeny class is easily detected among the yield (progeny virus that results from the replication of the parental viruses during a mixed or single infection) of the cross, which also contains the parental ts mutants (used to initiate mixed infections) and the tsA–tsB double mutant. The WT recombinant class is detected at NPT at which neither the ts parental viruses nor the double mutant recombinant can grow. The recombination test is used to divide the ts mutants of RV into groups. Mutations within a group fail to generate WT reassortants in mixed infection, but mutations in different groups generate WT reassortants at high frequency (4–8). Because genome segments of the Reoviridae contain single genes, all mutations on a single genome segment (reassortment group) reside in the same gene or in its cis-acting regulatory sequences.

1.2. Mixed Infections for Complementation Tests Complementation tests are performed to determine if two mutant viruses contain lesions in the same function. With ts mutants, the mixed infection of tsA with tsB is performed at NPT, so that the mutant phenotype is expressed. At NPT in the mixed infection, neither the tsA or tsB parental virus can replicate, unless tsB can provide the function that is mutant in tsA and tsA can provide the function that is mutant in tsB. This requires that the mutant function in tsA can be supplied in trans by the nonmutant function of tsB (and vice versa), and results both in growth of mutants and the production of a yield that is primarily the two mutants. Transcomplementation is detected by assaying the yield at PT, and documenting an increase in yield over the infection of each parent alone. Failure of the mixed infection to increase the yield indicates that both mutants contained lesions in the same function. The segmented nature of the RV genome, with each segment encoding a single gene product, suggests that the gene products function in trans, and that complementation should be efficient. However, complementation is not useful

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for assignment of RV mutants to functional groups (3), because results of complementation analyses are generally negative and irreproducible (5). The failure of RV mutants to complement is ascribed to the interference phenotype associated with all of the mutants (6,7).

1.3. Interference Tests Mixed infections between a ts mutant and WT at either PT or NPT are used to determine if the ts mutant has an interference phenotype (6,7). The fact that all ts mutants tested interfered with the growth of WT suggests that ts mutants also interfere with each other in mixed infections, which is thought to account for failure to complement (see Subheading 1.2.). Interference tests are performed to determine if a virus mutant interferes with growth of another virus. Interference tests are done between a ts mutant and WT virus, because interference can be easily detected as a decrease in yield of WT from the mixed infection, compared to a control infection of WT alone. It is much more difficult to determine the interference of one ts mutant, with another ts mutant, because neither parental virus can be easily scored. To determine the interference phenotype of a ts mutant, mixed infections are performed between the ts mutant and WT, together with control infections of the ts mutant and WT alone (6,7). The mixed infections are incubated at either 31 or 39°C, depending on the condition at which the interference phenotype is of interest. The yield is determined at 39°C, at which only WT (ts+) virus will grow. An interfering mutant reduces the growth of the WT virus in the mixed infection, compared to the control single infection of WT. Interference tests are often performed at equal and unequal multiplicity of infection (MOI), and an interfering mutant generally shows a stronger interference phenotype when it dominates the input mixture (6,7). (MOI is the number of plaque-forming units [PFU]/cell used to initiate an infection). PFUs are the number of infectious units present in a virus stock, measured by their ability to form plaques on cell monolayers; generally expressed as PFU/mL. The number of PFU/mL in a virus stock is the titer of that stock, and is nearly always determined at PT.)

1.4. Generation of Reassortant Viruses Mixed infections between WT RV strains are used to generate reassortant viruses that contain defined constellations of genome segments derived from the two parental strains. The resulting reassortants are useful in studies to map expression of various phenotypes to specific genome segments or constellations of genome segments (see refs. 2 and 3 for examples). The methods for generation and selection of reassortant viruses are described in detail elsewhere (9).

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1.4.1. Nonselective Mixed Infection for Random Isolation of Reassortant Progeny Viruses Reassortant viruses, with carefully defined constellations of genome segments derived from two different virus strains, are usually generated by mixed infection of cultured cells. For these infections, it is best to use WT, nonmutagenized viruses as parents, because viral mutants selected following mutagenesis often contain unknown mutations that can affect the phenotype of reassortants in unpredictable ways (10). Because the parental viruses are WT, both the mixed infections and subsequent plaque assays for isolation of reassortant progeny are performed at the nonselective temperature of 37°C. The absence of selective markers in the mixed infections, and the absence of selective pressures during isolation of progeny reassortant, means that a large number of randomly chosen progeny plaques must be screened to identify the reassortant with the segment constellation of interest. When screening progeny from a mixed infection for reassortants, the host cell used can affect the spectrum of reassortant constellations isolated. Evidence has been presented that the host cell used for plaque formation can affect the spectrum of reassortant constellations isolated (11,12). Thus, a failure to isolate the desired reassortant constellation from a large number of screened plaques may be remedied by changing the cell line from which the plaques for screening are picked. Alternatively, a genome segment that is absent or underrepresented may be increased in representation in progeny by the use of unequal MOI for the mixed infections (11).

1.4.2. Mixed Infection with Selection for Reassortants with Defined Segment Constellation Often, mixed infections are performed with the intention of isolating reassortants with a specific constellation of parental genome segments. This often occurs when generating RV reassortants with viral protein VP4 and VP7 neutralization proteins in heterologous combination, relative to either parental virus. In such a case, a standard mixed infection is performed using the nonselective temperature of 37°C. The plaque assay from which plaques will be picked is overlaid with agar containing a neutralizing monoclonal antibody (MAb) directed at the VP4 or VP7 that one does not want in the reassortants. This antibody (Ab) selection during plaque formation selects against one of the parental viruses, and against reassortants that contain the protein being selected against. This significantly reduces the number of progeny plaques that must be screened to identify the reassortant with the

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segment constellation of interest. However, caution must be exercised in the use of such selective techniques, because reassortant viruses may express phenotypes that are not consistent with parental origins of the proteins (3). In one study (13), in a cross between RV strains SA11-4F and B223, a MAb directed against VP4 of SA11-4F was used to select reassortants with VP4 from the B223 parent. However, all of the reassortants isolated from this selection contained VP4 from SA11-4F (13). When the reassortant of interest was isolated using nonselective conditions (e.g., without selecting Ab), it was found to be neutralized by the Ab directed against SA11-4F VP4, although the reassortant contained VP4 from B223. Conversely, the reassortant containing SA11-4F VP4 was not neutralized by the Ab as expected (13). This unexpected presentation of neutralizing epitopes was demonstrated to result from interactions between VP4 and VP7 outer capsid proteins of heterologous parental origin (14). This example emphasizes the problems that may be encountered in the use of selective conditions when making reassortants with specific constellations of genome segments.

1.5. Mapping RV Mutants Mixed infections between RV ts mutants (15), or between a ts mutant and WT virus of a different strain (16), are used to map the ts mutations to genome segments. The ts phenotype of the ts parent(s) is counterselected to simplify identification of the ts+ reassortant progeny useful for mapping (15,16).

1.6. Mixed and Single Infections for Kinetic Analyses If mixed or single infections are desired for a kinetic analysis, the protocol outlined here is easily adapted. The preparation of vials for infection and preparation of activated virus stocks for infection are scaled-up, and multiple vials are infected with each single virus or virus mix from a single inoculum preparation. Vials are then incubated at the appropriate temperature, and individual vials of each single or mixed infection are harvested at the appropriate time postinfection. Single infections harvested in kinetic fashion have been used to generate highly reproducible one-step growth curves, and mixed infections have been used to examine the kinetics of reassortment (5–7). 2. Materials The sources and catalog numbers in the following list indicate items and sources the author has used with consistent success over the past 20 yr. Only with the 2-dram vials is the specific source and catalog number important. BioWhittaker (Walkersville, MD) is the only commercial source for MA104 cells.

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2.1. Cells and Viruses 1. African Green monkey kidney MA104 cells; (BioWhittaker, cat. no. 75-104F). For optimal results, MA104 cells should be used at passages lower than 100–120. 2. High-titer stocks of RV ts mutants and WT.

2.2. Equipment 1. Tissue culture incubators (Forma [Marietta, OH], Model 3110), humidified and flushed with 5% CO2-air. One each at 39, 37, and 31°C. 2. Water bath at 37°C. 3. Probe-type sonication device equipped with a microprobe for small volumes (Braun-Sonic [Melsungen, Germany] Model U, equipped with 3 mm diameter microprobe). Probe is sterilized between uses by immersion in virus killer (0.5 gm KI, 1.0 gm I 2 , dissolved in 1000 mL 95% ethanol) for 10 s; excess virus killer wiped off with tissue, immersion in sterile H 2O to briefly rinse. 4. Vacuum aspirator with trap. 5. Inverted microscope. 6. Hemocytometer.

2.3. Reagents 1. Medium 199 + fetal bovine serum (FBS). (Medium 199 with Earle’s salts, Irvine Scientific [Santa Ana, CA], cat. no. 9466) Resuspended and sterilized according to manufacturer’s instructions. The medium is then completed by making the following additions: 0.3 g/L glutamine; 1.22 g/L NaHCO3; 0.16 g/L penicillin; 0.25 g/L streptomycin; and 5% [v/v] FBS. 2. Medium 199 without FBS. Medium 199 completed as above, except FBS is omitted. 3. 2× Medium 199. Medium 199 with Earle’s salts resuspended at twice the concentration recommended by the manufacturer. For use, 2× 199 is supplemented with 0.6 g/L glutamine; 2.0 g/L NaHCO3; 0.16 g/L penicillin; 0.25 g/L streptomycin; 0.025 g/L gentamicin; 5000 U/L nystatin; and pancreatin (135 trypsin-activity U/L). 2× 199 should be supplemented with NaHCO 3 immediately prior to use, because precipitates rapidly form as complete medium is stored. 4. 2% Bacto-agar (Difco, Detroit, MI, cat. no. 0140-01): 20 g/L in distilled H2O and sterilized by autoclaving. 5. Gelatin-saline: 8.0 g/L NaCl, 0.03 g/L CaCl2, 0.17 g/L MgCl2, 1.2 g/L H3BO3, 0.05 g/L Na2B4O7, 5.0 g/L gelatin (pH 7.2). 6. 1 mg/mL trypsin, (Worthington, Freehold, NJ) dissolved in 0.001 N HCl, and stored in single-use 1.0-mL aliquots at –80°C. 7. Pancreatin (Oxoid, London, UK) One tablet dissolved in 50 mL distilled H2O, filter sterilized. Trypsin activity determined and frozen in 1.0-mL aliquots until use. Added to 2× 199 at the rate of 135 trypsin activity U/L. 8. Gentamicin (Sigma) at a final concentration of 0.025 g/L in 2× 199. 9. Nystatin (Gibco-BRL, Gaithersburg, MD) at a final concentration of 5000 U/L in 2× 199.

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10. Two-dram vials (8 mL): flat bottom, borosilicate glass, with rubber-lined screw cap. (Wheaton, Millville, NJ, cat. no. 224884). Unwashed vials directly from manufacturer are loosely capped, wrapped in butcher paper, and sterilized by autoclaving for 20 min on a drying cycle. 11. Sterile 0.1 and 1.0 mL pipets or automatic pipeters with sterile tips. 12. Sterile Pasteur pipets. 13. Six-well sterile tissue culture cluster plates (Costar, Costar, NY, cat. no. 3506).

3. Methods The methods and procedures outlined here are specifically for reassortment tests using ts mutants. Variations in procedure for complementation tests appear as notes throughout Subheadings 3.3.–3.9., as appropriate. Variations in procedure for other tests using mixed infection are covered in Subheadings 3.10.1.–3.10.4.

3.1. General Considerations Several general considerations, taken into account in the development of this protocol, are outlined below. 1. Optimal mixed infections require mixed infection of all the cells in the population, so that the viral genomes and gene products have maximal opportunity for interaction. This requires knowledge of the number of cells present in the monolayer. The protocol below is designed to produce MA104 cell monolayers for infection, with, on average, 1 × 105 cells/monolayer. However, the number of cells in the average monolayer, under the specific conditions of the laboratory, should be confirmed by trypsinizing several monolayers, and counting the cells in a hemocytometer. The cell monolayers are formed on the flat bottoms of 2-dram vials, so that they contain a relatively small number of cells. The small number of cells minimizes the use of virus stocks, and allows the use of virus stocks with titers as low as 2 × 107 PFU/mL. 2. The prerequisite for efficient mixed infections is infection of each cell with both parental viruses. The percentage of cells mixed infected with two different parental viruses is given by the expression: (1 – e–mA)(1 – e–mB) × 100

(1)

where mA and mB are the MOI of parental viruses A and B. Theoretically, an MOI of 5 PFU/cell of each parental virus will infect 98.7% of the cells, and, as a practical matter, an MOI of 10 PFU/cell of each parental virus will mixed infect virtually 100% of the cells in the monolayer. An equal, high MOI of each parental virus should always be used for mixed infections. The use of lower MOI, so that not all cells are mixed and infected, reduces the opportunity for interaction between the parental viruses, and progressively reduces the frequency of reassortment as the MOI is decreased (7). The use of unequal MOI of the two parental viruses also reduces reassortment frequency (11; and R. F. Ramig,

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unpublished observations). This protocol is based on infection with an MOI of 10 PFU/cell of each parental virus in mixed infections. In control, single infections, the MOI is increased to 20, so that the total MOI remains constant, negating the multiplicity effects seen with some mutants. 3. The high MOI used for mixed infections should limit the infections to a single cycle. One-step growth determinations for WT and ts mutants indicate that the growth cycle is complete within 32 h at 31°C, and more rapidly at 37 or 39°C (5,6). Studies of reassortment kinetics (7) indicate that reassortant viruses can be detected as early as 12 h postinfection, and that maximal frequencies of reassortment are rapidly attained within 16–24 h postinfection. However, slight increases in reassortment frequency are observed at times later than 24 h postinfection. Studies with asynchronous mixed infections demonstrate that infection with the second virus can be delayed as much as 24 h, compared to infection with the first virus, and reassortants are still obtained (17). Asynchronous co-infection has even been shown to increase the recombination frequency in cases of one parent replicating to higher titers and forming larger plaques than the other parent (18). These observations emphasize the need to incubate the infections to completion, to allow optimal and complete interaction of the parental viruses. This protocol utilizes a 48-h period of incubation of mixed and control infections, to ensure that all viruses have completed their infectious cycle. 4. Components contained in FBS inhibit the growth of RVs. As a result, MA104 cell monolayers are formed in the presence of FBS, and the FBS must be washed out of the system before initiating the infections. This protocol minimizes manipulations by simple removal of the serum-containing medium and replacement with serum-free medium. The residual inhibitory serum components are metabolized by the cells during an 18–24 h incubation after the medium change.

3.2. Experimental Time-Scale Mixed infections require planning, so that MA104 cell monolayers in the proper medium, and at the proper cell density, are available at the appropriate time. The following is the schedule for cell preparation and performance of the mixed infections: 1. D – 2. MA104 cells in Medium 199 + FBS are seeded into 2-dram vials and incubated at 37°C to form monolayers. 2. D – 1. The medium on the MA104 cells is changed to Medium 199 without FBS, and the cells are incubated at 37°C. 3. D – 0. The monolayers are inoculated with virus mixtures to initiate the infections, and are incubated at the appropriate temperature (31°C for reassortment tests). 4. D + 2. The infected monolayers are harvested. Infected cells are harvested by tightly capping the vials, and freezing them at –20°C until they can be analyzed.

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3.3. Preparation of MA104 Cell Monolayers for Infection Monolayers of MA104 cells must be prepared in advance of infection. This protocol yields monolayers that contain, on average, 1 × 105 cells in medium without serum, which are ready for infection. 1. A confluent monolayer of MA104 cells in a 150-cm2 tissue culture flask is trypsinized and suspended in 100 mL Medium 199 + FBS, using standard tissue culture techniques (see Note 1). 2. Sterile 2-dram glass vials are seeded with 1.0 mL MA104 cell suspension, and loosely capped. Always seed 3–5 more vials than are needed for the number of crosses and controls to be performed. 3. The loosely capped vials are incubated at 37°C in an incubator with an atmosphere of 5% CO2-air, for approx 24 h, to allow the cells to form a confluent monolayer. Monitor the growth of the cells using an inverted microscope at low magnification. Removing the cap of the vial may facilitate viewing the monolayer (see Note 2). 4. When the cell monolayers are confluent (generally 24 h), the medium is removed from each vial by aspiration, using a sterile Pasteur pipet (see Note 3). 5. Each vial is re-fed with 1.0 mL Medium 199 without FBS, loosely capped, and returned to the 37°C, 5% CO2 incubator. 6. After 18–24 h incubation at 37°C, to allow serum components to be metabolized, the monolayers are inspected using an inverted microscope. If the monolayers are intact and confluent, they are ready to use for infections. 7. Before use, the monolayers in 2–3 vials should be trypsinized and counted in a hemocytometer, to establish the average number of cells/monolayer. Once the procedure is established, one assumes a constant number of cells/vial (see Note 4).

3.4. Preparation of Virus Stocks at Titers Necessary for Protocol This protocol assumes that: MA104 cell monolayers to be infected contain 1 × 105 cells; the desired MOI is 10 PFU/cell of each parental virus for mixed infections and 20 PFU/cell for single-infection controls; that virus mixes for mixed infection will be made with 0.1 mL of each parental virus, and 0.2 mL virus for single-infection controls; that virus mixes and single viruses will be activated with an equal volume (0.2 mL) of trypsin prior to infection; and that infections will be initiated with 0.2 mL activated virus or virus mix. 1. The number of PFU required for each parental virus to obtain the desired MOI: (No. cells)(PFU/cell) = (1 × 105 cells)(10 PFU/cell) = 1 × 106 PFU required (2) (see Notes 5 and 6). 2. Because the number of PFU desired must be in 0.1 mL, the titer of virus stock to contain that number of PFU in PFU/mL is calculated: (PFU/0.1 mL)(10) = (1 × 106 PFU/0.1 mL)(10) = 1 × 107 PFU/mL needed

(3)

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Table 1 Preparation of Virus Stocks at Proper Titer for Use in Mixed Infections For final titer of 2 × 10 7 PFU/mL Virus mutant

Mutant titer at PT (PFU/mL)

Volume mutant virus stock (mL)

Volume 199 w/o FBS (mL)

1.20 × 108 1.55 × 108 1.80 × 108 2.95 × 108 1.05 × 108 3.00 × 108

0.167 0.129 0.111 0.068 0.190 0.067

0.833 0.871 0.899 0.932 0.810 0.933

tsA(778) tsB(339) tsC(606) tsD(975) tsA(2289) WT

3. To adjust for dilution of the virus or virus mix with an equal volume of trypsin for virus activation, the required titer of stock virus must be multiplied by two: (PFU/mL)(2) = (1 × 107 PFU/mL)(2) = 2 × 107 PFU/mL required virus titer (4) (see Note 7). 4. RV stocks generally have titers higher than 2 × 10 7 PFU/mL and must be diluted to that concentration for use in this protocol. The amount of high-titer virus stock diluted to a total volume of 1.0 mL is calculated as follows (a sample virus dilution sheet is shown in Table 1): PFU/mL needed PFU/mL stock

=

2.0 × 10 7 PFU/mL needed 5.5 × 10 8 PFU/mL stock

=

0.037 mL diluted (5) to 1.0 mL total

3.5. Preparation and Activation of Virus Mixes and Single Viruses for Infections After virus stocks are prepared at the proper titer for use in the experiment, virus mixes and single viruses are prepared and activated as described below (a sample preparation sheet is shown in Table 2): 1. Small sterile tubes are numbered to match the control and mixed infections (Table 2), and chilled on ice. 2. Control viruses and virus mixes are prepared by adding the amounts of virus stock (prepared at a titer of 2.0 × 10 7 PFU/mL, as in Subheading 3.4.) to the individual tubes, as indicated in Table 2. Use a sterile pipet for each addition, to prevent cross-contamination of tubes. 3. Ten µg/mL trypsin is prepared by diluting 0.1 mL trypsin stock (1 mg/mL) into 9.9 mL Medium 199 without FBS. 0.2 mL 10 µg/mL trypsin is added to each of the control or virus mix tubes (Table 2) (see Note 8). Use a sterile pipet for each addition to prevent cross-contamination of tubes.

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Table 2 Preparation of Virus Mixes, Activation, and Inoculation Volumes for Performing Mixed and Control Infections Volume parental virus (2 × 10 7 PFU/mL) to make virus mixes

Tube 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17

Cross (Parent 1 ⊗ Parent 2)a

Parent 1 (mL)

Parent 2 (mL)

10 µg/mL trypsinb (mL)

Volume to inoculate (mL)

tsA(778) alone tsB(339) alone tsC(606) alone tsD(975) alone tsA(2289) alone WT alone tsA(778) ⊗ tsB(339) tsA(778) ⊗ tsC(606) tsA(778) ⊗ tsD(975) tsA(778) ⊗ tsA(2289) tsB(339) ⊗ tsC(606) tsB(339) ⊗ tsD(975) tsB(339) ⊗ tsA(2289) tsC(606) ⊗ tsD(975) tsC(606) ⊗ tsA(2289) tsD(975) ⊗ tsA(2289) Uninfected

0.2 0.2 0.2 0.2 0.2 0.2 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.2 c

– – – – – – 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.1 –

0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2

0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2

a

For viruses alone for control infections, the virus is parent 1; for virus mixes for mixed infections, the left-hand virus is parent 1 and the right-hand virus is parent 2. b After addition of trypsin, the virus mixes and single-control viruses are incubated at 37°C for 30 min, to activate infectivity. c For the uninfected cell monolayer, the mock inoculum consists of 0.2 mL 199 w/o FBS.

4. The tubes with virus and trypsin are incubated for 30 min in a water bath at 37°C, for activation of viral infectivity. 5. After activation, the tubes are chilled on ice until use for inoculation of MA104 cell monolayers. Activated viruses should be used for infection within 1–2 h after activation.

3.6. Control and Mixed Infection of Cell Monolayers Prior to initiating the infections, cell monolayers have been prepared and average cell number determined, as described in Subheading 3.3. The day the infections are to be performed, virus stock dilutions are prepared as described

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in Subheading 3.4. Finally, the virus mixtures are prepared and trypsin activated, as described in Subheading 3.5. When these steps have been completed, the infections can be initiated, as described below. 1. Vials containing monolayers of MA104 cells are numbered to match the numbers of the control and mixed infections (Table 2). The authors find it convenient to use 0.5-inch round self-adhesive labels (Avery, Diamond Bar, CA, cat. no. 05406 or equivalent) placed on the cap of each vial. 2. At room temperature, the medium is removed from a group of vials (approx 10 vials per group) by aspiration with a sterile Pasteur pipet (see Note 9). 3. A volume of 0.2 mL from the appropriate virus mix tube is added to the matched vial containing the MA104 cell monolayer, and the vial is loosely capped. Moderate care is taken to ensure that the bottom of the vial is level, so that the inoculum covers the monolayer. 4. Steps 2 and 3 are repeated until all the vials have been inoculated. 5. The inoculated vials are placed in a 31°C incubator with an atmosphere of 5% CO2-air, and incubated for 1 h to allow virus to adsorb to the cells. Again, care should be taken that the bottoms of the vials are relatively level (see Notes 10 and 11). 6. After the adsorption period, move the vials to the work space at room temperature. Again working with small groups of approx 10 vials, remove the inoculum from each vial by aspiration, using a sterile Pasteur pipet. Use a separate, sterile Pasteur pipet for each vial, to prevent cross-contamination (see Note 12). 7. To each individual vial, add 1.0 mL Medium 199 without FBS, using a separate, sterile pipet for each vial, and loosely cap the vial. 8. Steps 6 and 7 are repeated until all the vials have been fed with fresh medium (see Note 13). 9. Place the vials in an incubator with an atmosphere of 5% CO2-air at the proper temperature. With ts mutants as used in this example, the vials are incubated at 31°C for reassortment tests (see Note 14). 10. Incubate the vials with infected monolayers for 48 h. Cytopathic effects will generally be complete by 48 h, using this protocol. 11. Harvest the infected cells by tightening the cap of each vial, and place them in a freezer at –20°C. The frozen vials can be stored until it is convenient to titer the resulting infected cell lysates.

3.7. Analysis of Yields of Progeny Virus from Mixed and Control Infections The yields of progeny virus from mixed infections for reassortment tests with ts mutants are assayed for infectivity using plaque assays at 31 and 39°C. The protocols for plaque assays have been described several times (5,9,19,20), and will be described only briefly here. 1. MA104 cells (1 × 105 cells/mL) in Medium 199 + FBS are seeded into the wells of six-well cluster plates, 2 mL/well. The plates are incubated at 37°C in 5% CO2-air, until the cells form confluent monolayers.

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2. When the cells have formed confluent monolayers, the medium is removed from the wells and replaced with Medium 199 without FBS (2 mL/well), and incubated 12–18 additional hours, to allow metabolism of the residual serum components inhibitory to RV plaque formation. 3. Immediately before performing the plaque assay, the frozen yields of the mixed infections are thawed and briefly sonicated, to disrupt intact cells and virus aggregates. Serial 10-fold dilutions of the infected cell lysates are prepared using gelatin-saline as diluent (see Note 15). 4. The medium is removed from the wells of the serum-deprived confluent MA104 cell monolayers, and the wells are inoculated (0.1 mL/well) with appropriate dilutions of the mixed infection yield. 5. The plates are incubated at 31°C for 1 h, to allow virus adsorption. 6. After adsorption, the infected monolayers are overlaid with 3 mL/well of nutrient medium made by mixing equal volumes of 2% water agar and 2× 199. 7. The cluster plates are incubated at the appropriate temperature for the appropriate time in 5% CO2 incubators (see Note 16). 8. At the appropriate time, each well of the monolayer is overlaid a second time with 1.0 mL/well neutral red medium made by mixing 2% water agar with 2× 199, and adding 25 µg/mL neutral red. The plates are returned to the incubator for 12–18 h further incubation, to allow viable cells to take up the neutral red, revealing the plaques. 9. The plaques present on the plates are counted, and the counts recorded (see Table 3 for sample data).

3.8. Calculation of Results from Mixed Infections for Reassortment Assays Sample data from mixed infections between RV ts mutants, generated using the protocol described here, are shown in Table 3. This assay was performed to determine reassortment frequencies between the mutants shown in Tables 1 and 2. The crosses were performed at 31°C, so that the viruses could replicate and their genomes could interact. The progeny were assayed at 31 and 39°C, and plaque counts for the duplicate wells of each dilution plated are shown. The titer of the yield determined at 39°C for the mixed infections represents the sum of WT (ts+) reassortants, ts+ revertants, and ts+ pseudorevertants, for the single infection controls, the titer at 39°C represents ts+ revertants and ts+ pseudorevertants. Subtracting the 39°C titers of the two relevant single infection controls from the 39°C titer of the mixed infection provides a good estimate of the number of WT (ts+) reassortants in the yield of the mixed infection. The titer of the mixed infections at 31°C represents the total yield of the infection, and includes ts parentals, ts double-mutants, WT (ts+) reassortants, ts+ revertants, and ts+ pseudorevertants. The frequency of WT (ts+) reassortants is calculated from the following formula, in which the superscript indicates the temperature of the assay, to determine the titer of the progeny (see Note 17).

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Table 3 Titration Data and Recombination Frequencies

Plate

Cross (Parent 1 ⊗ Parent 2)

Dilution plated Temp (°C)

10–2

10–4

10–5

10–6

Titer (PFU/mL)

EOPd or % ts+ Recomb.e

92

1

tsA(778) alone

39 31

17/15a –

0/0 C/Cb

0/0 51/54

– 9/8

1.60 × 104 5.25 × 107

3.05 × 10–4

2

tsB(339) alone

39 31

0/4 –

0/0 C/C

0/0 32/37

– 2/3

2.00 × 103 3.45 × 107

5.80 × 10–5

3

tsC(606) alone

39 31

39/44 –

0/0 C/C

0/0 55/64

– 10/8

4.15 × 104 5.95 × 107

6.97 × 10–4

4

tsD(975) alone tsA(2289) alone

6

WT alone

7

tsA(778) ⊗ tsB(339)

8

tsA(778) ⊗ tsC(606)

9

tsA(778) ⊗ tsD(975)

C/C – C/C – C/C – C/C – C/C – C/C –

7/5 C/C 2/1 C/C C/C C/C 57/62 C/C 57/59 C/C 43/53 C/C

0/0 T/Tc 0/0 T/T 68/84 C/C 6/6 T/T 6/7 T/T 4/3 T/T

– 8/12 – 8/8 – 16/13 – 11/17 – 10/10 – 14/8

6.00 × 105 1.00 × 108 1.50 × 105 8.00 × 107 7.60 × 107 1.45 × 108 5.95 × 106 1.40 × 108 5.80 × 106 1.00 × 108 4.80 × 106 1.10 × 108

6.00 × 10–3

5

39 31 39 31 39 31 39 31 39 31 39 31

1.88 × 10–3 0.52 4.25 5.74

Ramig

3.80

tsA(778) ⊗ tsA(2289)

11

tsB(339) ⊗ tsC(606)

12

93

39 31 39 31

C/C – C/C –

3/2 C/C 33/28 C/C

0/0 T/T 2/3 72/54

– 14/9 – 6/10

2.50 × 105 1.15 × 108 3.05 × 106 6.30 × 107

0.07

tsB(339) ⊗ tsD(975)

39 31

C/C –

38/43 C/C

6/6 73/74

– 7/10

4.05 × 106 7.35 × 107

4.69

13

tsB(339) ⊗ tsA(2289)

39 31

C/C –

16/11 C/C

1/0 51/59

– 5/12

1.35 × 106 5.50 × 107

2.18

14

tsC(606) ⊗ tsD(975) tsC(606) ⊗ tsA(2289)

16

tsD(975) ⊗ tsA(2289)

17

Uninfected

C/C – C/C – C/C – 0/0 –

48/54 C/C 28/23 C/C 44/43 C/C 0/0 0/0

5/6 T/T 2/2 T/T 4/6 T/T 0/0 0/0

– 14/18 – 9/9 – 8/6 – 0/0

5.10 × 106 1.60 × 108 2.55 × 106 9.00 × 107 4.35 × 106 7.00 × 107 – –

2.79

15

39 31 39 31 39 31 39 31

a

No./no. indicates numbers of plaques in duplicate wells of six-well cluster plate.

b

C/C indicates so many plaques that monolayer was totally destroyed.

c

T/T indicates plaques too numerous to count because of overlap.

d

EOP is titer at 39°C/ titer at 31°C; shown in italics (Plates 1–6).

e%

4.77

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2.62 5.14 –

Recomb. indicates recombination frequency calculated as described in Subheading 3.8.; shown in bold (Plates 7–16).

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(6)

The following points should be noted from this reassortment data (Table 3): 1. Each of the mutants retains a ts efficiency of plating EOP < 0.1 during growth at 31°C for the control infections. 2. The WT control virus retains a ts+ EOP > 0.1 during growth at 31°C for the control infection. 3. The mixed-infection tsA(778) ⊗ tsA(2289) failed to yield a significant proportion of ts+ reassortant progeny virus. This indicates that tsA(778) and tsA(2289) reside on cognate genome segments of the parental viruses, so that reassortment cannot generate ts+ progeny. Indeed, data of this type is used to place both mutants in reassortment group A. All other crosses yield significant proportions of ts+ reassortant progeny, indicating that each mutant pair contains lesions on different genome segments, so that ts+ progeny can be generated by reassortment. Indeed, these mutants are assigned to reassortment groups A, B, C, and D, based on these results.

3.9. Calculation of Results from Mixed Infections for Complementation Assays If the crosses shown in Table 3 had been performed for complementation analysis, the mixed infections would have been incubated at 39°C, at which the mutant gene products of the mutants would have been expressed (see Note 18). Expression of mutant phenotype at 39°C would prevent the mutant viruses from growing, unless they were mutant in different transacting functions. If the mutant pair is mutant in different transacting functions, each could provide the function missing in the other, resulting in a yield of mutant virus. The progeny yields of the mixed infections are assayed at 39°C, at which ts+ reassortants, ts+ revertants, and ts+ pseudorevertants are the only progeny classes that would grow. Subtracting this background of wild phenotype from the total yield of the mixed infection determined at 31°C, provides a good estimate of the titer of mutant virus yielded by the mixed infection. This number is compared to the ability of each of the parental mutants to grow alone at 31°C. The complementation index is calculated as follows (where superscripts indicate the temperature of the titration): Complementation Index (CI)

=

(Titer A ⊗ B)31 – (Titer A ⊗ B)39 (Titer A)31 + (Titer B)31

(7)

Complementation analysis is of limited use with RV mutants (see Note 19). When pairs of mutants are subjected to complementation tests, the CI is quite low and irreproducible. The complementation data cannot be used to group RV

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mutants into self-consistent groups, regardless of the level of complementation assumed to be significant (3). 4. Notes 1. This monolayer preparation protocol is designed to allow convenient intervals between steps. If monolayers are not confluent within 24 h, a larger volume of cell suspension can be seeded, or the monolayers can be allowed more time for growth to confluence. 2. 24 h is the minimal time for formation of the monolayers. If shorter times are allowed for monolayer formation, the cells often release from the bottom of the vial when the inoculum-containing trypsin is added. 3. The monolayers can be washed free of serum by several changes of medium, and immediately used. However, this increases handling and the chances of contamination, compared to the protocol as outlined. 4. Cell monolayer preparation is identical, whether the cells will be used in reassortment, complementation, or other types of tests. 5. If a different MOI is desired, appropriate changes in calculations must be made. 6. A high MOI of 10 PFU/cell is used for reassortment, complementation, and other types of tests, unless MOI is an experimental variable. Occasionally, one encounters RV mutants that cannot be grown to the minimum titer of 2 × 107 PFU/mL required for the protocol as outlined here. In such cases, two alternative approaches can be taken. The first is preferable, because it allows crosses involving the mutant in question to be performed under conditions identical to other crosses, allowing direct comparison of the results. In this case, large 100–250 mL lysates of infected cells are prepared using standard techniques for virus propagation. The lysate is extracted with Genetron (Fisher, Pittsburgh, PA) to remove lipids, and the resulting supernatant is pelleted through a sucrose cushion. The pellet is then resuspended in a minimal volume (0.5–1.0 mL), titered, and, if the partially purified virus has a titer of 2 × 107 or greater, it is used to perform the cross, using the standard technique. If the low-titer mutant virus cannot be concentrated to sufficient titer, the second approach can be used. In this approach, asynchronous mixed infections are performed, infecting first with the low-titer mutant, and allowing several hours (1–4 h) for amplification of the virus before infecting with the second virus. The asynchronous infections are performed as described (17). The disadvantage of this alternative is that the low-titer virus is often used at an MOI significantly less than that required to infect all of the cells. The second virus is used at an MOI of 10, so that all cells infected with the first virus will be infected with the second virus. The result is that all cells infected with the low-titer virus are also infected with the high-titer virus, but, in addition, a significant number of cells are infected with the high-titer parental virus alone. The results obtained are, at best, qualitative, and should not be compared to infections performed at MOI 10 for each parent where quantitative differences may be meaningful. Control mixed infections can be performed (using mutants of sufficient titer to also be used in

96

7.

8.

9. 10.

11.

12.

13.

Ramig the standard protocol) at unequal MOI, using asynchronous infection of the two mutants, to determine the effects of the unusual format of the assay. This approach has been used in the closely related bluetongue virus (11). For the protocol presented here, with trypsin activation of viruses prior to infection, each virus stock is required at a titer of 2 × 107 PFU/mL. Stocks with higher titer must be diluted to a concentration of 2 × 10 7 PFU/mL prior to preparation of virus mixes. This protocol results in a final concentration of trypsin during activation of 5 µg/mL. This amount of trypsin is consistent with activation of the viruses, and does not cause the cells to release from the monolayer during the absorption period, as often occurs if higher trypsin concentrations are used for activation. With some virus strains, lower concentrations of trypsin can be used, and full activation of infectivity will still be obtained. Preparation and activation of virus mixes and controls is identical for reassortment, complementation, and other types of tests. Small groups of vials are infected at a time, to prevent drying of the monolayer during prolonged periods with no medium. Occasionally, the monolayer releases from the substratum and rounds up into a small ball of cells during the adsorption period. If this occurs, the experiment can be salvaged if the inoculum is removed very carefully, so that the ball of cells remains in the vial. The vial is then fed normally with medium, and cytopathic effects are observed to proceed as usual. The ts mutants in this laboratory collection were selected using a 1 h adsorption period at 31°C, followed by growth or plaque formation at 39°C, so that mutants with lesions in cell attachment would not be selected (5,6). Thus, 31°C should always be used for adsorption of virus to host cells, because any mutants with lesions in binding would fail to infect the cells. When removing the inoculum from the infected monolayer after adsorption (step 6), it is critical that a new sterile pipet be used to remove the inoculum from each vial to prevent cross-contamination of vials. For the same reason, a new sterile pipet should be used to feed each vial (step 7). In this protocol, unadsorbed virus is not removed by washing the monolayer or adding neutralizing Ab. The majority of the unadsorbed virus is removed when the inoculum is aspirated from the vial, and the residual unadsorbed parental virus does not introduce an unacceptable background to the experiment. If high backgrounds of unadsorbed virus cause unacceptable background, one can wash the monolayers prior to feeding and incubating for growth. The use of neutralizing Ab to remove unadsorbed virus is not recommended, because residual Ab may neutralize a significant proportion of the progeny virus from the infected cells. The medium used to feed the infected cell monolayers does not contain trypsin. Because the cells are infected at high MOI, cell-to-cell spread of virus is not required for the infection to proceed to completion, so addition of trypsin is not required. In practice, the authors find that some residual trypsin remains in the vial, so that the progeny virus yield is mostly activated without the addition of exogenous trypsin. This facilitates titration of the yields in subsequent plaque assays.

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14. For reassortment tests, the mixed infections are incubated at 31°C, to allow the mutants to replicate and increase the size of the genome pool available for interaction. For complementation tests, the mixed infections should be incubated at 39°C, so that gene products with mutant phenotype are expressed. 15. For mixed infections performed as described here, titration of the following dilutions has been found to yield wells with countable numbers of plaques. At 39°C, dilutions 10–2, 10–4, and 10–5 are plated. At 31°C, dilutions 10–4, 10–5, and 10–6 are plated. 16. The titrations at 39°C are incubated for 5 d, and then overlaid with neutral red late on d 5. They are counted on d 6 following inoculation. The titrations at 31°C are incubated for 9 d, and then overlaid with neutral red late on d 9. They are counted on d 10 following inoculation. 17. As with many quantitative assays in biology, reassortment tests are complicated by the issue of determining the level of significance. The reassortment behavior expected of viruses with segmented genomes is all-or-none. If reassortment is random, one would expect 25% of the progeny of a mixed infection to be ts+ and, as can be seen from the data in Table 3, this high frequency is not attained with RV mutants that reassort. The lower-than-expected frequency of reassortment in RVs has been ascribed to the interference property of the mutants with WT virus (3,6,7), so that less-than-expected amounts of ts+ progeny are generated. On the other hand, as can be seen in Table 3, when reassortment is not expected, the frequency is seldom zero, but some value greater than zero. In practice, reassortment tests yielding frequencies of ts+ reassortants between 0.2 and 1.0% are repeated several times. Nearly always, one obtains an average frequency of reassortment that is either less than 0.2% or greater than 1.0% when the mixed infections are repeated (5), and this has been used as the level of significance for negative (<0.2%) and positive (>1.0%) tests. Using this criterion, self-consistent grouping of RV mutants by reassortment has been possible (4–8). 18. For complementation tests, each of the ts mutants should be used in a mixed infection with WT, as well as alone and in mixed infection with other ts mutants. The mixed infection of ts mutant (WT serves as a control, to demonstrate that the ts mutant can be complemented in trans by the cognate gene product of WT. 19. Complementation tests are not particularly useful with RV ts mutants. The complementation indices obtained with RV SA11 ts mutants are low, highly irreproducible, and cannot be used to place mutants into self-consistent complementation groups (3,5,6). Interference tests performed by mixed infections of ts mutant and WT viruses showed that the ts mutants interfered strongly with the growth of WT virus (5,6). It is assumed that ts mutants also interfere with the growth of other ts mutants, so that complementation is not observed, although this has not been directly tested. If RV ts mutants lacking an interference phenotype were available, they would probably complement as expected. This was shown to be the case in the closely related reovirus, in which ts mutants that do not interfere show normal complementation, but those that do interfere fail to complement (21).

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Acknowledgments The work in the author’s laboratory, which led to development and refinement of this protocol, was supported by research grants from the National Science Foundation (PCM80002359), and from the Institute of Allergy and Infectious Diseases, National Institutes of Health (AI16687, AI21494, AI36385). References 1. Fields, B. N. (1996) Reoviridae, in Field’s Virology, 3rd ed., (Fields, B. N., Knipe, D. M., Howley, P. M., et al., eds.), Lippincott-Raven, Philadelphia, pp. 1553–1555. 2. Ramig, R. F. and Ward, R. L. (1991) Genomic segment reassortment in rotaviruses and other reoviridae. Adv.Virus Res. 39, 163–207. 3. Ramig, R. F. (1997) Genetics of the rotaviruses. Annu. Rev. Microbiol. 51, 225–255. 4. Greenberg, H. B., Kalica, A. R., Wyatt, R. W., Jones, R. W., Kapikian, A. Z., and Chanock, R. M. (1981) Rescue of noncultivable human rotavirus by gene reassortment during mixed infection with ts mutants of a cultivable bovine rotavirus. Proc. Natl. Acad. Sci. USA 78, 420–424. 5. Ramig, R. F. (1982) Isolation and genetic characterization of temperature-sensitive mutants of simian rotavirus SA11. Virology 120, 93–105. 6. Ramig, R. F. (1983) Isolation and genetic characterization of temperature-sensitive mutants that define five additional recombination groups in simian rotavirus SA11. Virology 130, 464–473. 7. Ramig, R. F. (1983) Factors that affect genetic interaction during mixed infections with temperature-sensitive mutants of simian rotavirus SA11. Virology 127, 91–99. 8. Faulker-Valle, G. P., Clayton, A. V., and McCrae, M. A. (1982) Molecular biology of rotaviruses. III. Isolation and characterization of temperature-sensitive mutants of bovine rotavirus. J. Virol. 42, 669–677. 9. Chen, D. and Ramig, R. F. (1994) Construction and characterization of rotavirus reassortants. Methods Mol. Genet. 4, 183–194. 10. Rubin, D. H. and Fields, B. N. (1980) Molecular basis of reovirus virulence: Role of the M2 gene. J. Exp. Med. 152, 853–868. 11. Ramig, R. F., Garrison, C., Chen, D., and Bell-Robinson, D. (1989) Analysis of reassortment and superinfection during mixed infection of Vero cells with bluetongue virus serotypes 10 and 17. J. Gen. Virol. 70, 2595–2603. 12. Graham, A., Kudesia, G., Allen, A. M., and Desselberger, U. (1987) Reassortment of human rotavirus possessing genome rearrangements with bovine rotavirus: nonrandomness and evidence for host cell selection. J. Gen. Virol. 68, 115–122. 13. Chen, D., Burns, J. W., Estes, M. K., and Ramig, R. F. (1989) The phenotypes of rotavirus reassortants depend upon the recipient background. Proc. Natl. Acad. Sci. USA 86, 3743–3747. 14. Chen, D., Estes, M. K., and Ramig, R. F. (1992) Specific interactions between rotavirus outer capsid proteins VP4 and VP7 determine expression of a crossreactive, neutralizing VP4-specific epitope. J. Virol. 66, 432–439.

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15. Gombold, J. L. and Ramig, R. F. (1987) Assignment of simian rotavirus SA11 temperature-sensitive mutant groups A, C, F, and G to genome segments. Virology 161, 463–473. 16. Gombold, J. L., Estes, M. K., and Ramig, R. F. (1985) Assignment of simian rotavirus SA11 temperature-sensitive mutant groups B and E to genome segments. Virology 143, 309–320. 17. Ramig, R. F. (1990) Superinfecting rotaviruses are not excluded from genetic interactions during asynchronous mixed infections in vitro. Virology 176, 308–310. 18. Tauscher, G. I. and Desselberger, U. (1997) Viral determinants of rotavirus pathogenicity in pigs: Production of reassortants by asynchronous coinfection. J. Virol. 71, 853–857. 19. Matsuno, S., Inouye, S., and Kono, R. (1977) Plaque assay of neonatal calf diarrhea virus and the neutralizing antibody in human sera. J. Clin. Microbiol. 5, 1–4. 20. Smith, E. M., Estes, M. K., Graham, D. Y., and Gerba, C. P. (1979) A plaque assay for the simian rotavirus SA11. J. Gen. Virol. 43, 513–519. 21. Chakraborty, P. R., Ahmed, R., and Fields, B. N. (1979) Genetics of reovirus: the relationship of interference to complementation and reassortment of temperature-sensitive mutants at nonpermissive temperature. Virology 94, 119–127.

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6 Pathogenesis and Animal Models Linda J. Saif and Lucy A. Ward 1. Introduction 1.1. Rationale for Use of Gnotobiotic Pigs in Rotavirus Research Because of the limitations in studying human rotavirus (HRV) pathogenesis and mucosal immunity in the natural host (infants and children), various animal models have been utilized to investigate rotavirus (RV) disease pathogenesis and immunity. Mice and rabbits serve as useful models to evaluate and dissect immune responses to RV (see Chapter 9). However, because older mice (>14 d) and rabbits are not susceptible to diarrhea after inoculation with either homologous (murine or lapine, respectively) or heterologous (human) RVs, assessment of protective immunity is restricted to prevention of virus shedding only (1–3). Gnotobiotic pigs are susceptible to infection and disease induced by several HRVs, until at least 6 wk of age (4,5). Moreover, a virulent strain of HRV (Wa) induced intestinal lesions (villous atrophy and crypt hyperplasia) in gnotobiotic pigs (6), resembling those seen in children with natural symptomatic RV infections (7). Further advantages of gnotobiotic pigs for HRV research include the following: Colostrum-deprived pigs are devoid of maternal antibodies (Abs) at birth (the placental type in swine acts as a barrier to the transfer of maternal Abs), but are immunocompetent, thus permitting an assessment of true primary immune responses to RV (4,5,8,9), and then determination of the impact of defined titers or specificities of maternal Abs on immunity to RV (10–12); the gnotobiotic environment ensures that exposure to extraneous RVs or other enteric pathogens is not a confounding variable; gnotobiotic piglets closely resemble human infants in gastrointestinal physiology (monogastrics), mucosal immune development, size, and milk diet. From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ

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1.2. Accomplishments in RV Research Resulting from Gnotobiotic Pig Studies The gnotobiotic pig model has contributed to several important advances in RV research, including adaptation of the first HRV (Wa strain) to cell culture after passage and amplification in piglets (13); delineation of the independent roles of the RV outer capsid viral proteins VP4 and VP7 in induction of neutralizing Abs and cross-protection (14); recognition of the potential role for a nonstructural protein (NSP4), besides VP4 and VP7, in RV virulence (15); and comparative pathogenesis of virulent and attenuated strains of HRV in the intestine, leading to the recognition that factors other than villous atrophy may contribute to the early diarrhea induced by virulent HRV in pigs (6) or homologous virulent RVs in gnotobiotic pigs (16) and calves (17). These latter findings, including a recently proposed role for RV NSP4 as a viral enterotoxin (18), may reveal new pathogenic mechanisms of viral diarrhea. In order to design and improve RV vaccination strategies, correlates of protective immunity against HRV-induced diarrhea have been analyzed in gnotobiotic pigs inoculated orally with virulent HRV (mimic natural infection), or live attenuated or inactivated HRV vaccines (4,5,8,9). Complete protection against RV diarrhea and virus shedding was induced by virulent RV and significant correlations were observed between the numbers of intestinal immunoglobulin A (IgA) antibody-secreting cells (ASC), intestinal IgA Ab titers and intestinal lymphoproliferative responses to RV, and the degree of protection. Such results agree with findings by others that natural symptomatic and asymptomatic RV infections in infants in the first year of life conferred a high rate of protection against subsequent diarrhea (19,20) and that pre-exposure fecal IgA Abs to RV in children remaining uninfected during a RV season were significantly higher than in symptomatically infected children (21). Thus, the gnotobiotic pig studies have established key parameters and correlates related to protective immunity, several of which were also observed in children, and have confirmed the applicability of this model to examine new strategies for the improvement of HRV vaccines for infants.

1.3. Pathogenesis of Group A HRV in Gnotobiotic Pigs Studies on RV pathogenesis need to be performed using well-defined inoculation doses expressed as the median infective dose (ID50) or median diarrheal dose (DD50) for a given virus strain within a given host species. The ID50 and DD50 of a virulent (infant stool passaged in pigs) HRV (Wa strain, P1A[8],G1) in gnotobiotic pigs was found to be extremely low (1 focus-forming unit [FFU]), indicating high susceptibility of the gnotobiotic pig to this virus (6). Inoculation doses of 105 ID50 (or 105 DD50) were found to elicit clinical disease in 100% of 3–5-d-old gnotobiotic pigs by post-inoculation h (PIH) 24,

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and were subsequently used for pathogenesis studies. Morphologic evaluation of intestinal and selected lymphoid tissues were done at the onset of diarrhea or PIH 13, during virus shedding (PIH 24–96), and after recovery from diarrhea or postinoculation d (PID) 7. Virulent Wa HRV induced diarrhea and morphologic lesions typical of homologous (porcine) RV infections (16,22,23). Macroscopic changes are characterized by mild thinning of intestinal walls, reduced chyle within mesenteric lymphatics, mesenteric lymph node enlargement, and reduction in large (spiral) colon mass. Microscopic changes include loss of normal mature absorptive epithelial cells, detachment of absorptive cells from their basement membranes (especially over the tips of the villi of the small intestine), and mild lymphoreticular hyperplasia within villous tip stroma, followed by villous atrophy and crypt hyperplasia. Lymphoreticular hyperplasia becomes widespread and more pronounced after PIH 72, with a concurrent, although transient, increase in the number of intestinal intraepithelial lymphocytes. By PID 7, diarrhea subsides and villous morphology is within normal histologic limits, with only lymphoid hyperplasia persisting. Virus antigen (Ag) distribution in intestinal tissues, like lesion distribution, is nonuniform. Few infected cells (<1–8%) are detected in the duodenum, but they are numerous (30–50%) within the jejunum and ileum. Small amounts of virus Ag are transiently present extraintestinally, within mesenteric lymph nodes at PIH 24, presumably as a result of Ag uptake by resident phagocytic cells at the peak of virus replication. The large amount of virus Ag and infectious virus within the intestines from PIH 24 to 72 results from lysis of the infected villous epithelial cells (leading to villous atrophy), which forms the basis of the proposed pathogenesis for virulent Wa HRV in gnotobiotic pigs (6). The number of cells that are infected, and presumably functionally impaired, is large and these cells are rapidly lysed. Thus, the mucosa is unable to efficiently compensate or recover sufficiently to maintain minimal capacity for normal digestion and absorption. Some absorption probably takes place in the large bowel, but it is not able to compensate for the functional loss incurred by the small intestine of the young gnotobiotic pig. During the initial phase of lesion progression, rapid cellular loss may also permit leakage of plasma proteins and electrolytes from the mucosa, which further contributes to the movement of fluid to the lumen, and to the subsequent diarrhea. The absence of significant morphologic changes at the onset of diarrhea, which also has been observed in pigs infected with porcine RVs (16,22,23), suggests that other factors (in addition to cell lysis and mucosal disruption) are contributing to the early disease expression following virulent Wa HRV infection in gnotobiotic pigs. A recent report suggests a possible role for NSP4 as a viral enterotoxin (18), and studies of RV reassortants in gnotobiotic pigs have shown that replacement of the NSP4 gene of a virulent porcine RV strain with

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the corresponding gene of an attenuated HRV strain yielded a reassortant that failed to induce diarrhea (15), thus implicating specific viral factors in disease expression. The role of host factors, such as activated T-cells and cytokines, in disease expression in gnotobiotic pigs is under active investigation by the authors. The pathogenesis of attenuated (serially passaged in cell culture) Wa strain HRV was also examined in gnotobiotic pigs (6). Because inoculation doses of >107 FFU resulted in shedding in only a few pigs (6%), and no diarrhea in any pigs, but seroconversion in 100% of pigs, the ID50 of this virus was calculated as the median seroconversion dose (SD50). Subsequently, morphologic changes associated with the attenuated virus were assessed following inoculation with 107 FFU (102 SD50). No macroscopic lesions were observed, and microscopic changes were minimal and transient, with a loss of normal absorptive cell vacuolation throughout the duodenum and jejunum from PIH 13 to 24. Lymphoid hyperplasia was also evident in mesenteric lymph nodes and Peyer’s patches from PIH 72 to PID 7. The authors’ findings suggest that infection takes place following oral inoculation with the attenuated virus strain; however, the inability to detect intestinal viral Ag, and the absence of significant histologic lesions, suggests that very few intestinal cells become infected initially, or that replication of the virus is inefficient or possibly even abortive. The attenuation process (which consisted of serial propagation in cell culture) probably leads to changes in the virus genome that ultimately influence virulence and pathogenicity. Several RV gene segments and their products have been implicated in the pathogenicity of RV in pigs, including the structural gene products, VP4, VP3, and VP7, and the nonstructural gene product, NSP4 (15,24–26). Clearly, more studies are needed to elucidate the mechanisms that account for the pathogenic differences seen between attenuated and virulent HRVs.

1.4. Studies of Immunity to HRVs in Gnotobiotic Pigs 1.4.1. Passive Immunity Gnotobiotic pigs were used to evaluate the efficacy of passively administered circulating (serum) and/or local colostrum/milk maternal (swine) or heterologous (bovine) Abs for preventing HRV-induced diarrhea (10–12,27). Cows were immunized with HRV serotypes P1A,G1 and P1B,G2 and simian RV SA11 (P[2],G3), and the immune colostrum was collected (27). Feeding of Ab concentrates from the immune bovine colostrum effectively reduced RV shedding and diarrhea, in a dose-dependent manner. Pigs protected against HRV-induced disease developed active immune responses in the presence of the heterologous, passive colostral Abs. In recent experiments, the effects of colostrum/milk (mimic breast-fed infants) and/or circulating serum maternal Abs (mimic non-breast-fed infants)

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from sows immunized with Wa HRV (immune, high Ab titer) or nonimmunized seropositive sows (control, low Ab titer), on passive protection and development of active immunity to RV were assessed in neonatal gnotobiotic pigs inoculated and challenged with HRV (10–12). At birth, pigs received immune or control circulating serum maternal Abs (via intraperitoneal injection), with or without daily immune or control colostrum/milk dietary supplements. The effect of maternal Abs on protection was assessed by analysis of diarrhea and virus shedding after inoculation and challenge, and on B-cell responses by quantitation of ASC by enzyme-linked immunospot (ELISPOT) and Abs by enzyme-linked immunosorbent assay (ELISA), and on T-cell responses by determination of lymphoproliferative assay (LPA) responses. Immune responses were examined in intestinal and systemic lymphoid tissues or secretions at selected PID and post-challenge days (PCD), as described previously (5,8). Pigs that received immune maternal Abs were at least partially protected from diarrhea and virus shedding after inoculation, compared to pigs receiving control or no maternal Abs, but had higher rates of diarrhea and virus shedding after challenge. Higher susceptibility to diarrhea after challenge coincided with lower numbers of IgG and IgA ASC in intestinal tissues, lower serum and intestinal IgA Ab titers, and lower LPA intestinal responses at challenge, compared to pigs receiving control or no maternal Abs. Thus, high titers of maternal Abs in the circulation or diet of neonatal pigs conferred at least partial passive protection against virulent HRV, but led to decreased active immune responses. Such decreased immune responses in the presence of high titers of maternal Abs could impact oral HRV vaccine efficacy, especially in developing countries.

1.4.2. Active Immunity Studies of immunity to HRV in gnotobiotic pigs have focused on the identification of correlates of homotypic protection (4,5,8,9). The viral inocula selected were designed to mimic immunity induced by natural infection (virulent Wa RV), or to assess immunity induced to live attenuated, or inactivated Wa RV vaccines. Neonatal piglets were orally inoculated with virulent, attenuated or inactivated (with 10% binary ethylenimine) RV and challenged at PID 21 with homologous virus. Pigs were examined daily for diarrhea and fecal RV shedding, by ELISA and cell culture immunofluorescence assays after inoculation and challenge. Correlates of protective immunity were evaluated by ELISPOT (B-cell responses) and LPA (T-cell responses) on systemic (blood, spleen) and intestinal (gut lamina propria, mesenteric lymph node) lymphoid tissues collected at selected PIDs and PCDs. After inoculation, almost all pigs exposed to virulent RV developed diarrhea, and all shed virus, and seroconverted with neutralizing Abs to Wa RV. None of the pigs given attenuated or inactivated RV developed diarrhea (moderate to severe) and only

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6% of pigs given attenuated RV shed virus, but 96–100% of pigs in both groups seroconverted. After challenge, most virulent virus-inoculated pigs were protected from diarrhea (87% protection rate) and virus shedding (100% protection), compared to protection rates against diarrhea of 31% for attenuated virus-inoculated pigs and 0 and 6% for inactivated virus-inoculated pigs (administered orally or parenterally, respectively). At challenge, the mean numbers of IgA ASC in the intestinal lamina propria were significantly greater in virulent virus-inoculated pigs (53 ASC/5 × 105 mononuclear cells [MNC]) than in attenuated (6 ASC/5 × 105 MNC) or inactivated (3.5 ASC/5 × 105 MNC) virus-inoculated pigs, and were significantly correlated with the degree of protection. Furthermore, the transient appearance of IgA ASC in the blood of virulent virus-inoculated pigs mirrored the IgA responses in the gut, suggesting that IgA ASC in the blood of children could serve as a surrogate marker for IgA ASC responses in the intestine after RV infection. High numbers of IgG ASC or memory IgG ASC, induced in the systemic lymphoid tissues of the inactivated virus-inoculated pigs at challenge, did not correlate with protection. Thus, inactivated RV administered orally or parenterally, was significantly less effective in inducing intestinal IgA ASC responses and conferring protective immunity than the live oral HRVs (virulent or attenuated). Based on these studies, gnotobiotic pigs should provide a useful model for examining more efficient mucosal delivery systems, mucosal adjuvants, and vaccination strategies to enhance induction of intestinal IgA ASC by candidate RV vaccines.

1.5. Derivation and Maintenance of Gnotobiotic Pigs An early description of the use of gnotobiotic pigs, in an isolator system to study disease pathogenesis, appeared in 1962, in a study of hog cholera virus (28). Since then, both gnotobiotic pigs and calves have been used as animal models to study the pathogenesis of HRV infections and immunity (4–6,8–15,27,29,30). Their prolonged susceptibility to diarrhea induced by HRV, which was not reproducible in other animal models, permitted challenge exposure to evaluate protection against disease.

1.6. Histopathology Studies The rate of autolytic change in the digestive tract is related to the microflora present within the intestine: Significant autolytic change is present in all sections of the conventional pig small intestine and colon within 24 min of death, whereas significant autolytic changes in these same tissues from gnotobiotic pigs are not observed until 1 hour after death (31). The small intestines should be among the first samples taken, because of the more rapid rate of intestinal (especially duodenal) autolysis, compared to other tissues, when microscopic

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examination is required (31). Exsanguination at euthanasia helps minimize hemorrhage artifacts, and enhances visualization of some lymphoid tissues (especially spleen). Agonal death, or application of prolonged electrical current (as during electrocution), may cause nonspecific lesions, such as petechiation in some tissues. Tissue collection should be performed rapidly and immediately following euthanasia in a cool environment. Placement of intestines on ice-filled trays, covered by plastic (cellophane wrap) or aluminum foil, helps to slow autolysis during collection of segments. Flushing the lumen of the intestinal segments collected with fixative, prior to immersion in the fixative, also reduces autolytic change and aids in fixation. A minimum of a 10:1 fixative:tissue volume ratio should be used for fixation of tissues, and tissue thickness should not exceed 0.5 cm for any specimen collected. Because lesions induced by RV may be scattered, multiple segments of intestine from each region of the intestine (duodenum, jejunum, ileum) should be collected for thorough histologic evaluation. Collection of nine 6-cm-long small intestinal segments from gnotobiotic pigs at equally spaced sites, beginning 5 cm distal to the pylorus and ending 5 cm proximal to the ileal–cecal junction, will give two to four representative segments from each region.

1.7. Pathophysiology Studies To the authors’ knowledge, no published studies exist of intestinal permeability during RV infection of gnotobiotic pigs. However, several reports evaluate the effect of RV infections on the absorption and transport of macromolecules within the intestine in species other than pigs. Readers are referred to these studies for applicable methodologies (32–37). It would appear from these studies that RV infection has diverse affects on intestinal absorption and transport of macromolecules. The uptake of polyethylene glycols (mol wt 282–1250), lactulose, and D -xylose are reduced during RV infection, but uptake of macromolecules, such as horseradish peroxidase (HRP) and L -rhamnose, are increased. This diversity may best be explained by differences between species in terms of the intestinal region involved in absorption and transport of a given macromolecule and the region most affected by the infecting RV strain. 2. Materials 2.1. Diluent and Growth Media 1. Minimal Essential Medium (Gibco-BRL, Grand Island, NY); 1% antibiotics: penicillin G (Pfizer, New York); dihydrostreptomycin sulfate (NB Biochemicals, Cleveland, OH); mycostatin (Squibb, Princeton, NJ); 1% sodium bicarbonate (Na HCO 3) (7% stock); 1% HEPES (Gibco-BRL).

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2.2. Fixatives 1. 10% neutral buffered formalin: 37–40% 100 mL formaldehyde; 900 mL distilled water; 4 g monobasic sodium phosphate; 6.5 g dibasic sodium phosphate. 2. 10% zinc formalin (Z-fix® Concentrate, Anatech, Battle Creek, MI): 4 gal distilled or deionized water; 1 gal Z-fix® concentrate. 3. Prefer (Prefer® Concentrate, Anatech): 11.6 L distilled or deionized water; 3.5 L ethanol or reagent buffer; 1.0 gal Prefer® Concentrate. Shake to mix thoroughly.

2.3. Clearants 1. Xylene. 2. Xylene substitutes: Propar ® clearant (Anatech); Clearium® (Surgipath, Richmond, IL).

2.4. Stains 1. Mayer’s hematoxylin (Sigma, St. Louis, MO). 2. Harris hematoxylin (Lerner, Pittsburgh, PA). 3. 1% eosin alcoholic stock: 1 g water soluble eosin Y; 20 mL distilled water; dissolve eosin in water, then add 80 mL 95% ethanol. 4. Eosin Y (working solution): 1 part 1% alcoholic eosin stock; 3 parts 95% ethanol. Prior to use, add 0.5 mL glacial acetic acid to each 100 mL stain, and stir.

2.5. Solutions 1. Tris-HCl buffer 0.05 M, pH 7.6: 3.03 g Trizma-HCl (Sigma), 0.69 g Trizma base (Sigma). Dissolve in 500 mL deionized water. 2. Tris - Triton-X buffer (TTBS-HS/Tx): 15.15 g Trizma-HCl (Sigma), 3.47 g Trizma base (Sigma), 1000 mL deionized water, 20.29 g NaCl, 5 mL 10 Triton-X (Sigma). 3. Pronase, 0.01%: 0.15 g Protease XIV (Sigma); 150 mL 0.05 M Tris-HCl buffer, pH 7.6. 4. Bluing agent: 0.2% NaH2CO3 and 0.04% lithium carbonate (LiCO3) in water.

2.6. Immunohistochemistry Reagents 1. 2. 3. 4.

Streptavidin-HRP (Dako, Carpinteria, CA). Biotinylated secondary Abs (Dako; Sigma). 3',3'-Diaminobenzidine (DAB) solution (Sigma). 3-Amino-9-ethylcarbazole (AEC) (Zymed, San Fransisco, CA).

3. Methods 3.1. Derivation of Gnotobiotic Pigs by Hysterectomy (38) 1. Healthy pregnant sows are selected for surgery between the d 110 and d 114 of gestation. 2. Ketamine HCl (11 mg/kg) is administered i.m., to sedate the sow.

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Fig. 1. Transfer isolator used to procure gnotobiotic piglets (see Subheading 3.1.). 3. The sow is elevated using a soft cotton rope around each posterior leg, and the abdomen is scrubbed with a germicidal solution (Prodine, Phoenix Pharmaceuticals, St. Joseph, MO). 4. The sow is rendered unconscious using CO 2 delivered into an inhalation bag, and at the end of the surgery the sow is euthanized. 5. An incision is made down the midline of the abdomen of the unconscious sow, and the entire uterus is removed and passed through a chlorinated liquid trap entry into a sterilized surgery isolator. 6. Using rubber gloves in the sides of the isolator for manipulations, the pigs are rapidly removed from the uterus and passed into a sterile transfer isolator, where the umbilical cords are tied. 7. The pigs are transferred into sterile rearing isolators maintained in a room at 35–37°C for the first few days, and thereafter maintained at room temperature. 8. The isolators are constructed of stainless steel rectangular tubs with flexible plastic bubble tops, fitted with two pairs of rubber gloves on the sides. Each tub contains a removable false bottom, metal partitions for 2–4 pigs (each pig in a separate compartment), feeders, and retaining grid. The air blower and filters are similar to ones described previously (38,39; see Fig. 1).

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3.2. Maintenance of Gnotobiotic Pigs 1. All pigs are fed 2–3× daily with a total of 240 mL liquid diet of pig milk replacer (SPF Lac, Borden, Hampshire, IL). 2. Microbiological monitoring is initiated at derivation, then at weekly intervals. Rectal swabs are cultured and examined microscopically for bacteria, yeast, and fungi. Blood agar and fresh thioglycolate broth are used for culture media, and held at least 2 wk before being judged negative.

3.3. Inoculation, Challenge, and Sample Collection 1. Pigs are immunized with various RV candidate vaccines, and challenged at 2–4 wk after the final booster immunization (5). Pigs are challenged orally with 10 6 ID 50 (the DD 50 was approx equal to the ID 50 ) of virulent Wa HRV (P1A[8],G1) or M strain HRV (P1A[8], G3), to consistently reproduce diarrhea and lesions in 100% of inoculated pigs (6). 2. Control pigs are given equal volumes of diluent alone, to monitor fecal consistency, and to collect control tissues and samples. 3. Pigs are examined daily for clinical signs, including anorexia and diarrhea (scored as fecal consistency of 0 [normal] to 3 [liquid]). 4. Blood (2–5 mL) is drawn weekly from the anterior vena cava by using a 20–22 gage, 1.5-in. needle, and serum is obtained by standard procedures. 5. Daily duplicate rectal swabs are collected at 0–10 PID and PCD into 8 mL diluent (see Subheading 2.1.).

3.4. Cell Culture Immunofluorescence (CCIF) for Titration of Infectious RV (40) 1. Monkey kidney (MA104) cells are grown for 6–9 d in flat-bottomed 96-well microtiter plates (Costar, Cambridge, MA) in growth media, but containing 10% fetal calf serum. 2. Wells are rinsed with diluent about 3 h prior to viral inoculation. Rectal swabs or fecal samples to be tested for virus are diluted 1 in 25 (two swabs in 8 mL), then 1 in 100, and serially thereafter in 10-fold dilutions in diluent, and 0.1 mL diluted viral specimens are added (in duplicate) to each well. Thirty microliters of a 1 in 200 dilution of stock pancreatin (1 in 10, 4XNF, Gibco-BRL) are added to each well. Virus positive and negative specimens are added to each plate. 3. The plates are placed in a centrifuge plate carrier and centrifuged at 1300g for 60 min. 4. After centrifugation, the plates are incubated at 37°C for 18 h in a 5% CO2-air atmosphere, and then the medium is aspirated and the wells rinsed with phosphate-buffered saline (PBS) (pH 7.2). 5. The cells are fixed with 0.2 mL of a solution containing 80% acetone and 20% distilled H 2 O for 10 min at room temperature. The fixative is aspirated, and plates are rinsed with PBS. 6. To stain, 25 µL diluted (1 in 100 in PBS) fluorescein isothiocyanate (FITC)conjugated hyperimmune anti-RV serum, prepared in gnotobiotic pigs, is added, and plates are incubated for 1 h at 37°C.

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7. Plates are rinsed with PBS, mounting media is added, and plates are examined, using a fluorescent microscope. The number of individual fluorescing cells/well are enumerated at the highest dilution with fluorescing cells, and the virus infectivity titer is calculated as FFU/mL.

3.5. Immunofluorescent (IF) Staining of RV-Infected Intestinal Epithelial Cells (41) 1. Remove small intestine from gnotobiotic pig immediately after euthanasia. 2. Cut approx 6-cm-long segments from the duodenum, jejunum, and ileum. Slit open each segment lengthwise, to expose the mucosal surface. 3. Gently rinse the mucosal surface with 0.85% saline, then use a scalpel to scrape the mucosal surface, and put the scrapings on an alcohol-cleaned microscope slide. Using a second alcohol-cleaned slide, press the two slides together, to make a mucosal impression smear. 4. Air-dry the smears, and fix in 100% acetone for 10 min. 5. Rinse in PBS (pH 7.2) for 5 min, then distilled H2O for 30 s. Air-dry and IF-stain, or store at –20°C until IF staining is done. 6. Apply 0.03 mL diluted (1 in 50–1 in 100 in PBS) FITC-conjugated anti-RV serum as described for the CCIF procedure (Subheading 3.4., step 6), and incubate for 30 min at 37°C. 7. Rinse slides using PBS, and leave in PBS for 10 min. Rinse with distilled H2O for 30 s and air-dry. 8. Add mounting medium and cover-slip, and examine the smears for fluorescing cells, using a fluorescent microscope. Calculate the approximate percentage of fluorescing cells in randomly selected fields. 9. Paraffin-embedded tissue sections may also be IF-stained (as described in Subheading 3.6.).

3.6. Processing of Tissues and Preparation of Paraffin Sections for IF Staining (40) 1. Flush the lumen of freshly collected small intestinal segments with 100% acetone, and immerse in two changes of acetone for 30 min to 1 h each. 2. Place acetone-fixed tissue directly in melted paraffin in a vacuum oven at 60°C and 25 in. (63.5 cm) Hg for 10 min. Embed and section as described in Subheading 3.7., steps 3–5. 3. Deparaffinize slides in two changes of Propar (or xylene) for 2 min each. 4. Rinse in 100% acetone for 5 min and allow to air-dry. 5. Wash in PBS for 10 min, followed by distilled H2O for 30 s. 6. Air-dry and resume IF-staining at Subheading 3.5., step 6.

3.7. Processing Paraffin-Embedded Tissues for Hematoxylin and Eosin (H&E) or Immunohistochemistry (IHC) Staining (6,42–44) 1. Process tissues through two changes of fixative, 60 min each; one change of 80%, two changes of 95%, and two changes of 100% ethanol, 30 min each;

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6.

Saif and Ward three changes of Propar, 60 min each, and two changes of melted paraffin, at least 60 min each. Paraffin-embed formalin-fixed tissues within 24–48 h of collection. Prefer-fixed tissues may stay in a fresh change of fixative for up to 2 wk before paraffinembedding. Section paraffin blocks at 4–5 µm, and float on a water bath containing only deionized or distilled water (no adhesives). Gelatin may be added to water bath for H&E sections. Pick up flattened sections on ProbeOn® Plus (Fisher Scientific, Pittsburgh, PA) slides (poly-L-lysine or chrome alum-coated slides are also acceptable), and bake in a 60°C oven for 1 h prior to deparaffinization (see Note 1). Label all slides using Secureline® (Precision Dynamics Corp., San Fernando, CA) pen (most other pens are soluble in alcohol and/or xylene, and will wash off during deparaffinization, dehydration, and clearing). Proceed with desired staining procedure.

3.7.1. H&E Staining Procedure (42) 1. Deparaffinize slides in three changes of Propar (or xylene) for 5 min each (see Note 2). 2. Transfer slides to 100% ethanol, two changes, 3 min each, followed by 95 and 70% ethanol for 3 min each. Rinse under running tap water for 1 min. 3. Transfer to Harris’ hematoxylin for 5 min (filter Harris’ just prior to use), and place under running tap water for 1 min. 4. Dip slide once in 0.25% citric acid, and rinse under running tap water for 1 min. 5. Dip slide 4× in bluing agent, and rinse under running tap water for 1 min. 6. Place in 95% and 70% ethanol for 1 min each. 7. Place in eosin Y (working solution) for 2 min. 8. Dip slide once each in 95% and 100% isopropanol. 9. Place in two changes of 100% isopropanol for 1 min each. 10. Remove slides from isopropanol, and immediately add Clearium (Surgipath, cat. no. 01100) mounting medium and cover-slip (wet mount).

3.7.2. IHC Staining Procedure (43,44) 1. Deparaffinize slides in two changes of Propar (or xylene) for 10 min each (see Note 2). 2. Transfer slides to 100% alcohol, two changes, for at least 2 min each. 3. Rinse slides in water, and place in PBS (see Note 3). 4. Load slides into a staining rack, and rinse again with PBS (a flat humid chamber may be used, instead of a rack system). 5. Apply 100 µL diluted primary Ab to each chamber (or flat slides in humid chamber), cover, and incubate 1 h at room temperature, or 30 min at 37°C, or overnight at 4°C. Next day, remove slides from the refrigerator and continue with (Step 6, see Notes 4 and 5) 6. Rinse slides in three changes of PBS, 2 min each, or rinse/blot 10× in TTBS-HS/Tx.

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7. Wipe slide again, and apply 100 µL diluted biotinylated secondary Ab (must be matched to species and/or isotype of the primary Ab). Allow to incubate at room temperature for 30 min or 37°C for 20 min (see Note 6). 8. Rinse slides in three changes of PBS, 2 min each, or rinse/blot 10× in TTBS-HS/Tx. 9. Apply 100 µL strepavidin/HRP (commercially available prediluted) to each slide, and incubate at room temperature 30 min. 10. Rinse slides in three changes of PBS, 2 min each. 11. Prepare DAB according to manufacturer’s specifications (see Note 7). 12. Carefully drain slides, and place them on a flat surface. Ensure the slides do not dry out. 13. Add approx 400 µL DAB solution to each slide, making sure all the tissue is covered by solution. Incubate approx 5 min. 14. Carefully pick up each slide in the order that DAB was applied, drain on paper towel, and place in staining rack in a dish of water (see Note 8). 15. Rinse slides well in water 3×. 16. Counterstain slides: Dip twice in Mayer’s hematoxylin (or stain for 1–2 min); rinse thoroughly in water; dip twice in bluing reagent or dilute ammonia water; rinse thoroughly in water. 17. Dehydrate in four changes of 100% ethanol, clear in 3–4 changes of Propar (or xylene) and cover-slip. Always allow slides to drain 15–20 s between containers, to prevent carryover from dish to dish.

4. Notes 1. Paraffin sections can be stored at room temperature prior to staining. 2. An endogenous peroxidase block may be used between steps 1 and 2. Incubate slides at room temperature in 0.03% H2O2 in PBS, or 3% H2O2 in absolute methanol, for 5 –10 min. Do not use a methanol blocking solution on frozen sections. 3. Pretreatment for Ag retrieval may be done between steps 2 and 3 by incubating in 0.01% Pronase for 2–10 min, 37° C. Rinse slides in running deionized water, 5 min, or rinse/blot in TTBS-HS/Tx 10×. DO NOT digest Prefer-fixed specimens. 4. The authors dilute all primary and secondary Abs in a 1.5% horse serum blocking buffer. Slides may be further blocked by incubating sections in 1–5% horse serum in PBS at room temperature for 10–30 min prior to application of the primary Ab (it is best to block with serum from the species in which the secondary is made). Do not rinse after blocking, but go straight to the primary Ab application. 5. If a humid chamber is used instead of the staining racks, the amount of Ab required to cover the section will usually be greater than 100 µL. 6. The authors generally detect the primary (monoclonal) Ab using the commercial product, Vectastain Elite ABC (Vector, Burlingame, CA), which includes a biotinylated horse secondary Ab and an avidin-biotin-peroxidase complex and replaces Subheading 3.7.2., steps 7–14. 7. Caution: DAB is a suspected carcinogen and must be handled with care. Always wear gloves.

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8. If the chromagen used is not stable in alcohol (such as AEC from Zymed), cover-slip the slides with aqueous mounting media and DO NOT dehydrate or clear slides as in Subheading 3.7.2. Step 17.

References 1. Burns, J. W., Krishnaney, A. A., Vo, P. T., Rouse, R. V., Anderson, L. J., and Greenberg, H. B. (1995) Analyses of homologous rotavirus infection in the mouse model. Virology 207, 143–153. 2. Conner, M. E., Estes, M. K., and Graham, D. Y. (1988) Rabbit model of rotavirus infection. J. Virol. 62, 1625–1633. 3. Ward, R. L., Bernstein, D. I., McNeal, M. M., and Sheridan, J. F. (1990) Development of an adult mouse model for studies of protection against rotavirus. J. Virol. 64, 5070–5074. 4. Saif, L. J., Ward, L. A., Rosen, B. I., and To, T. L. (1996) The gnotobiotic piglet as a model for studies of diseases pathogenesis and immunity to human rotaviruses. Arch. Virol. 12, 153–161. 5. Yuan, L., Ward, L. A., Rosen, B. I., To, T. L., and Saif, L. J. (1996) Systemic and intestinal antibody-secreting cell responses and correlates of protective immunity to human rotavirus in a gnotobiotic piglet model of disease. J. Virol. 70, 3075–3083. 6. Ward, L. A., Rosen, B. I., Yuan, L., and Saif, L. J. (1996) Pathogenesis of an attenuated and a virulent strain of group A human rotavirus in neonatal gnotobiotic pigs. J. Gen. Virol. 77, 1431–1441. 7. Barnes, G. L. and Townley, R. R. W. (1973) Duodenal mucosal damage in 31 infants with gastroenteritis. Arch. Dis. Childh. 48, 343–349. 8. Ward, L. A., Yuan, L., Rosen, B. I., and Saif, L. J. (1996) Development of mucosal and systemic lymphoproliferative responses and protective immunity to human group A rotavirus in a gnotobiotic pig model. Clin. Diag. Lab. Immunol. 3, 342–350. 9. Yuan, L., Kang, S., Ward, L. A., To, T. L., and Saif, L. J. (1998) Antibody-secreting cell responses and protective immunity assessed in gnotobiotic pigs inoculated orally or intramuscularly with inactivated human rotavirus. J. Virol. 72, 330–338. 10. Hodgins, D. C., Kang, S. Y., deArriba, L., Parreno, V., Ward, L. A., Yuan, L., To., T., and Saif, L. J. (1999) Effects of maternal antibodies on protection and the development of antibody responses to human rotavirus in gnotobiotic pigs. J. Virol. 73, 186–197. 11. Ward, L., Kang, S., deArriba, L., Parreno, V., Yuan, L., To, T., and Saif, L. (1998) Effects of maternal antibodies on human rotavirus (HRV)-specific T-cell responses in a neonatal pig model of HRV disease. American Society for Virology Annual Meeting, Vancouver, Abst. W2–6 July, 11–15. 12. Parreno, V., deArriba, L., Kang, W., Yuan, L., Hodgins, D., Ward, L., To, T., and Saif, L. J. (1999) Serum and intestinal isotype antibody responses to Wa human rotavirus in gnotobiotic pigs are modulated by maternal antibodies. J. Gen. Virol. 80, 1417–1428.

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13. Wyatt, R. G., James, W. D., Bohl, E. H., Theil, K. W., Saif, L. J., Kalica, A. R., Greenberg, H. B., Kapikian, A. Z., and Chanock, R. M. (1980) Human rotavirus type 2: cultivation in vitro. Science 207, 189–191. 14. Hoshino, Y., Saif, L. J., Sereno, M. M., Chanock, R. M., and Kapikian, A. Z. (1988) Infection immunity of piglets to either VP3 or VP7 outer capsid protein confers resistance to challenge with a virulent rotavirus bearing the corresponding antigen. J. Virol. 62, 744–748. 15. Hoshino, T., Saif, L. J., Kang, S. Y., Sereno, M., Chen, W. K., and Kapikian, A. Z. (1995) Identification of group A rotavirus genes associated with virulence of a porcine rotavirus and host range restriction of a human rotavirus in the gnotobiotic piglet model. Virology 209, 274–280. 16. Theil, K. W., Bohl, E. H., Cross, R. F., Kohler, and E. M., Agnes, A. (1978) Pathogenesis of porcine rotaviral infection in experimentally inoculated gnotobiotic pigs. Amer. J. Vet. Res. 39, 213–220. 17. Mebus, C. A. (1976) Reovirus-like calf enteritis. Amer. J. Digest. Dis. 21, 592-599. 18. Ball, J. M., Peng, T., Zeng, C. Q.-Y., Morris, A. P., and Estes, M. K. (1996) Age-dependent diarrhea induced by a rotaviral nonstructural glycoprotein. Science 272, 101–104. 19. Bernstein, D. I., Sander, D. S., Smith, V. E., Schiff, G. M., and Ward, R. L. (1991) Protection from rotavirus reinfection: 2-year prospective study. J. Infect. Dis. 164, 277–283. 20. Coulson, B. S., Grimwood, K., Hudson, I. L., Barnes, G. L., and Bishop, R. F. (1992) Role of coproantibody in clinical protection of children during reinfection with rotavirus. J. Clin. Microbiol. 30, 1678–1684. 21. Matson, D. O., O’Ryan, M. L., Herrera, I., Pickering, L. K., and Estes, M. K. (1993) Fecal antibody responses to symptomatic and asymptomatic rotavirus infections. J. Infect. Dis. 167, 577–583. 22. Collins, J. E., Benfield, D. A., and Duimstra, J. R. (1989) Comparative virulence of two porcine group-A rotavirus isolates in gnotobiotic pigs. Amer. J. Vet. Res. 50, 827–835. 23. McAdaragh, J. P., Bergeland, M. E., Meyer, R. C., Johnshoy, M. W., Stotz, I. J., Benfield, D. A. and Hammer, R. (1980) Pathogenesis of rotaviral enteritis in gnotobiotic pigs: A microscopic study. Amer. J. Vet. Res. 41, 1572–1581. 24. Tauscher G. I. and Desselberger, U. (1997) Viral determinants of rotavirus pathogenecity in pigs: production of reassortants by asynchronous coinfection. J. Virol. 71, 853–857. 25. Burke, B., Bridger, J. C., and Desselberger, U. (1994) Temporal correlation between a single amino acid change in the VP4 of a porcine rotavirus and a marked change in pathogenicity. Virology 202, 754–759. 26. Burke, B., McCrae, M. A., and Desselberger, U. (1994) Sequence analysis of two porcine rotaviruses differing in growth in vitro and in pathogenicity: Distinct VP4 sequences and conservation of NS53, VP6 and VP7 genes. J. Gen. Virol. 75, 2205–2212.

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27. Schaller, J. P., Saif, L. J., Cordle, C. T., Candler, E., Winship, T. R., and Smith, K. L. (1992) Prevention of human rotavirus induced diarrhea in gnotobiotic piglets using bovine antibody. J. Infect. Dis. 165, 623–630. 28. Weide, K. D., Waxler, G. L., Whitehair, C. K., and Morrill, C. C. (1962) Hog cholera in gnotobiotic pigs. J. Amer. Vet. Med. Assoc. 128, 94–98. 29. Mebus, C. A., Wyatt, R. G., Sharpee, R. L., Sereno, M. M., Kalica, A. R., Kapikian, A. Z., and Twiehaus, M. J. (1976) Diarrhea in gnotobiotic calves caused by the reovirus-like agent of human infantile gastroenteritis. Infect. Immun. 14, 471–474. 30. Torres-Medina, A., Wyatt, R. G., Mebus, C. A., Underdahl, N. R., and Kapikian, A. Z. (1976) Diarrhea in gnotobiotic piglets caused by the reovirus-like agent of human infantile gastroenteritis. J. Infect. Dis. 133, 22–27. 31. Cross, R. .F and Kohler, E. M. (1969) Autolytic changes in the digestive system of germfree, Escherichia coli monocontaminated, and conventional baby pigs. Can. J. Comp. Med. 33, 108–112. 32. Ijaz, M. K., Sabara, M. I., Frenchick, P. J., and Babiuk, L. A. (1987) Lactose malabsorption following rotavirus infection in young children. J. Pediatrics 99, 916–918. 33. Mavrovichalis, J., Evans, N., McNeish, A. S., Bryden, A. S., Davies, H. A., and Flewett, T. H. (1977) Intestinal damage in rotavirus and adenovirus gastroenteritis assessed by D-xylose malabsorption. Arch. Dis. Childh. 52, 589–591. 34. Stintzing, G., Johansen, K., Magnusson, K. E., Svensson, L., and Sundqvist, T. (1986) Intestinal permeability in small children during and after rotavirus diarrhoea assessed with different-size polyethyleneglycols (PEG 400 and PEG 1000). Acta Paediatr. Scand. 75, 1005–1009. 35. Noone, C., Menzies, I. S., Banatvala, J. E., and Scopes, J. W. (1986) Intestinal permeability and lactose hydrolysis in human rotaviral gastroenteritis assessed simultaneously by non-invasive differential sugar permeation. Eur. J. Clin. Invest. 16, 217–225. 36. Heyman, M., Cortheir, G., Petit, A., Meslin, J.-C., Moreau, C., and Desjeux, J.-F. (1987) Intestinal absorption of macromolecules during viral enteritis: an experimental study on rotavirus-infected conventional and germ-free mice. Pediatr. Res. 22, 72–78. 37. Hayhow, C. S. and Saif, Y. M . (1993) Experimental infection of specificpathogen-free turkey poults with single and combined enterovirus and group A rotavirus. Avian Dis. 37, 546–557. 38. Meyer, R. C., Bohl, E. H., and Kohler, E. M. (1964) Procurement and maintenance of germ-free swine for microbiological investigations. Appl. Microbiol. 12, 295–300. 39. Trexler, P.C. and Reynolds, L. I. (1957) Flexible film apparatus for rearing and use of germfree animals. Appl. Microbiol. 5, 406–412. 40. Bohl, E. H., Saif, L. J., Theil, K. W., Agnes, A. G., and Cross, R. F. (1982) Porcine pararotavirus: detection, differentiation from rotavirus, and pathogenesis in gnotobiotic pigs. J. Clin. Microbiol. 15, 312–319.

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41. Bohl, E. H., Kohler, E. M., Saif, L. J., Cross, R. F., Agnes, A. G., and Theil, K. W. (1978) Rotavirus as a cause of diarrhea in pigs. J. Amer. Vet. Med. Assoc. 172, 458–463. 42. Carson, F. L. (1997) Histotechnology: A Self-Instructional Text., American Society of Clinical Pathologists, Chicago. 43. Shoup, D. I., Swayne, D. E., Jackwood, D. J., and Saif, L. J. (1996) Immunohistochemistry of transmissable gastroenteritis virus antigens in fixed paraffin-embedded tissues. J. Vet. Diag. Invest. 8, 161–167. 44. Grooms, D. L., Ward, L. A., and Brock, K. V. (1996) Morphologic changes and the immunohistochemical detection of viral antigen in ovaries from cattle persistently infected with bovine viral diarrhea virus. Amer. J. Vet. Res. 57, 830–833.

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7 Immunologic Methods and Correlates of Protection Kristine K. Macartney and Paul A. Offit 1. Introduction 1.1. Immunologic Correlates of Protection After Natural Rotavirus Infection in Children and Adults Studies of natural rotavirus (RV) infection in children have shown that protection against subsequent RV disease occurs (1). Assessment of humoral immune responses has included study of the importance of circulating vs intestinal antibodies (Abs), serotype-specific vs group-specific Abs, and RV-specific immunoglobulins IgA, IgM, and IgG (1). Following natural RV infection, RV-specific IgM, followed by IgA and IgG, appear in serum and duodenal fluid or stool of young children (2). Protection against subsequent RV infection is predicted by the quantity of virus-specific IgA in the feces and serum (3,4). In addition, virus-specific antibody-secreting cells (ASC) of the IgA, IgM, and IgG isotypes have been detected in the blood of infants following RV infection (5), although correlation between the presence of ASCs and protection against subsequent disease has not been studied. Serum neutralizing antibodies (nAbs) occur after natural RV infection in children, and are serotype-specific (4,6). Overall, protection against subsequent RV infection is correlated with higher titers of nAb (4). Protection against infection has been correlated with homotypic nAb to the G1 serotype (4); however, other studies suggest that protection is not dependent on serotype-specific nAb (7). Studies of the role of virus-specific helper or cytotoxic T-cell (CTL) responses in RV infection have been limited by difficulty in obtaining adequate numbers of both T-cells from peripheral blood and major histocompatibility complex-compatible target cells. In adults infected during a RV outbreak, RV-specific proliferative T-cells were detected (8), and, in young children, From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ

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RV-specific T-helper cells have been detected following infection (9). The importance of T-cell immune responses in protection against infection in human subjects has not been evaluated.

1.2. Immunologic Correlates of Protection After RV Immunization in Children Unlike natural infection, studies of immunization of children, with either animal RVs or reassortants between animal and human rotaviruses (HRVs), demonstrate that the induction of serum IgA does not correlate with protection against subsequent disease (10). Unfortunately, extensive studies of the presence of virus-specific IgA in vaccine recipients’ feces, and correlation with protection against reinfection, have not been performed. This observed difference in the capacity of serum IgA to predict protection against disease following natural infection, but not following immunization, may be related to differences in host response to homologous and heterologous virus, or, alternatively, to the way in which Ab studies were performed. In studies of natural infection, regular collection of sera and feces were made throughout the RV season, so that the length of time between serum collection and onset of disease was usually less than 1 mo. However, in immunization studies, sera were usually collected within 1 mo after the final vaccine dose, which meant that the interval between determination of serum IgA and disease may have been as long as 5 mo (1). Because virus-specific IgA in serum may be short-lived, comparison of titers between these types of studies may not be valid. The presence of serotype-specific nAbs following immunization with animal–HRV reassortant vaccines has been shown to correlate with protection against some G-types of RV infection, but, no specific titer of any Ab was a reliable predictor of protection (11). An increase in the number of IgA and IgM ASCs after oral reassortant vaccine administration has also been shown to occur; however, that response was not correlated with protection against subsequent natural infection (12). Studies of helper or CTL responses in vaccine recipients have not been performed.

1.3. Immunologic Correlates of Protection After Infection or Immunization in Animal Models The development of different animal models of RV infection has allowed more extensive studies of the humoral and cellular immune response to infection, and the immunologic determinants associated with protection. Calves, lambs, pigs, rabbits, and mice (both immunocompetent and immunodeficient) have been used to study the response to heterologous and homologous host RVs, inactivated viruses, and purified viral proteins (VPs) (1). Following infection with homologous host RV, animals may develop RV isotype-specific Abs in

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intestinal secretions, lamina propria (LP), Peyer’s patch (PP), mesenteric lymph node (MLN), spleen, serum, and breast milk. Protection following challenge generally occurs in association with the presence of serum and intestinal IgA, but, these findings have varied, depending on the host animal and the host-specificity of the immunizing and challenge viruses (1). In murine and porcine models of infection, the location, frequency, and isotype of RV-specific Ab-secreting cells have been determined by enzyme-linked immunospot (ELISPOT) assay, and have been correlated with protection upon challenge (13,14). Type-specific nAbs develop in animals following oral and parenteral immunization, but immunization with one RV strain does not induce crossreactive nAbs to other strains to which the animal has not been exposed (15). It is unclear whether serum or fecal nAbs directed against the challenge virus are important in protection against infection. Studies of the role of T-cell responses in RV infection have been performed, including isolation and transfer of RV-specific CTLs from immunized mice to naïve suckling mice, who were subsequently protected against challenge (16). In addition, transfer of RV-specific CTLs to immunodeficient recipient mice, which were chronically shedding RV, resulted in termination of viral shedding (17).

1.4. Choice of Methods for Detection of Ab and T-Cell Responses RV-specific Abs can be detected by a number of assays that recognize different viral epitopes. Assays that primarily detect isotype-specific Abs directed against the highly conserved inner capsid proteins, specifically VP6, are enzyme-linked immunoborbent assay (ELISA) (18), radioimmunoassay (RIA) (19), complement-fixation assay (20), and immune-adherence hemagglutination assay (21). Assays that detect nAbs directed against the outer capsid proteins VP4 and VP7 are the hemagglutination-inhibition assay (22), plaque reduction neutralization assay (PRNA) (23), fluorescent-focus assay (6), the ELISA-based antigen-reduction neutralization assay (24), and the epitope-blocking assay (25). Another assay, radioimmunoprecipitation (26), may detect Abs directed against RV-specific structural and nonstructural proteins. Quantitation and isotype determination of virus-specific Ab-secreting plasma cells by the ELISPOT has been performed in animal models and in infants (5,12–14). Assays to detect T-cell responses to RV infection in human subjects include the lymphoproliferation assay to detect RV-specific proliferating cells (9). These cells may include virus-specific T-helper cells, virus-specific CTLs, and virus-specific B-cells. In addition, the 51Cr-release assay detects virus-specific CTLs (8). However, because it is difficult to obtain large numbers of virus-specific T-cells from the circulation of infants, these studies have been limited, and will not be described here.

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Because ELISA is the most commonly performed method for detection of isotype-specific Abs, a protocol for determination of IgA and IgG from human serum and fecal samples is described. The PRNA is considered as sensitive as newer techniques for detecting nAbs (24), and is also described in Subheading 3.1. In addition, a method for determination of the frequency and isotype of murine RV-specific ASC by ELISPOT assay is included. The choice of methods to be employed will depend on the experimental aims of the study and expertise of the lab; however, the following techniques should provide an opportunity to investigate various aspects of the immune response to RV infection. 2. Materials 2.1. Plaque Reduction Neutralization Assay 1. Fetal green monkey kidney MA104 cells. 2. 1000 mL of Dulbecco's Modified Eagle's Medium, 10% fetal bovine serum (FBS) (DMEM 10%), 100 mL tryptose phosphate broth, 120 mL heat inactivated FBS, 12.5 mL glutamine (200 mM), 2 mL penicillin/streptomycin (stock solution: 100 µ/mL streptomycin sulfate and 100 U/mL penicillin). Store at 4°C. 3. DMEM media, without FBS (DMEM/N) (1000 mL): 1000 mL DMEM media, 12.5 mL glutamine (200 mM), 2 mL penicillin/streptomycin. Store at 4°C. 4. Phosphate-buffered saline (PBS), pH 7.2. Sterile, stored at 25°C. 5. Minimal salt overlay (MSO): 615 mL sterile distilled H 2O, 200 mL 10X Earle’s Balanced Salt solution (EBSS), 20 mL 100X essential amino acids for basal medium eagle (w/o glutamine), 20 mL 100X vitamins for minimum essential medium, 20 mL glutamine (200 mM), 50 mL NaHCO 3 (5.6% solution), 2 mL penicillin/streptomycin. To decrease precipitate formation, 0.5 mL 1 N HCl/100 mL can be added. Store at 4°C. 6. Agarose 1% solution, stored at 4°C. 7. 1X Tryspin/EDTA: 625 mL of 2.5% tryspin (1 in 250), 100 mL of 0.53 mM Versene. 8. 2X EBSS, diluted in distilled water from 10X solution, stored at 4°C. 9. Neutral red solution, stored at 4°C.

2.2. ELISA for Human Serum RV-Specific IgA or IgG 1. 96-well, flat-bottom, high-binding ELISA plates (see Note 1). 2. Wash buffer: PBS (without calcium chloride or magnesium chloride) with 0.025% Tween-20 (PBS-T), stored at 25°C. 3. Coating agent: Single-shelled group A RV; purified by cesium chloride (CsCl) gradient, quantitated by spectrophotometry, diluted to 1 µg/µL in PBS, stored at –70°C. 4. Coating buffer: 0.1 M carbonate–bicarbonate solution, stored at 25°C. 5. Blocking buffer: Tris-buffered saline (TBS) with 1% heat-inactivated Carnation® skim milk (boiled for 5 min before use) with 0.05% Tween-20 (SK-T), stored at 4°C for <14 d.

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6. Positive control: human serum with known high-titer IgA or IgG to RV, respectively, stored at –70°C. 7. Negative control: from pool of human serum without Ab to RV, stored at –70°C (see Note 2). 8. Detector Ab: peroxidase-conjugated goat IgG fraction to human IgA (for IgA detection) or peroxidase-conjugated goat IgG fraction to human IgG (for detection of IgG), stored at –70°C, individual 100-µL aliquots stored at 4°C for <14 d. 9. Substrate: tetramethyl-benzidine peroxidase solution, stored at 4°C. 10. Reaction termination: 85% ortho-phosphoric acid, stored at 25°C. 11. ELISA plate washer (although manual washing is also satisfactory) and ELISA plate reader.

2.3. ELISA for Human Fecal RV-Specific and Total IgA and IgG 1. Coating agents: single-shelled RV; purified by CsCl gradient, quantitated by spectrophotometry (at wavelength of 2.60 nm), diluted to 1 µg/µL in distilled H 2O, stored at –70°C. 2. Goat-affinity purified Ab to human IgA (for total IgA detection) or goat-affinity purified Ab to human IgG (for total IgG detection), stored at –70°C, individual aliquots stored at 4°C for ≤14 d. 3. Blocking buffer: sterile PBS with 1% bovine serum albumin and 0.05% Tween-20 (BSA-T), stored at 4°C. 4. Positive controls: human fecal specimen with known high IgA or IgG titer to RV, diluted to 10% w/v in 1X EBSS. Store at –70°C. 5. Negative control: human fecal specimen negative for RV-specific IgA (diluted and stored as above).

2.4. Mononuclear Cell Isolation 1. 2. 3. 4. 5. 6. 7. 8.

Mice. Hanks’ Buffered Salt Solution (HBSS) containing 5% FBS. L-shaped (bent) 21-gage needle. Ammonium chloride-potassium (ACK) solution containing 4.3 g NH 4Cl, 0.5 g KHCO3 in 500 mL distilled H2O, and sterilized by filtration. Ficoll-Hypaque. Cotton column. Cell collector (sieve). Trypan blue.

2.5. ELISPOT Assay for RV-Specific ASCs 1. 1% heat-inactivated skim milk in TBS. Store at 4°C for ≤14 d. 2. Complete media (100 mL): 89 mL RPMI medium 1640, with 10 mL FBS (10% v/v), 1 mL glutamine (200 mM), and 100 µL HEPES (1 M). Store at 4°C. 3. Development solution: Dissolve 10 mg 3-amino-9-ethyl carbazole (AEC) in 1 mL N,N-dimethylformamide, using a glass pipet and glassware. Add 30 mL 0.1 M sodium acetate. Filter this solution through a 0.45-µm filter. Store at 4°C for up to 14 d. Immediately prior to use, add 15 µL 30% H 2O 2 to the filtered AEC solution.

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4. 0.1 M sodium acetate buffer (pH 5.0): Mix 148 mL solution A (0.2 M acetic acid = 11.55 mL glacial acetic acid made up to 1 L with distilled H 2 O) with 352 mL solution B (sodium acetate = 27.2 g sodium acetate made up to 1 L with distilled H 2 O). Add 400 mL distilled H 2O to this solution, and adjust the pH to 5.0. Then add distilled H 2O to make final volume up to 1 L. Store at 25°C.

3. Methods 3.1. PRNA of Serum This assay determines the dilution end point of a serum or stool suspension that neutralizes 50% of RV plaques. A plaque is defined as an area of focal cell degeneration produced by a single infectious virion in a cultured monolayer of cells. More than one virus can be used in the assay to determine cross-neutralization titers of a given serum or stool sample. For human samples from infants, the authors routinely assay nAb against up to five viruses per sample. The method employs constant concentrations of virus, incubated in equal volumes with varying concentrations of the serum or stool sample. The experiment is designed to provide 50 plaques/well in the control wells, in which virus is incubated in corresponding dilutions in the presence of media diluent only. The method used for serum and stool samples is identical, except in the preparation of the stool specimen (described in Subheading 3.2.). This method measures total nAb and does not allow for differentiation between IgA and IgG. 1. Prepare MA104 cell cultures (passage 59–69) in flat-bottom, six-well (35-mm) plates, by splitting a 175 cm2 flask of cultured cells (using 5 mL trypsin/EDTA) into five six-well plates. Use stock culture flasks no more than 7 d old. The cells are grown in DMEM/N 10% media (3 mL/well), and used 3–4 d after preparation. 2. Serum samples for assay are diluted 1 in 10 in PBS, and stored at –20°C. After thawing, heat-inactivate the samples in a 56°C water bath for 30 min. Samples are then prepared as serial five-fold dilutions in DMEM/N, starting with a 1 in 25 dilution (because the serum is originally diluted 1 in 10 in PBS, the first dilution can be made by adding 1 mL serum sample to 1.5 mL DMEM/N). Three dilutions are commonly used: 1 in 25, 1 in 125, and 1 in 625. 3. Dilute virus in DMEM/N in order to achieve a concentration of 5.0 × 10 2 PFU/mL (this should provide approx 50 plaques/control well). 4. Add 200 µL of each dilution of serum to 200 µL diluted virus in sterile Wasserman tubes. 5. For virus controls, add 200 µL diluted virus to 200 µL DMEM/N into Wasserman tubes. Make enough to inoculate 2–3 control wells for each virus used. Incubate all the Wasserman tubes in a 37°C water bath for 30 min. 6. Toward the end of the incubation period, prepare six-well plates of MA104 cells by washing them twice with 2 mL/well of warm PBS (see Note 3).

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7. Inoculate each well with 200 µL of either the serum/virus mixture or virus/DMEM/N control mixture. Adsorb by placing plates in an atmosphere of 5% CO2-air at 37°C for 30 min. 8. Toward the end of the adsorbtion step, prepare 1:1 agarose:MSO solution (see step 10). 9. Wash plates 2X with PBS (2 mL/well). During washing, aspirate PBS by starting at the well with the lowest serum dilution, and proceeding to the highest serum dilution. Change Pasteur pipets between series of dilutions. 10. Overlay wells with 2.5 mL/well of 1:1 agarose:MSO solution containing 0.5 µg/mL trypsin (original concentration of trypsin is 100 µg/mL). For example, to overlay five plates, a total volume of 75 mL overlay solution is needed (make a total of 80 mL). Therefore, use 40 mL each of 1% agarose and MSO solution, and 0.4 mL trypsin. To make the overlay solution, melt the agarose in a microwave, until bubbling. Do not add it to the MSO-trypsin until just before plating. The final solution should be plated when its container is warm to touch, but not uncomfortable (see Note 4). 11. Incubate plates for 3 d at 37°C in an atmosphere of 5% CO 2-air. 12. On the third day, stain the plates by overlaying with 1.5 mL/well of the following: 1:1 solution of 2X EBSS: 1% agarose, with 1:30 of neutral red. For example, to stain five plates, make 50 mL staining solution by combining 25 mL each of 2X EBSS and 1% agarose with 1.7 mL neutral red. 13. Count plaques 4 h poststaining, and daily thereafter. Continue to incubate plates and count plaques each day, until either the number of plaques remains static, or there is death of the cell monolayer. The titer will be calculated for each virus on the day on which the number of plaques in the corresponding virus control wells is closest to 50. There must be at least 20 plaques/well and no more than 100. 14. To calculate the plaque reduction neutralization (PRN) titer; determine the following values: for each virus: A, the average of the virus control plaque counts/well, divided by 2 (this is the 50% plaque reduction number); B, the plaque number in the well at serum dilution closest to, and less than, A; C, the plaque number at the serum dilution closest to, and greater than, A; D, the serum dilution that corresponds with B; and E, the serum dilution that corresponds with C. The 50% plaque reduction titer for the serum/virus equals: [(A – B)/(C – B) × (E – D) + D] × 2

(1)

For example, if the virus control equals 50 plaques, therefore A = 25, and the observations are: serum dilutions 1 in 25, 1 in 125, and 1 in 625; plaque count 5, 20, and 50, respectively. The 50% plaque reduction titer is: [(25 – 20)/(50 – 20) × (625 – 125) + 125] × 2

(2)

[(5 × 500)/30 + 125] × 2 = 416.67

(3)

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Macartney and Offit Round this to the nearest value of 5 = 415. This figure is the reciprocal of the serum dilution at which a 50% reduction in plaques occurred.

3.2. PRNA of Stool 1. Stool samples are collected and stored frozen at –20°C until ready for use (see Note 5). 2. After thawing, dilute samples 1 in 10 in PBS (1 gm/9 mL PBS). 3. Vortex each diluted sample 4×, for 15 s, until thoroughly mixed. 4. Centrifuge at 1800g for 20 min at 4°C. 5. Remove supernatant, and heat-inactivate it for 30 min in a 56°C water bath. 6. Discard the pellet. 7. The stool suspension supernatant can now be diluted and assayed (see Subheading 3.1.), as for serum specimens.

3.3. ELISA for Detecting RV-Specific Human Serum IgA or IgG Abs The following method is an indirect ELISA that has been used extensively for detection and/or titration of RV-specific Abs from serum. The specificity of the assay is directed by the antigen, which can be any single-shelled group A RV strain, although the authors most commonly use the simian strains SA11 or RRV, or the bovine strain, WC3, because they are easily propagated in cell culture. Single-shelled virus is formed prior to assembly of the viral outer capsid proteins, VP4 and VP7, thus, the exposed VP is predominantly VP6, which is highly conserved among all RV strains. Generally, however, the same single-shelled virus should be used in all comparative studies. The methods for detection of IgA or IgG differ only in the isotype of the detector Ab employed (see Subheading 2.2.). 1. All specimens from the same individual should be plated on the same day. Plates should be covered during all incubations. 2. Coat positive wells with 200 ng/well virus diluted in sterile PBS, using 100 µL/well (1 in 500 dilution of virus). Coat negative wells with 100 µL/well sterile PBS. Cover and place in a humidified chamber at 4°C overnight. 3. After overnight incubation, wash the plates 5× with PBS-T. Block all wells with 200 µL/well of SK-T. Incubate at 25°C for 1 h. 4. Wash the plates 5× with PBS-T. Dilute specimens and positive control in SK-T. Plate serial two-fold dilutions from 1 in 100, until specimen is determined to be negative. Dilute negative control 1 in 100, in SK-T. Add 100 µL of each dilution to one negative, and two positive-coated wells. Incubate at 25°C for 1 h. 5. Wash the plates 5× in PBS-T. Add 100 µL/well of detector Ab diluted 1 in 1000 in SK-T. Incubate at 25°C for 1 h. 6. Wash the plates 5× in PBS-T. Mix substrate–enzyme reagents 1:1 in a trough. Add 100 µL/well combined substrate–enzyme solution. Incubate at 25°C for 5 min. 7. Add 100 µL/well 85% ortho-phosphoric acid, and mix by agitating gently.

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8. Immediately after adding ortho-phosphoric acid, read the plates at 450 nm on an ELISA plate reader. 9. Data can be interpreted from the ELISA plates, if the end point titer of the positive control is within one dilution of its previously determined titer, and the negative control is negative. A specimen is considered positive at a given dilution if the mean optical density (OD) value of the positive wells is at least 2× greater than the value of the corresponding negative well and has a value of at least 0.1 OD; otherwise, a specimen is considered negative.

3.4. ELISA for Human Fecal IgA and IgG Abs to RV Determination of Ab concentrations in feces or intestinal secretions is complicated by variable dilution of Ab depending on the method of collection, water content of the specimen, entrapment of Ab by mucus, and degradation of Ab by intestinal proteases. To control for these factors, the authors determine virus-specific fecal IgA or IgG as a percentage of total fecal IgA or IgG. Two methods to express relative Ab concentrations can be used. Determination of the concentration of both virus-specific and total Ab in ng/mL protein, using OD values from a standard curve generated by testing known concentrations of highly purified human IgA or IgG, is preferred (see Subheading 3.4., step 10). Alternatively, the end point titer of virus-specific Ab can be expressed as a percentage of the end point titer of the total Ab (see Note 7). 1. Stool samples are stored at –20°C until ready for use. After thawing, dilute samples 1 in 10 in EBSS (1 g/9 mL EBSS). Vortex each diluted sample 4×, for 15 s each, until thoroughly mixed. Clarify the specimen by centrifugation at 1800g for 5 min at 4°C. The supernatant can be stored at –20°C until ready for use. 2. For detection of virus-specific IgA and virus-specific IgG, coat positive wells with 200 ng/well of virus diluted 1 in 500 in 0.1 M carbonate–bicarbonate buffer (use 100 µL/well). For detection of total IgA and standard, coat positive wells with antihuman IgA diluted 1 in 1000 in buffer (or, for detection of total IgG and standard, coat positive wells with antihuman IgG diluted 1 in 1000). Coat negative wells with 100 µL/well buffer. Cover and place in moisture chamber at 4°C overnight. 3. Wash the plates 5× with PBS-T. Block all wells with 200 µL/well of BSA-T, and incubate at 25°C for 1 h. 4. Wash the plates 5× with PBS-T. Dilute specimens and controls in BSA-T. For the detection of virus-specific IgA or IgG, plate serial two-fold dilutions of the specimens and positive control from 1 in 20, until the specimen is determined to be negative. For the detection of total IgA or IgG, plate serial two-fold dilutions of the specimens and positive control from 1 in 1000, until the specimen is determined to be negative. Add 100 µL of each dilution to one negative and two positive wells. Incubate at 25°C for 1 h.

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5. Wash the plates 5× with PBS-T. Add 100 µL/well detector Ab diluted 1 in 1000 in BSA-T. Incubate at 25°C for 1 h. 6. Wash the plates 5× with PBS-T. Mix substrate–enzyme reagents 1:1 in a trough. Add 100 µL/well of combined enzyme–substrate solution. Incubate for 5 min at 25°C. 7. Add 100 µL/well ortho-phosphoric acid, and mix. Read the plates immediately at a wavelength of 450 nm. 8. Data are interpreted as for the serum ELISA above, but a specimen is considered positive at a given dilution, if the OD value of the positive wells is both at least 0.1 OD units and 3× greater (not 2× greater, as for serum assay) than the mean value of the corresponding negative well. To calculate the absolute concentration of virus-specific and total fecal IgG and IgA, plot the net ODs obtained for each dilution of the commercially prepared IgA or IgG against the corresponding known concentration. Net ODs are calculated by subtracting the control value of the corresponding negative well from the OD value of the sample. From the plotted values, generate a best-fit curve. The correlation coefficient of the curve should be >0.9, and preferably >0.95. Plot the net OD values for each sample on the curve, to obtain the corresponding Ab concentrations (see Note 8).

3.5. Mononuclear Cell Isolation 1. Bleed mice to obtain serum. Dissect spleen, PP, MLN, small intestinal LP, from animals, and place them in separate Petri dishes on ice, containing 20 mL 5% FBS in HBSS. 2. Create single-cell suspensions from each tissue by homogenizing with an L-shaped (bent) 21-gage needle. 3. Spin cells and blood at 290g for 5 min. Aspirate supernatant, and resuspend cells in fresh media (except spleen and blood). 4. To spleen, add ACK solution, using 3 mL/spleen, to digest the red cells. Leave for 1 min on ice, then add fresh media and centrifuge at 290g for 5 min. Leave spleen cells on ice, again for 1–2 min, to allow large clumps of cell debris to sediment, collect supernatant, and centrifuge again. 5. To the pellet of cells obtained from spinning the blood sample, add 2.5 mL PBS. Add 4 mL Ficoll-Hypaque to a separate 15-mL conical tube, and then gently overlay this with the PBS–blood suspension. Centrifuge at 580g for 25 min. Harvest the gray-colored lymphocyte layer with a Pasteur pipet and bulb, and resuspend it in PBS. Centrifuge at 290g for 5 min, then resuspend the pellet in complete media. 6. Remove debris from PP cells by passing the cell suspension through a prewet cotton column. Remove debris from MLN by filtering through a cell collector (sieve). LP cells require no further treatment. 7. Assess viability of all the cell populations by trypan blue (use cells that are >90% viable), and adjust the cell concentrations prior to plating (see Subheading 2.5.).

3.6. ELISPOT Assay for Detection of Murine ASCs from Tissues and Circulation The ELISPOT assay relies on binding of virus-specific ASCs to a solid phase (virus-coated 96-well plate), in which a single virus-specific ASC is counted

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as an individual spot under magnification by light microscopy. In addition, the total number of ASCs in a specimen may be determined by coating plates with isotype-specific Abs (see Note 9). Cells to be assayed may be obtained from PP, MLN, small intestinal LP, spleen, or serum. After obtaining mononuclear cells, the procedure is performed as follows: 1. The day prior to the assay, coat individual wells of 96-well milliliter HA plates (Millipore, Bedford, MA), with 100 µL PBS or 200 ng CsCl purified RV, suspended in 100 µL PBS. Cover and place in moisture chamber at 4°C overnight. 2. Wash the plates 4× with PBS, then block for 1 h at room temperature with 100 µL/well 1% BSA diluted in PBS (stored at 4°C) for the following wells: spleen (IgA and IgG), MLN (IgA and IgG), PP (IgG), LP (IgG), and blood (IgA, IgM, and IgG). Alternatively, block for 1 h with 100 µL/well 1% skim milk in TBS, for cells from the following tissues: spleen (IgM), MLN (IgM), PP (IgM and IgA), and LP (IgM and IgA). 3. Wash the plates 4× with PBS. Dilute cells to suspensions containing 1 × 106, 2 × 105, or 4 × 104 cells in complete media. Add 100 µL of each cell suspension to positive and negative (PBS-coated) wells. Incubate for 4 h at 37°C in an atmosphere of 5% CO2. 4. Wash the plates 5× with PBS, then 5× with PBS-T. Add 100 µL/well of horseradish peroxidase (HRP)-labeled isotype-specific detector Abs diluted in 1% BSA-T. For goat antimouse IgG–HRP, dilute the Ab 1 in 1000 in BSA-T, for goat antimouse IgM–HRP, dilute 1 in 1000, and for goat antimouse IgA–HRP, dilute 1 in 2000. Incubate overnight at 4°C in a humidified chamber. 5. Wash the plates 4× with PBS. Add 100 µL/well development solution for 15 min (see Subheading 2.5.). Wash with distilled water, then dry the plates overnight at 25°C. 6. Examine the wells by light microscopy under 8–10-fold magnification. The average number of spots in duplicate negative wells (without antigen coating) are subtracted from the average number of spots in duplicate positive wells (coated with antigen). The number of virus-specific, isotype-specific ASCs are expressed per 106 mononuclear cells from the sample assayed. Alternatively, the number of virus-specific, isotype-specific ASCs can be expressed as a percentage of the total number of ASCs in the sample (see Note 9).

4. Notes 1. The choice of ELISA plates for determination of RV-specific IgA and IgG is important for consistent and high binding of RV antigen. The method described has been optimized using electroimmunoassay or RIA 96-well plates from Costar (Cambridge, MA), cat. no. 3590. 2. To obtain serum that does not contain RV-specific IgA and IgG, for use as the negative control in the serum ELISA (2.2. g), screening the sera of infants likely to be RV Ab negative is performed. Choose nonbreast-fed infants who are between 6–9 mo of age at the end of the summer or fall months. These infants are

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4.

5. 6. 7.

8.

9.

Macartney and Offit less likely to have been exposed to natural RV infection, have not acquired breast milk RV-specific Abs, and are likely to have lost maternal transplacentally transferred RV-specific Abs. After identifying seronegative samples, pool these to use as the negative control for serum IgA and IgG determinations. During the PRNA, care should be taken during all washing steps of the cell monolayer to aspirate the solution from the same position in the well, for example, at the 6 o’clock position, because disruption of the monolayer by the Pasteur pipet tip may be misinterpreted as a plaque. In the PRNA, preparation of the agarose–MSO–trypsin solution for overlay of inoculated MA104 cells is important to optimize, in order to maintain the integrity of the cell monolayer over the next 3 d. The agarose can be added to the MSO–trypsin while still bubbling, however, the mixture should be cooled to 37°C before plating. A mixture that is too hot will cause cell death, and therefore an inaccurate or no plaque count will ensue. Stool and serum samples for PRNA and ELISA can be stored for up to 6 mo at –20°C, after which degradation of Ab may occur. ELISA plates may be rotated on a platform for more uniform Ab binding during the addition of the detector Ab, in the case of inconsistent results. In the ELISA for fecal Ab determinations, RV-specific Ab to total Ab concentrations can also be expressed as a percentage. Calculate this by dividing the reciprocal of the end point titer of virus-specific IgA or IgG, by the reciprocal of the end point titer of the corresponding total IgA or IgG. ODs of individual sample dilutions, obtained by the fecal IgA and IgG ELISA, should be plotted on the best fit curve only when their OD reading corresponds to the linear segment of the curve. Because binding at increasing Ab concentrations (generally >100 ng/mL) commonly occurs in an exponential relationship, determining Ab concentrations at higher ODs (when the relationship between these values is nonlinear) is not accurate. The ELISPOT assay can also be used to determine the total number of Ab-secreting cells in a sample of mononuclear cells. To determine the total number of IgA-secreting cells, coat additional wells with 100 µL per well goat-affinity purified Ab to mouse IgA, diluted 1 in 1000 in PBS. Likewise, for total IgG and IgM-ASC determinations, coat wells with the same dilution of the isotype-specific purified Ab. The remainder of the assay is the same.

References 1. Offit, P. A. (1994) Immunologic determinants of protection against rotavirus disease. Curr. Top. Microbiol. and Immun. 85, 229–254. 2. Davidson, G. P., Hogg, R. J., and Kirubakaran, C. P. (1983) Serum and intestinal response to rotavirus enteritis in children. Infect. Immun. 40, 447–452. 3. Coulson, B., Grimwood, K., Hudson, I., Barnes, G., and Bishop, R. (1992) Role of coproantibody in clinical protection of children during reinfection with rotavirus. J. Clin. Microbiol. 30,1678–1684.

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4. O’Ryan, M. L., Matson, D. O., Estes, M. K., and Pickering, L. K. (1994) Anti-rotavirus G-type specific and isotype-specific antibodies in children with natural rotavirus infection. J. Infect. Dis. 169, 504–511. 5. Kaila, M., Isolauri, E., Soppi, E., Virtanen, E., Laine, S., and Arvilommi, H. (1992) Enhancement of the circulating antibody secreting cell response in human diarrhea by a human lactobacillus strain. Ped. Res. 32, 141–144. 6. Coulson, B. S., Fowler, K. J., Bishop, R. F., and Cotton, R. G. H. (1985) Neutralizing monoclonal antibodies to human rotavirus and indications of antigenic drift among strains from neonates. J. Virol. 54, 14–20 7. Ward, R. L., Clemens, J. D., Knowlton, D. R., Rao, M. R., Van Loon, F. P. L., Huda, N., Ahmed, F., Schiff, A. M., and Sack, D. A. (1992) Evidence that protection against rotavirus diarrhea after natural infection is not dependent on serotype-specific neutralizing antibody. J. Infect. Dis. 166, 1251–1257. 8. Totterdell, B. M., Patel, S., Banatvala, J. E., and Chrystie, I. L. (1988) Development of a lymphocyte transformation assay for rotavirus in whole blood and breast milk. J. Med. Virol. 25, 27–36 9. Offit, P. A., Hoffenberg, E. J., Pia, E. S., Panackal, P. A., and Hill, N.L. (1992) Rotavirus-specific helper T cell responses in newborns, infants, children and adults. J. Infect. Dis. 165, 1107–1111. 10. Ward, R. and Bernstein, D. (1995) Lack of correlation between serum rotavirus antibody titers and protection following vaccination with reassortant RRV vaccines. Vaccine 13, 1226–1232. 11. Ward, R. L., Knowlton, D. R., Zito, E. T., Davidson, B. L., Rappaport, R., and Mack, M. E. (1997) Serologic correlates of immunity in a tetravalent reassortant rotavirus vaccine trial. J. Infect. Dis. 176, 570–577. 12. Isolauri, E., Joensuu, J., Suomalainen, H., Loumala, M., and Vesikari, T. (1995) Improved immunogenicity of oral D × RRV reassortant rotavirus vaccine by Lactobacillus casei GG. Vaccine 13, 310–312. 13. Khoury, C. A, Brown, K. A., Kim, J. E., and Offit, P. A. (1994) Rotavirus-specific intestinal immune response in mice assessed by enzyme-linked immunospot assay and intestinal fragment culture. Clin. Diag. Lab. Immunol. 1, 772–728. 14. Chen, W. K., Campbell, T., vanCott, J., and Saif, L. J. (1995) Enumeration of isotype-specific antibody-secreting cells derived from gnotobiotic piglets inoculated with porcine rotaviruses. Vet. Immunol. Immunopath. 45, 265–84. 15. Bridger, J. C. and Oldham, G. (1987) Avirulent rotavirus infections protect calves from disease with and without inducing high levels of neutralizing antibody. J. Gen. Virol. 68, 2311–17. 16. Offit, P. A. and Dudzik, K. I. (1990) Rotavirus-specific cytotoxic T lymphocytes passively protect against gastroenteritis in suckling mice. J. Virol. 64, 6325–6328. 17. Dharakul, T., Rott, L., and Greenberg, H. B. (1990) Recovery from chronic rotavirus infection in mice with severe combined immunodeficiency: virus clearance mediated by adoptive transfer of immune CD8+ T lymphocytes. J. Virol. 64, 4375–4382. 18. Yolken, R. H., Wyatt, R. G., Kim, H. W., Kapikian, A. Z., and Chanock, R. M. (1978) Immunologic response to infection with human reovirus-like agent: measurement of

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anti-human reovirus-like agent immunoglobulin G and M levels by the method of enzyme-linked immunosorbant assay. Infect. Immun. 19, 540–546. Babiuk, L. A., Acres, S. D., and Rouse, B. T. (1977) Solid-phase radioimmunoassay for detecting bovine (neonatal calf diarrhea) rotavirus antibody. J. Clin. Microbiol. 6, 10–15. Gust, I. D., Pringle, R. C., Barnes, G. L., Davidson, G. P., and Bishop, R. F. (1977) Complement fixing antibody response to rotavirus infection. J. Clin. Microbiol. 5, 125–130. Matsuno, S., Inouye, S., Hasegawa, A., and Kono, R. (1982) Assay of human rotavirus antibody by immune adherence hemagglutination with a cultivable human rotavirus as antigen. J. Clin. Microbiol. 15, 163–165. Martin, M. L., Gary, G. W. Jr., and Palmer, E. L. (1979) Comparison of hemagglutination-inhibition, complement-fixation and enzyme-linked immunosorbant assay for quantitation of human rotavirus antibodies. Arch. Virol. 62, 131–136. Matsuno, S., Inouye, S., and Kono, R. (1977) Plaque assay of neonatal calf diarrhea virus and the neutralizing antibody in human sera. J. Clin. Microbiol. 5, 1–4. Knowlton, D. R., Spector, D. M., and Ward, R. L. (1991) Development of an improved method for measuring neutralizing antibody to rotavirus. J. Virol. Methods 33, 127–134. Shaw, R. D., Fong, K. J., Losonsky, G. A., Levine, M., Maldonado, Y., Yolken, R., Tilores, J., Kapikian, A. Z., Vo., P. T., and Greenberg, H. B. (1987) Epitope-specific immune responses to rotavirus vaccination. Gastroenterology 93, 941–50. Offit, P. A., Clark, H. F., and Plotkin, S. A. (1984) Response of mice to rotaviruses of bovine or primate origin assessed by radioimmunoassay, radioimmunoprecipitation, and plaque-reduction neutralization. Infect. Immun. 42, 293–300.

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8 In Vivo Study of Immunity to Rotaviruses Selected Methods in Mice Manuel A. Franco and Harry B. Greenberg 1. Introduction Rotaviruses (RVs) are important human pathogens. The murine model of RV infection has been very useful in clarifying the mechanisms that mediate clearance of primary RV infection, and the mechanisms that mediate immunity to reinfection. The use of immunodeficient strains of mice, immunodepletion studies with specific monoclonal antibodies (MAbs), and passive transfer of purified cells are three basic, complementary experimental approaches that have been used for this purpose, and are the subject of this chapter. These experimental approaches analyze the outcome of RV infection under artificial conditions; thus, the relevance of the results obtained, to the physiological immune response of immunocompetent mice or humans, is at times difficult to establish. For example, immunodeficient strains of mice frequently develop compensatory immune mechanisms that are potentially absent or nonfunctional in immunocompetent mice. Immunodepletion experiments introduce into the experimental animal high (nonphysiological) levels of antibodies (Abs) that potentially have other immunomodulatory effects different from the desired one, and, many times, depletion strategies fail to completely eliminate the target cell population. Passive cell transfer experiments analyze the antiviral capacity of a specific cell population (many times abnormal in number), independent of other cells with which it may normally interact, and in an environment to which it is at least partially alien. Because of these and other limitations of such experimental approaches, one should be very careful in selecting adequate controls, and cautious in the interpretation of the results, by taking into account results obtained with two or three of the approaches and From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ

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analysis of the characteristics of the immune response in normal animals. The combination of two or three strategies (for example, passive cell transfer into immunodeficient hosts and immunodepletion of selected cell populations in immunocompetent hosts) have proven particularly useful in the study of immunity to RV. RV replication is almost exclusively limited to mature enterocytes (1). For this reason, the anti-RV immune response is typical of an immune response generated in the intestinal mucosa. The characteristics of the immune response against RV have recently been reviewed (2), and will only be briefly mentioned here. The acquired immune system is necessary for clearance of primary RV infection, because, in most cases, T- and B-immunodeficient SCID and Rag-2 knockout (–/–) mice become chronically infected with murine RV (3,4). Clearance of primary RV infection has been shown to be mediated by CD8+ T-cells in mice with a C57BL/6 background (5). In the absence of CD8+ cells clearance of primary RV infection is delayed, compared to immunocompetent mice, but other mechanisms quickly compensate and mediate viral clearance. The CD8+ cells seem to mediate their antiviral effect independently of perforin, fas, and interferon (IFN)-γ (6). In BALB/c mice, in addition, clearance of primary infection also depends on a CD4+ T-cell activity (7). Protection from viral reinfection after primary infection with murine RV is dependent on (Ab), most probably local RV-specific IgA (4,8,9). Hybridomas that produce nonneutralizing Abs against the internal structural viral protein (VP)6 or the surface protein VP4 can mediate immunity to RV in vivo (10). Upon transfer into chronically infected Rag-2 –/– mice, α4β7(high) memory phenotype CD8+ T-cells are highly efficient at clearing RV infection. In contrast, α4β7– memory CD8+ T-cells are inefficient or ineffective, depending on the cell numbers transferred. These experiments indicate that functional memory for RV resides primarily in memory cells that display the mucosal homing receptor α4β7 (11). What follows introduces each of the three experimental approaches for the study of RV immunity in the mouse, followed by selected protocols that have proven useful in these studies. Many protocols have been adapted from other investigators employing approaches known to be effective in other murine models of viral infection (12,13).

1.1. Use of Immunodeficient Strains of Mice Table 1 lists the immunodeficient strains of mice that have been used to study immunity to RV in mice. With the exception of the SCID, nude, and lpr mice, these mice have been generated by gene-targeted mutation. The basic assumption of experiments using these immunodeficient strains of mice is that

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Table 1 Immunodeficient Strains of Mice Use to Study Immunity to RV Mouse strain

Deficiency

Genetic background

Ref.

SCID Rag-2 Nude

T- and B-cells T- and B-cells Thymus dependent T-cell

(3,5) (4,6) (5,27)

αβ TCR γδ TCR αβ/γδ TCR JH D µMt β2–Microglobulin

αβ TCR T-cell γδ TCR T-cell αβ/γδ TCR T-cells B-cell B-cell Class 1 restricted T-cells

Perforin 1pr IFN-γ

Perforin-dependent CTL Fas-dependent CTL IFN-γ

BALB/c, C57BL/6 C57BL/6 × 129a BALB/c, C57BL/6, outbred C57BL/6 C57BL/6 C57BL/6 × 129 C57BL/6 × 129 C57BL/6 C57BL/6, C57BL/6 × 129 C57BL/6 × 129 C57BL/6 BALB/c, C57BL/6

a

(5) (5) (5) (4,8,9) (7,9) (4) (6) (6) (6)

Incompletely backcrossed to 129.

any difference in immunity to RV between a normal mouse, and the immunodeficient mouse can be attributable, in some way, to the gene missing in the immunodeficient mouse. Because of this, every effort should be made to include control immunocompetent mice in each experiment that do not have other potential genetic differences, or differ in other variables (age, diet, and so on) from the deficient mice. When the immunodeficient strain of mice under study is not in an inbred genetic background, less-optimal controls are mice that have been derived from the same parental strains as the immunodeficient mice, or that are heterozygote for the particular gene deletion. Other important considerations to be addressed when studying RV infection in immunodeficient mice are: 1. As previously stated, many of these mice develop compensatory immune mechanisms that may not be relevant to the immune response of normal mice. Knock-out mice with inducible gene mutations have been produced in an attempt to obviate this problem (14). Some examples of known compensatory immune mechanisms in immunodeficient strains of mice used for the study of immunity to RV are: a. SCID mice have been shown to develop natural killer cells with increased antiviral activity (15). b. αβ T-cell receptor (TCR) –/– and Rag-2 –/– mice develop a TCR–, CD3+ population of cells that is not present in normal mice (16,17).

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2. Some of these mice have incomplete deficiencies: a. Nude mice have T-cells, especially γδ T-intraepithelial lymphocytes (18). Some of these T-cells develop in situ in the gut (18,19). b. β2-Microglobulin –/– mice have some residual class 1 molecule expression on the cell surface (20). Certain class 1 molecules (e.g., Ld) have been shown to present at the cell surface in the absence of β2-microglobulin (20). For this reason, these mice have low levels of class 1 restricted CD8+ T-cells. Administration of an anti-CD8 MAb to β2-microglobulin knock-out mice efficiently eliminates this residual population of T-cells (4). c. SCID mice develop low levels of Abs, especially of the immunoglobulin (Ig)M class (21). The presence of these Abs varies, depending on the genetic background and the age of the mice (21). Rag-2 –/– mice seem to develop fewer Abs than SCID mice, but also have some early pro B-cells (22). d. The αβ and αβ/γδ TCR knock-out mice develop inflammatory bowel disease (23). 3. There are also differences in the response of the mice according to their genetic background: a. Although SCID mice on a BALB/c background invariably become chronically infected with murine RV, 40% of C57BL/6 SCID mice clear primary RV infection (5). b. Although BALB/c adult nude mice invariably have a delay in clearing primary RV infection (antigen [Ag]-shedding can last up to 9–10 d), compared to immunocompetent controls, C57BL/6 and outbred nude mice often can shed Ag for only 1–2 d (5).

1.2. Immunodepletion Studies In these studies, mice are treated with an Ab, usually a MAb that blocks or eliminates a specific component of the immune system. Any difference in the outcome of RV infection between a normal mouse and the treated mouse is then attributed to the component blocked or eliminated by the specific MAb. An advantage of this experimental system over studies with immunodeficient strains of mice is that mice receiving this kind of treatment do not have a long time to develop elaborate compensatory mechanisms for the deficiency created. Two types of immunodepletion studies have been done in studies of RV immunity in mice: depletion of selected populations of cells, such as CD4, CD8, and γδ TCR+ T-cells (4–9); and depletion of cytokines: IFN-γ (6). In what follows, the authors present protocols useful for production of the MAb ascites used in the immunodepletion studies and the depletion protocols that have been useful in study of RV immunity. Finally, a protocol for verification of the depletion experiments is provided.

1.3. Verification of Cell Depletion On the day of RV infection, depleted and nondepleted mice should be killed, and cells from relevant organs should be stained for the marker of interest, and subsequently analyzed by fluorescent-activated cell sorter (FACS). Typically,

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if mice are depleted of CD8 or γδ T-cells, it is sufficient to evaluate depletion of intraepithelial lymphocytes (IEL) (60–90% of IELs are CD8+) and spleen. If CD4 cells have been depleted, analysis should be extended to the lamina propria (LP) where these cells predominate. The target cells should be stained with an Ab that recognizes a different epitope than the Ab used for the depletion. This precaution is taken, so that residual Ab used for the depletion will not mask the possibility of detecting nondepleted cells by the Ab used for staining. Table 2 gives the names of hybridoma clones producing MAbs that have been useful for the depletion and subsequent cell staining to verify the depletion. What follows presents a protocol for the isolation of IEL and LP lymphocytes (see Subheadings 3.4. and 3.5.).

1.4. Cell Transfer Experiments The passive cell transfer experiments that have been performed to study RV immunity in mice are shown in Table 3. The recipient mice in these experiments include immunocompetent naive adult and infant mice, and immunodeficient SCID and Rag-2 –/– mice that are chronically infected with murine RV. The cells that have been transferred include hybridomas producing RV-specific IgA or IgG MAbs, spleen cells that are selectively depleted of certain cell populations and specific cell populations, purified by FACS. The authors present a protocol for the transfer of a purified population of mouse cells to chronically infected Rag-2 –/– mice (6) (see Subheadings 3.6 and 3.7). 2. Materials 2.1. Production of Ascites Fluid 1. 2. 3. 4. 5.

6–8-wk-old nude mice. Pristane (2,6,10,14-tetramethyl-pentadecane). Hybridoma cells (see Notes 1 and 2). Large-gage needle. Centrifuge.

2.2. Cell Depletion Protocol 1. FACS machine.

2.3. Cytokine Depletion Protocol 1. Anti-IFN-γ (clone XMG1.2 American Type Culture Collection [ATCC], Rockville, MD).

2.4. Isolation of IELs 1. RPMI 1640 cell culture medium (Gibco-BRL, Rockville, MD). 2. Heat-inactivated fetal bovine serum (hiFBS). 3. 100 µg/mL streptomycin sulfate.

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Table 2 MAbs Used for Depletion and Verification of Cell Depletion MAb specificity CD8 CD4 γδ T-cells

Depleting MAb

Verifying MAb

Ref.

2.43 GK 1.5 UC7-13D5

53 6.7 RM4-5 GL3

(4–9) (5,7) (5,8)

Hybridomas used to produce MAb for depletion can be obtained from the ATCC. Labeled MAb can be obtained from Pharmingen (San Diego, CA).

Table 3 Summary of RV Specific Passive Cell Transfer Experiments Cell transferred IgA hybridomas Splenocytes depleted of CD8 and Thy 1+ cells Spleen and IEL CD8+ cells Recombinant protein specific CD8+ cells Perforin –/– CD8+ cells α4β7 high- and low-memory CD8+ cells α4β7 high- and low-memory B cells

Recipient host Immunocompetent and SCID BALB/c mice C57BL/6 and BALB/c mouse pups

Ref. (10)

SCID BALB/c SCID BALB/c

(28) (29) (30)

Rag-2–/–

(6)

Rag-2–/– Rag-2–/–

(11) (31)

4. 100 U/mL penicillin. 5. Glass wool column (12-mL plastic syringe, loosely packed with glass wool, and equilibrated with RPMI 1640 medium). 6. 40% and 75% Percoll (Pharmacia, Uppsala, Sweden).

2.5. Isolation of LP Lymphocytes 1. 2. 3. 4. 5. 6.

Phosphate-buffered saline (PBS), pH 7.2. PBS, pH 7.2, containing 5 mM ethylenediaminetetraacetic acid (EDTA). Dulbecco’s modified Eagle’s medium (DMEM) medium. Dispase (1.5 mg/mL). Water bath. Centrifuge.

2.6. Preparation of Donor Cells 1. Iso-osmotic medium (77 mosM Hanks’ buffered salt solution containing 16.8 mM HEPES, 0.246 M sorbitol, 38.5 mM glucose). 2. RPMI 1640 cell culture medium. 3. HEPES buffer.

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2.7. Enrichment of CD8+ Cells Prior to Sorting 1. 2. 3. 4. 5. 6. 7. 8.

Polystyrene plates (100-mm diameter). Rabbit antimouse IgG serum (Sigma, St Louis, MO). Anti-CD4 MAb GK1.5. PBS, pH 7.2. RPMI 1640 medium containing 10% and 20% FBS. DMEM containing 1% FBS. FACS machine. Rag-2 –/– or SCID mice.

3. Methods 3.1. Production of Ascites Fluid 1. Inoculate 6–8-wk-old outbred nude mice with 0.5 mL pristane intraperitoneally (ip). 2. One to 2 wk later, inoculate these mice with 2 × 106 – 107 of the appropriate hybridoma cells ip (see Notes 1 and 2). 3. One wk after the inoculation of cells, start periodic follow-up of mice, until the mouse’s abdomen is dilated. 4. When the mouse is ready for collection of ascites fluid, using appropriate sedation, puncture the abdomen with a large-gage needle in the lower right quadrant, and let the ascitic fluid flow freely into a 15-mL tube (see Notes 3 and 4). 5. Spin the ascites fluid at 1000g for 10 min at 4°C, then take the ascitic fluid, and discard the pellet. 6. Ascites from several mice are pooled, aliquoted, and stored at –70°C until use. 7. Before inoculation of the experimental mice, the ascites fluid is thawed and respun again at 1000g for 10 min at 4°C, and the pellet discarded (see Note 5).

3.2. Cell Depletion Protocol 1. Inoculate experimental mice ip with 0.5 mL ascites fluid 5, 4, and 3 d before RV infection, on the day of RV infection, and on d 3, 6, and 9 after infection. 2. On the day of RV infection, depleted and nondepleted control mice are killed, to verify depletion of the appropriate population of cells in spleen, IELs and LP by FACS analysis (see Notes 6 and 7).

3.3. Cytokine Depletion Protocol The only cytokine that has been depleted in studies of RV immunity is IFN-γ. For depletion of IFN-γ, mice are administered 2 mg purified anti-IFN-γ MAb (hybridoma clone XMG1.2) on d –1, 1, 4, and 7 after infection (see Notes 8 and 9).

3.4. Isolation of IELs 1. Dissect the small intestine from one mouse (from duodenum to the cecum), and place it in a Petri dish with RPMI media.

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2. Place the intestine on a humid surface and meticulously remove Peyer’s patches and trim attached residual mesentery. Peyer’s patches are identified as discrete protrusions on the antimesenteric border of the intestine. 3. Open the small intestine longitudinally, carefully remove fecal material, and wash the intestine with fresh RPMI media. 4. Cut the intestine into 2-cm segments. 5. The pieces of intestine are then transferred to a 50-mL tube with 20 mL RPMI media containing 2% hiFBS, 100 µg/mL streptomycin sulfate, and 100 U/mL penicillin, and incubated for 30 min at 37°C, with vigorous shaking every 10 min. 6. The medium containing the IEL is then separated with a sieve from the pieces of intestine (see Note 10). 7. The medium containing IELs is passed over a glass-wool column. 8. The eluent of the column, containing IELs, is centrifuged, and the pellet is resuspended in 4 mL 40% Percoll, layered onto 2 mL 75% Percoll, and centrifuged at 25°C for 20 min at 600g. 9. Collect IELs at the interface of the 75/40% layers, and wash once. 10. Cells are adjusted to an appropriate volume for subsequent use.

3.5. Isolation of LP Lymphocytes 1. Pieces of intestine from Subheading 3.4., step 6, are washed once with PBS (without Ca and Mg ions), and then placed in 50-mL tubes with 20 mL PBS (without Ca and Mg ions) containing 5 mM EDTA for 20 min, with vigorous shaking every 10 min. 2. Discard the supernatant, and add fresh 5 mM EDTA/PBS, and repeat the same process. 3. Wash the pieces of intestine twice with 30 mL PBS, and once with 10 mL DMEM; medium should be clear after the wash. If the medium is not clear, repeat the washes. 4. Place the pieces of intestine in a 50-mL tube with 10 mL DMEM containing 1.5 mg/mL Dispase (without FBS), and incubate for 20 min in a 37°C water bath (see Notes 11 and 12). 5. Separate tissues from supernatant. Add 0.5 mL FBS to the supernatant to inactivate the Dispase, and immediately centrifuge the cells at 4°C, and keep on ice. 6. Repeat the above procedure a total of 3× with the tissue fragments, and pool the cells obtained at each cycle (see Note 13). 7. Continue with a Percoll gradient, as for the IEL (see Subheading 3.4., step 8).

3.6. Preparation of Donor Cells 1. Prepare a single-cell suspension from the desired organ of donor mice. 2. To avoid cell aggregates, incubate cells in low-ionic-strength iso-osmotic medium for 10 min on ice. A stock of this buffer can be sterilized, aliquoted, and frozen at –20°C, to be thawed when needed. 3. Wash and resuspended the cells in RPMI 1640 containing 10% hiFBS and 20 mM HEPES.

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4. At this point, cells can be stained and sorted by FACS, or can be enriched prior to sorting. The decision about what procedure to follow depends on the concentration of the target cell in the cell suspension: As a general rule, B-cells (40–60% of spleen cells) can be directly sorted, CD4 and CD8 T-cells (11% and 24% spleen cells, respectively) require further enrichment to reduce the long, potentially harmful, and expensive sorting time. The next protocol presents, as an example, a procedure for CD8+ T-cell enrichment from spleen donor cells before purification by FACS.

3.7. Enrichment of CD8+ Cells Prior to Sorting 1. Overnight at 4°C, coat 100-mm polystyrene plates with 100 µg/mL rabbit antimouse IgG serum and ascites fluid (1/8 dilution) from the anti-CD4 MAb GK 1.5 in 4 mL sterile PBS. 2. Wash the plates 3× with sterile PBS. Do not discard the PBS with the Abs because it can be reused to coat another plate in a similar fashion. 3. Adjust the donor cell suspension to 4 × 107 cells/mL (see Subheading 3.6., step 4). 4. Add 4 mL cell suspension to each plate, and incubate for 20 min at room temperature on a level surface. 5. Gently swirl the Petri dish, and remove the nonadherent cells. 6. Add the nonadherent cells to a fresh-coated Petri dish, and repeat the procedure (see Note 14). 7. Centrifuge the cells at 4°C, and resuspend them at a concentration of 1 – 2 × 108 cells/mL of RPMI containing 10% FBS. If a biotinylated MAb is used for staining the cells prior to sorting, replace the RPMI 10% FBS (high content of biotin) with DMEM 1% FBS. 8. Add the appropriate amount of labeled Ab (determined by previous titration) to stain the cells, and incubate for 20 min on ice, and then wash them with the medium used for the staining step. 9. Double-sort the cells on the FACS and recover the sorted cells in RPMI 20% fetal calf serum (FCS) in order to increase cell viability after the sort (see Notes 15 and 16). 10. Inoculate, ip, the desired amount of cells to Rag 2 –/– or SCID mice chronically infected with RV. 11. After the transfer, stool and serum samples from the Rag-2 –/– mice are collected and analyzed for RV Ag and RV-specific Abs by enzyme-linked immunosorbent assay (ELISA) (see Note 17).

4. Notes 1. The development of ascites in the mice inoculated with this protocol may vary, depending on the hybridoma cell being used. A good strategy to increase the yield of mice producing ascites is to inoculate mice with 0.2 mL fresh (immediately after being harvested) ascitic fluid from another mouse. Female nude mice appear to take hybridoma cells easier than male nude mice. 2. The protocol given above has several advantages: Nude mice will only produce low levels of endogenous Abs in the ascites; nude mice will not reject xenogenic

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3.

4.

5.

6.

7.

8.

9.

Franco and Greenberg hybridoma cells; outbred nude mice have a relatively low cost, and are sturdier than, for example, SCID mice. Ascitic fluid must be manipulated using sterile technique. To control for sterility, pelleted hybridoma cells from step 5 can be cultured in media without antibiotics. After verification of sterility, these cells can be frozen down for reserve, or reinoculated into nude mice to produce more ascites fluid. Yield of ascitic fluid per mouse varies from 2 to 15 mL. According to guidelines for the use of laboratory animals, mice should never be tapped more than twice. An interval of 1–3 d between the first and the second tap is adequate. Before use, the ascites fluid should be checked for specificity and adequate titer of MAb. This can be done by staining of appropriate Ag-expressing target cells with serial dilutions of the ascites fluid and a second stage antiserum labeled with a fluorocrome, and subsequent FACS analysis of the cells. Batches of ascites fluid effective for in vivo immunodepletion studies of selected cell populations will usually be able to stain cells at dilutions above 1 in 105. The efficiency of depletion obtained with the above protocol varies, depending on the cell type being depleted, the strain of mice that is being used, and the Ab employed. The above protocol is very efficient: over 99 and 85% for depleting CD8 (IEL) and CD4 T-cells (LP), respectively, in immunocompetent mice (5). In β 2 microglobulin –/– mice, depletion of the residual CD8+ T-cells (mostly γδ IELs) is less efficient (78%) (4). For γδ T-cells, the protocol has been shown to deplete over 80% of IEL in αβ TCR –/– mice. In contrast, the administration of the anti-γδ MAb depleted approx one-half of the γδ IELs present in the JHD –/– mice (8). The remaining γδ T-cells in the JHD –/– mice showed a reduction in the intensity of staining. A similar finding has been reported by others (24) using a different anti-γδ TCR MAb, suggesting that administration of anti-γδ TCR MAb can modulate TCR expression, rather than deplete the cell population. The effect of immune depletion (given similar depletion efficiencies) probably varies, depending on factors such as the strain of murine RV used to infect the mice and the strain of mice infected. For example, C57BL/6 mice, depleted of CD4 cells, clear primary infection with the EC strain of murine RV in a fashion similar to nondepleted mice (5). In contrast, a low percentage of BALB/c mice, depleted with the same MAb and protocol, shed low levels of viral Ag for a prolonged time after infection with the EW strain of murine RV (7). Protocols for the depletion of a cytokine in general require that the Ab preparation used for the depletion be purified, to permit careful control of the dose administered, and to avoid Ab aggregates that may influence the outcome of the experiment (25,26). Control of depletion of cytokine in the serum of the mice can be done by measuring the cytokine involved, with methods such as ELISA or bioassays (25,26). Control of depletion of a cytokine in intestinal tissues, which is required for studies of immunity to RV infection, poses a more complicated problem. A partial solution to this problem is to verify that the same preparation of MAb has

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10.

11. 12.

13.

14. 15. 16.

17.

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a reproducible effect on the infectious process of another intestinal mucosal pathogen (6). The IEL yield with this protocol varies, depending on the strain and age of the mouse used. With a young (8-wk-old) C57BL/6 mouse, typical yields vary between 2 and 8 × 106 cells per mouse, with more than 90% viable. Cell viability is lowered if, after digestion, the Dispase is not immediately inactivated by the addition of FCS, and by keeping the cells at 4°C. After digestion of the tissue with Dispase, the cells tend to form aggregates. Before putting cells on the Percoll gradient, aggregates should be disrupted by gentle pipeting to avoid disruption of the gradient. Cell yield with this protocol varies, depending on the age and strain of mouse used. With an adult C57BL/6 mouse, 0.8–2 × 106 cells (over 90% viable) are recovered. After the enrichment protocol described above, CD8+ cells pass from a concentration of 10% to 40–60%. Double-sorting of the cells is necessary to obtain purity above 99.7%, which will generally assure that the population transferred is pure. An alternative method for enriching a cell population prior to FACS purification is to use commercially available magnetic beads coated with diverse MAbs (31). This method, although more expensive, is more efficient. When it is desired to sort rare (<10%) populations of cells, this method is clearly advisable. As for the immunodepleting experiments, after passive cell transfer, verification should be made that the transferred animals only have the transferred population of cells.

Acknowledgments: This work was supported by grants R37AI21632 and DK38707 from the NIH and by a V.A. Merit Review grant. HBG was a medical investigator at the Palo Alto Veterans Administration Medical Center. MAF is supported by a Walter and Idun Berry Fellowship. References 1. Greenberg, H. B., Clark, H. F. P., and Offit, A. (1994) Rotavirus pathology and pathophysiology. Curr. Top. Microbiol. Immunol. 185, 255–283. 2. Franco, M. A., Feng, N., and Greenberg, H. B. (1996) Molecular determinants of immunity and pathogenicity of rotavirus infection in the mouse model. J. Infect. Dis. 174 Suppl 1, S47–S50. 3. Riepenhoff-Talty, M., Dharakul, T., Kowalski, E., Michalak, S., and Ogra, P. L. (1987) Persistent rotavirus infection in mice with severe combined immunodeficiency. J. Virol. 61, 3345–3348. 4. Franco, M. A. and Greenberg, H. B. (1995) Role of B cells and cytotoxic T lymphocytes in clearance of and immunity to rotavirus infection in mice. J. Virol. 69, 7800–7806.

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5. Franco, M. A. and Greenberg, H. B. (1997) Immunity to rotavirus in T cell deficient mice. Virology 338, 169–179. 6. Franco, M. A., Tin, C., Rott, L., VanCott, J. L., McGhee, J. R., and Greenberg, H. B. (1997) Evidence for CD8+ T cell immunity to murine rotavirus in the absence of perforin, fas and gamma interferon. J. Virol. 71, 479–486. 7. McNeal, M. M., Rae, M. N., and Ward, R. L. (1997) Evidence that resolution of rotavirus infection in mice is due to both CD4 and CD8 cell-dependent activities. J. Virol. 71, 8735–8742. 8. Franco, M. A. and Greenberg, H. B., (1997) CD8+ T cells can mediate almost complete short term and partial long term immunity to rotavirus in mice. J. Virol. 71, 4165–4180. 9. McNeal, M. M., Barone, K. S., Rae, M. N., and Ward, R. L. (1995) Effector functions of antibody and CD8+ cells in resolution of rotavirus infection and protection against reinfection in mice. Virology 214, 387–397. 10. Burns, J. W., Siadat, P. M., Krishnaney, A. A., and Greenberg, H. B. (1996) Protective effect of rotavirus VP6-specific IgA monoclonal antibodies that lack neutralizing activity [see comments]. Science 272, 104–107. 11. Ros, J. R., Williams, M. B., Rott, L. S., Butcher, E. C., and Greenberg H. B., (1998) Expression of the mucosal homing receptor alpha4beta7 correlates with the ability of CD8+ memory T cells to clear rotavirus infection. J. Virol. 72, 726–730. 12. Leist, T. P., Cobbold, S. P., Waldmann, H., Aguet, M., and Zinkernagel R. M. (1987) Functional analysis of T lymphocyte subsets in antiviral host defense. J. Immunol. 138, 2278–2281. 13. Sarawar, S. R., Sangster, M., Coffman, R. L., and Doherty, P. C. (1994) Administration of anti-IFN-gamma antibody to beta 2-microglobulin-deficient mice delays influenza virus clearance but does not switch the response to a T helper cell 2 phenotype. J. Immunol. 153, 1246–1253. 14. Schwenk, F., Kuhn, R., Angrand, P. O., Rajewsky, K., and Stewart, A. F. (1998). Temporally and spatially regulated somatic mutagenesis in mice. Nucleic Acids Res. 26, 1427–1432. 15. Taterka, J., Cebra, J. J., and Rubin, D. H. (1995) Characterization of cytotoxic cells from reovirus-infected SCID mice: activated cells express natural killer- and lymphokine-activated killer-like activity but fail to clear infection. J. Virol. 69, 3910–3914. 16. Mombaerts, P., Mizoguchi, E., Ljunggren, H. G., Iacomini, J., Ishikawa, H., Wang, L., Grusby, M. J., Glimcher, L. H., Winn, H. J., Bhan, A. K., et al. (1994) Peripheral lymphoid development and function in TCR mutant mice. Int. Immunol. 6, 1061–1070. 17. Shinkai, Y. and Alt, F. W. (1994) CD3 epsilon-mediated signals rescue the development of CD4+CD8+ thymocytes in RAG-2–/– mice in the absence of TCR beta chain expression. Int. Immunol. 6, 995–1001. 18. Lin, T., Matsuzaki, G., Kenai, H., Nakamura, T., and Nomoto, K. (1993) Thymus influences the development of extrathymically derived intestinal intraepithelial lymphocytes. Eur. J. Immunol. 23, 1968–1974.

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19. Saito, H., Kanamori, Y., Takemori, T., Nariuchi, H., Kubota, E., TakahashiIwanaga, H., Iwanaga, T., and Ishikawa, H. (1998) Generation of intestinal T cells from progenitors residing in gut cryptopatches [see comments]. Science 280, 275–278. 20. Quinn, D. G., Zajac, A. J., Hioe, C. E., and Frelinger, J. A. (1997) Virus-specific, CD8+ major histocompatibility complex class I-restricted cytotoxic T lymphocytes in lymphocytic choriomeningitis virus-infected beta2-microglobulin-deficient mice. J. Virol. 71, 8392–8396. 21. Nonoyama, S., Smith, F. O., Bernstein, I. D., and Ochs, H. D. (1993) Strain-dependent leakiness of mice with severe combined immune deficiency. J. Immunol. 150, 3817–3824. 22. Young, F., Ardman, B., Shinkai, Y., Lansford, R., Blackwell, T. K., Mendelsohn, M., Rolink, A., Melchers, F., and Alt, F. W. (1994) Influence of immunoglobulin heavy- and light-chain expression on B-cell differentiation Genes Dev. 8, 1043–1057 (published erratum appears in Genes Dev. 9, 3190). 23. Mombaerts, P., Mizoguchi, E., Grusby, M. J., Glimcher, L. H., Bhan, A. K., and Tonegawa, S. (1993)ß Spontaneous development of inflammatory bowel disease in T cell receptor mutant mice. Cell 75, 274–282. 24. Kaufmann, S. H., Blum, C., and Yamamoto, S. (1993) Crosstalk between alpha/beta T cells and gamma/delta T cells in vivo: activation of alpha/beta T-cell responses after gamma/delta T-cell modulation with the monoclonal antibody GL3. Proc. Natl. Acad. Sci. USA 90, 9620–9624. 25. Sheehan, K. C., Ruddle, N. H., and Schreiber, R. D. (1989) Generation and characterization of hamster monoclonal antibodies that neutralize murine tumor necrosis factors. J. Immunol. 142, 3884–3893. 26. Finkelman, F. D., Katona, I. M., Mosmann, T. R., and Coffman, R. L. (1988) IFN-gamma regulates the isotypes of Ig secreted during in vivo humoral immune responses. J. Immunol. 140, 1022–1027. 27. Eiden, J., Lederman, H. M., Vonderfecht, S., and Yolken, R. (1986) T-cell-deficient mice display normal recovery from experimental rotavirus infection. J. Virol. 57, 706–708. 28. Offit, P. A. and Dudzik, K. I. (1990) Rotavirus-specific cytotoxic T lymphocytes passively protect against gastroenteritis in suckling mice. J. Virol. 64, 6325–6328. 29. Dharakul, T., Rott, L., and Greenberg, H. B. (1990) Recovery from chronic rotavirus infection in mice with severe combined immunodeficiency: Virus clearance mediated by adoptive transfer of immune CD8+ lymphocytes. J. Virol. 64, 4375–4382. 30. Dharakul, T., Labbé, M., Cohen, J., Bellamy, A. R., Street, J. E., Mackow, E. R., Fiore, L., Rott, L., and Greenberg, H. B. (1991) Immunization with baculovirusexpressed recombinant rotavirus proteins VP1, VP4, VP6, and VP7 induces CD8+ T lymphocytes that mediate clearance of chronic rotavirus infection in SCID mice. J. Virol. 65, 5928–5932. 31. Williams, M. B., Rosé, J. R., Rott, L. S., Franco, M. A., Greenberg, H. B., and Butcher, E. C. (1998) The memory B cell subset responsible for the secretory IgA response and protective humoral immunity to rotavirus expresses the intestinal homing receptor, alpha4beta7. J. Immunol. 161, 4227–4235.

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9 Evaluation of Rotavirus Vaccines in Small Animal Models Max Ciarlet and Margaret E. Conner 1. Introduction The high morbidity and mortality of rotavirus (RV) infections has spurred the development of RV vaccines (1–13). Although children naturally infected with RV commonly undergo multiple infections, primary infections in children generally induce disease, and children are normally protected against severe disease during subsequent infections (1,2,6–8,14). For RV, the immunologic mechanisms responsible for protection are poorly understood, but antibody (Ab) in the intestine appears to be the primary mechanism of protection (2,15,16). Because RV is a localized enteric infection, and induction of intestinal mucosal immune responses was expected to be required for protection, live orally administered vaccines were pursued first. Vaccine development of the live attenuated vaccines proceeded to clinical trials in humans without prior animal testing. In August 1998, RotashieldTM, a three dose, live attenuated tetravalent (TV), rhesus rotavirus (RRV) vaccine produced by Wyeth Lederle Vaccines and Pediatrics (West Henrietta, NY), was licensed. This vaccine shows promise, eliciting ~80% protection against severe disease (6,7,12,17–19). The recent detection or emergence of new RV serotypes in humans suggests that incorporation of additional P- and G-serotypes into this vaccine may be necessary in the future (20–22). Additional concerns with the use of live attenuated vaccines include interference of vaccine replication by other enteric pathogens (common in children from the underdeveloped world); neutralization by maternal Ab; limited replication competence of animal strains, because of the host range restriction observed with RVs; and safety, because of the possibility of producing new virulent virus, emerging by reassortment of circulating wild-type (WT) virus with the vaccine virus. From: Methods in Molecular Medicine, Vol. 34: Rotaviruses and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ

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Development and testing of nonreplicating RV vaccines have also been pursued, and will be the focus of this chapter. The use of nonreplicating immunogens presents additional challenges beyond that of developing RV vaccines effective in young children against potential infection by multiple serotypes of RV. The nonreplicating immunogen must be able to induce protective immune responses against the target virus. Traditionally, nonreplicating vaccines have been thought to be poor inducers of mucosal immune responses and protection of the mucosa. Without amplification of the vaccine virus by replication, a high dose of nonreplicating immunogen may be required. To enhance the immune response to nonreplicating immunogens, development and testing of new adjuvants and/or delivery systems, and alternative routes of immunization to boost immunogenicity and protective efficacy, are needed. If administered orally, the nonreplicating immunogens must be stable in the digestive environments of the stomach and intestine. The application of molecular biology to vaccine research has opened new vistas for development and production of subunit vaccines, including peptides, live vectors that express the proteins of interest at the site of infection, expression or purification of viral proteins (VPs), expression of virus-like particles (VLPs), or immunization with DNA. All of these approaches have been tried for development of a RV vaccine with variable success (1–4,23–35). To date, little success has been reported with VP4 or VP7 peptides, individually expressed proteins, or RV proteins in bacterial or viral expression vectors (1,2,5). Lack of success with these approaches probably results from the fact that Abs induced to soluble peptides or proteins do not react with the protective conformational epitopes on the virus. RV VLPs maintain the structural and functional characteristics of native particles, and are particulate immunogens that induce both systemic and mucosal immunity (23,28). VLPs have been shown to be highly protective when administered parenterally, orally, or intranasally to animals, and they can induce passive, lactogenic, or active protection (2,3,20,23,32,36). Recently, immunizations with DNAs encoding RV proteins have been tested in mice, with variable success (24–27,29,30,35). Testing of potential subunit vaccines has relied on the use of animal models to generate immunogenicity and efficacy data. Studies in animals are set up to determine efficacy and safety in animal models prior to U. S. Food and Drug Administration approval of clinical trials in humans due to the high cost of human trials. The use of animal models has also been essential to increase basic understanding of RV infection, pathology, disease, and immunity, including correlates of protection (see Chapter 7), ultimately aiding vaccine development. Currently, there are two routinely used small animal models of RV infection, rabbit and mouse. The rabbit model was the first small animal model developed to examine active humoral immunity and protection (37–42), followed by the development of the adult mouse model (43–46). Both of these

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models are infection models, as will be discussed in more detail in Subheading 3. The only disease animal model is in gnotobiotic piglets (see Chapter 6). Its use has been limited because of high cost, limited number of animals available, limited time that animals can readily be maintained in isolators (8 wk), and the need for specialized equipment, facilities, and staff; however, the work has yielded very interesting data (see Chapter 6). The authors’ laboratory has evaluated the induction of protective active immunity, using live- or nonreplicating, inactivated RV vaccine or subunit RV vaccines consisting of different formulations of VLPs, delivery systems, and adjuvants administered parenterally, orally, or intranasally to rabbits or mice (1,3,4,20,23,31,33,34,47,48).

1.1. Immunogen Production and Adjuvant Usage to Evaluate RV Vaccines In evaluation of RV vaccines in animals, production and purification of RV in tissue culture may be required to provide a control immunogen in animals, challenge virus for animals, and antigen (Ag) for assays to detect Ab responses. Protective efficacy of a nonreplicating vaccine should be compared to protective efficacy of live virus administered orally. An additional control that can be included is inactivated virus administered by the same route as the test vaccine. Alternatively, inactivated virus can be tested as a vaccine. Inactivated virus administered parenterally to rabbits and mice induces intestinal immune responses and protection from RV infection (23,32,39,47,49). Production and characterization of a nonreplicating test vaccine are also required. This laboratory has tested VLPs of different formulations.

1.2. Expression of RV Structural Proteins in the Eukaryotic Baculovirus–Insect Cell System Baculoviruses (BVs) (family Baculoviridae) belong to a group of viruses that possess a large double-stranded circular DNA genome (80–200 kbp for different species) that infect a great variety of insects. The BV most widely studied as a vector is the Autographa californica nuclear polyhedrosis virus (AcNPV), which is useful as an expression system, because some of its nonessential genes (such as polyhedrin, or gene p10) can be replaced by a substantial amount of foreign DNA, without substantially affecting the virus replication cycle. AcNPV grows readily in cells derived from ovarian tissue of a clonal isolate of Spodoptera frugiperda IPLB-SF21-AE (S. frugiperda 9 [Sf9] cells), and recombinant proteins can be produced at various levels of expression of the total VPs (50–52). All 11 RV proteins have been successfully cloned and expressed in eukaryotic insect cells with BV recombinants, using the BV AcNPV as an

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expression vector. Individually expressed RV proteins have been used to study protein function, virus morphology, protein–protein interactions, and protein–RNA interactions (8,28,53–56; see Chapter 3). Upon co-expression of the recombinant RV structural proteins in insect cells, VLPs self-assemble (28,53,54,57). Co-expression of different combinations of the RV structural proteins results in the assembly of VLPs with different structures. VLPs self-assemble following expression of VP2 alone, yielding core-like particles (53), following co-expression of VP2 and VP6 alone (yielding single-shelled subviral particles), or with VP4 (double-layered 2/6- or 2/4/6-VLPs, respectively), or following co-expression of VP2, VP6, and VP7, with or without VP4 (yielding triple-layered 2/6/7- or 2/4/6/7-VLPs, respectively) (28). All VLPs maintain the structural and functional characteristics of native particles, and are particulate immunogens that induce both systemic and mucosal active, passive, or lactogenic immunity when administered to animals (2–4,20,23,32– 34,36,47,57). As vaccines, VLPs offer several advantages, including safety (no possibility of reversion to virulence), purity, stability (>6 yr at 4°C), and the ability to alter protein content and type (2/6-, 2/6/7-, or 2/4/6/7-VLPs) to permit determination of the minimal protein content that affords protection. BV recombinants containing the gene(s) of interest can be produced using one of a number of published methods or commercial kits (50–52). Subheading 3.2. provides a general outline of the methodology to produce, purify, and characterize RV VLPs (28,31,57; see also Chapter 3).

1.3. Adjuvants and Delivery Systems Another important consideration when testing RV vaccines is the choice of adjuvant or delivery system, which varies according to the route of immunization. For RV vaccines, a good adjuvant should augment the level of the mucosal immune response, compared to immunogen alone. Typically, systemic immune responses are then also augmented. Choices of adjuvants that are currently approved for used in humans is limited to parenteral administration of aluminum phosphate (AlP) and aluminum hydroxide (AlOH). No mucosal adjuvants are currently approved. Development of new safe adjuvants or delivery systems, especially for mucosal administration is under intense investigation. New adjuvants and delivery systems that have been evaluated with RV vaccines show promise. The choice of adjuvant or delivery system for future RV vaccine evaluations should be based on current available information.

1.3.1. Parenteral Adjuvants Although parenteral immunizations were initially considered unlikely to induce protective immunity to RV, parenteral immunization has proven quite successful in small animal models (1–4,20,23,32,39,47,49,57). Parenteral

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immunization with inactivated virus and VLPs have been evaluated with Freund’s adjuvant, AlP, AlOH, and QS-21 (3,20,23,32,39,47,57). Freund’s adjuvant is widely used in initial vaccine testing in animals, because it elicits excellent humoral and cell-mediated immunity. However, Freund’s adjuvant cannot be used for human or veterinary vaccines because of its toxicity. Parenteral administration of live or inactivated virus or VLPs in Freund’s adjuvant have elicited the highest levels of RV-specific serum and mucosal Abs and high levels of protection (20,23,39). In rabbits and mice, the immune response or protective efficacy elicited by RV or VLPs in AlP or AlOH is variable, but appears to be lower than that elicited by Freund’s adjuvant in other studies (3,20,23,32,39,57). A more recently developed adjuvant that has undergone field trials in humans is QS-21, a saponin adjuvant (58–60), which is a purified triterpene glycoside obtained from extracts of the bark of a South American Molina tree, Quillaja saponaria. In mice, VLPs in QS-21 elicited significantly higher Ab responses or protection than VLPs in AlOH (32,57). In rabbits, live- or inactivated-SA11 RV or VLPs in QS-21 elicited significantly higher immune responses than AlP, but not necessarily higher protective efficacy (2,3,20,23). If parenteral vaccines are pursued, then QS-21 or other promising adjuvants that are efficacious in eliciting a protective immune response, without toxicity or adverse reactions, need to be tested further.

1.3.2. Mucosal Adjuvants Although parenteral administration of RV vaccines induce mucosal immunity, direct administration of a vaccine to a mucosal surface is expected to be more effective at stimulating mucosal immunity. Until recently, it was thought that mucosal adjuvants would be absolutely required to induce immunity to nonreplicating immunogens administered mucosally. However, a mucosal adjuvant was found not to be necessary for the induction of immune responses to Norwalk VLPs, although the response was more consistent with cholera toxin (CT) adjuvant (61). In contrast, RV VLPs administered intranasally without adjuvant were poorly immunogenic or protective, but were highly protective when administered with CT (33,34). No mucosal adjuvants are currently licensed for use in humans. Two mucosal toxins, CT and Escherichia coli heat-labile toxin (LT), are strong mucosal adjuvants, but are unacceptable for use in humans because the doses required to obtain adjuvanticity also elicit diarrhea (62,63). Many groups (64–68) are pursuing mutants of both CT and LT with reduced toxicity that retain adjuvanticity. LT-R192G, a reduced-toxicity mutant of LT, is a highly effective adjuvant for RV VLPs administered intranasally: This adjuvant induced protection equivalent to VLPs in native CT or LT (33,34). In safety studies in human volunteers, no adverse reactions were noted at 5-, 25-, or 50-µg doses of LT-R192G; a 25-µg dose of LT-R192G induced maximal Ab responses (69). These reduced-toxicity

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mutants offer promising alternatives as mucosal adjuvants, but before these and other new mucosal adjuvants are approved, they need to be extensively tested for safety and efficacy in humans. 1.3.3. Delivery Systems Delivery systems are being devised and tested to increase the delivery, uptake, or long-term release of intact immunogen at mucosal sites (70–74). Immunogens can be microencapsulated by using different formulations of biodegradable polymers, which were selected based on previous history of safety in humans. The efficiency of encapsulation varies with each immunogen, and commercial feasibility remains to be determined. Alteration of size and formulation of the microcapsules control their uptake and the timing of release of the immunogen. Increased uptake, immunogenicity, and protection have been obtained with lower doses of microencapsulated, live attenuated or inactivated RV administered orally or intramuscularly (im) compared to free virus (70–74). 1.4. Small Animal Models Both rabbit and mouse models have been instrumental in the development of new vaccination strategies, by providing systems to dissect systemic and local immunity following infection or immunization. However, both rabbit and mouse models are infection, not diarrhea, models. All-age rabbits and mice are readily infected by RV and shed high levels of RV in their stools, but diarrhea is age-restricted to animals of <2 wk of age (43,44,46,75). Therefore, protective efficacy in the rabbit and mouse is based on reduction of RV Ag shedding, not on amelioration of disease. Nevertheless, reduction of virus Ag shedding is likely to be a more stringent measure of protective efficacy than protection from disease because diarrhea does not occur without infection. Both animal models have another limitation: Only homologous strains (isolated from the same species) replicate efficiently and spread horizontally to uninoculated control animals, whereas heterologous virus strains (isolated from a different species) do not (this includes human RVs) (45,46,48,75). When new vaccines are being evaluated, a number of parameters will affect the results, including choice of immunogen, route of immunization, adjuvant, and animal model. Experiments to compare and optimize different RV vaccines in different animal models may provide conflicting results regarding the constituents of a protective immune response or protective efficacy. Whether vaccine evaluation data obtained in any of the animal models will accurately reflect results in children is not clear because none of the subunit vaccine candidates has progressed to testing in humans.

1.5. Propagation and Titration of WT RV Strains in Rabbits and Mice RVs that are propagated strictly by animal-to-animal passage are referred to as WT RV strains. WT murine or lapine RVs have been propagated in their

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respective species of origin (45,46,75). WT RVs are often used as the challenge virus in the mouse model, but have had limited use in the rabbit model. Upon tissue culture adaptation, attenuation of the virus may occur, rendering it less virulent or nonvirulent, compared to WT virus in the original host species from which it was isolated. In the mouse, adaptation of several WT mouse RV strains to tissue culture resulted in an increase in the infective dose in mice, compared to WT virus (46). Similar comparisons have not been performed in the rabbit model.

1.6. Rabbit Model of RV Infection The rabbit model was the first small animal model to be developed to examine basic parameters of active immunity, immunogenicity, and protective efficacy of RV vaccines (23,37–39,41,42,48,75). Compared to large animal models (pigs and cows) available at the time of development of the rabbit model, the use of rabbits offered advantages of cost-effectiveness, the ability to incorporate large numbers of animals in studies, the feasibility of isolation of large numbers of infected animals, short gestation, multiparous births, and the availability of RV naïve animals. Group A lapine RV strains (LRV, R-2, 82/311F, ALA, C-11, BAP-2) have been isolated worldwide, and those that have been characterized belong to the VP7 serotype G3 and VP4 genotype P[14] (41,76–80). Rabbits can be productively infected with homologous lapine RV strains up to at least 5 yr of age, which allows examination of active long-term immunity for vaccine studies (37–42,48,75). Recently, evaluation of heterologous (nonlapine) RV strains demonstrated for the first time that human RV strains and other animal heterologous strains replicate in rabbits (48); among these strains, only RRV replicates efficiently in rabbits (48,81). For all vaccine studies, this laboratory utilizes the homologous lapine RV ALA as challenge virus, to measure protective efficacy against a replication-efficient RV. Although natural RV infection has been reported to cause diarrhea, following experimental infection with lapine RV, rabbits develop limited or no diarrhea (2,5,23,37–42,48,75,79,82–84). Recently, the authors determined that RV-induced diarrhea in rabbits is age-restricted, and that both WT and tissue-culture-adapted lapine RV strains are virulent in 1-wk-old rabbits (75). Also, rabbits of >2 wk of age infected with lapine ALA RV developed marked intestinal histopathological lesions, irrespective of disease status, indicating that age-dependent compensatory mechanisms may mask disease manifested in the small intestine (75). When diarrhea was observed in RV-infected rabbits of >2 wk of age, it was probably caused by other pathogens, such as Clostridium spiroforme (75,84–89).

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1.6.1. Immunization of Rabbits Following are a series of methods for performing immunizations and virus challenge for vaccine evaluation in rabbits (see Subheading 3.5.). Prior to the start of an experiment, serum and fecal samples from all animals should be tested by enzyme-linked immunosorbent assay (ELISA) to ensure RV Ab-negative status. For all immunizations, group doses can be prepared in one syringe or tube. Group doses should contain at least one extra dose to provide extra material. Vaccines and all buffers or water for dilution are tested for endotoxin prior to administration to animals. Maximum endotoxin levels are 0.05 µg/dose.

1.6.2. Collection of Rabbit Serum and Fecal Samples Serum and fecal samples are collected to measure RV-specific Ab responses (0, 28, and 56 d postvaccination [DPV], and at 0 and 28 days postchallenge [DPC]). Fecal samples are collected from 0 to 14 DPC to measure virus Ag shedding.

1.7. Adult Mouse Model of RV Infection RV infections in mice were first described in the 1950s as epizootic diarrhea of infant mice (EDIM) (90,91) although the identification of the causal agent as RV was not made until more than a decade later (92–95). Neonatal mice (<2 wk of age) are readily infected with murine and some heterologous RVs and develop RV-induced diarrhea (45,46,96,97). Diarrhea is age-restricted in the mouse, so, diarrhea as an end point for a vaccine study has mostly been used for the study of passive immunity (96,98–101). Early studies reported that RV shedding is limited in amount and duration in mice >2 wk of age (92,94,95), but later it was observed that RV infection can be readily established in adult mice (43–46,49,97). The adult mouse model, like the rabbit model, is an infection, not a disease, model. Compared to the rabbit model, the use of mice offers advantages in cost, availability of inbred, outbred, and genetically altered mice (see Chapter 8), and availability of more specific reagents. A number of different strains of murine RVs have been isolated (EC, EW, EB, EL, EHP, YR-1), and all strains characterized belong to VP7 serotype G3, and either VP4 genotype P[16] or P[18] (102,103). Strains EW, EC, and EL are supposed to be direct descendants from the original EDIM isolate (46,90,91,93). Different murine strains have different 50% infective doses (ID50) or shedding doses (SD50) in mice, and the SD50 also can vary in different-age mice (45,46). Tissue-culture-adapted murine strains generally do not grow to high titer in vitro, and have higher SD50 in vivo than the analogous WT virus. Mice are susceptible to RV infection to >1 yr of age (97,104). Mice are usually not productively infected with heterologous (nonmurine) RV (45,48).

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For active immunity and vaccine studies, both WT murine RVs (EC, EW, and EDIM) and tissue-culture-adapted strains are used to challenge mice (29,33,45,46,74). However, it should be noted that, although some inbred strains of mice are not routinely susceptible to infection with tissue-culture-adapted RV strains, they are susceptible to infection with WT murine strains. Therefore, measurement of protective efficacy against a replication-efficient RV may require use of WT murine RV as a challenge virus. Mice, unlike other species infected with RV, develop either no or limited histopathologic changes in the small intestine at any age (2,105).

1.7.1. Immunization of Mice Following are a series of methods for performing immunizations and virus challenge for vaccine evaluation in mice (see Subheading 3.7.). Prior to the start of an experiment, all animals should be tested to ensure RV Ab-negative status. For all immunizations, group doses can be prepared in one syringe or tube. Group doses should contain at least 1–2 extra mouse doses to provide extra material. Vaccines and all buffers or water for dilution are tested for endotoxin prior to administration to animals. Maximum endotoxin levels are 0.05 µg/dose.

1.7.2. Processing of Mouse Fecal Samples To allow rigorous comparison of virus Ag shedding or Ab titers between samples, all samples are processed based on weight, not pellet number or standard volume size of the fecal processing buffer (see Subheading 3.8.2.1.). GuerinDanan et al. (106) recently showed that virus Ag-shedding results can be altered dramatically, if they are not normalized based on the amount of fecal sample. During fecal processing, samples should be kept on ice or at 4°C at all times. Individual sample processing should be completed within 1 d. Freeze-thawing of original and processed samples and storage time at 4°C should be minimized.

1.8. RV Detection in Mouse or Rabbit Fecal Samples Following virus infection or virus challenge of immunized rabbits and mice, the amount of virus-shedding is measured in daily fecal samples. By comparing the amount of virus shed from mock-immunized animals to immunized animals, the protective efficacy of the different immunogens is determined. Protective efficacy can be based on infectious virus shed or virus Ag shed. Infectious virus titers are determined by fluorescent focus assay (FFA) (see Subheading 3.9.2.) or plaque assay (see Subheading 3.9.3.). However, if the virus used to inoculate the animals is a WT virus, infectious virus assays may not detect, or may underestimate, the amount of infectious virus shed because the WT virus may have limited ability to replicate in vitro without adaptation. Many challenge studies in mice use WT murine RVs that are not readily

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detected in fecal samples from infected mice by infectious virus assays. Therefore, little attempt has been made in the mouse model to correlate virus-shedding as detected by ELISA and infectious assays. Ag detection is most often performed by ELISA because of ease of assay, assay time, and assay cost (see Subheading 3.9.1.). In rabbits inoculated with tissue-culture-adapted ALA RV that replicates efficiently in rabbits and in vitro, a good correlation exists between the infectious virus titer and level of virus Ag shed (23). How protective efficacy based on Ag detection correlates with protection from disease is not known, and cannot currently be measured in small animal models. Methods used for Ag detection in rabbits and mice are similar, so general methods are described, and any species-specific steps are noted (see Subheading 3.9.).

1.9. RV-Specific Ab Detection in Mouse or Rabbit Serum or Fecal Samples A number of assays can be used to measure neutralizing antibody (nAb) or virus-binding Ab responses in both models (23,38,48). Although nAb can be protective, the presence or titer of nAb does not always correlate with protection (2,15,16,107–109). The recently licensed live attenuated TV RV vaccine contains VP7s of the four serotypes of RV that most commonly infect children worldwide (G1–G4). The vaccine is TV because early clinical trials with monovalent vaccines indicated that nAb to multiple serotypes of virus is needed to achieve high levels of efficacy (6,7,18). However, at least in the small animal models, mice and rabbits immunized with 2/6-VLPs or double-layered virus, in the absence of nAb to either VP4 or VP7, can be partially or totally protected from RV challenge (23,33,34,47). There are several types of samples and methods to measure nAb or virus-binding Ab responses at mucosal sites. For sequential measurement of virus Ab in individual animals, which is present in the lumen of the intestine at the time of sampling, the authors routinely collect fecal samples. Although oral gavages can also be used, gavage techniques are technically more difficult and time-consuming (23,38,39). There is probably some degradation of Ab during transit in the intestine. In paired lavage and fecal samples, the authors found that Ab titers in lavage samples were approximately fourfold higher than Ab titers in fecal samples. Fragment cultures and enzyme-linked immunospot (ELISPOT) assay are other methods that are routinely used to measure Ab responses in the intestine (see Chapters 6 and 7). Fragment cultures and ELISPOT assay measure Ab production or number of Ab-secreting cells (ASC), respectively, present in the lamina propria at the time of tissue harvest. Both methods require euthanasia of animals to obtain samples, so sequential measurements of Ab responses in the same individual animal are not possible. Whether all the Ab produced in the lamina propria reaches the intestinal lumen

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is not known. ELISPOT measures the number of ASCs but does not directly measure the amount or the affinity of Ab. Isotype-specific and subclass-specific responses can also be measured by this method. Methods used for Ab detection in rabbits and mice are similar, so general methods are described in Subheading 3.10. Any steps specific to each species are noted.

1.10. RV Vaccine Evaluation 1.10.1. Protection from Live RV Challenge Both the rabbit and mouse model are infection, not disease, models (see Subheading 1.4.). Therefore, protection from RV challenge is evaluated by comparing the amount of virus Ag shed or infectious virus shed in vaccinated animals vs mock-vaccinated control animals. Protective efficacy can be evaluated by comparing single parameters, such as mean duration of shedding, day of onset, or mean amount of shedding per day (2,5,37–39,48). However, the authors now routinely measure protective efficacy of RV vaccines based on both the duration and the amplitude of virus Ag or infectious virus shed (23,33,48). The results of infectious virus (data are log-transformed) or viral Ag detection assays are graphed, and the area under the curve (AUC) for each animal is calculated. Protective efficacy in individual animals is expressed as the percent reduction in virus Ag or infectious virus shedding found by comparing the AUC for individual animals to the mean AUC for the mock-vaccinated control group. The protective efficacy in each group is then determined by calculating the mean AUC for each vaccine group, and comparing it to the mean AUC for the mock-vaccinated group (23,33). Prevention of infection may be a more stringent measurement of protection than protection from disease because virus Ag shedding occurs in the absence of disease (2,3,5,23,33,39,48,75). Although a significant reduction in infectious virus or virus Ag-shedding indicates a reduction in the replication of the challenge virus (23), it is still impossible to predict how a reduction of virus Ag or infectious virus shedding, observed in the adult rabbit and mouse models, will relate to protection from mild or severe RV disease in children. Unfortunately, sufficient data are not currently available to make this determination because the measurement of the protective efficacy of live attenuated RV vaccines in humans has relied on measurements of the severity of disease, based on clinical scoring, not on virologic or immunologic parameters (6,10).

1.10.2. Immune Response Following Live RV Challenge In the small animal models, when protective efficacy is high, the levels of the different immune responses measured before and after challenge are generally maintained (23,33,34,38,39,48,75). No or very low anamnestic

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increases in serum or intestinal Ab titers are observed because the challenge virus did not replicate sufficiently in the animal’s intestine to boost the vaccine-induced immune response. Besides analyzing protection based on virus shedding, protection can therefore also be confirmed by comparing immune responses before and after challenge. If an animal is protected from challenge, significant changes in Ab responses usually do not occur. If an animal is not totally protected from challenge, changes in the immune response can include increases (>4-fold) in serum Ab responses following challenge, an increase (or appearance) of an intestinal immunoglobulin (Ig)A or IgG response, or development of an Ab response to RV nonstructural proteins. Measurement of Ab responses may be a more sensitive measure of protection than measuring Ag shedding for detecting very low or abortive replication. Increases in Ab responses following virus challenge can be detected in the absence of detectable virus Ag shedding, suggesting that the animal was not completely protected from infection (23). When an animal is not protected from challenge, a rise in the serum and fecal Ab titers can be detected as early as 2–3 d postchallenge. It is important to remember that lack of significant increases in mean Ab titers for vaccine groups does not preclude significant increases in Ab responses in individual animals in the group.

1.11. Summary Small animal models have proven useful to test the preclinical protective efficacy of new candidate RV vaccines and to gain new insights into immunologic mechanisms that function to protect animals from RV infection. Preclinical results in small animal models, evaluating use of inactivated RV, parenteral and intranasal routes of administration, subunit VLP vaccines, DNA vaccines, and new adjuvants, and delivery methods (microencapsulation) are promising and warrant further investigation. The limitation of small animal models is that protection from disease cannot be measured. Therefore, the question remains, how much of a reduction in virus Ag or infectious virus shedding is necessary to result in significant protection against disease. The answer to this question awaits vaccine evaluation in the piglet disease model (see Chapter 6), and in children. Based on the results in these latter studies, we will be able to determine whether the small animal models predict success (protection from disease) in humans. This information is important because it will allow appropriate selection of animal models for future RV vaccine studies. No RV vaccine that has undergone preclinical testing in any animal model has yet been tested in children. 2. Materials 2.1. Propagation and Inactivation of Tissue Culture-Adapted RVs 1. African green monkey kidney, MA104, cells (BioWhittaker, Walkerville, MD). 2. Growth medium: Medium 199 with Earle’s salts (Irvine Scientific, Santa Ana, CA) supplemented with 5% (v/v) fetal bovine serum (FBS), 0.3 g/L glutamine, 1.22 g/L sodium bicarbonate (NaHCO3), 0.16 g/L penicillin, and 0.25 g/L streptomycin.

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3. Porcine trypsin (type IX; Worthington) (1 mg/mL) dissolved in 0.001 N HCl. 4. Maintenance medium: 199 Medium with Earle’s salts (Irvine Scientific), supplemented with 1 µg/mL porcine trypsin (type IX; Worthington), 0.3 g/L glutamine, 1.22 g/L NaHCO3, 0.16 g/L penicillin, and 0.25 g/L streptomycin. 5. Phosphate-buffered saline (PBS, pH 7.4): 140 mM NaCl, 10 mM Na 2 HPO 4 , 2 mM KH 2PO 4 . 6. Tris-sodium-calcium (TNC) buffer: 10 mM Tris-HCl [pH 7.4], 140 mM NaCl, 10 mM CaCl2. 7. 50 mM Tris-HCl, pH 8.0. 8. 10 mM disodium ethylenediamine tetraacetate (EDTA), pH 8.0. 9. Cesium chloride (CsCl), optical grade. 10. Fresh formalin (37% formaldehyde, Aldrich Chemical Co., Milwaukee, WI). 11. 0.35% solution NaHSO3. 12. 1 mg/mL 4'-aminomethyltrioxalin-hydrochloride (Psoralen) (Lee Biomolecular), dissolved in filter-sterilized 1:1 ethanol and water solution. 13. 0.1 N NaOH. 14. 0.1 N HCl. 15. Genetron (trichlorotrifluoroethane). 16. T-150 flasks (150 cm2). 17. Roller bottles (450 or 900 cm2). 18. 250-mL centrifuge tubes. 19. Sterile dram vials. 20. Pasteur pipets. 21. Ultracentrifuge with medium and high maximum speed (Beckman, SorvallDuPont, or others). 22. Rotors and tubes for centrifuges. 23. Sonicator. 24. Black-ray lamp, longwave UV-366 nm, 115 V, 0.16 Amps (Model UVL-21, UVP, Upland, CA). 25. Incubator (37°C). 26. Shaker incubator (37°C). 27. Spectrophotometer.

2.2. Propagation and Titration of WT RVs 1. 2. 3. 4.

WT RV stock. RV-free and pathogen-free mice or rabbits. Homogenizer. Materials for oral inoculations and fecal collection and processing (see Subheadings 2.3. and 2.5.).

2.3. Rabbit Model of RV Infection 1. RV Ab-free New Zealand white rabbits of either sex and any age, purchased from a specific pathogen free barrier-maintained rabbit facility. 2. Biohazard level (BL)2 animal housing.

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3. Isolation cages for rabbits. 4. High fiber autoclave feed for rabbits (Purina). Water is autoclaved municipal supply. 5. Tranquilizer anesthesia: acepromazine maleate (10 mg/mL). 6. Combination anesthesia: mixture of xylazine, ketamine hydrochloride, and acepromazine maleate (8 mg/kg, 40 mg/kg, and 2 mg/kg, respectively). 7. Alcohol and cotton balls. 8. Vacutainer ® brand blood collector set (butterfly) with a 21–23-gage, threequarter-inch needle 9. 10-mL serum separator Vacutainer tubes. 10. Commercial or proprietary adjuvants. If Freund’s adjuvant is used, the first dose is usually given in complete Freund’s adjuvant, and the second dose in incomplete Freund’s adjuvant. 11. Challenge virus: either WT or tissue culture-adapted lapine strain, for which the ID50 or virus Ag SD50 is known. 12. Sterile, blunt-end feeding 16- or 22-gage needles with ball ends (Popper & Son, New Hyde Park, NY). 13. 1-, 3-, 5-, and 10-mL syringes with 18-, 22-, and 25-gage needles of varying length. 14. Micropipeter and pipet tips. 15. Container (10 mL) to collect fecal pellets. 16. Autoclaved window screening (1/cage), cut to fit over top of rabbit litter pan, with approx 1–2-inch overlap. Two to three sets of screens are necessary for daily changeout of screens. 17. Autoclaved binder clips (4/cage) to attach screen to litter pan of cage.

2.4. Processing of Rabbit Fecal Samples 1. 2. 3. 4. 5.

Sterile metal spatulas and/or sterile wooden applicator sticks, 1/sample. Centrifuge tubes (15 or 50 mL). Balance. Medium-speed centrifuge. Fecal Ag buffer: Tris-buffered saline (TBS) (50 mM Tris-HCl [pH 7.4], 14 mM NaCl), supplemented with 0.32 g/L penicillin and 0.5 g/L streptomycin, or with 50 µg/mL gentamicin. Store for up to 8 wk at 4°C. 6. Fecal Ab buffer: PBS (pH 7.4) (140 mM NaCl, 10 mM Na2HPO4, 2 mM KH2PO4), supplemented with 0.1% Tween-20 (PBS-T), 0.1 mg/mL soybean trypsin inhibitor, and 0.1 mg/mL merthiolate. Store for up to 4 wk at 4°C. 7. Sterile plastic or glass dram vials. 8. Vortex mixer.

2.5. Mouse Model of RV Infection 1. Outbred or inbred strains of mice that are RV Ab-negative and mouse pathogen-free. Ab status should be confirmed prior to start of experiment. 2. Microtainer serum-separation tubes (Becton Dickinson, Franklin Lakes, NJ).

Evaluation of Rotavirus Vaccines in Small Animal Models 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

13. 14. 15. 16. 17.

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Sterile scalpel blades. Alcohol and cotton balls. Heat lamp. Broome style mouse restraint. Combination anesthesia: ketamine hydrochloride (3.75 mg/mouse), xylazine (0.19 mg/mouse), and acepromazine maleate (0.037 mg/mouse). Fecal collection cage. PBS, pH 7.4: 140 mM NaCl, 10 mM Na2HPO4, 2 mM KH2PO4, endotoxin-free. 5% sodium bicarbonate buffer diluted in endotoxin-free PBS. Commercial or proprietary adjuvants. Challenge virus, either WT or tissue-culture adapted-murine RV for which the ID 50 or virus Ag SD 50 has been determined. Maximum volume per mouse is 250 µL. Sterile 22-gage feeding needles with ball ends (1 mm) (Popper & Son). Micropipeter and pipet tips. Tuberculin syringes with 25-gage needles. Sterile plastic or glass dram vials. Microcentrifuge.

2.6. Processing of Mouse Fecal Samples 1. 2. 3. 4.

5. 6. 7. 8.

Sterile metal spatulas and sterile wooden applicator sticks, 1/sample. Sterile screw cap tubes (1.5 mL). Centrifuge tubes (15 mL). Fecal processing buffer: Filter-sterilized TNC (10 mM Tris-HCl [pH 7.4], 140 mM NaCl, 10 mM CaCl 2), supplemented with 0.05% Tween-20, 5 mM sodium azide, 1 mM benzamidine, (1 µL/mL) aprotinin, (10 µg/mL) leupeptin, and (10 µg/mL) pepstatin A. Store for up to 4 wk at 4°C. 0.5 M disodium EDTA, pH 8.0. Balance. Vortex mixer. Low-speed centrifuge.

2.7. Detection of RV Ag Shedding 1. Samples: 10% fecal suspensions (processed to detect fecal-Ag, see Subheadings 3.6.3. and 3.8.2.1.). 2. RV tissue culture passaged stock. 3. Flat-bottom polyvinylchloride microtiter plates (96 wells) (Dynatech). 4. Carbonate–bicarbonate buffer (CBB), pH 9.6: 15 mM Na2CO3, 35 mM NaHCO3. 5. PBS pH 7.4: 2 mM KH2PO4, 140 mM NaCl, 10 mM Na2HPO4. 6. PBS-T (0.05%). 7. BLOTTO (5% [w/v] Carnation nonfat dry milk in PBS). 8. Hyperimmune serum to RV prepared in two different species. 9. Normal serum (of same species as used for Ab coating of plate) or fetal calf serum (FCS) free of Ab to RV.

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10. Enzyme-conjugated Ab against immunoglobulin of species source used as the detector Ab. 11. Appropriate substrate for enzyme-conjugated Ab. 12. Appropriate stopping reagent for substrate. 13. ELISA plate reader with appropriate wavelength filter for the substrate. 14. Incubator (37°C). 15. Micropipet.

2.8. Detection of Infectious RV Shedding 1. Sterile 96-well plates (FFA) or 6-well plates (PFA) seeded with confluent MA-104 cells. 2. Sterile PBS, pH 7.4: 2 mM KH 2PO 4 , 140 mM NaCl, 10 mM Na 2 HPO 4 . 3. Samples: 10% fecal suspensions (processed to detect fecal Ag; see Subheadings 3.6.3. and 3.8.2.1.). 4. RV stock of known titers by FFA and PFA. 5. Porcine trypsin (type IX; Worthington): 1 mg/mL dissolved in 0.001 N HCl. 6. 5% CO 2 incubator. 7. Cold (–20°C) methanol. 8. Multichannel pipet and reagent reservoir. 9. Anti-RV hyperimmune serum. 10. Antispecies-specific immunoglobulins conjugated to fluorescein isothiocyanate (FITC) or other fluorescent dye. 11. Fluorescence microscope with proper filters for selected dye and proper objectives, with adequate focal length for examining 96-well plates. 12. 199 medium with Earle’s salts (Irvine Scientific), supplemented with 0.6 g/L glutamine, 2.44 g/L NaHCO3, 0.32 g/L penicillin, and 0.5 g/L streptomycin. 13. Solution A: 2X 199 medium with Earl’s salts (Irvine Scientific), supplemented with 0.6 g/L glutamine, 2.44 g/L NaHCO 3, 0.32 g/L penicillin, and 0.5 g/L streptomycin warmed to 37°C in a water bath. 14. Solution B: Sterile 1.2% Seakem LE (FMC Bioproducts, Rockland, ME) agarose that has been autoclaved and can be melted again in a microwave, and allowed to equilibrate in a 56°C water bath. 15. Solution C: Gibco pancreatin 4XNF (25 mg/mL). 16. Solution D: dieithylaminoethyl DEAE-dextran (10 mg/mL) (Pharmacia). 17. Solution E: Filter-sterilized 1% neutral red solution in H 2 O. 18. First overlay: 1:1 mixture of solutions A and B. Add C (final concentration of 0.5 µg/mL), and D (final concentration of 1 mg/mL) to 1:1 mixture of A and B warmed in a water bath at 56°C. 19. Second overlay: 1:1 mixture of solutions A and B. Add E (final concentration of 0.02%) to 1:1 mixture of A and B warmed in a water bath at 56°C. 20. Incubator (37°C).

2.9. Measurement of RV-Specific nAb 1. 96-well plates (fluorescent focus neutralization assay [FFNA]) or 6-well plates (plaque reduction neutralization assay [PRNA]) seeded with confluent MA104 cells.

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2. Virus stocks (as needed). 3. PBS, pH 7.4: 2 mM KH 2 PO 4 , 140 mM NaCl, 10 mM Na 2 HPO 4 . 4. Anti-RV hyperimmune serum for detection of Ag in FFNA, following fixation of cells (virus serotype not important). 5. FITC-conjugated antispecies-specific immunoglubulins to species from which anti-RV hyperimmune serum was obtained in FFNA. 6. Anti-RV hyperimmune sera raised against different virus serotypes to be tested in PRNA (specificity of the species in which the serum was raised is not important). 7. Medium 199 with Earle’s salts (Irvine Scientific), supplemented with 0.6 g/L glutamine, 2.44 g/L NaHCO 3, 0.32 g/L penicillin, and 0.5 g/L streptomycin. 8. Tubes (4-mL) to make dilutions. 9. Solution A: 2X Medium 199 with Earle’s salts (Irvine Scientific), supplemented with 0.6 g/L glutamine, 2.44 g/L NaHCO 3, 0.32 g/L penicillin, and 0.5 g/L streptomycin, warmed to 37°C in a water bath. 10. Solution B: Sterile 1.2% Seakem LE agarose that has been autoclaved, and can be melted again in a microwave oven and allowed to equilibrate in a 56°C water bath. 11. Solution C: Gibco pancreatin 4XNF (25 mg/mL). 12. Solution D: DEAE-dextran (10 mg/mL) (Pharmacia). 13. Solution E: Filter sterilized 1% neutral red solution in H 2 O. 14. First overlay: 1:1 mixture of solutions A and B. Add C (final concentration of 0.5 µg/mL), and D (final concentration of 1 mg/ml) to 1:1 mixture of A and B warmed in a water at 37°C. 15. Second overlay: 1:1 mixture of solutions A and B. Add E (final concentration of 0.02%) to 1:1 mixture of A and B warmed in a water at 37°C. 16. Incubator (37°C).

2.10 Measurement of RV Total-, Isotype-, or Subclass-Specific Ab 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13.

Flat-bottom polyvinylchloride microtiter plates (96 wells) (Dynatech). RV tissue culture stock previously titrated for use as Ag in this assay. CBB, pH 9.6: 15 mM Na 2CO 3, 35 mM NaHCO3 . PBS, pH 7.4: 2 mM KH 2 PO 4 , 140 mM NaCl, 10 mM Na 2 HPO 4 . PBS-T (0.05%). BLOTTO (5% [w/v] Carnation nonfat dry milk in PBS). Normal serum from same species as hyperimmune serum used for coating plate, or FCS that does not contain anti-RV Ab. Enzyme-conjugated Ab made to species being tested. Appropriate substrate for enzyme-conjugated Ab. Appropriate stopping reagent for substrate. ELISA plate reader with appropriate wavelength filter for the substrate. Micropipets. Incubator (37°C).

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3. Methods

3.1. Method for Propagation of RV in Tissue Culture 1. Grow MA104 cells in growth medium in T-150 flasks or roller bottles. 2. Upon confluency of MA104 cells, pour growth medium off cell monolayers, and wash cell monolayers once by adding maintenance medium overnight, or, alternatively, cells are washed 3× with PBS or maintenance medium (containing no trypsin) prior to virus infection. The washing must be done to remove residual serum from the cell monolayers prior to infection. 3. Sonicate the virus inoculum for 3 min at 4°C. 4. To increase infectivity of RV, proteolytic cleavage of VP4 is required. Therefore, the virus inoculum is pretreated with 10 µg/mL Worthington trypsin for 30 min at 37°C. 5. Determine the desired multiplicity of infection (MOI) (see Note 1). Usually a low MOI (0.01) is used to prepare virus stocks; a high MOI (1–40) is used for preparing virus for purification. To determine how much virus is needed to obtain the selected MOI, use the following formula: mL virus = (MOI) (number of cells in vessel) / (titer of virus stock). 6. Pour off the medium in the flask or roller bottle, and add 5 mL appropriately diluted trypsin-activated virus stock to the flask or roller bottle, and allow the virus to adsorb for 1 h at 37°C. 7. At the end of the absorption period, pour off the virus inoculum, and add 20 mL maintenance medium per flask or roller bottle. 8. Roll the bottles or incubate flasks at 37°C until the cytophatic effect is 85–90% (normally 1–5 d, depending on the RV strain). Check each day postinoculation. 9. Freeze and thaw the flasks or roller bottles 3×. 10. Pool virus from individual vessels and sonicate for 3 min at 4°C. Aliquot into centrifuge tubes if virus is going to be used for purification (see Subheading 3.1.1.), or into dram vials if virus will be used as stock. 11. Store virus stocks at –80°C.

3.1.1. Method for RV Purification (see Notes 2 and 3) 1. Clarify virus preparation by centrifugation at 4000g for 30 min at 4°C. 2. Discard the cell pellet, and retain the supernatant. 3. Concentrate the virus from the supernatant by centrifugation at 300,000g for 1 h at 4°C. 4. Pour off the supernatant, and dry the tubes, being careful not to disrupt pellets. 5. Using a total of 1 mL TNC, suspend all pellets with a Pasteur pipet. 6. Repeat 4× (final volume of pooled suspended pellets should be approx 4 mL). 7. Add 0.52 g CsCl/mL TNC, and mix by gentle inversion. The refractive index (RI) should be 1.3690. 8. Bring the total volume up to 10 mL using a premade CsCl/TNC mix (RI = 1.3690), and transfer to the ultracentrifuge tube(s). 9. Centrifuge at 280,000g for 18 h at 4°C.

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10. Collect the virus particles by side puncture of centrifuge tubes. Virus particles will separate or band in the CsCl according to different densities. Each particle type can be collected individually or pooled. Typically, there are two major bands, triple-layered particles (TLPs) (density 1.36 g/cm3) and double-layered particles (DLPs) (density 1.38 g/cm3). The DLPs should be approximately in the middle of the gradient, and the TLPs should be directly above the band containing the DLPs. 11. Dilute the collected bands containing CsCl in TNC (triple-layered virus) or 50 mM Tris-HCl, pH 8.0 (double-layered virus). Alternatively, the bands can be dialyzed to remove the CsCl. 12. Pellet virus particles by ultracentrifugation at 280,000g for 2 h at 4°C. 13. Resuspend pellets in TNC or 50 mM Tris-HCl, pH 8.0, accordingly, and measure the optical density (OD) at a wavelength of 260 nm in a spectrophotometer. 14. Determine the virus concentration by multiplying OD 260 × dilution factor × 185 µg/mL. 15. Store at –80°C.

3.1.2. Method for Formalin Inactivation of RVs 1. Treat virus-infected MA104 cell lysates with formalin to a final concentration of 0.00925% at 37°C in a shaker incubator, letting the virus–formalin mix swirl without creating froth. 2. Collect sample aliquots of virus at 0, 10, 20, 30, and 60 min, and at 2, 4, 6, 12, 24, 48, and 72 h after the addition of formalin. 3. Inactivate residual formalin by adding sodium bisulfite (150 µL/mL of a 0.35% solution) to each sample taken at each time point, and to the remaining virus preparation. 4. A control virus-infected cell lysate (not treated with formalin, but incubated at 37°C in the shaker) should be performed. 5. Store formalin-inactivated and control heat-treated preparation at 4°C until use. 6. Titrate virus in all samples collected (including the control heat-inactivated and original virus preparation) by plaque assay (see Subheading 3.9.3.), to establish a kill curve. Plot log titer vs. time in hours. 7. Passage an aliquot of formalin-treated virus at least 3× in tissue culture or MA104 cells, followed by an infectivity assay to confirm complete viral inactivation. 8. Evaluate inactivated virus by ELISA using neutralizing monoclonal antibodies (MAbs), to confirm that critical epitopes in the outer capsid proteins VP4 and VP7 were maintained during the inactivation procedure (39). Integrity of inactivated virus should be analyzed by electron microscopy (EM).

3.1.3. Method for Psoralen-UV Inactivation of RVs (see Note 4) 1. Start with purified desired RV at approx 300 µg/mL in TNC buffer or 50 mM Tris-HCl, pH 8.0, depending on whether virus is triple- or double-layered, respectively. 2. Sonicate virus 2 min to disrupt aggregates. 3. Mix virus and psoralen, final concentration of psoralen is 40 µg/mL.

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4. Add mixture (~200–400 µL) to a well of a six-well tissue culture plate in the tissue culture hood. 5. Place plate on ice for 15 min. 6. Wrap foil around the bottom and sides of the six-well plate, and place back in ice. 7. Position the UV light 10 cm above the plate, with the plate lid removed. 8. UV-irradiate for 40 min (see Note 5), and then remove virus, and store at 4°C. 9. Plaque assays should be performed on the UV-psoralen-treated virus, and the virus should be passaged at least 3× in MA104 cells, followed by an infectivity assay to ensure complete inactivation. 10. Evaluate inactivated virus by ELISA, using neutralizing MAbs to confirm that critical epitopes in the outer capsid proteins VP4 and VP7 were maintained during the inactivation procedure (110). Integrity of inactivated virus should be analyzed by EM.

3.2. Recombinant BV Stocks 1. Once the recombinant baculovirus (rBV) is obtained and confirmed, it is plaque-purified 3× to ensure the authenticity of the clone. 2. To prepare a rBV stock, Sf9 insect cells are infected (usually at an MOI = 0.1) with the plaque-purified recombinant virus (see Note 6). 3. After an incubation of 5–7 d, the recombinant virus stock is obtained by harvesting and clarifying the infected cells (see Note 7). 4. The recombinant virus is titered (by plaque assay), and the presence of the protein(s) of interest are confirmed by Western blotting. (For further details of BV production, see Chapters 3 and 4).

3.2.1. Production and Purification of RV VLPs 1. Large volumes of insect cells (>108 cells) are infected with the rBV stock(s) at an MOI = 5–10 (optimal conditions may vary), and are incubated for 5–7 d in spinner flasks (see Note 8). 2. The cells are then harvested and clarified by low- and medium-speed centrifugation, and the supernatant containing the VLPs is kept for purification. 3. The VLPs are pelleted through a sucrose cushion, and purified through a single gradient of CsCl. 4. The bands on the CsCl gradient (containing the VLPs) are collected by side puncture, concentrated by pelleting, resuspended in TNC buffer, and stored at 4°C in the dark (generally as described in Subheading 3.1.1.).

3.2.2. Characterization of RV VLPs (see Note 9) VLPs are characterized for integrity, purity, protein composition, concentration, serotype, and endotoxin concentration, as detailed previously (23,28). 1. To assess the level of intact particles and the general purity of the preparation, VLPs are negatively stained with 1% ammonium molybdate, and examined by EM. Only VLP preparations, with >90% intact particles without excessive debris, are used for animal inoculations.

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2. Protein composition of the VLPs is confirmed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (see Chapter 3) and silver nitrate staining and Western blotting. To ensure adequate detection of all proteins by Western blotting, multiple hyperimmune sera may be pooled. The authors commonly use both polyclonal mouse anti-RV hyperimmune serum and a VP4-specific MAb to detect all the structural proteins. 3. Protein concentration is determined with the Bio-Rad protein assay, using IgG as the standard. Because results can differ using different reagents, protein concentrations are always determined by this procedure. 4. To ensure that immune responses to VLPs are not caused by nonspecific enhancement of the immune response by contaminating endotoxin, endotoxin levels of all VLP preparations are determined using the LAL endotoxin assay (Associates of Cape Cod). VLPs are not used for animal inoculations if the dose of endotoxin per inoculation exceeds 0.05 endotoxin units (see Notes 10 and 11).

3.3. Method for Propagation and Titration of WT RVs 1. Isolate new virus directly from stool sample or obtain WT RV stock from another investigator. 2. To prepare a laboratory stock, orally inoculate multiple animals with the WT stock. In the mouse, 5-d-old pups are inoculated, the intestines are harvested at 2 d postinfection (DPI), pooled, frozen and thawed once, homogenized with media (10% suspension w/v), aliquoted, and frozen at –80°C (96). A similar method can be used in nursing rabbit kits less than 6 wk of age. In weanling or older rabbits, rabbits can be orally inoculated and large quantities of fecal samples harvested 3–5 DPI, when the peak of virus shedding is typically observed (23,37,48,75). Ten percent fecal suspensions are made, aliquoted, and frozen at –80°C. 3. Once a laboratory WT stock has been produced, it is necessary to determine the titer of the stock, so that the challenge dose administered to immunized animals is known. In rabbits and mice, the infectivity of WT RV strains is measured and expressed as the presence or absence of RV-specific diarrhea (feasible only in rabbits and mice <2 wk of age) [50% diarrhea dose (DD50)], the presence or absence of virus Ag shedding in fecal samples [50% shedding dose (SD 50)], or the presence or absence of RV shedding [50% infective dose (ID50)] or an RV-specific immune response. 4. The titer of the WT stock is determined in animals, using standard methodology. Briefly, 10-fold dilutions of the stock are administered to multiple animals (≥4). Infection is scored in individual animals by the chosen method, and the total number of infected and uninfected animals at each dilution is determined. The infectivity dose of the WT stock is calculated using either the Reed-Munch or Spearman Karber formula.

3.4. Husbandry 1. Care should be taken to prevent accidental exposure of mice and rabbits to RV. Because RVs are ubiquitous in nature, the rooms in which the animals will be

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Ciarlet and Conner housed must be disinfected multiple times with Clidox (Pharmacal Research, Naugatuck, CT) (10% bleach solution with detergent) prior to introduction of the animals. The animal rooms where challenges are performed should be maintained under negative pressure, and should not house other animals. All materials entering the rooms should be heat- or chemically sterilized. During the challenge portion of the experiment, all materials removed from the room must be heat- or chemically sterilized to prevent spread of RV to other animals. CAUTION: All persons entering the rooms or handling the animals must wear protective clothing and double gloves. These supplies should be maintained in the rooms. If RV-naïve and RV-infected mice or rabbits cannot be separated in two rooms, always handle RV-naïve animals first, and never handle or open their cages after handling or opening cages of RV-infected rabbits. If two rooms are available, RV-naïve animals should be cared for first, and that room should not be reentered once you have been in the room with infected animals. As a rule, always feed and water animals prior to any collection of samples or handling of rabbits. During the immunization phase of an experiment, rabbits can be housed in open cages, and mice are housed in microisolator cages, either in a BL2 containment room or in a room under positive pressure. During the challenge or infection portion of an experiment, rabbits should be housed in individual negative-pressure isolation units, to prevent cross-contamination between rabbits, and mice are housed in microisolator cages, preferably inside a Hepa-filtered isolation rack. The homologous lapine and murine RV strains spread readily from rabbit to rabbit or mouse to mouse, respectively. All animals are housed and cared for under BL2 conditions.

3.4.1 General Design of Animal Experiments for RV Vaccine Testing 1. All experiments should be performed with a minimum of five rabbits or mice per group. Although smaller numbers of animals are sometimes used, using larger groups of animals provides greater power for statistical comparisons. 2. Several doses of the immunogen should be tested for immunogenicity and protective efficacy. 3. Parenteral immunizations are performed at 28-d intervals in rabbits and 21-d intervals in mice. Intranasal and oral immunizations of mice are performed at 14-d intervals. 4. Challenge viruses can be either WT or tissue-culture-adapted strains. Typically, challenge viruses are homologous viruses, because they productively infect the host species in a dose-independent manner. However, protective efficacy may be dependent on challenge dose. 5. Negative control rabbits or mice are mock-immunized with buffer and adjuvant. These rabbits or mice should be totally susceptible to challenge virus, and serve as the standard for determining percent protection for the immunized groups.

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6. Positive control rabbits or mice are orally infected with live RV at the time of the last immunization. These rabbits or mice should be totally protected, not shed virus, after challenge. 7. To measure RV-specific Ab responses, serum and fecal samples are collected on each day of immunization or at 21- or 28-d intervals thereafter (e.g., 0, 28, and 56 DPV, and at 0 and 21 or 28 DPC. 8. To measure virus Ag shedding, fecal samples are collected from 0 to 10–14 DPC. 9. All samples are collected from individual rabbits or mice, and processed separately. Most testing is performed on individual samples. Testing may be performed on pooled samples, but statistical comparison of results is then difficult. 10. Comparison of percent reduction in virus shedding and Ab titers should be compared statistically. If results obtained are not normally distributed, then log transformation of results is necessary before performing Student’s t-test or nonparametric tests, such as Mann-Whitney U-test, Wilcoxon signed ranks test, or Kruskal-Wallis test followed by Mann-Whitney U-test, should be performed. 11. Results should be shown to be repeatable in subsequent experiments.

3.5. Immunization of Rabbits 3.5.1. Oral Immunization or Challenge 1. Place inoculum (usually 1–3 mL) in a sterile syringe, and attach a blunt-end feeding needle (16 gage for adult rabbits and 20 gage for rabbits <2 wk of age). 2. Place the rabbit on cart or table. 3. With one forearm, hold rabbit against your body and gently insert the feeding needle on the side of the mouth against the tongue. Once the feeding needle has reached the back of the throat, slowly release the inoculum. To make sure the rabbit swallows the inoculum, hold its mouth tightly closed and gently stroke its neck after administering immunogen until swallowing reflex is induced (see Note 12). 4. Rabbits are challenged with lapine RVs because these strains are highly infectious in rabbits. Heterologous RVs generally do not replicate well in rabbits (48). The author’s standard challenge dose with tissue-culture-adapted ALA virus is approx 103 ID50 (~105 plaque-forming units [PFU]).

3.5.2. Parenteral Immunization 1. Mix virus or vaccine inoculum with appropriate adjuvant. 2. Place inoculum into syringe and inject vaccine im in the quadriceps muscle of the leg, with a maximum volume of 0.5 mL. Alternate legs for different doses. 3. Check animal periodically after administration of vaccine for any sores or lesions from injection (especially if Freund’s adjuvant is used).

3.5.3. Intranasal Immunization 1. Prior to inoculation, anesthetize rabbits by subcutaneous administration of the combination anesthesia. The rabbit must be deeply anesthetized. 2. Administer vaccine drop by drop into alternating nares waiting until previous drop has been inhaled before placing next drop in alternate nares (see Note 13).

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3. Monitor rabbits until consciousness is regained and rabbits are sternally recumbant.

3.6. Collection of Rabbit Serum and Fecal Samples 3.6.1. Serum Samples 1. Inject rabbit subcutaneously with 0.1–0.2 mL/kg acepromazine maleate, to slightly tranquilize rabbit and cause vasodilatation. Wait approx 10–20 min after injection until rabbit is noticeably tranquil. 2. Swab ear over medial artery with alcohol. 3. Bleed from the medial ear artery using a Vacutainer brand blood collector set (butterfly) with a 21–23-gage, three-quarter-inch needle. 4. Collect 4–10 mL blood in Vacutainer tubes with separator. The total blood volume collected should not exceed 6% body wt over a 3-wk interval. 5. To separate serum, spin Vacutainer tubes with blood at 1000g for 10 min. 6. Collect serum, aliquot, label, and store at –80°C to –20°C.

3.6.2. Fecal Samples 1. Place autoclaved screens on litter pans 24 h prior to first fecal collection, using four clips, and pull screen tightly to prevent sagging in the middle. The screening will collect fecal samples, and allow urine to flow through to litter pan. 2. On the day of collection, collect fecal pellets from several places from the top of the screen into a 10-mL sterile screw-top container. Even when collected on the same day, separate fecal samples should be collected for Ag and Ab detection. 3. Once the sample is collected, remove the screen, dump the remaining pellets in the litter pan, place dirty screen in container for autoclaving, and place a fresh screen over litter pan. Daily changing of the screens during the challenge phase of the study diminishes cross-contamination of samples. Dirty screens should be removed from the room daily, decontaminated, washed, and autoclaved for future use. 4. Process samples daily on day of collection (see Subheading 2.4.), or freeze samples at –20°C (fecal Ag) or –80°C (fecal Ab) for later processing.

3.6.3. Processing of Fecal Samples to Detect RV Ag 1. Crush fresh or thawed fecal pellets using a metal spatula or wooden applicator stick (see Note 14). 2. Weigh the desired amount of fecal sample into a sterile tube, and dilute the fecal sample in fecal Ag buffer, to yield a final 10% (w/v) suspension. 3. Vortex thoroughly. 4. Centrifuge at 30,000g for 10 min at 4°C. 5. Harvest and aliquot the supernatant into sterile tubes using sterile pipets without disturbing the pellet. 6. Store the processed samples at –80°C to –20°C (see Note 15).

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3.6.4. Processing of Fecal Samples to Measure RV-Specific Ab 1. Crush fresh or thawed fecal pellets, using a metal spatula or wooden applicator stick. 2. Weigh the desired amount of fecal sample into a sterile tube, and dilute the fecal sample in fecal Ab buffer, to yield a final 5% (w/v) suspension (see Note 16). 3. Perform steps 3–5 of fecal Ag sample processing. 4. Store the processed samples at –80°C.

3.7. Immunization of Mice 3.7.1. Oral Immunization or Challenge 1. Prepare 5% bicarbonate buffer and inoculum for immunization or challenge in separate sterile syringes, and attach sterile bent feeding needle to each syringe. Maximum inoculum volume in the adult mouse is 250 µL (see Note 17). 2. Just prior to oral inoculation with immunogen or challenge virus, orally inoculate each mouse with 40 µL 5% bicarbonate buffer, to neutralize stomach acid (see Note 18). 3. To prevent contamination of control mice, perform all mock-inoculations before inoculating any mice with virus. Once control mice have been inoculated, inoculate remainder of mice. 4. To inoculate a mouse, hold the mouse by the scruff of the neck and by the tail, and gently and slowly insert the feeding needle into the mouth. Initially guide the feeding needle along the roof of mouth and when needle is at the back of the throat, either tilt feeding needle up or mouse down, so that the feeding needle can be inserted down the esophagus. Do not force the needle past resistance. If resistance is encountered, back the feeding needle out and start again.

3.7.2. Intranasal Immunization 1. Prepare inocula for immunizations (20 µL/mouse). 2. Anesthetize mice intraperitoneally with combination anesthesia. 3. Once a mouse is anesthetized, inoculum is administered in approx 2 µL vol (one drop) instilled into alternating outer nares. Successive doses should not be administered to alternate nares until previous dose has been inhaled. Volume of inoculum should not exceed 20 µL. Smaller inoculum volumes are preferred, to avoid respiratory compromise. 4. Mice should be monitored closely until consciousness is regained.

3.7.3. Parenteral Immunization 1. Prepare inocula (immunogen and adjuvant) for immunization in 1 mL syringes with 25-gage needles (≤50–100 µL/mouse). 2. Inject mice im with vaccine. Every effort should be made to minimize the volume of the inoculum, so that each dose can be administered to one leg (quadriceps muscle), with a maximum dose of 50 µL/leg. Use alternate legs for additional immunizations (see Note 19). 3. Check animal periodically after administration of vaccine for any sores or lesions from injection, especially when using Freund’s adjuvant.

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3.8. Collection of Mouse Serum and Fecal Samples 3.8.1. Serum Samples 1. Mice are tail bled for routine sequential serum samples (see Note 20). 2. All the mice from one cage are placed in a box with a heat lamp, until the mice begin to groom. 3. The mice are then removed one at a time, and placed in a Broome-style restraint. 4. The tail is swabbed with alcohol, a scalpel blade is used to lightly knick a tail vein, and the blood is allowed to drip into a BD microtainer serum separator collection tube. 5. Following blood collection, pressure is applied to vein with cotton ball until bleeding stops entirely. 6. The blood is allowed to clot for 30 min and the serum is separated by microcentrifuge at 6000g for 10 min. 7. The serum is harvested, aliquoted, labeled, and stored at –20°C (see Note 21).

3.8.2. Fecal Samples 1. Fecal samples for Ag detection are collected 0–10 DPI or DPC with RV. If onset of virus shedding is delayed, fecal samples should be collected until at least 14 DPI or DPC. 2. Fecal samples for Ab detection are typically collected prior to any immunizations or virus infections, on the day of any additional immunizations, on the day of RV challenge and 28 DPC. Additional samples can be collected at intervals between any of the afore mentioned time points. 3. Fecal samples for Ag shedding or Ab detection are collected from individual mice. Mice are placed for 3–4 h in individual sections of a wire collection cage, which is kept inside a microisolator cage. A collection pan underneath the wire collection cage allows flowthrough of urine, while keeping fecal samples from individual mice separated (see Note 22). 4. Mice are removed from the fecal collection cage and placed back in the microisolator cage. Fecal pellets from individual mice are collected from the pan underneath the cage, and placed in properly labeled tubes. 5. Fecal Ab samples are stored at –80°C and fecal Ag samples are stored at –20°C.

3.8.2.1. PROCESSING OF FECAL SAMPLES TO DETECT RV AG 1. Process samples on day of collection or thaw fecal pellets previously collected and frozen at –20°C. 2. Weigh the desired amount of fecal material using wooden sticks to get fecal pellets out of original tube and into a 1.5-mL tube. Place a maximum of 0.1 g into each tube. Prepare one tube per sample, and refreeze the remainder of the sample, or, alternatively, process the entire sample in a 15-mL centrifuge tube. 3. Add fecal processing buffer to each tube to make a 10% (w/v) suspension (e.g., 900 µL/0.1 g sample) (see Note 23). 4. Crush pellets using metal spatulas, one spatula per sample.

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5. 6. 7. 8.

Vortex thoroughly. Let sit on ice for 45 min. Freeze at –20°C. On the day samples are needed for an assay, thaw the samples, and clarify all samples by centrifugation for 1 min at 4°C. 9. Aliquot a sufficient amount (typically 100 µL) of clarified sample into a new tube or into wells (50 µL/well) on assay plate, and refreeze remaining clarified sample for future use.

3.8.2.2. PROCESSING OF FECAL SAMPLES TO MEASURE RV-SPECIFIC AB 1. 2. 3. 4.

Perform steps 1–7 of fecal Ag procedure (see Subheading 3.8.2.1.). Following 45 min incubation on ice, clarify sample by centrifugation for 5 min at 4°C. Harvest and aliquot the supernatant into sterile 1.5-mL screw-top tubes. Store the labeled aliquots at –80°C.

3.9. Detection of RV Shedding 3.9.1. ELISA to Detect RV Ag Shedding 1. Coat plates with 50 µL/well hyperimmune anti-RV serum diluted in CBB, and incubate plates overnight at room temperature (see Notes 24 and 25). 2. Block plates with 200 µL/well BLOTTO for 2 h at 37°C. 3. Wash plates 3× with PBS-T. 4. Add 50 µL/well 10% (w/v) fecal samples to duplicate wells. Serial dilutions in 1/10 BLOTTO of the RV stock to achieve end point titration must be included on each plate as a positive control for the assay. Incubate 1 h at 37°C (see Note 26). 5. Wash 5× with PBS-T. 6. Add 50 µL/well hyperimmune anti-RV serum diluted in 1/10 BLOTTO, and incubate for 1 h at 37°C. To avoid false-positive results, this hyperimmune anti-RV serum should be produced in a different species from the one used to produce the coating antiserum. 7. Wash 5× with PBS-T. 8. Add 50 µL/well appropriate enzyme-conjugated antispecies-specific immunoglobulin diluted in 1/10 BLOTTO, and incubate for 1 h at 37°C. To reduce background, dilute the conjugated serum with 5–10% (final concentration) normal (non-RV Ab containing) serum obtained from the same species as the coating Ab or with FCS. 9. Wash 5× with PBS-T. 10. Add 100 µL/well of appropriate substrate for enzyme, according to manufacturer’s instructions, and read results after standard incubation time, or, if necessary add stopping reagent (see Note 27). 11. Read plates in an ELISA reader at appropriate wavelength. 12. Sample is considered positive for Ag when the mean OD value of the sample minus the OD value of the negative control is ≥0.1 and at least two standard deviations above the negative control.

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13. End point titration of positive control must be standardized and consistent between assays. If titer of positive virus control varies by more than (plus or minus) one twofold dilution, the assay is not considered valid and must be repeated.

3.9.2. Fluorescent Focus Assay (FFA) for Infectious RV Shedding 1. Trypsin-activate 10% fecal samples, and control virus by adding 10 µg/mL (final concentration) porcine trypsin (type IX; Worthington) to samples and virus, and incubate for 30 min at 37°C. 2. Make 10-fold dilutions of trypsin-activated samples and virus in Medium 199. 3. Gently wash, once, the 96-well plates containing the confluent MA104 cell monolayers with Medium 199. 4. Inoculate MA104 cell monolayers in duplicate wells with 100 µL/well of each sample or virus dilution, and incubate 1 h at 37°C in 5% CO2. 5. Decant inoculum, wash cells once, and add 100 µL/well Medium 199. Incubate overnight at 37°C in 5% CO2. 6. On the next day, wash wells once with 200 µL/well PBS. 7. Fix the cells at room temperature with 100 µL/well cold methanol (–20°C). 8. Wash wells once with 200 µL/well PBS. 9. Add 50 µL/well diluted (usually 1/300 or 1/500) anti-RV hyperimmune serum in PBS, and incubate for 2 h at 37°C. 10. Wash wells once with 200 µL/well PBS. 11. Add 50 µL/well appropriately diluted (PBS) antispecies-specific immunoglobulins conjugated to FITC, and incubate for 2–4 h at 37°C. 12. Count fluorescent foci in an inverted fluorescence microscope. Determine the infectious virus titer by averaging the number of foci in duplicate wells of each sample and virus dilution. Multiply the average foci at the dilution that yields between 100 and 200 foci by the dilution factor. This number is then adjusted according to the volume of the inoculum to allow determination of the titer as focus-forming units (FFU) per milliliter (FFU/mL).

3.9.3. Plaque-Forming Assay (PFA) to Detect Infectious RV Shedding 1. Trypsin-activate all 10% fecal samples, and control virus by adding 10 µg/mL (final concentration) porcine trypsin (type IX; Worthington) to samples and virus, and incubate for 30 min at 37°C. 2. Make 10-fold dilutions of trypsin-activated samples and control virus in Medium 199. 3. Gently wash, once, the six-well plates containing the confluent MA104 cell monolayers with Medium 199. 4. Inoculate duplicate wells of six-well plates with 200 µL/well trypsin-activated sample or virus control, and incubate for 60 min at 37°C in 5% CO 2 . Every 15 min, the plates should be removed from the incubator and rocked gently, not swirled, to ensure that no part of the monolayer dries out during absorption. 5. About 30 min before the absorption is finished, prepare first overlay, and keep warm at 56°C.

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6. After absorption of sample or virus inoculum, gently wash cells once with Medium 199, and add 3 mL/well first overlay. Once the overlay is solid, invert the plates before incubating for 2–4 d at 37°C in 5% CO2. 7. After the 2–4 d incubation, add 2 mL/well second overlay, invert the plates when the overlay is solid, wrap with aluminum foil, and incubate at 37°C in a 5% CO2. The vital dye neutral red is light-sensitive; therefore, after addition of the second overlay, the plates are wrapped with foil to reduce light exposure. 8. Plaques should be visible within 1 d after the addition of the second overlay. Count plaques for up to five consecutive days, or until plaques are no longer visible. 9. Determine the infectious virus titer by averaging the number of plaques in duplicate wells of each sample and virus dilution. Multiply the average plaque number, at the dilution that yields between 20 and 100 plaques by the dilution factor. This number is then multiplied by a factor adjusted according to the volume of the inoculum to allow determination of the titer as PFU/mL.

3.10. Ab Detection in Rabbits and Mice 3.10.1. FFNA to Measure RV-Specific nAb 1. Trypsin-activate virus stock of known titer by adding 10 µg/mL (final concentration) porcine trypsin (type IX; Worthington) to virus, and incubate for 30 min at 37°C. 2. In a 96-well plate, make twofold dilutions of samples or control Abs in Medium 199. 3. Add equal volumes of the appropriate trypsin-activated virus dilution (100–200 fluorescent focus units/well) to each of the sample or control Ab dilutions, mix gently, and incubate for 1–2 h at 37°C. The final volume of the Ab–virus mixture required is at least 150 µL. To serve as virus control, virus should be mock-diluted with 199 media in a similar manner. 4. Gently wash, once, the 96-well plates containing the confluent MA104 cell monolayers with Medium 199. 5. Transfer 100 µL Ab–virus mixture or virus control into each of two duplicate wells of the plates with monolayers of MA104 cells. Incubate for 1–2 h at 37°C. 6. Gently wash the plates once with Medium 199 to remove inoculum. 7. Add 100 µL Medium 199 to each well, and incubate overnight at 37°C. 8. Perform steps 6–11 of FFA (Subheading 3.9.2.). 9. Count fluorescent foci in an inverted fluorescence microscope. Average the number of fluorescent foci for all virus control wells and in duplicate wells of samples. The end point of the nAb titer is the reciprocal of the dilution of each sample, which results in >66% reduction in the average number of fluorescent foci in the virus control wells.

3.10.2. PRNA to Measure RV-Specific nAb 1. Trypsin-activate virus stock of known titer by adding 10 µg/mL (final concentration) porcine trypsin (type IX; Worthington) to virus, and incubate for 30 min at 37°C.

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2. In 4-mL tubes, make twofold dilutions of samples and control hyperimmune serum in Medium 199 (250 µL/tube). 3. Add equal volume (250 µL) of the appropriate trypsin-activated virus dilution (one that should give approx 50–100 PFU/well) to each of the Ab dilutions, and mix gently. Control virus should be mock-diluted with Medium 199 in a similar manner. Incubate for 1 h at 37°C. 4. Gently wash, once, six-well plates containing the confluent MA104 cell monolayers with Medium 199. 5. Transfer 200 µL/well Ab–virus mixture into each of two duplicate wells of the MA104 cell seeded plates. In a similar manner, inoculate six wells each with mock-diluted control virus. Incubate plates for 1 h at 37°C in 5% CO2. Every 15 min, the plates should be removed from the incubator, and rocked gently, not swirled, to ensure that no part of the monolayer dries out during absorption. 6. Perform steps 5–8 of PFA method (see Subheading 3.9.3.). 7. Count plaques and average the number of plaques for all virus control wells and in duplicate wells of samples. The end point of the nAb titer is the reciprocal of the dilution of each sample, which results in >60% reduction in the average number of plaques in the virus control wells (see Notes 28–30).

3.10.3. ELISA to Measure RV Total, Isotype-, or Subclass-Specific Ab 1. Coat plates with 100 µL/well hyperimmune anti-RV serum diluted in CBB overnight at room temperature (see Note 31). 2. Block plates with 200 µL/well BLOTTO for 2 h at 37°C. 3. Wash plates 3× with PBS-T. 4. Add 100 µL/well tissue-culture-passaged RV, diluted in 1/10 BLOTTO, to alternating rows or columns of the 96-well plate. Add 100 µL mock-infected MA104 cell lysate, diluted in 1/10 BLOTTO, to the alternate rows or columns of the plate. This negative control Ag is diluted to yield an equivalent protein concentration to the total protein concentration of the diluted RV Ag. Alternatively, 100 µL/well 1/10 BLOTTO can be added as the negative control. The first column of the plate is always used as blank. Incubate for 1 h at 37°C. 5. Wash 5× with PBS-T. 6. Add 100 µL test sample (serum or fecal suspensions processed for Ab detection), serially diluted twofold on the plate in 1/10 BLOTTO, and incubate for 1 h at 37°C. Twofold serial dilutions (in 1/10 BLOTTO) of a positive control serum must be included on each plate. Sufficient dilutions of the positive control and sample serum are needed to obtain end points. 7. Wash 5× with PBS-T. 8. Add 100 µL/well enzyme-conjugated antispecies-specific immunoglobulin (to the species from which the samples were collected), diluted in 1/10 BLOTTO containing 5% normal serum obtained from the same species as the hyperimmune serum used to coat the plate, and incubate for 1 h at 37°C (see Note 34).

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9. By using different Ab conjugates, the assay can measure total Ab (IgM, IgG, IgA), individual isotype-specific responses (IgM, IgG, or IgA), or, in mouse samples, IgG subclass responses (IgG1, IgG2a, IgG2b, or IgG3) (see Notes 35–39, and Chapter 7). 10. Wash 5× with PBS-T. 11. Develop with appropriate substrate (100 µL/well) for standard time, and add stopping reaction, as appropriate. 12. Read results in an ELISA plate reader, at the appropriate wavelength. 13. The end point titer of the positive control serum (run on each plate) must be within one dilution of an established standard, for the assay to be acceptable. 14. A positive reaction is defined as an OD value of ≥0.1 after subtraction of the OD value of the no-Ag well from those of the Ag well. End point titer is expressed as the reciprocal of the highest dilution that was positive.

4. Notes 1. To determine MOI, the average number of MA104 cells in a confluent monolayer in each vessel size needs to be determined in each laboratory, using basic cell counting procedures. 2. If only DLPs are desired, prior to purification the virus preparation can be treated for 30 min at room temperature with 10 mM EDTA, pH 8.0 to remove the outer capsid proteins VP4 and VP7. 3. To purify WT RV, prepare a 10 or 20% (w/v) suspension from a stool sample in TNC buffer. Add an equal volume of Genetron to the virus suspension, and homogenize for 1 min at 4000g. Keep on ice, and let it settle for 1 min, and repeat the homogenization. Pour Genetron–virus suspension into a centrifuge tube, and spin for 10 min at 2000g at 4°C. Pipet the aqueous–virus containing phase (top phase) into tubes and follow steps 3–14. 4. The psoralen inactivation procedure described above is modified from a published protocol (110). 5. For each virus preparation, a psoralen–UV inactivation time-course should be performed, to determine the amount of time necessary to inactivate all virus. 6. A good recombinant stock will have a titer of ≥ 1 × 108 PFU/mL. 7. To determine the best time to harvest the stock, a time-course experiment is performed to compare cell viability and titer. 8. To reduce the possibility of contamination by endotoxin, VLPs used for vaccine studies are made in glassware that is dedicated for vaccine use, and all reagents are endotoxin free. 9. Each preparation of VLPs is characterized independently. If several preparations need to be pooled to obtain sufficient VLPs for one animal experiment, all characterizations are repeated on the pooled preparations. If necessary, the serotype of VP7 on the VLPs can be confirmed by immunogold labeling or ELISA with serotype-specific MAbs. 10. Animal experiments are not started until sufficient VLPs are prepared to perform all scheduled inoculations. 11. A portion of each preparation or pool used for all animal experiments is retained at 4°C for any future additional characterizations.

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12. Neutralization of gastric acidity prior to oral inoculation of rabbits is not necessary. 13. Keep inoculum dose small (<100 µL), to avoid respiratory complications. 14. Keep samples on ice at all times. 15. Complete individual sample processing within 1 d. 16. Degradation of Ab upon storage will occur more rapidly if sample is processed without protease-inhibitors in processing buffer, or if the samples are stored above –80°C. 17. Most mice are easier to inoculate without anesthesia, but an occasional mouse needs to be subdued by light anesthesia, using Metofane (methoxyflurane) (Pitman-Moore, Mundelein, IL), prior to inoculation. 18. Neutralization of stomach acidity is not necessary prior to administration of high doses (104 SD50) of ECwt virus, but this has not been tested prior to immunization with lower doses of virus or with VLPs. 19. The authors perform parenteral immunizations intramuscularly (i.m.). Although im is not a preferred route of immunization for mice, it is the most analogous route for comparison for parenteral immunization of children. 20. If serum collection is needed at the completion of experiment, it can be collected by cardiac puncture. Mice are completely anesthetized with a combination anesthesia (see Subheading 2.5.). Blood is collected into a syringe following cardiac puncture, and transferred into the BD microtainer tubes for serum separation. 21. To prevent degradation of Abs, serum samples should not be repeatedly frozen and thawed. 22. Even when collected on the same day, separate samples should be collected for Ag and Ab detection. 23. If samples are to be run in an ELISA, the sensitivity of the ELISA may be increased by treating the sample with EDTA (final concentration, 0.025 M). This treatment removes the outer capsid layer of the virus, exposing the middle capsid layer composed of VP6. Most Ab in an immune serum to virus is directed to VP6. 24. The authors use flexible 96-well polyvinyl microtiter plates, because they are substantially less expensive, but still have high binding affinity. Other plates can be used. 25. Optimal dilutions of all reagents should be predetermined by checkerboard titration. 26. Dilutions of fecal samples for Ag detection are not routinely performed. Only the OD value of each 10% sample is determined and compared. 27. Assay can be performed with any combination of enzyme-conjugated Ab– substrate, and read at the appropriate wavelength. 28. If first overlay is too hot, it will kill cells, and if it cools too quickly, it will gel before it can be added to the plates. 29. If wells have dried out during absorption, there will usually be holes in the monolayer in the middle of the plate. Holes in monolayer around the edge of the plates usually indicate that pipeting was too vigorous. 30. Optimal length of incubation after addition of first monolayer is virus-dependent. Generally, human RV strains have relatively longer replication cycles than

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31.

32.

33.

34.

35.

36.

37.

38. 39.

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animal RV strains, requiring additional incubation times before second overlay is added. Sensitivity of assay is greatest when coating Ab and Ag used in assay are homologous for the virus that was used to immunize or infect the animals. However, any RV strain and any hyperimmune Ab can be used in the assay. All assays are standardized with a positive-control serum to RV on each plate, to ensure that results obtained between plates and over time can be compared. The assay is not considered valid unless the end point titer of the positive-control serum is within one dilution (plus or minus) of the standard established titer of the positive-control serum. Initially, samples from every animal, or several samples from each group of animals, can be screened for dilutions appropriate to perform in the ELISA. Samples are diluted 1 in 100, and tested in the assay. Based on the OD value of the 1 in 100 dilution, appropriate dilution ranges can be chosen. All new reagent concentrations are determined by checkerboard titration to yield the standard positive control serum titer. For example, when a new Ag preparation is needed for the ELISA, various dilutions are tested to determine the dilution needed to obtain the standard titer of the positive-control serum. If the authors are performing new experiments and do not know what isotypespecific response has been induced, total (IgM, IgG, IgA) Ab responses will be measured first. If sample sizes are limited, this assay will also be run first to ensure that Ab titers are obtained for all samples, before proceeding with individual immunoglobulin isotype- or subclass-specific responses. The authors routinely use horseradish peroxidase (HRP) conjugates and the substrate, 3,3',5,5'-tetramethylbenzidine (TMB) (Kirkegaard & Perry, Gaithersburg, MD). Assays performed with these reagents are sensitive, not requiring the use of toxic or carcinogenic substrates. To increase the sensitivity of the ELISA, avidin–biotin systems can be used. The authors find the extra sensitivity of avidin–biotin is needed to detect rabbit IgA responses. We use a biotinylated mouse antirabbit IgA monoclonal conjugate. This step is followed by five washes with PBS-T, incubation with HRP–avidin for 1 h at 37°C, five washes with PBS-T, and 10 min incubation with TMB at room temperature. Sensitivity and specificity of each conjugate should be confirmed prior to use. Many commercial reagents vary widely in sensitivity and specificity. When comparing IgG subclass responses, the sensitivity of each conjugate should be determined. For the assay, the dilution of the individual conjugates is chosen, so that all conjugates have similar sensitivities.

Acknowledgments The authors are grateful to Sue E. Crawford and Mary K. Estes for critical reading of this manuscript. This work was supported by the Public Health Service grants AI 24998 and AI 16687 from the National Institute of Allergy and Infectious Diseases, Advanced Technology Program grant 0049-49-029 from

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the Texas Higher Education Coordinating Board, Public Health Service Training Grant T32-DK07664, World Health Organization grant MIMV 2718130 and VA Merit Review. References 1. Conner, M. E., Matson, D. O., and Estes, M. K. (1994) RV vaccines and vaccination potential. Curr. Top. Microbiol. Immunol. 185, 286–337. 2. Conner, M. E., Estes, M. K., Offit, P. A., Clark, H. F., Franco, M. A., Feng, N., and Greenberg, H. B. (1996) Development of a mucosal RV vaccine, in Mucosal Vaccines (Kiyono, H., Ogra, P. L., and McGhee, J. R., eds.), Academic Press, San Diego, CA, pp. 325–344. 3. Conner, M. E., Zarley, C. D., Hu, B., Parsons, S., Drabinski, D., Greiner, S., Smith, R., Jiang, B., Corsaro, B., Barniak, V., Madore, H. P., Crawford, S. E., and Estes, M. K. (1996) Virus-like particles as a rotavirus subunit vaccine. J. Infect. Dis. 174, S88–S92. 4. Conner, M. E., Crawford, S. E., Barone, C., O’Neal, C., Zhou, Y.-J., Fernández, F., Parwani, A., Saif, L. J., Cohen, J., and Estes, M. K. (1996) Rotavirus subunit vaccines. Arch. Virol. 12(Suppl.), 199–206. 5. Conner M. E. and Ramig, R. F. (1996) Enteric Diseases, in Viral Pathogenesis (Nathanson, N., Ahmed, R., González-Scarano, F., Griffin, D. E., Homes, K. V., Murphy, F. A., and Robinson, H. L., eds.), Lippincott-Raven, Philadelphia, pp. 713–743. 6. Estes, M. K. (1996) Rotaviruses and their replication in, Fields Virology (Fields, B. N., Knipe, D. M., and Howley, P. M., eds.), 3rd ed. Lippincott-Raven, Philadelphia, pp. 1625–1655. 7. Clark, H. F., Borian, F. E., Bell, L. B., Modesto, K., Gouvea, V., and Plotkin, S. A. (1988) Protective effect of WC3 vaccine against rotavirus diarrhea in infants during a predominantly serotype 1 rotavirus season. J. Infect. Dis. 158, 570–587. 8. Clark, H. F., Offit, P. A., Ellis, R. W., Krah, D., Shaw, A. R., Eiden, J. J., Pichichero, M., and Treanor, J. J. (1996) WC3 reassortant vaccines in children. Arch. Virol. 12(Suppl.), 187–198. 9. Kapikian, A. Z., Hoshino, Y., Chanock, R. M., and Pérez-Schael, I. (1996) Jennerian and modified Jennerian approach to vaccination against rotavirus diarrhea using a quadrivalent rhesus rotavirus (RRV) and human-RRV reassortant vaccine. Arch. Virol. 12(Suppl.), 163–175. 10. Kapikian A. Z. and Chanock, R. M. (1996). Rotaviruses, in Fields Virology, 3rd ed. (Fields, B. N., Knipe, D. M., and Howley, P. M., eds.), Lippincott-Raven, Philadelphia, pp. 1657–1708. 11. Vesikari, T. (1993) Clinical trials of live oral rotavirus vaccines: the Finnish experience. Vaccine 11, 255–261. 12. Vesikari, T. (1996) Trials of oral bovine and rhesus rotavirus vaccines in Finland: a historical account and present status. Arch. Virol. 12(Suppl.), 177–186. 13. Vesikari, T. (1997) Rotavirus vaccines against diarrhoeal disease. Lancet 350, 1538–1541. 14. Velázquez, F. R., Matson, D. O., Calva, J. J., Guerrero, M. L., Morrow, A. L., Carter-Campbell, S., Glass, R. I., Estes, M. K., Pickering, L. K., and Ruiz-

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28. Crawford, S. E., Labbé, M., Cohen, J., Burroghs, M. H., Zhou, Y.-J., and Estes, M. K. (1994) Characterization of virus-like particles produced by the expression of rotavirus capsid proteins in insect cells. J. Virol. 68, 5945–5952. 29. Herrmann. J. E., Chen, S., Fynan, E., Santoro, J., Greenberg, H., and Robinson, H. (1996) DNA vaccines against rotavirus infections. Arch. Virol. 12(Suppl.), 207–215. 30. Herrmann. J. E., Chen, S., Fynan, E., Santoro, J., Greenberg, H., Wang, S., and Robinson, H. (1996) Protection against rotavirus infections by DNA vaccination. J. Infect. Dis. 174, S93–S97. 31. Jiang, B., Barniak, V., Smith, R., Sharma, R., Corsaro, B., Hu, B., and Madore, H. P. (1998) Synthesis of rotavirus-like particles in insect cells: comparative and quantitative analysis. Biotechnol. Bioeng. 60, 369–374. 32. Jiang, B., Estes, M. K., Barone, C., Barniak, V., O’Neal, C., Ottaiano, A. Madore, H. P., and Conner, M. E. (1999) Heterotypic protection from rotavirus infection in mice vaccination with virus-like particles. Vaccine 17, 1005–1013. 33. O’Neal, C., Crawford, S. E., Estes, M. K., and Conner, M. E. (1997) Rotavirus VLPs administered mucosally induce protective immunity. J. Virol. 71, 8707–8717. 34. O’Neal, C., Clements, J. D., Estes, M. K., and Conner, M. E. (1998) Rotavirus 2/6-VLPs administered intranasally with cholera toxin, E. coli heat labile toxin and LT-R192G induce protection from rotavirus challenge. J. Virol. 72, 3390–3393. 35. Suradhat, S. Yoo, D., Babiuk, L., Griebel, P., and Baca-Estrada, M. E. (1997) DNA immunization with a bovine rotavirus VP4 gene induces a Th1-like immune response in mice. Viral Immunol. 10, 117–127. 36. Fernández, F. M., Conner, M. E., Hodgins, D. C., Parwani, A., Nielsen, P., Crawford, S. E., Estes, M. K., and Saif, L. J. (1998) Passive immunity to bovine rotavirus in newborn calves fed colostrum supplements from cows immunized with recombinant SA11 rotavirus core-like particles (CLP) or virus-like particle (VLP) vaccines. Vaccine 16, 507–516. 37. Conner M. E., Estes, M. K., and Graham, D. Y. (1988) Rabbit model of rotavirus infection. J. Virol. 62, 1625–1633. 38. Conner M. E., Gilger, M. A., Estes, M. K., and Graham, D. Y. (1991) Serologic and mucosal immune response to rotavirus infection in the rabbit model. J. Virol. 65, 2562–2571. 39. Conner M. E., Crawford, S. E., Barone, C, and Estes, M. K. (1993) Rotavirus vaccine administered parenterally induces protective immunity. J. Virol. 67, 6633–6641. 40. Conner, M E., Graham, D. Y., and Estes, M. K. (1997) Determination of the duration of a primary immune response and the ID50 of ALA rabbit rotavirus in rabbits. Arch. Virol. 142, 2281–2294. 41. Thouless, M. E., DiGiacomo, R. F., Deeb, B. J., and Howard, H. (1988) Pathogenicity of rotavirus in rabbits. J. Clin. Microbiol. 26, 943–947. 42. Hambraeus, B. A. M., Hambraeus, L. E. J., and Wadell, G. (1989) Animal model of rotavirus infection in rabbits-protection obtained without shedding of viral antigen. Arch. Virol. 107, 237–251.

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71. Brown, K. A., Moser, C., Speaker, T., Khoury, C., Kim, J., and Offit, P. A. (1995) Enhancement by microencapsulation of rotavirus-specific intestinal immune responses in mice assessed by enzyme-linked immunospot assay and intestinal fragment culture. J. Infect. Dis. 171, 1334–1338. 72. Khoury, C., Moser, C., Speaker, T., and Offit, P. A. (1995) Oral inoculation of mice with low doses of microencapsulated, noninfectious rotavirus induces virus-specific antibodies in gut-associated lymphoid tissue. J. Infect. Dis. 172, 870–874. 73. Moser, C., Speaker, T., Berlin, J. A., and Offit, P. A. (1996) Aqueous-based microencapsulation enhances virus-specific humoral immune responses in mice after parenteral inoculation. Vaccine 14, 1235–1238. 74. Moser, C., Speaker, T., and Offit, P. A. (1998) Effect of water-based microencapsulation on protection against EDIM rotavirus challenge in mice. J. Virol. 72, 3859–3862. 75. Ciarlet, M., Gilger, M. A., Barone, C., McArthur, M., Estes, M. K., and Conner, M. E. (1998) Rotavirus disease, but not infection and development of intestinal histophatological lesions, is age-restricted in rabbits. Virology 251, 343–360. 76. Castrucci, G., Ferrari, M., Frigeri, M., Cilli, V., Perecca, L., and Donelli, G. (1985) Isolation and characterization of cytopathic strains of rotavirus from rabbits. Arch. Virol. 83, 99–104. 77. Ciarlet, M., Estes, M. K., and Conner, M. E. (1997) Comparative amino acid sequence analysis of the outer capsid protein VP4 from four lapine rotavirus strains reveals identity with genotype P[14] human rotaviruses. Arch. Virol. 142, 1059–1069. 78. Tanaka, T. N., Conner, M. E., Graham, D. Y., and Estes, M. K. (1988) Molecular characterization of three rabbit rotavirus strains. Arch. Virol. 98, 253–265. 79. Petric, M., Middleton, P. J., Grant, C., Tam, J. S., and Hewitt, C. M. (1978). Lapine rotavirus: preliminary studies on epizoology and transmission. Can. J. Comp. Med. 42, 143–147. 80. Sato, K., Inaba, Y., Miura, Y., Tokuhisa, S., and Matsumoto, M. (1982) Antigenic relationships between rotaviruses from different species as studied by neutralization and immunofluorescence. Arch. Virol. 73, 45–50. 81. Ciarlet, M., Estes, M. K., and Conner, M. E. (1999) Simian rhesus rotavirus (RRV) is a unique heterologous (non-lapine) rotavirus strain capable of productive replication and horizontal transmission in rabbits, submitted. 82. DiGiacomo, R. F., and Thouless, M. E. (1986) Epidemiology of naturally occurring rotavirus infection in rabbits. Lab. Anim. Sci. 36, 153–156. 83. Schoeb, T. R., Casebolt, D. B., Walker, V. E., Potgieter, L. N. D., Thouless, M. E., and DiGiacomo, R. F. (1986) Rotavirus-associated diarrhea in a commercial rabbitry. Lab. Anim. Sci. 36, 149–152. 84. Thouless, M. E., DiGiacomo, R. F., and Deeb, B. J. (1996) Effect of combined rotavirus and E. coli infections in rabbits. Lab. Anim. Sci. 46, 381–385. 85. Borriello, S. P. and Carman, R. J. (1983) Association of iota-like toxin and Clostridium spiroforme with both spontaneous and antibiotic-associated diarrhea and colitis in rabbits. J. Clin. Microbiol. 17, 414–418. 86. Evans, D. G., Cabada, F. J., and Evans, Jr., D. J. (1982) Correlation between intestinal immune response to colonization factor antigen I and acquired resistance

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Ciarlet and Conner to enterotoxigenic E. coli diarrhea in an adult rabbit model. Eur. J. Clin. Microbiol. 1, 178–185. Moon, H. W. (1994) Pathophysiology of viral diarrhea, in Viral Infections of the Gastrointestinal Tract (Kapikian, A. Z., ed.), Marcel Dekker, New York. pp. 27–52. O’Neal, C. M., Ciarlet, M., Carman, R., and Conner, M. E. (1999) Immune Response to Clostridium spiroforme and C. spiroforme toxoid in rabbits. Manuscript in preparation. Peeters, J. E., Pohl, P., and Charlier, G. (1984) Infectious agents associated with diarrhea in commercial rabbits: a field study. Ann. Rec. Vet. 15, 335–340. Kraft, L. M. (1957) Studies on the etiology and transmission of epidemic diarrhea of infant mice. J. Exp. Med. 106, 743–755. Adams, W. R. and Kraft, L. M. (1963) Epizootic diarrhea of infant mice: identification of the etiologic agent. Science 141, 359–360. Eydelloth, R. S., Vonderfecht, S. L., Sheridan, J. F., Enders, L. D., and Yolken, R. H. (1984) Kinetics of viral replication and local and systemic immune responses in experimental rotavirus infection. J. Virol. 50, 947–950. Greenberg, H. B., Vo, P., and Jones, R. (1986) Cultivation and characterization of three strains of murine rotavirus. J. Virol. 57, 585–590. Wolf, J., Cukor, G., Blacklow, N., Dambrauskas, R., and Trier, J. (1981) Susceptibility of mice to rotavirus infection: effects of age and administration of corticosteroids. Infect. Immun. 33, 565–574. Riepenhoff-Talty, M., and Lee, P.-C. (1982) Age-dependent rotavirus-enterocyte interactions. Proc. Soc. Exp. Biol. Med. 170, 146–152. Ramig, R. F. (1988) The effects of host age, virus dose, and virus strain on heterologous rotavirus infection of suckling mice. Microb. Pathog. 4, 189–202. McNeal, M. M., Broome, R. L., and Ward, R. L. (1994) Active immunity against rotavirus infection in mice is correlated with viral replication and titers of serum IgA following vaccination. Virology 204, 642, 650. Offit, P. A., Clark, H. F., Kornstein, M. J., and Plotkin, S. A. (1984) A murine model for oral infection with a primate rotavirus (simian SA11). J. Virol. 51, 233–236. Offit, P. A. and Clark, H. F. (1985) Protection against rotavirus-induced gastroenteritis in a murine model by passively acquired gastrointestinal but not circulating antibodies. J. Virol. 54, 58–64. Offit, P. A. and Clark, H. F. (1985) Maternal antibody-mediated protection against gastroenteritis due to rotavirus in newborn mice is dependent on both serotype and titer of antibody. J. Infect. Dis. 152, 1152–1158. Offit, P. A., Shaw, R. D., and Greenberg, H. B. (1986) Passive protection against rotavirus-induced diarrhea by monoclonal antibodies to surface proteins VP3 and VP7. J. Virol. 58, 700–703. Dunn, S. J., Burns, J. W., Cross, T., Vo, P., Ward, R. L., Bremont, M., and Greenberg, H. B. (1994) Comparisons of VP4 and VP7 of five murine rotavirus strains. Virology 203, 250–259.

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103. Ushijima, H., Morikawa, S., Mukoyama, A., and Nishio, O. (1995) Characterization of VP4 and VP7 of a murine rotavirus (YR-1) isolated in Japan. Jpn. J. Med. Sci. Biol. 48, 237–247. 104. Shaw, R. D., Merchant, A., Groene, W., and Cheng, E. H. (1993) Persistence of intestinal antibody response to heterologous rotavirus infection in a murine model beyond 1 year. J. Clin. Microbiol. 31, 188–191. 105. Greenberg, H. B., Clark, H. F., and Offit, P. A (1994). Rotavirus pathology and pathophysiology, in Rotaviruses (Ramig, R. F., ed.), Springer-Verlag, Berlin, pp. 255–283. 106. Guerin-Danan, C., Meslin, J. C., Lambre, F., Charpilienne, A., Serazat, M., Bouley, C., Cohen, J., and Andrieux, C. (1998) Development of a heterologous model in germfree suckling rats for studies of rotavirus diarrhea. J. Virol. 72, 9298–9302. 107. Ward, R. L., Bernstein, D., Shukla, R., Young, E., Sherwood, J. R., McNeal, M. M., Walker, M., and Schiff, G. M. (1989) Effects of antibody to rotavirus on protection of adults challenged with a human rotavirus. J. Infect. Dis. 159, 79–88. 108. Ward, R. L., Knowlton, D. R., Greenberg, H. B., Schiff, G. M., and Bernstein, D. (1990) Serum-neutralizing antibody to VP4 and VP7 proteins in infants following vaccination with WC3 bovine rotavirus. J. Virol. 64, 2687–2691. 109. Ward, R. L., McNeal, M. M., Sander, D. S., Greenberg, H. B., and Bernstein, D. (1993) Immunodominance of the VP4 neutralization protein of rotavirus in protective natural infections of young children. J. Virol. 67, 464–468. 110. Groene, W. S. and Shaw, R. D. (1992) Psoralen preparation of antigenically intact noninfectious rotavirus particles. J. Virol. Methods 38, 93–102.

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10 Methods of Rotavirus Detection, Sero- and Genotyping, Sequencing, and Phylogenetic Analysis Miren Iturriza Gómara, Jon Green, and Jim Gray 1. Introduction 1.1. Virus Detection The clinical symptoms associated with rotavirus (RV) gastroenteritis are not sufficiently characteristic to distinguish between RV infection and other causes of gastroenteritis. Therefore, laboratory procedures, including electron microscopy (EM) (1), enzyme-linked immunosorbent assays (ELISA) (2), passive particle agglutination tests (PPAT) (3,4), polyacrylamide gel electrophoresis (PAGE) (5), or reverse transcription-polymerase chain reaction (RT-PCR) (6–8), are necessary to confirm a clinical diagnosis of RV gastroenteritis. Virus-shedding in the feces of RV-infected individuals, with up to 1011 virus particles/mL feces at the peak of diarrhea, coincides with the duration of the diarrhea. However, virus-shedding can continue once symptoms have ceased (9). Laboratory diagnosis is relatively easy, either through the visualization of virus particles by EM , the detection of virus antigen (Ag) by ELISA or PPAT, or the detection of the virus genome by PAGE or RT-PCR. The specimen of choice for the diagnosis of RV infection is feces, but rectal swabs and soiled diapers can also be used, if feces is not available. Specimens should be collected between d 1 and 4 of illness and stored at 4°C until tested. Long-term storage should be at –70°C.

1.2. Detection of RV Particles by EM Direct EM examination of feces after negative staining reveals the characteristic morphology of RV particles (2) (Fig.1). Complete particles are ~75 nm in diameter and have a double-shelled icosahedral capsid, consisting of an outer and an inner shell (new structural data show that there are three From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ

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Fig. 1. Electron micrograph, showing the morphology of RVs after staining with PTA.

layers; see Chapter 2). Single-shelled particles are ~70 nm in diameter, through the loss of the smooth outer shell (10). Single-shelled particles have a rough appearance as a result of the capsomeres of the inner capsid forming shallow projections on the surface of the virus particle.

1.3. Immune EM Immune electron microscopy (IEM) can be used to increase the sensitivity of EM, although EM will be sufficiently sensitive to detect RVs in a fecal specimen collected <4 d after the onset of illness (1). In RV diagnosis, IEM is used to differentiate between the morphologically identical group A, B, and C RVs.

1.4. Detection of RV Ag by ELISA RVs can be detected in fecal specimens by ELISA (2), using broadly reactive antibodies (Abs) against epitopes of viral protein 6 (VP6) (the intermediate and inner shell protein), shared among group A RVs.

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1.5. Detection of RV Ag by PPAT Latex particles (3) or red blood cells (4) coated with RV Abs will agglutinate in the presence of RV Ag to produce macroscopically visible aggregates. Although agglutination tests may be performed more rapidly than EM or ELISAs, they are less sensitive and prone to nonspecific reactivity (4).

1.6. Detection of the RV Genome Genomes of RVs can be easily isolated from stool samples using the guanidinium isothiocyanate/silica method (11), based on the nucleic acidbinding properties of silica in the presence of a chaotropic agent, such as guanidinium isothiocyanate, which also inhibits ribonuclease activity. The extracted genomes can be separated directly by PAGE and visualized by silver staining (5), or can be used for group-specific RT-PCR. Group A RVs have been detected by RT-PCR of the gene segment 9, using oligonucleotide primers complementary to the 5'- and 3'-ends of the gene, which are conserved among members within the group (7). Group B and C RVs are morphologically identical to group A strains, and are not detected by commercial ELISAs or PPATs used for the diagnosis of RV infection. They can, however, be detected by RT-PCR using oligonucleotide primer pairs specific for group B (gene segment 8) and group C (gene segment 6) (8). RVs of different groups can also be recognized by their different RNA profiles on PAGE. These include differences in the relative migration of the RNA segments 10 and 11 of group A RVs, known as short and long electropherotypes. Atypical RNA profiles of group A RVs with genome rearrangements (12) limit the usefulness of group identification by recognition of RNA migration patterns.

1.7. Serotyping of RVs RVs are classified into groups (A–E), and there may be two additional groups (F,G); RV subgroups are determined by serological reactivities of VP6, the inner capsid protein, which contains common epitopes shared among group A RVs, and either zero, one, or several forms of the subgroup-specific epitopes. Classification within group A is by subgroups (I, II, I + II, and non-I,non-II), and G- and P-sero- and genotypes (G1–G14 and P1–P20, respectively). G- and P-types are defined by the reactivity of Ab to the two outer capsid proteins, VP7 and VP4, respectively. Monoclonal Abs (MAbs), specific for subgroup Ags (15) or G- and P-serotypes, are available for characterizing RV isolates in ELISAs (16–19). Most G-genotypes, but not P-genotypes, are serologically confirmed as serotypes (13). Therefore, for P-types, the genotype is indicated in square brackets and the serotype without brackets. To characterize a RV strain by its surface components, a double nomenclature

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has been introduced (14), e.g., the human RV strain Wa is defined as G1P1A[8] (see also Chapter 1).

1.7.1. G-Typing (VP7) All G- and P-typing ELISAs require the presence of intact double-shelled virus particles. Approximately 70–85% RVs in stool samples are typable by VP7–ELISA (20–23). A panel of neutralizing MAbs against different monotypes is necessary in order to detect all members of a particular G-serotype because of the existence of monotypes (intraserotypic differences in neutralization epitopes) within human RV G-serotypes (24–26).

1.7.2. P-Typing (VP4) MAbs to VP4 have proved to be more difficult to produce, and fewer have been raised to VP4 than to VP7. Many of the MAbs to VP4 are only reactive to a subset of strains within a P-type, can crossreact between P-serotypes, or give high background readings (18–19). MAbs reactive to VP8*, the smaller subunit of VP4 generated by proteolytic cleavage, are more suitable for VP4 typing by ELISA (26).

1.8. Genotyping of RVs G- and P-genotyping can be performed by RT-PCR using viral RNA extracted from the fecal samples. G-genotyping involves the amplification of fragments of the gene encoding VP7 (RNA segment 7, 8, or 9, depending on the strain), using oligonucleotide primers complementary to six variable regions that are very distinct between different G-types, but highly conserved within each G-type (7). Similarly, P-genotyping RT-PCR is performed with oligonucleotide primers complementary to four variable regions in the gene segment 4 (encoding VP4), which are highly conserved within each P-type, and distinct between different P-types (27).

1.8.1. G-Typing (VP7) The G-typing-PCR can be performed directly from the cDNA obtained after random priming-RT (28), either as a single round multiplex PCR or as a seminested PCR. In the nested PCR, the first-round amplifies the whole length of the VP7 gene, and the second round is a multiplex reaction which includes six different G-type-specific oligonucleotide primers, specific for G1, G2, G3, G4, G8, and G9, and the common primer (see Fig. 2)(7).

1.8.2. P-Typing (VP4) Similar to G-typing, P-typing can be performed as a multiplex single-round PCR or as a seminested PCR. In the seminested PCR, the first-round consensus

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Fig. 2. Diagrammatic representation of the VP7 gene, showing the location of the consensus regions within group A RVs, the regions conserved within types, and the location of the oligonucleotide primers used for RV RNA detection and genotyping.

oligonucleotide primers amplify a fragment of the gene segment 4 of all group A RVs. The second-round PCR includes the 3'-end consensus primer and four different oligonucleotide primers complementary to variable regions, which are specific for types P1A[8], P1B[4], P3[9], and P2A[6] (see Fig. 3)(27).

1.9. Sequencing PCR Amplicons There are several situations in which sequencing of the nucleotides (nt) of the first-round or second-round PCR amplicons may be necessary. Phylogenetic analysis of RV strains, monitoring changes within genotypes (genomic drift), investigating vaccine failures, or monitoring the emergence of new types, all rely significantly on sequence analysis. Genotyping PCRs may fail to identify the genotype in approx 10% of cases in which a first-round PCR product is obtained, but the second-round typing PCR does not reveal a specific type (28). In these cases, sequencing the first-round amplicons is necessary, in order to genotype the virus. PCR amplicons can be sequenced with or without prior cloning; methods for both are described in Subheadings 3.3.3. and 3.3.4. The use of automated fluorescence-based sequencing has largely replaced conventional sequencing, using 35S-labeled dideoxynucleotide chain terminators, and the method described in Subheading 3.3.4. refers to these. A key step in all sequencing methods is the careful preparation of template (i.e., PCR amplicon) to adequate concentration and purity.

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Fig. 3. Diagrammatic representation of the VP4 gene, showing the location of the consensus regions within group A RVs, the regions conserved within types, and the location of the oligonucleotide primers used for genotyping of five P types.

1.9.1. Purification of PCR Products Prior to the sequencing reaction, the unincorporated nts and primers remaining after the PCR must be removed or inactivated, because these interfere with the sequencing reaction. In most cases, this can be done simply by using a commercial spin column that allows the separation of amplicons and nucleotide/primers on the basis of size (e.g., GeneClean, Anachem, Luton, UK). Excellent results have also been obtained using the Exonuclease I/Shrimp Alkaline Phosphotase Treatment Kit (Amersham-Pharmacia Biotech, Herts, UK, product code US70995). The DNA to be sequenced must contain only a single-reaction product. It is necessary to first gel-purify the specific band when amplification bands, in addition to the specific amplicon, are visualized by agarose gel electrophoresis.

1.9.2. Cloning of PCR Products PCR products can be cloned prior to sequencing. This may be necessary when the PCR product yields are low, or when multiple PCR products are amplified. In the latter case, the specific amplicon should be gel-purified prior to cloning. Cloning of PCR products is a simple, rapid procedure, and is facilitated by the availability of commercial kits. The Invitrogen TOPO TA Cloning Kit (Invitrogen BV, The Netherlands) has been used widely and successfully in the authors’ laboratories; the method is described in Subheading 3.3.3.

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1.9.3. DNA Quantitation and Sequencing Reaction Cycle-sequencing reaction kits provide an adequate means to obtain sequence information from PCR products, provided the template DNA is adequately pure, and is present in sufficient quantity. For amplicons of 500–1000 bp in size, approx 75–100 ng purified template is required for reliable results. Accurate determination of DNA concentrations by spectrophotometry is not necessary: Estimation of the concentration by comparison with a known concentration of DNA, run simultaneously on an ethidium-bromide-stained agarose gel, is adequate. The use of the ABI Prism Dye Terminator Cycle Sequencing Kit, containing AmpliTaq DNA polymerase FS (Perkin-Elmer, Warrington, UK), which has given reliable results in the authors’ laboratories, and gives data of the order of 450 bp/run, is described in Subheading 3.3.4. Alternatively, the BigDye Terminator Kits (for use on ABI Prism 310, ABI373 with BigDye filter wheel, and ABI Prism 377 sequencing machines) may give data of longer amplicons.

1.9.4. Purification of Extension Products Unincorporated dye terminators must be removed before the samples can be analyzed by PAGE. Excess dye terminators in sequencing reactions confound the data, particularly in the first part of the sequence. Several protocols for purification of extension products are available, and the protocol of choice will vary, depending on the cycle-sequencing chemistry used. Spin column methods remove all excess terminators, but are relatively costly, compared to precipitation methods, which are also faster, but can be less efficient. The precipitation method described in Subheading 3.3.5. has worked reliably in the authors’ laboratories (see ref. 29 for alternative precipitation methods).

1.9.5. Polyacrylamide Gel Electrophoresis The concentration and volume of sample to load onto a gel depends on many factors, including the sequencing chemistry used, the quality and nature of the DNA template (DNA amplicon), oligonucleotide primer performance, and instrument configuration. Because of this, it is inappropriate here to give details for the generic preparation of sequencing gels, sample loading conditions, and optimal electrophoresis conditions. The authors therefore recommend consulting specific information relevant for the local sequencing equipment in use, and refer to the Perkin-Elmer–applied Biosystems manual, Automated DNA sequencing, available from the Perkin-Elmer web site (30). Many institutions and commercial companies now offer a sequencing gel service, in which the products of sequencing reactions, in a desiccated form, are submitted for PAGE. Sequence data are then returned on floppy disk or by

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Table1 Software Sequence Analysis Packages and Programes Software

Format Mac/PC

Suppliers’ URLs

Sequencher Lasergene (DNAStar) GeneJockey DNAsis

Mac/WinNT/Win 95 Mac/Win Mac Mac/Win

MacVector

Mac

http://www.genecodes.com/ http://www.dnastar.com/ http://www.biosoft.com/genejock.htm http://www.hitachi-soft.com/gs/ dnasis/jrney25.pdf http://deamon.oxmol.co.uk/prods/ macvector/

E-mail. This is often a cost-efficient solution when in-house facilities are not available, and/or, in some cases, when extra capacity is required. Availability of such services is advertised in the scientific press and via the Internet.

1.9.6. DNA Sequence Analysis A variety of sequence software packages and programs are now available for the importation of sequence data and their subsequent manipulation, alignment, and phylogenetic analysis. A far-from-comprehensive list of commonly used software packages is given in Table 1. These packages all have particular strengths, and selection of a package compatible with a laboratory’s needs requires evaluation within that laboratory. Demonstration versions of the software mentioned are available from the uniform resource locations (URLs) given in Table 1. All software programs are subject to the GIGO rule (garbage in; garbage out): the input of poor quality sequence data results in inaccurate subsequent analysis. Generally, if the software is unable to determine the sequence from the raw data, it is far better to reevaluate the sequencing method, particularly the template preparation, and resequence, rather than attempt to read ambiguous bases/regions by eye. Having obtained good-quality sequence data, it is necessary to compare them with other published sequences stored in EMBL (European Bioinformatics Institution, Hinxton, Cambridge, UK) or Genbank (National Center for Biotechnology Information, Bethesda, MD). Genomic sequences can be accessed from CD-ROMs distributed by these centers, or by Internet access (30–32). The authors strongly recommended that new sequences obtained should be submitted for deposition in these databases, in order to make them available to fellow researchers. Most journals do not accept manuscripts with sequence data before deposition in one of the databases has been confirmed.

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2. Materials 2.1. Detection

2.1.1. Electron Microscopy 1. 10% extract of feces prepared in balanced salt solution (BSS) 199 (Sigma, Dorset, UK). 2. 3% phosphotungstic acid (PTA), adjusted to pH 6.3 with potassium hydroxide. 3. Waxed slide or card. 4. Carbon-formvar-coated copper grids. 5. EM forceps (Agar Aids, Scientific Ltd., Stanstead, UK). 6. Transmission electron microscope.

2.1.2. Immune Electron Microscopy 1. 2. 3. 4. 5. 6. 7. 8.

Fecal extract (see Subheading 2.1.1.). RV group A-, B- and C-specific Abs. 3% PTA.KOH, pH 6.3. Waxed slides or cards. Carbon-formvar-coated copper grids. 500 µg/mL protein A solution (Sigma). Incubator. Transmission electron microscope.

2.1.3. RV Ag ELISA 1. 96-well flat-bottomed microtiter plate (Becton Dickinson, Oxnard, CA). 2. 0.1 M sodium carbonate–bicarbonate buffer (CBB), pH 9.6. 3. 0.1 M phosphate-buffered saline (PBS), pH 7.3 (Oxoid, Hampshire, UK), containing 0.05% Tween-20 (PBS-T). 4. Rabbit antigroup A RV Abs (Dako, Ely, Kent, UK). 5. 5 and 1% solutions of skimmed milk powder (Oxoid) in PBS. 6. Fecal extract (see Subheading 2.1.1.). 7. Anti-RV mouse MAb. 8. Horseradish peroxidase-labeled antimouse Abs (Sigma). 9. Substrate buffer: 0.05 M phosphate-citrate buffer, pH 6.0 (Sigma). 10. 0.1 mg/mL tetramethylbenzidine (TMB; Sigma) containing 0.014% hydrogen peroxide (H2O2) in substrate buffer. 11. 2 M sulfuric acid. 12. Spectrophotometer (e.g., Rosys Anthos, Hombrechticon, Switzerland). 13. Microtiter plate-washer (Nunc, Denmark). 14. Incubator.

2.1.4. Nucleic Acid Extraction 1. Screw-capped 1.5-mL microcentrifuge tubes. 2. Lysis buffer L6: Add 120 g guanidinium isothiocyanate (Fluka, Dorset, UK) to 100 mL 0.1 M Tris-HCl, pH 6.4; 22 mL 0.2 M EDTA, pH 8.0; and 2.6 g

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3.

4.

5. 6. 7.

8. 9. 10. 11.

Iturriza, Green, and Gray Triton X-100 (Sigma). Stir overnight in the dark, to dissolve. Store in the dark at room temperature. Washing buffer L2: Add 120 g guanidinium isothiocyanate to 100 mL 0.1 M Tris-HCl, pH 6.4, and stir overnight in the dark to dissolve. Store at room temperature in the dark. Size-fractionated silica: Add 60 g silicon dioxide (Sigma) to 500 mL distilled H 2O in a 500-mL glass measuring cylinder, and allow to stand for 24 h at room temperature. Aspirate 430 mL supernatant fluid, and resuspend the solids in 500 mL distilled H 2 O. Allow the solids to settle at room temperature for 5 h, and aspirate 440 mL supernatant fluid. Add 600 µL concentrated HCl, pH 2.0, and aliquot into 1.5-mL vol. Sterilize by autoclaving at 15 lb for 15 min, and store in the dark. Ethanol (70%) in distilled H2O. Acetone. RNase-free distilled H 2 O. To make RNase-free distilled H 2 O, add 100 µL diethylpyrocarbonate (DEPC;Sigma) in a fume cupboard to 100 mL tissueculture-grade distilled H 2 O, and incubate for >12 h at 37°C. Autoclave at 15 lb for 15 min to inactivate the DEPC. RNasin (Promega, Madison, WI). Vortex. Microcentrifuge. Incubator.

2.1.5. Polyacrylamide Gel Electrophoresis 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16.

Acrylamide (AA;Sigma). bis-Acrylamide (bisAA;Sigma). 1.5 M Tris-HCl buffer, pH 8.8. Sodium dodecyl sulfate (SDS) (Sigma). N,N,N',N'-tetramethylethylenediamine (TEMED) (Sigma). 20% ammonium persulfate (APS) (Sigma). Ethanol. Ether. Water-saturated butan-2-ol (butanol). To make 10X running buffer (0.25 M Tris-glycine); dissolve 30 g Tris and 144 g glycine in 1 L distilled H2O. To make loading buffer; add 2.5 mL 0.5 M Tris-HCl, pH 6.8, 4.0 mL glycerol, 1 mL 10% SDS, and 0.1 mL 0.1% bromophenol blue to 2.5 mL distilled H2O. To make fixing solution, add 125 mL methanol and 50 mL glacial acetic acid to 325 mL distilled H2O. 0.19% aqueous silver nitrate. To make developer, dissolve 14 g NaOH and 8 mL 37% formaldehyde in 1 L distilled H2O. Glass plates (20 cm2), spacers, and combs (1 mm thick). Power pack (e.g., Anachem).

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17. Vertical gel tank (Anachem). 18. Vacuum pump.

2.1.6. Reverse Transcription-Polymerase Chain Reaction 1. Random primers, Pd(N)6 (Pharmacia Biotech): 20 mU/mL. 2. Extracted RNA. 3. 10X PCR buffer II (Life Technologies, Paisley, Scotland): 200 mM Tris-HCl buffer, pH8, 500 mM KCl. 4. 50 mM MgCl2 (Life Technologies). 5. 10 mM equimolar mix of deoxynucleotide triphosphates (dNTPs) (Life Technologies). 6. 200 U/µL M-MLV reverse transcriptase (Life Technologies). 7. 5 U/µL Taq polymerase (Life Technologies). 8. 20 pmol/µL RV VP7 (for group A) and VP6 (for groups B and C) oligonucleotide primers. Group A RV oligonucleotide primers (7): Sense 1: 5' GGC TTT AAA AGA GAG AAT TTC CGT CTG G 3' (nt 1–28) Antisense 2: 5' GGT CAC ATC ATA CAA TTC TAA TCT AAG 3' (nt 1062–1036) Product size: 1062 bp Group B RV oligonucleotide primers (8): Sense 1: 5' CTA TTC AGT GTG TCG TGA GAG G 3' (nt 18–40) Antisense 2: 5' CGT GGC TTT GGA AAA TTC TTG 3' (nt 506–485) Product size: 489 bp Group C RV oligonucleotide primers (8): Sense 1: 5' CTC GAT GCT ACT ACA GAA TCA G 3' (nt 994–1016) Antisense 2: 5' AGC CAC ATA GTT CAC ATT TCA TCC 3' (nt 1349–1325) Product size: 356 bp 9. RNase-free distilled H2O. 10. Nusieve 3:1 agarose (Flowgen, Staffs, UK). 11. TAE buffer, pH 8.3: 0.04 M Tris-acetate, 0.001 M EDTA. 12. TE buffer pH 8.0: 10 m2M Tris-HCl, 1 mM EDTA. 13. Loading buffer TE buffer containing 10% Ficoll Sigma and 0.25% Orange G Sigma. 14. 10 mg/mL ethidium bromide (Sigma). 15. DNA ladder 100 bp (Pharmacia). 16. Horizontal gel tank (Anachem). 17. Power pack (Anachem). 18. Thermocycler (Genetic Research Instruments, Essex, UK). 19. Ultraviolet (UV) transiluminator (Sigma). 20. Microwave oven (domestic).

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2.2. Serotyping 2.2.1. Subgrouping ELISA 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

96-well flat-bottomed microtiter tray. 0.1 M CBB, pH 9.6. Rabbit anti-RV polyclonal Abs (Dako). 0.1 M PBS, pH 7.2, containing 0.05% (v/v) Tween-20 (PBS-T). Fecal extract (see Subheading 2.1.1.). 5 and 2.5% (w/v) solutions of skimmed milk powder in PBS-T. RV subgroup-specific mouse MAbs. HRP-labeled antimouse IgG (Sigma). 0.05 M phosphate-citrate buffer, pH 6.0 (substrate buffer) (Sigma). TMB (see Subheading 2.1.3.). H2O2 (see Subheading 2.1.3.). 2 M sulfuric acid.

2.2.2. Serotyping ELISAs 1. As in Subheading 2.2.1., but using specific MAbs against G1 (KU-4 or RV4:2), G2 (S2-2G10 or RV5:3), G3(YO-1E2 or RV3:1), and G4(ST-2G7 or ST3:1) types (14–15) (see Note 1). 2. As in Subheading 2.2.1., but using P-type specific MAbs: 2F2 (P1B-specific) YO-2C2 and KU- 4D7 (VP4 crossreactive).

2.3. Genotyping 2.3.1. G-Typing PCR As in Subheading 2.1.4. for the RNA extraction, and in Subheading 2.1.6. for the RT-PCR, including group A-VP7 consensus primers 1 and 2 for the first-round PCR, and the G-type-specific primers for the multiplex PCR. G-type-specific primers (7) (20 pmol/µL): G1 sense: 5' CAA GTA CTC AAA TCA ATG ATG G 3' (nt 314–335) G2 sense: 5' CAA TGA TAT TAA CAC ATT TTC TGT G 3' (nt 411–435) G3 sense: 5' CGT TTG AAG AAG TTG CAA CAG 3' (nt 689–709) G4 sense: 5' CGT TTC TGG TGA GGA GTT G 3' (nt 480–498) G8 sense: 5' GTC ACA CCA TTT GTA AAT TCG 3' (nt 178–198) G9 sense: 5' CTA GAT GTA ACT ACA ACT AC 3' (nt 757–776) Common antisense: 5' GGT CAC ATC ATA CAA TTC T 3' (nt 1062–1044)

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2.3.2. P-Typing PCR As in Subheading 2.1.5. for the RNA extraction, and as in Subheading 2.1.6. for the RT-PCR, but using the following primers: VP4 consensus primers (27) (20 pmol/µL): Sense 1 5' TGG CTT CGC CAT TTT ATA GAC A 3' (nt 11–32) Antisense 2 5' ATT TCG GAC CAT TTA TAA CC 3' (nt 887–868) P-typing primers (27) (20 pmol/µL): P1A[8]: antisense 5' TCT ACT TGG ATA ACG TGC 3' (nt 339–356) P1B[4]: antisense 5' CTA TTG TTA GAG GTT AGA GTC 3' (nt 474–494) P2A[6]:antisense 5' TGT TGA TTA GTT GGA TTC AA 3' (nt 259–278) P3[9]: antisense 5' TGA GAC ATG CAA TTG GAC 3' (nt 385–402) P4[10]: antisense 5' ATC ATA GTT AGT AGT CGG 3' (nt 575–594)

2.4. Sequencing PCR Amplicons 2.4.1. Direct Purification of PCR Products 1. Amplicon. 2. GeneClean spin kit (Anachem). 3. Microcentrifuge.

2.4.2. Purification of PCR Products from Agarose Gel 1. 2. 3. 4. 5.

Reagents as for gel electrophoresis listed in Subheading 2.1.6.; steps 9–17. GeneClean spin kit (Anachem). Long-wavelength UV transilluminator (366 nm). Heat block or water bath at 50°C. Microcentrifuge.

2.4.3. Cloning of PCR Products 1. 2. 3. 4.

Post-PCR mix containing specific amplicon. Invitrogen TOPO TA cloning kit (Invitrogen BV). Taq polymerase (Life Technologies). Selective agar: Luria-Bertani (LB) plates containing 50 µg/mL ampicillin (Sigma) or kanamycin (Sigma), two per transformation. 5. 40 mg/mL 5-bromo-4-chloro-3-indolyl–β-D-galactopyranoside (X-gal) (Sigma) in dimethylformamide (Sigma).

2.4.4. DNA Sequencing Reaction 1. Approximately 200 ng purified DNA template. 2. An aliquot of each sequencing primers (i.e., those used to generate the PCR product) at 3.2 pmol/µL.

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3. ABI Prism Dye Terminator Cycle Sequencing Kit (Perkin-Elmer). 4. Reaction tubes appropriate for the thermal cycler. For the Perkin-Elmer Thermal cycler 480, use 0.6 mL double-snap-cap microcentrifuge tubes. For the GeneAmp 9600 (Perkin-Elmer) and the GeneAmp 2400, use 0.2 mL Geneamp tubes.

2.4.5. Purification of Extension Products 1. 95% ethanol (ACS reagent grade, BDH, Dorset, Kent, UK).

3. Methods 3.1. Detection

3.1.1. Electron Microscopy 1. Place 10 µL PTA solution onto a waxed slide. 2. Emulsify a small portion of the fecal extract into the PTA, to achieve a slightly opaque suspension. 3. Float the EM grid onto the fecal suspension, and leave for 1 min. 4. Place 10 µL distilled H2O onto the waxed slide, adjacent to the specimen. 5. Remove the grid with sterile forceps, and remove the excess liquid by touching the side of the grid to a piece of blotting paper. 6. Float the grid onto the distilled H2O and leave for 1 min. 7. Remove the grid, and blot as before. 8. Allow the grid to air-dry while held in the forceps. 9. Place the grid in a clean Petri dish, and irradiate with UV light for 1 min. 10. Examine the grid by EM at ×30,000–50,000 magnification.

3.1.2. Immune Electron Microscopy 1. Float three carbon-formvar-coated copper grids onto three drops (10 µL) protein A solution. 2. Incubate in a moist chamber at room temperature for 20 min. 3. Immediately, on a second waxed card, mix 5 µL fecal extract with 5 µL RV group-specific Ab (see Note 2). 4. Place the card containing the virus–Ab mixture in a moist chamber at room temperature. 5. Remove the grids from the protein A solution, and blot by touching the edge of the grid to a piece of filter paper. 6. Float the grid onto a 10-µL drop of distilled H2O, and blot as before. 7. Repeat the washing step once. 8. Float the grid onto the virus–Ab mixture, and incubate in a moist chamber at 37°C for 1 h. 9. Repeat the wash procedure (two washes with distilled H2O). 10. Blot to remove excess liquid, and float the grid onto a 10 µL drop of PTA, and leave for 1 min. 11. Blot, and allow the grid to air-dry.

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12. Place the grid under UV light for 1 min. 13. Specific virus–Ab interactions are characterized by the presence of immune complexes spread evenly across the grid when viewed by EM.

3.1.3. RV Antigen ELISA 1. Coat each well of a 96-well flat-bottomed microtiter plate with 100 µL predetermined concentration (~10 µg/µL) of rabbit antigroup A RV Abs in CBB, pH 9.6, for 2 h at 37°C, or overnight at 4°C. 2. Aspirate the coating solution, and wash the wells 3× with 300–400 µL PBS-T. 3. Add 200 µL PBS containing 5% skimmed milk powder to each well, and incubate at 37°C for 30 min, to block the uncoated sites. 4. Remove the blocking agent by aspiration. 5. Add 25 µL 10% extract of feces in BSS 199 and 75 µL PBS-T containing 2.5% skimmed milk powder to the well, and incubate at 37°C for 30 min. Include three negative and one positive controls in each assay plate. 6. Wash the wells 5× with PBS-T, as before. 7. Add 100 µL mouse anti-RV Ab to each well, and incubate at 37°C for 30 min. 8. Wash the wells 5× with PBS-T, as before. 9. Add 100 µL HRP-labeled antimouse Abs to each well, and incubate at 37°C for 30 min. 10. Wash the wells 5× with PBS-T, as before. 11. Add 100 µL TMB/H 2 O 2 in substrate buffer, pH 6.0, to each well, and incubate at room temperature for 15–30 min. 12. Add 100 µL 2 M sulfuric acid to each well to stop the reaction. 13. Measure the optical density (OD) values at a wavelength of 450 nm. Specimens with an OD value of >3 standard deviations (SD) from the mean of the negative controls are regarded as RV Ag positive.

3.1.4. Nucleic Acid Extraction 1. Add 100 µL 10% fecal extract to 500 µL L6 buffer and 10 µL size-fractionated silica in a screw-capped, 1.5-mL microcentrifuge tube. 2. Vortex the tube for 10 s, and incubate at room temperature for 15 min. 3. Centrifuge at 11,000g for 15 s, and collect the supernatant for disposal (see Notes 3 and 4). 4. Wash the silica pellet with 500 µL L2 buffer twice, 500 µL 70% ethanol twice, and 500 µL acetone once. Centrifuge for 15 s at 11,000g after each wash, and collect the supernatant for disposal (see Note 3). 5. Remove the acetone, and place the tube, with the lid open, at 56°C in a dry heating block for 5 min. 6. Add 30 µL RNase-free distilled H 2 O containing 20–40 U ribonuclease inhibitor RNasin. 7. Vortex the tube, and incubate at 56°C for 15 min. 8. Centrifuge the tube at 11,000g for 2 min, and extract the supernatant. The extracted nucleic acid can be stored at 4°C overnight, or at –70 °C for longer.

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3.1.5. Polyacrylamide Gel Electrophoresis 1. Prepare a resolving gel by mixing 12.5 mL AA/bisAA (30% AA, 0.8% bisAA), 10 mL 1.5 M Tris-HCl, pH 8.8, 17.5 mL distilled H2O, and 400 µL 10% SDS. 2. Degas the solution for 20–30 min (see Note 5). 3. Add 50 µL TEMED and 75 µL 20% APS, to polymerize the gel. 4. Fill the solution between the two glass plates, previously cleaned with 70% ethanol and ether, and containing 1-mm-thick spacers at the bottom and right- and left-hand sides. 5. Overlay with 2 mL H2O-saturated butan-2-ol, and leave at room temperature for at least 30 min, to allow polymerization. 6. Prepare the collecting gel by mixing 1.5 mL AA/ bisAA (30% AA, 0.8% bisAA), 2.5 mL 0.5 M Tris-HCl, pH 6.8, 100 µL 10% SDS, and 6.0 mL distilled H2O. 7. Degas the solution for 20–30 min (see Note 5). 8. Remove the butanol from the resolving gel, and wash out the gel surface 3× with distilled H2O. 9. Add 30 µL TEMED and 30 µL 20% APS to the collecting gel, and pour 6 mL between the glass plates. 10. Immediately position the comb, and leave for at least 30 min. 11. Remove the comb and bottom spacer. 12. Add the running buffer containing 1 mL 10% SDS/100 mL buffer to the bottom reservoir of the vertical electrophoresis tank. 13. Insert the glass plate into the tank, and add running buffer to the top reservoir. 14. Remove air bubbles from the bottom of the gel, and wash the slots with running buffer. 15. Load the extracted RNA in 15 µL loading buffer. 16. Run the gel at 1 mA/cm for 18–20 h. 17. Discard the running buffer, and remove the gel from the glass plates. 18. Fix the gel for 30 min in the fixing solution. 19. Remove the fixative, and stain the gel with 0.19% silver nitrate solution for at least 30 min at room temperature in the dark. 20. Wash the gel 3×, 15 s for each wash, with distilled H2O. 21. Add the developer, and agitate for 1 min. Leave for up to 10 min, checking for visualization of the RNA segments. 22. Decant the developer and add the fixative. 23. Leave the gel for 10 min, then seal in a plastic bag.

3.1.6. RT-PCR for Detecting Group A RVs 1. Add 20 µL extracted nucleic acid and 1 µL random primers to a 0.2-mL PCR tube. 2. Incubate the tubes at 97°C for 5 min, to denature the double-stranded RNA. 3. Incubate the tubes at 70°C for 5 min. 4. Chill the tubes on ice for 2 min. 5. Prepare the reverse transcription mix (RT-mix) as follows (volumes indicated as per specimen):

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13. 14.

205

Component Volume µL 10X PCR buffer II 3.5 50 mM MgCl 2 3.5 dNTPs (10 mM mix) 1.0 M-MLV reverse transcriptase 1.0 (200 U/µL) RNase-free distilled H2O 5.0 Total 14.0 Add 14 µL RT-mix to each tube containing the RNA random primer mix. Incubate the tubes at room temperature for 10 min. Transfer the tubes to 42°C for 1 h (water bath or heating block). Transfer the tubes to a 95°C dry heating block for 5 min. Chill the tubes on ice for 2 min. The cDNA can be used directly in the PCR, or stored at –20°C for future use. Prepare the PCR reaction mix for each specimen as follows (volumes indicated as per specimen): Component Volume mL 10X PCR buffer II 4.5 2.0 50 mM MgCl2 dNTPs (10 mM mix) 1.0 Taq polymerase (5 U/mL) 0.2 Primer 1 (20 pmol/mL) 1.0 Primer 2 (20 pmol/mL) 1.0 RNase-free distilled H2O 35.3 Total 45.0 Add 45 µL PCR mix and 5 µL cDNA to a 0.2-mL PCR tube. Add the tubes to the thermocycler, and run the following programe: No. cycles Temperature (°C) Time (min) 1 94 2 94 1 30 42 2 72 1 1 72 7 1 4 Hold

}

}

3.1.6.1. AGAROSE GEL ELECTROPHORESIS 1. Add 3 g Nusieve (Flourgen) 3:1 agarose to 100 mL TAE buffer. 2. Melt in a microwave oven at full power for 1–2 min. 3. Cool to 45°C, then pour into a horizontal gel plate fitted with a 16-slot (40 µL/ slot) comb. 4. Allow the gel to set. 5. Add 20 µL PCR product to 10 µL TE buffer and 10 µL loading buffer in a V-well microtiter tray. 6. Remove the comb from the gel, and add 40 µL prepared mixture (see step 5 above) to the appropriate well. Include MN markers (Pharmacia).

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7. 8. 9. 10. 11. 12.

Place the gel plate in the gel tank, and add TAE buffer level with the gel. Run the products into the gel for 5 min at a constant voltage of 10–12 V/cm. Flood the gel with TAE, buffer, making sure it is fully submerged. Run at 10–12 V/cm for 1.5 h. Remove the gel plate from the electrophoresis tank. Remove the gel from the plate, and stain in a tank with 300 mL TAE buffer containing 100 µL ethidium bromide stock solution (final concentration 0.3 µg/mL) (see Note 6 for disposal). 13. Allow to stain for 30 min. 14. Examine on a UV transilluminator for bands of the predicted size (1062 bp).

3.1.6.2. RT-PCR FOR DETECTING GROUP B AND C RVS

The group-specific primer pairs can be used in individual PCR assays, or they can be pooled together with the group A primers in a multiplex PCR for the simultaneous detection of the three RV groups A, B, and C (8). Random priming, reverse transcription, and PCR are performed as described above (Subheadings 3.1.4. and 3.1.6.), but group B and group C RV-specific oligonucleotide primers are used, and the volume of water used in the PCR reaction mix is reduced, to take account of the increased primer volume.

3.2. Serotyping 3.2.1. Subgrouping ELISA 1. Coat each well of a 96-well flat-bottomed microtiter plate with 100 µL predetermined concentration of rabbit antigroup A RV Abs in 0.1 M CBB, pH 9.6, for 2 h at 37°C, or overnight at 4°C. 2. Aspirate the coating solution, and wash the wells 3× with 300–400 µL of PBS-T. 3. Add 200 µL PBS containing 5% skimmed milk powder to each well, and incubate at 37°C for 30 min, to block the uncoated sites. 4. Remove the blocking agent by aspiration. 5. Add 25 µL 10% fecal extract in BSS 199 and 75 µL PBS-T containing 2.5% skimmed milk powder to each of two wells (1 well/subgroup MAb to be tested against), and incubate overnight at 4°C. Include one positive and three negative controls in each assay plate. 6. Wash the wells 5× with PBS-T, as before. 7. Add 100 µL mouse antisubgroup I RV Ab to one well, 100 µL mouse antisubgroup II to the second well, and incubate at 37°C for 2.5 h (see Note 7). 8. Wash the wells 5× with PBS-T, as before. 9. Add 100 µL of HRP-labeled antimouse Abs to each well, and incubate at 37°C for 1.5 h. 10. Wash the wells 5× with PBS-T, as before. 11. Add 100 µL TMB (0.1 mg/mL)/ H2O2 (0.014%) in substrate buffer, pH 6.0, to each well, and incubate at room temperature for 15–30 min.

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12. Add 100 µL 2 M sulfuric acid to each well, to stop the reaction. 13. Measure the OD values at a wavelength of 450 nm. 14. Specimens with an OD of above >3 SD the mean of the negative controls are regarded as reactive.

3.2.2. Serotyping ELISA 3.2.2.1. G-TYPING ELISA

The ELISA is performed as in Subheading 3.2.1., but each sample (10% fecal extract) is added to four wells (or more), and antiserotype G1-, G2-, G3-, or G4specific MAbs (or other G-type-specific MAbs) are each added to one well. Specimens with an OD of >3 SD above the mean of the negative controls are regarded as reactive. A fecal sample is considered positive for a particular serotype, if its OD against the Ab specific for that serotype is at least 2× the mean of the OD of that sample against Abs to the other serotypes tested. 3.2.2.2. P-TYPING ELISA

The P-typing ELISA is performed in the same way as G-typing ELISA, in Subheading 3.2.2.1., but using Abs specific for the different P-types.

3.3. Genotyping 3.3.1. G-Typing PCR 3.3.1.1. MULTIPLEX SINGLE-ROUND G-TYPING RT-PCR 1. RNA extraction is performed as in Subheading 3.1.4. 2. Random priming and reverse transcription are performed as in Subheading 3.1.6. 3. Prepare the PCR reaction mix for each specimen as follows (volumes indicated as per specimen): Component Volume µL 10X PCR buffer II 4.5 50 mM MgCl2 2.0 dNTPs (10 mM mix) 1.0 Taq polymerase (5 U/ µL) 0.2 Primer G1 (20 pmol/ µL) 1.0 Primer G2 (20 pmol/ µL) 1.0 Primer G3 (20 pmol/ µL) 1.0 Primer G4 (20 pmol/ µL) 1.0 Primer G8 (20 pmol/ µL) 1.0 Primer G9 (20 pmol/ µL) 1.0 Common primer (20 pmol/ µL) 1.0 RNase-free distilled H2O 30.3 Total 45.0 4. Add 45 µL PCR mix and 5 µL cDNA to a 0.2-mL PCR tube.

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5. Add the tubes to the thermocycler, and run the following program: No. cycles Temperature (°C) Time (min) 1 94 2 94 1 30 42 2 72 1 1 72 7 1 4 Hold 6. Analyze the PCR products by agarose gel electrophoresis, as in Subheading 3.6.1., for bands of the predicted sizes: G1 (749 bp), G2 (652 bp), G3 (374 bp), G4 (583 bp), G8 (885 bp), and G9 (306 bp) (see Fig. 2.)

}

}

3.3.1.2. NESTED PCR RT-PCR

The first round is performed as in Subheading 3.1.6. The second round is a multiplex PCR reaction, as in Subheading 3.2.3., but using 1 µL first-round product as template, and reducing the number of amplification cycles to 15.

3.3.2. P-Typing PCR 3.3.2.1. SEMINESTED RT-PCR 1. RNA extraction is performed as in Subheading 3.1.4. 2. Random priming and reverse transcription are performed as in Subheading 3.1.6. 3. Prepare the first-round PCR reaction mix for each specimen as follows: Component Volume µL 10X PCR buffer II 4.5 50 mM MgCl2 2.5 dNTPs (10 mM mix) 1.0 Taq polymerase (5 U/ µL) 0.2 Primer 1 (20 pmol/ µL) 0.5 Primer 2 (20 pmol/ µL) 0.5 RNase-free distilled H2O 35.8 Total 45.0 4. Add 45 µL PCR mix and 5 µL cDNA to a 0.2-mL PCR tube. 5. Add the tubes to the thermocycler and run the following programe: No. cycles Temperature (°C) Time (min) 1 94 2 94 1 40 50 1 72 2 1 72 7 1 4 Hold 6. Prepare the second-round PCR mix for each specimen as follows: Component Volume (µL) 10X PCR buffer II 4.9 50 mM MgCl2 2.5

}

}

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dNTPs (10 mM mix) 1.0 Taq polymerase (5 U/µL) 0.2 Primer P1A[8] (20 pmol/µL) 1.0 Primer P1B[4] (20 pmol/µL) 1.0 Primer P3[9] (20 pmol/µL) 1.0 Primer P4[10] (20 pmol/µL) 1.0 Primer P2A[6] (20 pmol/µL) 1.0 Primer 1 (20 pmol/µL) 1.0 RNase-free distilled H2O 34.4 Total 49.0 7. Add 49 µL PCR mix and 1 µL first-round PCR product to a 0.2-mL PCR tube. 8. Add the tubes to the thermocycler, and run the following programe: No. cycles Temperature (°C) Time (min) 1 94 2 94 1 25 45 2 72 1 1 72 7 1 4 Hold 9. Analyze the PCR products by agarose gel electrophoresis, as in Subheading 3.1.6.1., for bands of the predicted sizes. First-round product (876 bp), P4[10] (583 bp), P1B[4] (483 bp), P3[9] (391 bp), P1A[8] (345bp), and P2A[6] (267 bp) (see Fig. 3). An example of a representative ethidium bromide stained agarose gel is shown in Fig. 4.

}

}

3.3.2.2. MULTIPLEX SINGLE-ROUND RT-PCR

The single-round P-typing PCR is performed using the same PCR mix as for the second-round multiplex in Subheading 3.2.4.1., adding 45 µL PCR mix and 5 µL cDNA obtained after random priming-RT. The PCR programe is as in the second-round VP4-PCR, but with an increase in the number of PCR cycles to 40.

3.4. Sequencing PCR Amplicons 3.4.1. Purification of PCR amplicons 1. Shake to mix GeneClean Spin Glassmilk, and add 400 µL to Spin Filter. 2. Add 50 µL amplicon, and incubate at room temperature for 5 min with agitation, if possible. If not, invert the tube every minute to prevent settling of the glassmilk. 3. Centrifuge in a microcentrifuge at 11,000g for 1 min, to spin the liquid out of the Spin Filter into the catch tube, and discard the liquid in the catch tube. 4. Add 500 µL of GeneClean Spin new wash to the Spin Filter and centrifuge for 30 s (or until the Spin Filter is emptied of wash). 5. Empty the catch tube, and spin for 1 min to dry the pellet in the Spin Filter.

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Fig. 4. Agarose gel stained with ethidium bromide, showing the results of G- and P-typing PCRs of 12 fecal specimens. Lanes 1 and 26: size marker (Pharmacia 100-bp ladder). Lanes 2–13: G-types. The majority of the specimens were typed as G1 (amplicon size 749 bp; lanes 2–4, 6, 7, 8, 10, and 11), one G2 (amplicon size 652 bp; lane 9), two G3 (amplicon size 374 bp; lanes 7 and 12), one G4 (amplicon size 583 bp; lane 5), and one G9 (amplicon size 306 bp; lane 13). Lanes 14–25: P-types. One specimen was typed as P1B[4] (amplicon 483 bp; lane 21), and was the G2 specimen, (lane 9). The remaining 11 specimens were typed as P1A[8] (amplicon size 345 bp; lanes 14–20 and 22–25). 6. Transfer Spin Filter to clean catch tube, add 30 µL sterile distilled H2O, and resuspend the glassmilk by flicking the tube or by gently vortexing for 2 s. 7. Centrifuge for 30 s, to transfer eluted DNA to the catch tube, discard the Spin Filter and cap the tube.

3.4.2. Purification of PCR Products from Agarose Gel 1. Gel electrophoresis of 25 µL PCR product is as previously described (see Subheading 3.1.6.1.), except that the bands are visualized under longwave UV (see Note 9). 2. Excise the specific amplicon band from the gel using a scalpel (use a new scalpel for each sample to avoid cross-contamination), and transfer to a sterile 1.5-mL microcentrifuge tube. 3. Add 750 µL NaI buffer, and place in a water bath at 50°C until the gel is dissolved.

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4. Shake to mix GeneClean Spin Glassmilk, and add 5 µL Glassmilk to the dissolved agarose/NaI solution. 5. Vortex the tube, and then chill on ice for at least 10 min. 6. Centrifuge at 11,000g for 15 s, and discard the supernatant. 7. Add 500 µL new wash buffer, and resuspend the Glassmilk pellet thoroughly. 8. Repeat steps 6 and 7 twice, and then remove all the supernatant. If necessary, spin the tube again to collect residual wash buffer, and then discard the supernatant. 9. Resuspend the pellet in 30 µL sterile, distilled H2O, and place in a water bath for 3 min at 50°C. 10. Centrifuge at 11,000g for 30 s, and remove the supernatant to a sterile tube.

3.4.3. Cloning of PCR Products 1. Check the PCR amplicon to be cloned by agarose gel electrophoresis. If other bands, in addition to the specific band, are visible, gel-purify the specific band (see Subheading 3.3.2.). 2. Use the Invitrogen TOPO TA cloning kit: Thaw a vial of SOC medium (kit box 2), and bring to room temperature. 3. Warm LB plates containing 50 µg/mL ampicillin or 50 µg/mL kanamycin at 37°C for 30 min. 4. Spread 40 µL 40 mg/mL X-gal on each LB plate, and incubate at 37°C until ready for use. 5. Thaw, on ice, 1 vial of One Shot™ cells for each transformation. Place the 2-mercaptoethanol on ice. 6. Set up the following 5 µL TOPO cloning reaction: 1 µL fresh PCR product; 1 µL TOPO vector; 4 µL sterile H2O; 5 µL final volume. 7. Mix gently and incubate for 5 min at room temperature (~25°C). Do not leave for more than 5 min, or the transformation and cloning efficiencies will decrease (see Note 8). 8. Briefly centrifuge, and place on ice. 9. Proceed immediately to next step. Add 2 µL 0.5 M 2-mercaptoethanol to each vial of competent cells, and mix gently. 10. Add 2 µL TOPO cloning reaction into a vial of One ShotTM cells, and mix gently. 11. Incubate on ice for 30 min. 12. Heat-shock the cells for 30 s at 42°C, without shaking. 13. Immediately transfer the tubes to ice, and incubate for 2 min. 14. Add 250 µL SOC medium at room temperature. 15. Cap the tube tightly, and shake the tube horizontally at 37°C for 30 min (ampicillin selection), or 1 h (kanamycin selection), then place on ice. 16. Spread 50 µL from each transformation onto selective plates. 17. Incubate overnight at 37°C. 18. Pick ~10 white colonies, and inoculate into 1 mL LB broth, and, using the residual colony on the loop, inoculate into a PCR mix containing the M13 forward and reverse primers.

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19. Amplify using the following programe: No. cycles Temperature (°C) Time (min) 1 94 2 94 1 30 55 1 72 1 1 72 10 1 4 Hold 20. Visualize products by agarose gel electrophoresis, as described in Subheading 3.1.6.1.

}

}

3.4.4. DNA Sequencing Reaction 1. For each reaction, mix the following reagents in a labeled tube: Component Volume µL Terminator Ready Reaction mix (Perkin-Elmer) 8.0 Template (~75–100 ng purified amplicon) 3–6 Primer (3.2 pmol) 1.0 Sterile distilled H2O: add to final volume of 20.0 2. Overlay the reaction mixture with mineral oil if necessary (according to thermocycler instructions). 3. Place the tubes in the thermocycler, and begin cycle sequencing with the following thermal profile : No. Cycles Temperature (°C) Time 25 Rapid thermal ramp to 96°C 96 30 s Rapid thermal ramp to 50°C 50 15 s Rapid thermal ramp to 60°C 60 4 min Rapid thermal ramp to 4°C Hold

3.4.5. Purification of Extension Products 1. For each reaction, add 35 µL 95% ethanol to a 1.5-mL microcentrifuge tube. 2. Transfer all 20 µL of each extension reaction to the tubes, and mix the contents by vortexing. 3. Incubate the tubes on ice for 10 min (see Note 10). 4. Place the tubes in a microcentrifuge, mark their orientations, and centrifuge the tubes for 20 min at 11,000g. Proceed to the next step immediately (see Note 11). 5. Carefully aspirate the supernatants with a separate pipet tip for each sample, and discard. Pellets may or may not be visible. 6. Add 250 µL 70% ethanol to the tubes, and vortex them briefly (see Note 12). 7. Place the tubes in the microcentrifuge in the same orientation as in step 4, and centrifuge for 10 min at 11,000g. 8. Aspirate the supernatant carefully. 9. Dry the samples in a vacuum centrifuge for 10–15 min, or to dryness (see Note 13).

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Fig. 5. Phylogenetic tree of the VP7 genes (determining G-types) of different group A RVs (G1: strain G192B, Genbank accession no. HRD113; G2: strain G292A, Genbank accession no. V73947, and strain G292B, Genbank accession no. V73948; G3: Genbank accession no. D86264, Genbank accession no. D86271, and SA11, Genbank accession no. K02028; G4: strain ST4, Genbank accession no. A01321; G8: strain D68, Genbank accession No. AF034852; G9: apathogenic neonatal strain I321, Genbank accession no. L07658, and G13: equine RV, Genbank accession no. D13549). Two RV group B VP7 sequences (human and bovine group B strains, Genbank accession nos. M33872 and V84141, respectively) were used as outliers. The phylogenetic tree was calculated using the Clustal method (DNAStar, Lasergene, Madison, WI). The horizontal calibration bar indicates the degree of nt divergence. Vertical distances are not numerical, but purely presentational.

3.4.6. Analysis of Sequences 1. Establish sequence, and analyze using programs, as indicated in Table 1. An example of a phylogenetic tree is shown in Fig. 5.

4. Notes 1. These MAbs have been successfully used to serotype RVs from stool samples in this laboratory. Other MAbs have been described in the literature (23,24). 2. A series of Ab dilutions can be used to achieve equivalence between Ag and Ab in order to maximize the formation of immune complexes. 3. Guanidinium isothiocyanate waste must be decanted into a container in which 50% of the volume is 10 N NaOH. 4. Important: Resuspend the silica thoroughly after each wash step. 5. Prepare the gel in a conical flask with a side arm, and degas under vacuum to remove oxygen that inhibits polymerization. 6. Ethidium bromide containing waste must be decontaminated by filtration through an extractor device consisting of an activated carbon matrix (Sigma), which is disposed of after use, by incineration. 7. The working dilution of each MAb is established by titration against negative and positive sample controls. MAbs are diluted in a 2.5% solution of skimmed milk powder in PBS-T. 8. If a delay is absolutely necessary, the tubes can be left on ice for up to 30 min.

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9. The amplicons are visualized under long-wavelength UV light (rather than short wave), to minimize UV damage to the amplicon DNA. 10. For the BigDye terminator reaction, precipitation of 15 min at room temperature is recommended. 11. If it is not possible to proceed, then spin the tubes for 2 min immediately prior to performing the next step. 12. Wash step: Optional for the ABI Prism Dye terminator kit reactions, but recommended for the BigDye reactions. 13. Alternatively, place the tubes, with the lids open, in a heat block or thermocycler at 90°C for 1 min.

References 1. Morinet, F., Ferchal, F., Colimon, R., and Perol, Y. (1984) Comparison of six methods for detecting human rotavirus in stools. Eur. J. Clin. Microbiol. 3, 136–140. 2. Birch, C. J., Lehmann, N. I., Hawker, A. J., Marshall, J. A., and Gust, I. D. (1979) Comparison of electron microscopy, enzyme-linked immunosorbent assay, solidphase radioimmunoassay, and indirect immunofluorescence for detection of human rotavirus Ag in faeces. J. Clin Pathol. 32, 700–705. 3. Sanekata, S., Yoshida, Y., and Okada, H. (1981) Detection of rotavirus in faeces by latex agglutionation. J. Immunol. Methods 41, 377–385. 4. Gray, J. J., Vasquez, A. M., Kewley, D. R., and Coombs, R. R. A. (1990) Reverse passive haemagglutination with freeze-dried stabilized antibody-coupled red cells for detecting rotavirus in faecal samples. Serodiag. Immunother. Infec. Dis. 4, 143–149. 5. Herring, A. J., Inglis, N. F., Ojeh, C. K., Snodgrass, D. R., and Menzies, J. (1982) Rapid diagnosis of rotavirus infection by direct detection of viral nucleic acid in silver-stained polyacrylamide gels. J. Clin. Microbiol. 16, 473–477. 6. Xu, L., Harbour, D., and McCrae, M. A. (1990) The application of polymerase chain reaction to the detection of rotaviruses in faeces. J. Virol. Methods 27, 29–38. 7. Gouvea, V., Glass, R. I., Woods, P., Taniguchi, K., Clark, H. F., Forrester, B., and Fang, Z-Y. (1990) Polymerase chain reaction amplification and typing of rotavirus nucleic acid from stool specimens. J. Clin. Microbiol. 28, 276–282. 8. Gouvea, V., Allen, J. R., Glass, R. I., Fang, Z-Y., Bremont, M., Cohen, J., McCrae, M. A., Saif, L. J., Sinarachatanant, P., and Caul, E. O. (1991) Detection of group B and C rotaviruses by polymerase chain reaction. J. Clin. Microbiol. 29, 519–523. 9. Richardson, S., Grimwood, K., Gorrell, R., Palombo, E., Barnes, G., and Bishop, R. (1998) Extended excretion of rotavirus after severe diarrhoea in young children. Lancet 351, 1844–1848. 10. Prasad, B. V., Wang, G. J., Clerx, Y. P., and Chiu, W. (1988) Three-dimensional structure of rotavirus. J. Mol. Biol. 199, 269–275. 11. Boom, R., Sol, C. J. A., Salismans, M. M. M., Jansen, C. L., Wertheim-van Dillen, P. M. E., and van den Noordaa, J. (1990) Rapid and simple method for the purification of nucleic acids. J. Clin. Microbiol. 28, 495–503. 12. Desselberger, U. (1996). Genome rearrangements of rotaviruses. Adv. Virus Res. 46, 69–95.

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13. Kapikian, A. Z. and Chanock, R. M. (1996) Rotaviruses, in Fields Virology, 3rd ed., (Fields, B. N, Knipe, D.M., and Howley, P. M., et al., eds.), Lippincott-Raven, Philadelphia, pp. 1657–1708. 14. Estes, M. K. (1996) Rotaviruses and their replication in Fields Virology, 3rd ed., (Fields, B. N, Knipe, D. M., and Howley, P. M., et al., eds.), Lippincott-Raven, Philadelphia, pp. 1625–1656. 15. Greenberg, H. V., McAuliffe, V., Valdesuso, J., Wyatt, R., Flores, J., Kalika, A., Hoshino, Y., and Singh, N. (1983) Serological analysis of the subgroup protein of rotavirus using monoclonal Abs. Infect. Immun. 39, 91–99. 16. Taniguchi, K., Urasawa, T., Morita, Y., Greenberg, H. B., and Urasawa, S. (1987) Direct serotyping of human rotavirus in stools by an enzyme-linked immunosorbent assay using 1-, 2-, 3-, and 4-specific monoclonals antibodies to VP7. J. Infect. Dis. 155, 1159–1166. 17. Coulson, B. S., Unicomb, L. E., Pitson, G. A., and Bishop, R. F. (1987) Simple and specific enzyme immunoassay using monoclonal antibodies for serotyping human rotaviruses. J. Clin. Microbiol. 25, 509–515. 18. Coulson, B. (1993) Typing of human rotavirus VP4 by an enzyme immunoassay using monoclonal antibodies. J. Clin. Microbiol. 31, 1–8. 19. Padilla-Noriega, L., Werner-Eckert, R., Mackow, E. R., Gorziglia, M., Larralde, G., Taniguchi, K., and Greenberg H. B. (1993) Serologic analysis of human rotavirus serotypes P1A and P2 by using monoclonal antibodies. J. Clin. Microbiol. 31, 622–628. 20. Beards, G. M., Desselberger, U., and Flewett, T. H. (1989) Temporal and geographical distributions of human rotavirus serotypes, 1983 to 1988. J. Clin. Microbiol. 27, 2827–2833. 21. Bishop, R. F., Unicomb, L. E., and Barnes, G. L. (1991) Epidemiology of rotavirus serotypes in Melbourne, Australia, from 1973 to 1989. J. Clin. Microbiol. 29, 862–868. 22. Woods, P. A., Gentsch, J., Gouvea, V., Mata, L., Simhon, A., Santosham, M., Bai, Z-S., Urasawa, S., and Glass, R. I. (1992). Distribution of serotypes of human rotavirus in different populations. J. Clin. Microbiol. 30, 781–785. 23. Noël, J. S., Beards, G. M., and Cubitt, W. D. (1991) Epidemiological survey of human rotavirus serotypes and electropherotypes in young children admitted to two children’s hospitals in Northeast London from 1984-1990. J. Clin. Microbiol. 29, 2213–2219. 24. Coulson, B. S. and Kirkwood, C. (1991) Relation of VP7 amino acid sequence to monoclonal antibody neutralisation of rotavirus and rotavirus monotype. J. Virol. 65, 5968–5974. 25. Pushker, R., Matson, D. O., Coulson, B. S., Bishop, R. F., Taniguchi, K., Urasawa, S., Greenberg, H. B., and Estes, M. K. (1992) Comparisons of rotavirus VP7-typing monoclonal antibodies by competition binding assay. J. Clin. Microbiol. 30, 704–711. 26. Coulson, B. S. (1996) VP4 and VP7 typing monoclonal antibodies. Arch. Virol. 12(Suppl.), 113–118.

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27. Gentsch, J. R., Glass, R. I., Woods, P., Gouvea, V., Gorziglia, M., Flores, J., Das, B. K., and Bhan, M. K. (1992) Identification of group A rotavirus gene 4 types by polymerase chain reaction. J. Clin. Microbiol. 30, 1365–1375. 28. Iturriza-Gómara, M., Green, J., Brown, D. W. G., Desselberger, U., and Gray, J. J. (1999) Comparison of specific and random priming in the reverse transcription polymerase chain reaction for genotyping group A rotaviruses. J. Virol. Methods 72, 93–103. 29. Perkin Elmer Applied Biosystems. Precipitation Methods to remove Residual Dye Terminators from Sequencing Reactions User Bulletin; P/N 4304655. Warrington, Cheshire, UK, WWW site(http://www2.perkinelmer.com/ab/techsupp/pdf/ga/ub/Precipitation_UB.pdf) 30. Perkin-Elmer Web site. HYPERLINK, http://www2.perkinelmer.com/ab/ techsupp/doclib/ga/multi/refguide/pdf_ /DNASeqCG.pdf). 31. EMBL can be accessed via http://www.ebi.ac.uk/. 32. Genbank can be accessed via HYPERLINK http://www.ncbi.nlm.nih.gov//.

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11 Epidemiology of Group A Rotaviruses Surveillance and Burden of Disease Studies Mary Ramsay and David Brown 1. Introduction Human infection has been reported with groups A, B, and C rotaviruses (RVs). Of these, Group A RVs are the most important, being a major cause of severe gastroenteritis (GE). Each year, Group A RVs are estimated to cause approx 870,000 deaths worldwide in children less than 5 years (yr) of age, mostly in developing countries (1). This chapter will describe the epidemiological features of Group A RV infections, and will critically review the current surveillance strategies used to define the burden of disease. 2. Epidemiology of Human RV Infection 2.1. Age and Sex RV infections occur throughout life, but most recurrent infections are mild or asymptomatic (2). A prospective 29-month (mo) study in the United States (US) detected RV infection in 28% of children and 16% of adults under study (3). Serological data suggest that a high proportion of children have been infected at least once, but mostly several times by the age of 3 yr (4). Symptoms usually accompany primary infection, which is followed by protection against subsequent symptomatic infection (5). For this reason, the ratio of symptomatic to asymptomatic infection decreases with age (6). In prospective studies, symptomatic infection rates were highest in children under 2 yr, and lowest in those over 45 yr of age (6,7). This finding has been confirmed in studies conducted during outbreaks (8,9). In the newborn, RV infection is common (10), but is often associated with only mild disease or is asymptomatic disease (11,12). The mild manifestation of RV infection at this From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ

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age may be caused by the protection provided by passively acquired maternal antibody (Ab), or to infection with attenuated “nursery” strains (2,12). Like other childhood infections, severe diarrhea caused by RV is more common in males than females (3,13–15). Adult RV infection also occurs, and frequent asymptomatic adult infections may be important in maintaining the transmission of infection in the community (16). High levels of serum Ab detected in adults of child-bearing age suggest that secondary infection occurs from contact with young children (17). Outbreaks have been described in closed adult communities, particularly those for the elderly (18,19), as have waterborne outbreaks and cases of traveller’s diarrhea (16). A similar pattern of infection is seen in the developing world, but, because of a greater intensity of transmission, the peak age of infection is often very early in life (20). A recent study from Mexico showed that the majority of children had experienced four RV infections by the age of 5 yr (21).

2.2. Morbidity Symptomatic RV disease predominantly affects young children (2), with the peak attack rate for severe diarrhea between 6 and 23 mo of age in the developed world. In a prospective community study in the US, the annual incidence of symptomatic infection was estimated to be: 11/100 person-years in children under 12 mo; 40/100 person-years in children aged 12–23 mo; 13/100 person-years in children aged 24–35 mo; 8/100 person-years in children aged 36–59 mo; 6/100 person-years in children aged 5–15 yr; and 5/100 person-years in adults (3). In studies of children admitted with GE in Australia, RV was detected in 55% of those aged 12–23 mo, and in 15% of those aged under 6 mo (22). A small proportion of all cases of RV diarrhea require admission to hospital, and disease severity is strongly related to age (23). The incidence of hospital admission is greatest in children under 2 yr, and then declines rapidly (23). The annual admission rate was estimated to be 3.7/1000 in children under 1 yr, and 2.2/1000 in children aged 1–2 yr (23). Because of these high admission rates, RV is a major cause of childhood admissions for GE worldwide. The organism was identified in 15–55% of hospital admissions of children with diarrhea in various countries (15,23–35; Table 1). RV infection is also a common cause of medical outpatient consultation for diarrhea in children. RV diarrhea leads to dehydration more commonly than other causes of childhood GE (36) and therefore tends to represent a smaller proportion of childhood diarrhea managed in the community than among hospitalized cases. In an Australian study, 32% of children seen and discharged from the accident and emergency department had RV in their stools, compared to 51% of those

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Table 1 Association of RV Infection with Childhood Hospital Admissions Worldwide Location Finland UK Venezuela Hong Kong Singapore Africa Trinidad Italy Spain Bangladesh New Zealand Australia

Percent hospital admissions with RV GE

Ref.

49 39 30–50 35 27 24 23 22–27 21 20 19 15–55

(24) (15) (25) (26) (27) (28) (29) (30) (31) (32) (33) (22,34,35)

admitted (35). In Venezuela, RV was found to be responsible for 30–50% of hospitalized diarrhea, compared to only 10% of diarrhea in the community (25). In the United Kingdom (UK), RV was identified in 28 (37%) of those with a causative organism in general practice consultations for childhood GE (37).

2.3. Risk Factors Case control studies indicate that risk factors for RV infection include household contact with children under 2 yr of age (38), a low-fat milk diet (39), and formula milk feeding (40). Outbreaks of RV infection are common in daycare centers (42,43), and in hospitals, mostly on neonatal units and pediatric wards (10,44–48), and in geriatric wards (18,19,49,50), perhaps reflecting waning protection against symptomatic infection in the elderly. Outbreaks of symptomatic infection are less commonly described in primary schools (51).

2.4. Temporal and Geographical Distribution RV has a worldwide distribution. In a review of 34 studies from different countries, RV was detected in a median of 33% (11–71%) of children with diarrhea (52). In temperate zones, RV infections are said to show a marked excess of infections in the winter months (2,53,54). In an 8-yr follow-up study in Washington, DC, Brandt et al. (54) have observed a “clockwise precision” of the appearance of RV infections in hospitals during the winter months, but seasonal peaks have also been described in autumn and spring (52). Within a single season, the spread of RV infections has been described across North

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America from southwest in the autumn to northeast in spring (55). In Europe, an epidemic of childhood diarrhea was described moving across France from west to east (56), and the seasonal RV increase was detected 2 mo later in the Netherlands (57). The peak incidence of RV infection appears to occur later in the season, moving from the south and west to the east and north of Europe (58).

2.5. Transmission RV is mainly spread by the feco-oral route (59). Certain epidemiological features (the seasonal excess in cooler months and the rapid spread in both temperate and tropical zones) have led to the suggestion that RV may also be spread by the respiratory route. Although RVs do cause infection in a wide variety of animals, Group A RV infection does not appear to be predominantly a zoonosis. The possibility of reassortment occurring between animal and human strains has been suggested, and evidence of transmission between humans and animals is accumulating (60,61). Interspecies transmission of genes, however, appears to be a rare event, and the importance of such strains in human epidemiology is not clear (2).

2.6. RV Diversity Group A RVs have been identified in many animal and bird species: They are antigenically complex, and multiple serotypes infect humans. Protection from disease and from reinfection with RV are not fully understood, but protection correlates with the presence of neutralizing Abs to two surface proteins, the viral proteins VP4 and VP7, and with the presence of intestinal RV-specific immunoglobulin (Ig)A and neutralizing copro-Abs (62–64). VP7 has been identified as the major neutralizing antigen, and VP4 as the minor neutralizing antigen, although the latter can be immunodominant (5,63,65). A binary classification system is now used to describe group A RV strains, and serotypes are defined as P (protease-sensitive)- and G (glycoprotein)-types, based on variations of VP4 and VP7, respectively. In developed countries, >90% of group A RV strains that have been typed are of serotypes G1, G2, G3, or G4, with, on average, 8% of non-G1, G2, G3, or G4 strains in published studies (58,66). A limited number of P-types have been described (58,63). The percentage of nontypeable strains may fluctuate from one year to another, and has been as high as 18% in one study in the UK (67). Fewer studies have been published from the developing world, and these have shown a more diverse picture, with G9 strains commonly identified in India (60) and G5 strains in Brazil (68). This, and the more recent identification of unusual G9 strains in the US (69), indicates the need for a systematic study of RV diversity.

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3. Surveillance of RV Infection and Disease Surveillance information is required for the planning, implementation, and assessment of disease control and prevention (70). Population surveillance has been defined as “ongoing scrutiny, generally using methods distinguished by their practicability, uniformity and, frequently, their rapidity, rather than complete accuracy” (71). Surveillance systems therefore include the collection of data, analysis, interpretation, and feedback of information (72). When the disease under surveillance is vaccine-preventable, surveillance should encompass all aspects of the vaccination program: disease incidence, vaccine coverage, seroprevalence of immunity, and adverse events attributable to the vaccine (73). The objectives of surveillance of vaccine-preventable disease will depend on the stage of the vaccination program (74). Prior to the implementation of a vaccination program, policymakers need to estimate the burden of disease and to decide upon the appropriate vaccination strategy. When the vaccination program has been implemented, disease incidence data are required to monitor the effectiveness of that strategy. When the aim of the vaccination strategy is the elimination or eradication of the infection, surveillance is required to identify remaining pockets of susceptibility, and to modify the existing strategy to provide the maximum impact on the transmission of infection. In the US, a tetravalent (TV), rhesus RV (RRV)-based, human reassortant vaccine has been shown to prevent severe RV disease (75–77) and is now recommended for routine use, with the aim of reducing the burden from severe RV diarrhea (78). As yet, RV vaccines have not been integrated into the vaccination program in any other country (79). In most countries, therefore, vaccine policymakers are attempting to accurately estimate the true burden of disease caused by RV infection, and to decide on the choice of the most appropriate vaccination strategy (80). A baseline period of data collection is desirable, prior to vaccine implementation, and data collection should continue to monitor the impact of the vaccination program on the incidence of infection.

3.1. Sources of Data on RV GE 3.1.1. Clinical Reporting Schemes for Notifications of GE In many countries, routine sources of data on vaccine-preventable disease include notifications to the local public health authorities. This is often a legal requirement for the clinician making the diagnosis, and the primary objective is to provide an opportunity for local public health action (81). Weekly or monthly summaries are sent to a national office to provide timely data on national trends. Unlike other vaccine-preventable diseases, however, GE is rarely notifiable, except when the infection is considered to be food- or waterborne (82). In contrast to other causes of GE, however, RV is not

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predominantly transmitted by this route, and therefore notification of food poisoning is not likely to be a useful source of data on RV infection.

3.1.2. Sentinel Physician Reporting Schemes Sentinel physician reporting involves the establishment of a group of motivated medical practitioners who agree to report certain infections on a regular basis. Completeness of reporting can be enhanced by stimulating the practitioner to report, and by requiring a response, even if no case has been seen. Many of these schemes have a defined catchment population as a denominator, which allows estimation of age-specific incidence rates. In this type of scheme, only a small population can be surveyed, and this approach is therefore most useful where the infection is fairly common. General practice reporting schemes have been established in the UK (83), the Netherlands (84), Belgium (85), and France (86). Many of these schemes monitor the incidence of GE, based on outpatient consultations with symptoms and signs that conform to a clinical case definition (56,84). The epidemiological data collected include the age and sex of the patient, and the location of the practice. These schemes provide timely data on the age, sex, and geographical location of cases of GE (87,88); however, because most schemes are based in primary care, laboratory confirmation of the infection is rarely performed routinely. These schemes can be enhanced by ensuring that samples are taken from patients for bacteriological and virological investigation (57,84,89). Alternative approaches to attributing the etiology of GE in general practice include ad hoc surveys (37) or exploiting the seasonality of RV infection, to estimate the proportion attributable to RV (90).

3.1.3. Laboratory Reporting Schemes In many countries, laboratory-based surveillance of RV is performed as part of a generic reporting system for laboratory-confirmed infections (14,15,91,92). For example, in England and Wales, laboratories report a minimum data set of organism, patient identifier (used to avoid duplicate reporting), age, sex, date of sample, type of specimen, method of confirmation, and the location of laboratory for all confirmed infections (93). Additional microbiological, clinical, and epidemiological features are reported for certain infections (94). To monitor RV circulation in the US, the Centers for Disease Control, together with state and local public health laboratories, have established the laboratory-based National Respiratory and Enteric Virus Surveillance System (NREVSS)(95). In 1991–1996, 5343 of 22,912 fecal specimens examined were positive for RV (95). The Public Health Laboratory Service in England receives reports of confirmed infec-

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Table 2 Quarterly RV Laboratory Reports in England and Wales, 1992–1997 Year

First quarter

Second quarter

Third quarter

Fourth quarter

Total

1992 1993 1994 1995 1996 1997 1998

10,176 7879 8892 6788 4577 5436 4370

3799 5148 4603 8825 7438 7261 7131

823 776 906 1064 1332 1403 1909

1641 1835 1003 485 709 792 1364

16,439 15,638 15,404 17,162 14,056 14,892 14,774

Laboratory reports to Communicable Disease Surveillance Center, Public Health Laboratory Service, London; http://www.phls.co.uk/facts/rota

tions at higher numbers (ref. 96; Table 2), compared to NREVSS, and calculations on the overall disease burden this signifies are beginning to emerge from such data (15,90). Alternative models of laboratory surveillance include collation of data in a single reference laboratory, where samples are referred for confirmation (97), or reporting from a small sentinel group of laboratories. The latter approach is an option where the infection is fairly common, so that sufficient cases are detected to allow valid extrapolations from the data (15,98–101). All laboratory reporting schemes are subject to the same problems of underreporting as clinical schemes. In addition, the routine laboratory investigation of GE may significantly underestimate the number of RV infections, for various reasons. For example, the test for RV may not actually be performed, or samples may be taken late in the course of infection (101). Despite these reservations, laboratory surveillance is felt to be a good indicator of trends in disease incidence (72). In addition to a confirmatory role in surveillance, reference laboratories can perform specialized testing to describe qualitative features of the organism (72). Regarding RV infection, these tests will include sero- and/or genotyping of the virus (68,69,97,101,102). Reassortant RV vaccines are now available that confer protection against severe illness caused by RV serotypes G1–G4 (75–77). Laboratory surveillance can be used to estimate the proportion of symptomatic RV infection attributable to the strains covered by the current vaccine, or to detect different types that might escape from protection through vaccination. Vaccine failures will also be detectable. Because different RNA segments code for VP4 and VP7, various combinations of P- and G-types can be found in natural isolates (103).

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3.1.4 Medical Care Records Statistics on hospital admissions for diarrhea are becoming increasingly available, often in a computerized format (104). To ensure consistency, the European Community has recommended a minimum data set for recording hospital morbidity (105). This minimum dataset will include age, sex, geographical location, date of admission, and reason for hospitalization. Hospitalized cases reflect the symptomatic and more severe infections, but data sources are usually exhaustive, and therefore fairly complete. One of the problems with these kinds of data is that patient identifiers may be removed; multiple episodes of admission of the same individual may occur, and theoretically lead to double-counting of cases, although this should be infrequent with RV disease (104). In addition, hospital admissions are usually coded for the reason of admission using established coding schemes. Not all commonly used coding schemes, however, contain a code for RV infection. According to the ninth revision of International Classification of Diseases (ICD9; 106), a diagnosed RV infection should be coded to viral GE resulting from nonspecific cause (13,15,107). Previous studies suggest that miscoding of RV diarrhea to less specific codes commonly occurred in episodes coded under ICD9 in both the US (107) and the UK (15). This evidence suggests that miscoding would also remain under systems with specific codes for RVGE, e.g., using the tenth revision of the International Classification of Diseases, 1992 (108,109).

3.1.5 Death Certificates In most developed and many developing countries, a physician completes a death certificate outlining the likely cause of death for each patient who dies in his or her care. The certificates are then registered, and the cause of death coded, using established schemes (106,108). The coding of cause of death is variable between countries, but death certification is complete in most developed countries. Differentiating among deaths from the various causes of GE may be difficult, but a case series from Toronto suggests that RV can lead to death by dehydration in the developed world (110). Similarly, in Australia, the excess of deaths from dehydration caused by GE in the winter has been attributed to RV infection (111).

3.1.6 Seroprevalence Data Serum IgA and IgG to RV rise in response to acute infection, and remain detectable for up to 6 mo (2). In primary infection, RV-specific IgM can be detected, and its presence is correlated with severity of disease, indicating that primary infections are more clinically severe than subsequent infections (112).

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Passively acquired maternal serum Abs have declined by 1 yr of age (4), and IgG detected after this age can be used as a marker of past infection of the child (2). Age-dependent seroprevalence studies have been performed in several countries (17,113), which studies show that the prevalence of Ab and Ab titers increase as the incidence of infection increases (4). One study demonstrated high titers of Ab in women of child bearing age: This was felt to be consistent with secondary infection from their infants (17).

3.1.7. Special Surveys In addition to routine surveillance, special surveys can be performed on an ad hoc or a regular basis. For acute infections, prospective cohort studies in a defined population can be used to estimate the incidence of infection. Such cohort studies can avoid underestimation of the disease incidence by establishing morbidity actively, e.g., by regular interviews of the study population. For a common childhood infection, such as with RVs, this may involve weekly or fortnightly contact. Because RV disease shows a marked seasonality in many communities, any studies should extend over at least 12 mo, starting and finishing in the low-incidence period. RV infection can result in a range of clinical presentations, from asymptomatic infection to severe dehydrating disease. The case definition for reporting of childhood diarrhea caused by RV should be chosen to reflect severe disease that would be preventable by vaccination (75–77). The data collected on each case should include age, sex, breast feeding, number of siblings, date of onset, and the clinical features of the disease. A clinical scale for the severity of illness has been described (Table 3; 114), and such standard definitions for severity should be used to allow comparison within and between study groups. Questionnaires and patient information leaflets for such studies are available upon request.

3.2. Analysis and Interpretation of Surveillance Data Basic analysis of surveillance data should include the description of cases in terms of time, place, and person (72,104). Analysis by real time will allow assessment of seasonal trends, and support the early detection of outbreaks (115). Interpretation of such trends requires a detailed knowledge of the source of the surveillance data (72). Changes in incidence can be related to the introduction of public health interventions, to changes in clinical and/or laboratory practice, and to changes in the population. The interpretation of trends in laboratory surveillance data is aided by collection of the denominator of tests performed (98). Analysis by time and place is also vital in the detection of clusters or outbreaks, and to the early introduction of control measures (72). When the infection is vaccine-preventable, additional data collected should include the vaccination status of the case.

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Table 3 Numerical Scoring for Severity of RV Diarrhea (114) Symptom or sign Duration of diarrhea (d)

Maximum no diarrheal stools/24 h

Duration of vomiting (d)

Maximum no vomiting episodes/24 h

Fever (°C)

Dehydration

Treatment

Grouping

Score

1–4 5 ≥6 1–3 4–5 ≥6 1 2 ≥3 1 2–4 ≥5 <37.0 37.1–38.4 38.5–38.9 ≥39 None 1–5% ≥6% None Rehydration Hospitalization

1 2 3 1 2 3 1 2 3 1 2 3 0 1 2 3 0 2 3 0 1 2

3.3. Evaluation of Surveillance Schemes An ideal surveillance system should be simple, flexible, acceptable, complete, sensitive, specific, representative, and timely (116). The chief emphasis of the surveillance system will depend on the stage of the vaccination program. Prior to the introduction of vaccination, accurate estimates of disease burden are required. Completeness is therefore important, so that disease burden is not underestimated. If the extent of underascertainment is known, however, corrections to burden estimates can be made. Representativeness is also important, to ensure that vaccination is targeted to the appropriate groups. Underascertainment in surveillance systems can be assessed by comparison with special surveys, or by matching of different data sources. A statistical method known as capture–recapture has been developed to estimate the underreporting, when there are two or more independent reporting schemes for the same infection (117). After vaccination is introduced, the specificity of the case definition in the reporting scheme becomes very important. As the incidence of the infection

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declines, the positive predictive value of the case definition will fall, and more false-positive clinical cases will occur. Studies to estimate the proportion of clinically diagnosed cases that can be confirmed by laboratory investigation will be required (74).

3.4. Dissemination of Surveillance Information The dissemination of information from surveillance is vital in maintaining the motivation of reporting personnel (72), and can therefore help to improve the accuracy and completeness of the data. The regular outputs of surveillance systems need to be targeted at clinicians, laboratories, public health control officers, and decision-makers. Feedback of information may be by regular publication or in electronic format (94). Such information may be important in encouraging awareness of an infection, and hence the appropriate management among clinicians. Information may also be important in demonstrating the need for control measures. In particular, surveillance data can be used to obtain political commitment to mass vaccination programs. 4. Uses of RV Surveillance Data

4.1. Estimation of Disease Burden Choice of Vaccination Strategy Prior to a decision on whether to introduce mass vaccination, estimates of the burden of disease are required. In addition, before vaccines are introduced, it is necessary to establish the diversity of RVs in the target population, to ensure that the proposed vaccine will cover the prevalent serotypes in that population, and to establish a baseline for future surveillance strategies. Prior to licensing, the efficacy of the TV, RRV-based human ressortant vaccine has been established against various case definitions (75–77). By estimating the incidence of disease of a similar severity, the burden of disease potentially preventable by mass vaccination can be established, and economic analyses can be performed (80,118–120). The economic impact of a vaccination program will include the savings of direct medical costs (e.g., health service costs of treating acute disease) and of social costs (e.g., loss of earnings of parents caring for children) (120). Cost–benefit studies, comparing the cost of providing vaccination against the potential benefits of prevention of GE, are often required to influence vaccine policymakers (80,119). Rates of disease can be established in prospective cohort studies (25), or by community-based surveillance in stable communities. Surveillance data, if complete and linked to denominator information on the population under surveillance, can then be extrapolated to estimate the national burden of infection. For RVs, the population under surveillance would normally be all children under 5 yr of age resident in one or more defined areas. Sentinel physician schemes may be useful for assessing the burden of disease, because they are

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expected to be complete, and can usually be linked to age-specific denominator information (72). Ideally, the population should be sociodemographically representative of the total childhood population, including both rural and urban populations. Because the medical practitioners participating in sentinel schemes usually volunteer, the population under surveillance may not be fully representative of those in other practices (121) and in the general population. Because one of the primary aims of RV vaccination is to prevent diarrhea that leads to medical intervention, exhaustive data sets of medical care records can also provide data on the burden of disease. Hospital admission for RV infection is common, so hospital records have been used to estimate the burden of disease on several occasions (13–15,32,97,107,109,122). To circumvent issues with nonspecific coding of admissions, three approaches have been tried. The first approach simply used the winter excess of diarrhea in children under 5 yr to attribute the admissions thought to be attributable to RV infections (13,97,107). Second, a more sophisticated modeling technique was applied to obtain accurate estimates of the seasonal excess by combining weekly data on laboratory-confirmed infections with weekly data on hospitalizations (14,15). The third approach used prospective, cross-sectional, or retrospective surveys in small numbers of hospitals in order to obtain estimates of the proportion of childhood GE caused by RV infection, and to apply these estimates to the national number of hospital admissions (32,122). In addition, by using a standard severity scale (Table 3) to assess the severity of disease requiring hospitalization (114), estimates of the number of preventable hospital admissions can be made in countries with different cultural and health care practices, and the data become comparable (101). Deaths also potentially contribute to the assessment of disease burden, and to any cost–benefit analysis of vaccination. In the developed world, however, the numbers of deaths caused by childhood diarrhea are small (15,91,123). In the US, analysis of deaths by season was able to estimate those attributable to RV (123), but indicated that mortality from RV diarrhea has been falling (123,124). The numbers of deaths caused by childhood diarrhea are also small in Finland (97) and the UK (15). To accurately assess the true incidence of infection from surveillance data, age-specific adjustments must be made for asymptomatic disease. For other vaccine-preventable infections, age-specific seroprevalence rates have been used to estimate the incidence of infection (125). With the current candidate RV vaccine, however, efficacy against mild or asymptomatic infection is low (75–77). The chief aim of vaccination is to prevent severe disease, and estimates of the incidence of asymptomatic infection are therefore not urgently required. Age-specific seroprevalence studies are not likely to prove useful for assessing the burden of disease attributable to RV infection

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4.2. Monitoring Effectiveness of Vaccination Program After a vaccine is introduced, the case definition used in the surveillance system must be specific for the preventable infection. RV vaccine will not prevent GE caused by other organisms, and therefore reductions in the numbers of cases of clinical GE will underestimate the true impact of vaccination. To accurately monitor the effectiveness of the vaccination program, a laboratory diagnosis of RV must be obtained in clinical reporting schemes, or a laboratory reporting system must be established. To quickly detect the effects of vaccine introduction, or of changes in vaccine coverage, surveillance data must be also be timely. Laboratory surveillance of confirmed RV infection can provide timely and accurate data to demonstrate the impact of a vaccination program. Such data can be combined with estimates of vaccine coverage, and with information on the vaccination status of cases, in order to estimate the efficacy of the vaccine under field conditions (126). Laboratory surveillance can also be used to establish the impact of vaccination on the diversity of endemic RV infection. Present vaccines confer protection against severe disease with serotypes G1–G4, and it is conceivable that vaccination will result in the selection of RV serotypes for which the vaccine might be less effective. It has been shown that RVs can evolve by genetic drift (resulting from accumulation of point mutations), and genetic shift (resulting from reassortment of gene segments) (63,103). In addition, there is some evidence of zoonotic infection with animal viruses, and that reassortment may occur between human and animal RVs in vivo, thus further increasing the genetic diversity of human group A RV strains (127). Concern about the development of vaccine-escape mutants has been expressed regarding other vaccine-preventable infections (127,128) and has necessitated the establishment of sophisticated surveillance based on modern molecular techniques. Jin et al. (129) analyzed RV strains found in vaccinated children during one of the vaccine trials, and found no consistent genetic change between the VP7 genes of serotype 1 strains isolated from vaccine recipients and those receiving a placebo. With increasing vaccine coverage, however, this situation may change. By starting surveillance before vaccine programs are introduced (130), and by analysis of RV vaccine failures, such genetic changes may be detected at an early stage. Hospital admission data could also be used to monitor the impact of RV vaccination (109). A fall in the total numbers of hospital admissions and the loss of the winter seasonal excess, outside normal fluctuations seen yearby-year, will be good indicators of the impact of vaccination. Similarly, changes in the incidence of GE may be monitored in sentinel practitioner schemes.

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Without data on the proportion of consultations or admissions due to RV, however, the magnitude of the effect attributable to vaccine will be difficult to determine, as thresholds for admission and for consultation may change. A group of sentinel hospitals, in which all admissions for childhood diarrhea are examined for laboratory evidence of RV infection, has been suggested as a means of monitoring the effect of vaccine (109,123). Alternatively, incidence estimates could be available from computerized inpatient and outpatient data collected for the defined catchment population such as that covered by a health maintenance organization (109). The impact of the vaccine on herd immunity, providing protection from infection, can be modeled using mathematical techniques (131), and the age-specific susceptibility can be assessed using seroprevalence studies (132). The mathematical approach is more complex, however, for antigenically variable infectious agents (128). Because of this, and because the current vaccine is unlikely to lead to the development of significant herd immunity, such approaches may not be essential for surveillance of RV infection. References 1. Bern, C., Martines, J., de Zoysa, I., and Glass R. I. (1992) The magnitude of the global problem of diarrhoeal disease: a ten-year update. Bull. WHO 70, 705–714. 2. Bishop R. F. (1996) Natural history of human rotavirus infection. Arch. Virol. 12(Suppl.), 119–128. 3. Rodriguez, W. J., Kim, H. W., Brandt, C. D., et al. (1987) Longitudinal study of rotavirus infection and gastroenteritis in families served by a pediatric medical practice: clinical and epidemiologic observations. Ped. Infect. Dis. J. 6, 170–176. 4. Brüssow, H., Werchau, H., Liedtke, W., et al. (1988) Prevalence of antibodies to rotavirus in different age-groups of infants in Bochum, West Germany. J. Infect. Dis. 157, 1014–1022. 5. Ward, R. L. (1996) Mechanisms of protection against rotavirus in humans and mice. J. Infect. Dis. 174(Suppl. 1), S51–S58. 6. Bernstein, D. I., Sander, D. S., Smith, V. E., Schiff, G. M., and Ward, R. L. (1991) Protection from rotavirus reinfection: 2-year prospective study. J. Infect. Dis. 164, 277–283. 7. Koopman, J. S. and Monto, A. S. (1989) The Tecumseh Study. XV: Rotavirus infection and pathogenicity. Am. J. Epidemiol. 130, 750–759. 8. Friedman, M. G., Galil, A., Sarov, B., et al. (1988) Two sequential outbreaks of rotavirus gastroenteritis: evidence for symptomatic and asymptomatic reinfections. J. Infect. Dis. 158, 814–822. 9. Linhares, A. C., Pinheiro, F. P., Freitas, R. B., Gabbay, Y. B., Shirley, J. A., and Beards, G. M. (1981) An outbreak of rotavirus diarrhoea among a nonimmune, isolated South American Indian community. Am. J. Epidemiol. 113, 703–710. 10. Cameron, D. J., Bishop, R. F., Veenstra, A. A., and Barnes, G. L. (1978) Noncultivable viruses and neonatal diarrhoea: fifteen-month survey in a newborn special care nursery. J. Clin. Microbiol. 8, 93–98.

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11. Grillner, L., Broberger, U., Chrystie, I., and Ransjo, U. (1985) Rotavirus infections in newborns: an epidemiological and clinical study. Scand. J. Infect. Dis. 17, 349–355. 12. Bishop, R. F. (1994) Natural history of human rotavirus infections, in Viral Infections of the Gastrointestinal Tract (Kapikian, A. Z., ed.), New York, Marcel Dekker, pp. 131–167. 13. Ho, M. S., Glass, R. I., Pinsky, P. F., and Anderson, L. J. (1988) Rotavirus as a cause of diarrheal morbidity and mortality in the United States. J. Infect. Dis. 158, 1112–1116. 14. Visser, L. E., Cano Portero, R., Gay, N. J., and Martinez Navarro, J. F. (1999) The impact of rotavirus disease in Spain: an estimate of hospital admissions due to rotavirus. Acta Paediatr. Scand. 88(Suppl. 426), 72–76. 15. Ryan, M. J., Ramsay, M., Brown, D., Gay, N. J., Farrington, C. P., and Wall, P. G. (1996) Hospital admissions attributable to rotavirus infection in England and Wales. J. Inf. Dis. 174(Suppl. 1), S12–S18. 16. Hrdy, D. B. (1987) Epidemiology of rotaviral infection in adults. Rev. Infect. Dis. 9, 461–469. 17. Holdaway, M. D., Kalmakoff, J., Schroeder, B. A., Wright, G. C., Todd, B. A., and Jennings, L. C. (1982) Rotavirus infection in Otago: a serological study. New Zeal. Med. J. 95, 110–112. 18. Cubitt, W. D. and Holzel, H. (1980) An outbreak of rotavirus infection in a longstay ward of a geriatric hospital. J. Clin. Pathol. 33, 306–308. 19. Halvorsrud, J. and Orstavik, I. (1980) An epidemic of rotavirus-associated gastroenteritis in a nursing home for the elderly. Scand. J. Infect. Dis. 12, 161–164. 20. Kapikian, A. Z. (1997) Viral gastroenteritis, in Viral Infections of Humans: Epidemiology. (Evans, A. S. and Kaslow, R. A., eds.), New York, Plenum, pp 285–343. 21. Velazquez, F. R., Matson, D. O., Calva, J. J., et al. (1996) Rotavirus infections in infants as protection against subsequent infections. N. Engl. J. Med. 335, 1022–1028. 22. Crawley, J. M., Bishop, R. F., and Barnes, G. L. (1993) Rotavirus gastroenteritis in infants aged 0–6 months in Melbourne, Australia: implications for vaccination. J. Paediatr. Child. Health 29, 219–221. 23. Rodriguez, W. J., Kim, H. W., Brandt, C. D., et al. (1980) Rotavirus gastroenteritis in the Washington, DC, area: incidence of cases resulting in admission to the hospital. Am. J. Dis. Child. 134, 777–779. 24. Vesikari, T., Maki, M., Sarkkinen, H. K., Arstila, P. P., and Halonen, P. E. (1981) Rotavirus, adenovirus, and non-viral enteropathogens in diarrhoea. Arch. Dis. Childh. 56, 264–270. 25. Pérez-Schael, I. (1996) The impact of rotavirus disease in Venezuela. J. Infect. Dis. 174(Suppl. 1), S19–S21. 26. Biswas, R., Lyon, D. J., Nelson, E. A., Lau, D., and Lewindon, P. J. (1996) Aetiology of acute diarrhoea in hospitalized children in Hong Kong. Trop. Med. Int. Health 1, 679–683. 27. Vijayan, V., Quak, S. H., and Wong, H. B. (1990) Incidence, clinical features and epidemiology of rotavirus gastroenteritis in hospitalized children. Ann. Trop. Paediatr. 10, 179–183.

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28. Cunliffe, N. A., Kilgore, P. E., Bresee, J. S., et al. (1998) Epidemiology of rotavirus diarrhoea in Africa: a review to assess the need for rotavirus immunization. Bull. WHO 76, 525–537. 29. Hull, B. P., Spence, L., Bassett, D., Swanston, W. H., and Tikasingh, E. S. (1982) The relative importance of rotavirus and other pathogens in the etiology of gastroenteritis in Trinidadian children. Am. J. Trop. Med. Hyg. 31, 142–148. 30. Ruggeri, F. and Declich, S. (1999) Rotavirus infection among children with diarrhoea in Italy. Acta Paediatr. Scand. 88(Suppl. 426), 66–71. 31. Velasco, A., Mateos, M., Mas, G., Pedraza, A., Diez, M., and Gutierrez, A. (1984) Three-year prospective study of intestinal pathogens in Madrid, Spain. J. Clin. Microbiol. 20, 290–292. 32. Unicomb, L. E., Kilgore, P. E., Faruque, S. G., et al. (1997) Anticipating rotavirus vaccines: hospital-based surveillance for rotavirus diarrhea and estimates of disease burden in Bangladesh. Pediatr. Infect. Dis. J. 16, 947–951. 33. Kuttner, N. S. (1985) Electron microscopic findings in faeces from children with gastroenteritis. New Zeal. Med. J. 98, 801–804. 34. Barnes, G. L., Uren, E., Stevens, K. B., and Bishop, R. F. (1998) Etiology of acute gastroenteritis in hospitalized children in Melbourne, Australia, from April 1980 to March 1993. J. Clin. Microbiol. 36, 133–138. 35. Pitson, G. A., Grimwood, K., Coulson, B. S., et al. (1986) Comparison between children treated at home and those requiring hospital admission for rotavirus and other enteric pathogens associated with acute diarrhea in Melbourne, Australia. J. Clin. Microbiol. 24, 395–399. 36. Rodriguez, W. J., Kim, H. W., Arrobio, J. O., et al (1977) Clinical features of gastroenteritis associated with human reovirus-like agents in infants and young children. J. Pediatr. 91, 188–193. 37. Isaacs, D., Day, D., and Crook, S. (1986) Childhood gastroenteritis: a population study. Brit. Med. J. 293, 545–546. 38. Engleberg, N. C., Holburt, E. N., Barrett, T. J., et al. (1982) Epidemiology of diarrhea due to rotavirus on an Indian reservation: risk factors in the home environment. J. Infect. Dis. 145, 894–898. 39. Koopman J. S., Turkisk, V. J., Monto, A. S., Thompson, F. E., and Isaacson, R. E. (1984) Milk fat and gastrointestinal illness. Am. J. Publ. Health 74, 1371–1373. 40. Koopman, J. S., Turkish, V. J., and Monto, A. S. (1985) Infant formulas and gastrointestinal illness. Am. J. Publ. Health 75, 477–480. 41. Matson, D. O. (1994) Viral gastroenteritis in day-care settings: epidemiology and new developments. Pediatrics 94, 999–1001. 42. O’Ryan, M. L., Matson, D. O., Estes, M. K., Bartlett, A. V., and Pickering, L. K. (1990) Molecular epidemiology of rotavirus in children attending day care centers in Houston. J. Infect. Dis. 162, 810–816. 43. Bartlett, A. V., III, Reves, R. R., and Pickering, L. K. (1988) Rotavirus in infanttoddler day care centers: epidemiology relevant to disease control strategies. J. Pediatr. 113, 435–441.

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44. Pacini, D. L., Brady, M. T., Budde, C. T., Connell, M. J., Hamparian, V. V., and Hughes, J. H. (1987)Nosocomial rotaviral diarrhea: pattern of spread on wards in a children’s hospital. J. Med. Virol. 23, 359–366. 45. Gaggero, A., Avendano, L. F., Fernandez, J., and Spencer, E. (1992) Nosocomial transmission of rotavirus from patients admitted with diarrhea. J. Clin. Microbiol. 30, 3294–3297. 46. Ford-Jones, E. L., Mindorff, C. M., Gold, R., and Petric, M. (1990) The incidence of viral-associated diarrhea after admission to a pediatric hospital. Am. J. Epidemiol. 131, 711–718. 47. Berner, R., Schumacher, R. F., Harmeister, S., and Forster, J. (1999) Occurrence and impact of community-acquired and nosocomial rotavirus infections: a hospital-based study over 10 year. Acta Paediatr. Scand. 88(Suppl. 426), 48–52. 48. Raad, I. I., Sherertz, R. J., Russell, B. A., and Reuman, P. D. (1990) Uncontrolled nosocomial rotavirus transmission during a community outbreak. Am. J. Infect. Contr. 18, 24–28. 49. Abbas, A. M. and Denton, M. D. (1987) An outbreak of rotavirus infection in a geriatric hospital. J. Hosp. Infect. 9, 76–80. 50. Lewis, D. C., Lightfoot, N. F., Cubitt, W. D., and Wilson, S. A. (1989) Outbreaks of astrovirus type 1 and rotavirus gastroenteritis in a geriatric in-patient population. J. Hosp. Infect. 14, 9–14. 51. Oishi, I., Maeda, A., Kurimura, T., and Kimura, M. (1979) Epidemiological and virological studies on outbreaks of acute gastroenteritis associated with rotavirus in primary schools in Osaka. Biken J. 22, 61–69. 52. Cook, S. M., Glass, R. I., Lebaron, C. W., and Ho, M. S. (1990) Global seasonality of rotavirus infections. Bull. WHO 68, 171–177. 53. Haffejee, I. E. (1995) The epidemiology of rotavirus infections: a global perspective. J. Pediatr. Gastroenterol. Nutr. 20, 275–286. 54. Brandt, C. D., Kim, H. W., Rodriguez, W. J., et al. (1983) Pediatric viral gastroenteritis during eight year of study. J. Clin. Microbiol. 18, 71–78. 55. Torok, T. J., Kilgore, P. E., Clarke, M. J., Holman, R. C., Bresee, J. S., and Glass, R. I. (1997) Visualizing geographic and temporal trends in rotavirus activity in the United States, 1991 to 1996. Pediatr. Infect. Dis. J. 16, 941–946. 56. Flahault, A., Garnerin, P., Chauvin, P., et al. (1995) Sentinel traces of an epidemic of acute gastroenteritis in France. Lancet 346, 162–163. 57. Borgdorff, M., Koopmans, M., Simone, E., Goosen, M., and Sprenger, M. (1995) Surveillance of gastroenteritis. Lancet 346, 842–843. 58. Koopmans, M. and Brown, D. (1999) The seasonality and diversity of Group A rotaviruses in Europe. Acta Paediatr. Scand. 88(Suppl 426), 14–19. 59. Kapikian, A. Z., Wyatt, R. G., Levine, M. M., et al. (1983) Studies in volunteers with human rotaviruses. Dev. Biol. Standard 53, 209–218. 60. Ramachandran, M., Das, B. K., Vij, A., et al. (1996) Unusual diversity of human rotavirus G and P genotypes in India. J. Clin. Microbiol. 34, 436–439. 61. Nakagomi, O. and Nakagomi, T. (1993) Interspecies transmission of rotaviruses studied from the perspective of genogroup. Microbiol. Immunol. 37, 337–348.

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112. Brüssow, H., Werchau, H., Lerner, L., et al. (1998) Seroconversion patterns to four human rotavirus serotypes in hospitalized infants with acute rotavirus gastroenteritis. J. Infect. Dis. 158, 588–595. 113. Brüssow, H., Clark, H. F., and Sidoti, J. (1991) Prevalence of serum neutralizing antibody to serotype 9 rotavirus WI61 in children from South America and central Europe. J. Clin. Microbiol. 29, 208–211. 114. Ruuska, T. and Vesikari, T. (1990) Rotavirus disease in Finnish children: use of numerical scores for clinical severity of diarrhoeal episodes. Scand. J. Infect. Dis. 22, 259–267. 115. Farrington, C. P. and Beale, A. D. (1993) Computer-aided detection of temporal clusters of organisms reported to the Communicable Disease Surveillance Centre. Commun. Dis. Rep. 3, R78–R82. 116. Klaucke, D. N., Buehler, J. W., Thacker, S. B., et al. (1988) Guidelines for evaluating surveillance systems. MMWR 37, 1–18. 117. Hook, E. B. and Regal, R. R. (1995) Capture-recapture methods in epidemiology: methods and limitations. Epidemiol. Rev. 17, 243–264. 118. Hardy, A. M., Lairson, M., and Morrow, A. L. (1994) Costs associated with gastrointestinal illness among children attending daycare centres in Houston, Texas. Pediatrics 94, 1091–1093. 119. Takala, A. K., Koskenniemi, E., Joensuu, J., Makela, M., and Vesikari, T. (1998) Economic evaluation of rotavirus vaccinations in Finland: randomized, doubleblind, placebo-controlled trial of tetravalent rhesus rotavirus vaccine. Clin. Infect. Dis. 27, 272–282. 120. Tucker, A. W., Haddix, A. C., Bresee, J. S., Holman, R. C., Parashar, U. D., and Glass, R. I. (1998) Cost-effectiveness analysis of a rotavirus immunization program for the United States. JAMA 279, 1371–1376. 121. Lobet, M. P., Stroobant, A., and Mertens, R. (1987) Tool for the validation of the network of sentinel general practitioners in the Belgian health care system. Internat. J. Epidemiol. 16, 612–618. 122. Noël, J. S., Parker, S. P., Choules,W. D. (1994) Impact of rotavirus infection on a paediatric hospital in the east end of London. J. Clin. Path. 47, 67–70. 123. Glass, R. I., Kilgore, P. E., Holman, R. C,. et al. (1996) The epidemiology of rotavirus diarrhea in the United States: surveillance and estimates of disease burden. J. Infect. Dis. 174(Suppl. 1), S5–S11. 124. Kilgore, P. E., Holman, R. C., Clarke, M. J., and Glass, R. I. (1995) Trends of diarrheal disease—associated mortality in US children, 1968 through 1991. JAMA 274, 1143–1148. 125. Gay, N. J. (1996) A model of long-term decline in the transmissibility of an infectious disease: implications for the incidence of hepatitis A. Internat. J. Epidemiol. 25, 854–861. 126. Orenstein, W. A., Bernier, R. H., and Hinman, A. R. (1988) Assessing vaccine efficacy in the field. Further observations. Epidemiol. Rev. 10, 212–241. 127. Gentsch, J. R., Woods, P. A., Ramachandran, M., Das, B. K., Leite, J. P., Alfieri, A. et al. (1996) Review of G and P typing results from a global collection of strains: implication for vaccine development. J. Infect. Dis. 174(Suppl. 1), S30–S36.

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12 Future Rotavirus Research Ulrich Desselberger and Mary K. Estes 1. Introduction Since the discovery of animal rotaviruses (RVs) in the 1960s (1,2), and of human rotaviruses (HRVs) 25 yr ago (3,4), much has been learned about virus structure, classification, evolution, replication, pathogenesis, and specific immune responses, and their correlation with protection and epidemiology. As pathogens that infect the gastrointestinal tract, these viruses continue to serve as useful models to understand mucosal virus–cell interactions. Recently, a RV vaccine to prevent disease in children has been licensed in the United States (US); this vaccine is expected to be applied worldwide. The chapters of this book have reviewed some of the research on RVs, with an emphasis on molecular methodological approaches. This summary chapter draws attention to some topics on which ongoing research is likely to produce significant results in the years to come, but also on which more research is needed. 2. RV Receptors and Mechanisms of Virus Entry into Cells For some viruses, the cellular receptors and co-receptors mediating viral absorption and cell entry have been characterized (for review, see refs. 5 and 6). For RVs of animal origin, N-acetyl-neuraminic (sialic) acid has been crudely implicated as a component of the cellular receptor (7), but infection of tissue culture cells by HRVs is resistant to neuraminidase treatment of cells, indicating that infection is initiated by a sialic acid-independent mechanism (8). An analysis of a larger number of viruses has indicated that the majority of animal and HRVs initiate infection by a sialic acid-independent mechanism (9). Cellular entry may occur by direct penetration through the plasma membrane (10,11), by receptor-mediated endocytosis, or by both mechanisms. RV entry is not affected by drugs that affect pH-dependent cell entry, but this From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ

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does not exclude the possibility that RVs may enter cells by a pH-independent mechanism that involves receptor-mediated endocytosis. Virus-like particles (VLPs) of defined protein composition, and with mutations in regions of the spike protein, viral protein 4 (VP4), involved in attachment to cells, can be used to study the entry of particles (see refs. 12–14, and Chapter 4). More than one cellular protein might be involved as a receptor. Cloning of the RV receptor gene(s) remains to be accomplished. 3. RV Structure and Replication Cryo-electron microscopy (cryo-EM) and computer image processing have been instrumental in providing three-dimensional (3-D) structurual characterization of these large and complex viruses. The work of various groups (those of Cohen, Estes, Greenberg, Patton, Prasad, Ramig, Rey, Yeager and others) has substantially helped to provide an insight into specific events of RV replication (reviewed in ref. 15; for basic information, see Chapter 1). However, a number of issues remain to be further investigated. 1. The RNA-dependent RNA polymerase function is provided by VP1, but other viral gene products (VP3 [a guanylyl transferase and methylase], VP2 [the main core protein], and possibly VP6) are also involved in transcription and replication functions (16). The study of interactions among the VP1–VP3 proteins has just begun, and their functional interactions remain to be sorted out (17). Other enzyme activities, as found in reoviruses (phosphohydrolase, poly[A] polymerase, NTPase), need to be assigned to specific genes. 2. The study of transcription and replication of RV RNA has been greatly facilitated by finding an electrophoretic system that allows separation of plus and minus strands of RV RNA (18), and by an in vitro replication system (19), but the regulation of these processes is still poorly understood. 3. A structural basis for understanding the mechanisms that regulate the complex process of RV transcription is beginning to be established by the use of cryo-EM and image processing, as well as by X-ray crystallography. The large size of RVs has made crystallization of these particles difficult, but, instead, these particles have been useful in developing the methods of cryo-EM and image processing (20–23; see Chapter 2). Recent studies have shown that at least 25% of the viral genomic RNA is ordered inside particles, and viral transcripts emerge from channels at the fivefold axes, where the enzymatic complexes containing VP1 and VP3 are localized (24,25; see Chapter 2). An atomic resolution structure of the middle-shell protein VP6, in combination with cryo-EM reconstructions, is beginning to explain how the proteins in the different layers interact (26). Future high-resolution structural studies are anticipated, involving both X-ray crystallography and cryo-EM of intact particles, subassemblies, VLPs and individual structural and nonstructural proteins: such studies will provide a detailed basis for understanding the molecular mechanisms of uncoating, viral transcription, morphogenesis, and particle antigenicity. Morphogenesis is of

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particular interest, because it involves the acquisition of a third protein shell on particles that mature by budding through the intracellular endoplasmic reticulum membrane, a process that is unique to RVs. The two viral glycoproteins involved in this process (VP7 and the nonstructural protein 4 [NSP4]) are unique in themselves, and understanding their functions may unravel new mechanisms of glycoprotein trafficking and interactions with cellular proteins. Confocal microscopy of virus-infected cells should also provide additional insight into these processes. 4. The RV genome codes for five NSPs, NSP1–NSP5, which are variously involved in replication and morphogenesis (NSP1–3, 5), but also act as an intracellular receptor (NSP4) for double-layered particles. All the NSPs except NSP4 interact with nucleic acid. NSP4 has recently been characterized as a viral enterotoxin affecting intracellular Ca2+ levels and leading to chloride secretion (27,28). Other studies suggest that NSP4 alters plasma membrane permeability and is cytotoxic (29). The effects and necessity of the NSPs for viral replication remain to be studied in more detail. For example, NSP1 (VP5) seems to be nonessential for virus replication, at least in cultured cells (30), yet some studies have implicated this gene as a virulence factor in animals (31). RV NSP3 binds to the 3'-end of viral mRNA in infected cells, and functions in a manner similarily to cellular poly(A) binding protein (32–34). By interacting with the translation initiation factor eIF4GI, NSP3 evicts the poly(A) binding protein from eIF4F; furthermore, NSP3 apparently shuts off cellular protein synthesis, by a previously unrecognized mechanism (34). NSP5 is a kinase that autophosphorylates itself, interacts with NSP2, and is associated with viroplasms (35–38). The interaction of viral with cellular proteins (as shown for replication-facilitating proteins of influenza viruses [39–41]) is just beginning to be studied. 5. A distinctive feature of RV morphogenesis is that subviral particles assemble in cytoplasmic aggregates called viroplasms. In those subviral particles, there is a very strict packaging of one genome set per particle, because mature virions have been shown to have a particle infectivity ratio of close to 1 (42), and purified preparations of RV particles liberate equimolar amounts of the 11 segments. How the tight packaging mechanism of RVs is regulated remains to be elucidated. By contrast, influenza virus particles package between 12 and 15 segments, i.e., an excess of the eightsegment complement is necessary to make an infectious particle.

4. Virus–Cell Interactions and Viral Pathogenesis RVs replicate primarily in the differentiated enterocytes in the intestinal epithelium. Most studies of RV replication have been performed in monkey kidney cells (MA104 and BSC-1 cells), which have been useful for investigating many aspects of the virus replication cycle, but which do not fully represent the virus–cell interactions that occur in vivo. This idea is supported by recent studies, which have shown that RVs induce chemokines and signaling pathways in cells that mimic intestinal cells (HT29 and CaCO2 cells),

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and virus may not be as cytolytic in differentiated polarized cells as in MA104 cells (43–46). Detailed knowledge of the responses of intestinal cells to RV infection will be needed to fully understand viral pathogenesis. Therefore, dissection of the interaction between specific RV genes or their products and components of intestinal cells merits further research. The effects of signaling by RV-infected cells on adjacent cells in the intestinal mucosa may influence the outcome of an infection, as well as the immune response, yet this crosstalk is currently poorly characterized. 5. Reverse Genetics For a number of single-stranded RNA viruses of positive or negative polarity of the genomic RNA, reverse genetic systems have been reproducibly achieved by constructing infectious cDNA clones, which are active in the absence or presence of various supporter functions and/or helper viruses: poliovirus (47); retroviruses (48); influenza viruses (49); rabies virus (50); measles virus (51); respiratory syncytial virus (52); vesicular stomatitis virus (53); Sendai virus (54); bunyavirus (55); parainfluenza virus (56); paramyxovirus SV5 (57) and potentially Marbury virus (57a). These systems allow exact manipulation of RNA genomes at the cDNA level (by site-directed mutagenesis, deletion/insertion, rearrangement, and so on), and have led to a plethora of significant new knowledge relating to replication, biological characteristics, and pathogenesis of these viral genera and families (for reviews, see refs. 58 and 59). For double-stranded (ds) RNA viruses, such achievements have so far been restricted to the two-segmented birnaviruses (infectious bursal disease virus [60]; infectious pancreatic necrosis virus [61]), and to the trisegmented ϕ6 bacteriophages (62–64). Early reports of success with reoviruses (65) have been difficult to reproduce by others, for unknown reasons, although the system has been used to construct reovirus double temperature sensitive (ts) mutants (66). The very tight control of packaging the exact number of genome segments per virus particle seems to be a major obstacle for acceptance of a foreign gene. Infectiousness of cDNA and RNA of bacteriophage ϕ6 may be caused by the presence of very strong packaging signals (pac) on each gene (67). The work of Ramig’s group (68,89) in trying to understand what is needed for RV particles to be infectious has yielded interesting results. Infectious particles have been reconstituted in vitro from isolated cores and the separately purified capsid components, VP6, VP7, and VP4 (68,69), but attempts to reconstitute cores from its component proteins, VP1, VP2, and VP3, and viral RNA, which would, in turn, become infectious after addition of the inner and outer capsid components in vitro, have so far been unsuccessful. A working system may require cell lines expressing RV genes acting in trans, as was successfully used for other viruses (51,70). Once one, or several reverse genetics systems for

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RVs are successfully and reproducibly established, not only could many open questions of replication and pathogenesis be definitely answered, but RVs would also become very interesting as viral vectors, having been shown to possess the capacity of packaging up to 10% of their genome in rearranged forms, without apparent negative effects on replication or packaging(71). 6. Correlates of Protection The immune response to RV infection is complex: Humoral immunoglobulin (Ig)M, IgG, and IgA antibody (Ab) titers develop and can be measured in serum and feces (in the latter, mostly IgA and IgG) (72). Cell-mediated immune responses also arise (72); they can be specifically directed against various VPs (VP7, VP4, VP6, and others). Correlation with protection (72,73), often demonstrated by passive transfer experiments in animal models, has been found with VP7-specific IgG and IgA Ab (74,75); VP4-specific IgG and IgA Ab (74,75); VP6-specific IgA Ab (76,77); and VP1-, VP3-, VP4-, VP6and VP7-specific cytotoxic T-cells (CTLs) (CD8+) (78–84). Increasingly, mucosal IgA in the gut (copro-IgA) is recognized as the most significant correlate of protection following live virus infections (85–89). Fecal IgG is seen as a major correlate of protection following immunization with nonreplicating virus or VLP vaccines (90–93, see Chapter 10). However, it remains unclear which VP specificity is most important for eliciting protective immunity (94,95). Longitudinal studies of natural infections (96–98) seem to suggest a VP7 serotype specificity of protection after primary infection; however, in subsequent infections, cross-protection increasingly develops, which is not fully explained by VP7 serotype-specific Abs from previous infections. This cross-protection may be mediated by VP6-specific IgA (76,77), possibly via intracellular neutralization, as shown in cell culture systems for influenza virus-specific IgA Abs (99,100), or by local CTLs. VP4-specific Ab may be less important in vivo than VP7-specific Ab (101). Infections following two natural infections in childhood are normally asymptomatic (98). The contribution of avirulent strains to immunity remains to be elucidated. Some asymptomatic RV infections in human neonates by “nursery” strains were initially characterized by their having an unusual VP4 type (102). However, RV strains carrying the VP4 of the nursery strains have also been observed in older infants and children with diarrhea (103,104); this finding may result from maternal Abs protecting neonates and infants during their first 2–3 mo of life. Another question of great importance in the context of vaccination is why sterilizing immunity is induced by live virus infection in several animal models (90,93,105,106), but apparently not in children (98). A better understanding of the mechanisms that regulate protective mucosal immunity in general is still needed (92,107).

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RV infections in immunocompromised children are known to result in long-term shedding of virus, apparently caused by an inability to clear virus (42,108). RV has been reported to spread extraintestinally in such children, and to replicate in nonintestinal cells (109–112). Viruses excreted from immunocompromised children often display unusual genomic RNA patterns, because of genome rearrangements (42,108,113), suggesting a unique evolution of virus in the absence of immunologic control. Understanding of what regulates such infections may become increasingly important as live attenuated vaccine is administered in areas where HIV infection is endemic. In fact, the response of immunocompromised children to live attenuated vaccine may be abnormal; this issue needs to be evaluated. 7. Epidemiology Molecular epidemiological studies in various parts of the world have found co-circulation of RVs of different G/P constellations (for nomenclature, see ref. 16). In Europe, North America, and Australia, G1P1A[8], G2P1B[4], G3P1A[8], and G4P1A[8] strains are found in 95% of cases of children hospitalized with diarrhea (e.g., refs. 114–116), but, more recently, other serotypes have been increasingly found to circulate as the predominant strains in several parts of the world (117–122). Protection against non-G1–G4 serotypes may be achieved by cocktail vaccines containing G1–G4 viruses, but what extent remains uncertain (see Subheading 8.), and continuous surveillance of wild-type (WT) RV circulation during the implementation of vaccination programs is required (see Chapter 11). Some of the unusual virus strains isolated from children in several parts of the world possess genes that are commonly found in animal RVs (117–119). This suggests that interspecies transmission of virus may occur, but this has never been documented directly. Understanding whether epizootic infections occur is an important question that needs to be answered, to begin to appreciate what is the driving force behind RV diversification and evolution. During 1997–1998, sporadic cases of human infections with group B RV have been detected in Calcutta (123). Group B RVs caused large outbreaks of diarrhea among children and adults in China during the 1980s (124), but were never detected as cause of human infections outside China, although these viruses are widespread in mammalian hosts (sheep, goats, rats, and so on). The origin of the human group B RV isolates have never been directly elucidated, although serological data pointed at animals as a possible source. The Calcutta isolates provide a new chance to make progress in this area. 8. RV Vaccines A tetravalent, rhesus RV-based human reassortant vaccine has now been tested on over 6000 children in developed and semideveloped countries

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(125–128), and has been found to be effective in preventing 80% of severe RV disease (severe diarrhea and dehydration requiring hospitalization), but protection from infection is only moderate (between 40 and 57%). This vaccine was licensed by the Food and Drug Administration in the United States in August 1998, and detailed recommendations for the use of the vaccine in routine immunization of infants in the United States have been made (128a). Licensure for Europe is pending. During the postlicensure period, vaccine manufacturers are required to report any adverse event possibly attributable to the vaccine to the Vaccine Adverse Event Reporting System (VAERS). In the period of September 1, 1998, to July 7, 1999, during which time approximately 1.5 million doses of the vaccine were administered in the United States, gut intussusception (a bowel obstruction in which one segment of bowel becomes enfolded within another segment) were observed in 15 infants, of whom 13 developed the condition after the first dose of the 3-dose RRV-TV vaccine series, and 12 developed the symptoms within 1 wk of any dose (129b, with further details). Although the number of reported cases was within the expected range by chance during the week following receipt of any dose, the well-known incompleteness of reporting through VAERS led the US Centers for Disease Control (CDC) to recommend postponing administration of RRV-TV to infants scheduled to receive the vaccine between July and November, 1999. By this time it is hoped that a more complete analysis of the data will be available to decide if vaccination should continue. This recent setback has refocused attention on other approaches of vaccine development that remain to be explored more widely. Parental application of the virus, either naturally derived (90) or as VLPs constructed from baculovirus recombinant-expressed RV proteins (12,91–93,129), is being studied. Other areas of exploration relate to the use of adjuvants like QS-21 (90,92), microencapsidation (130) to increase immunogenicity, and the use of DNA-based vaccines (77,131,132). Further development of other oral vaccines based on live attenuated viruses, e.g., the bovine WC3 strain (133), or a neonatal naturally attenuated (“nursery”) strain (134) will much depend on the outcome of the VAERS investigation relating to the RRV-TV vaccine. Expression of RV proteins in plants (potatoes, bananas) from vectors downstream of plant-specific promoters is also under investigation as a procedure to obtain potential vaccine candidates. Such “edible vaccines” have already been shown to express viral and other microbial proteins that were antigenic, and, in some cases, immunogenic (e.g., rabies virus GP expressed in tomatoes [135]; human HBsAg expressed in tobacco [136,137]; Norwalk virus capsid protein expressed in tobacco and potatoes [138]; foot-and-mouth disease virus VP1 expressed in Arabidopsis [139]; Escherichia coli heat-labile enterotoxin expressed in tobacco and potatoes [140–142]; for review, see ref. 143). Combinations of live attenuated vaccines and VLPs may be especially useful for countries where interference of live

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virus replication by other enteric pathogens or maternal Ab is discovered to be a problem. Such combinations would follow early work with combined live and inactivated poliovirus vaccines given in developing countries (144). 9. Population Dynamics of HRV Infections and Their Change by Widespread Vaccination It has been shown that WT RVs of different G- and P-types and different G/P-type combinations co-circulate, and that the relative incidence of different strains occurring simultaneously at different geographical sites, or subsequently at the same site, varies in an unpredictable way. Mathematical modeling of the epidemiology of virus infections has been successfully carried out in a number of cases, e.g., for measles virus (145). For RVs, the transmission dynamics in populations is difficult to describe, and will require knowledge of the following parameters: annual human birthrate in the area of study; duration of the presence of maternal Abs after birth; age-related susceptibility; age-related incidence of RV infection, and of prevalence of specific Abs, specified by G- and P-type; transmission effectiveness; reduction of susceptibility in nonprimary infections; and animal reservoirs. The co-circulation of different RV types, possible superinfections, and the occurrence of asymptomatic infections will have to be taken into account. Thus, mathematical modeling of the transmission dynamics will be very complex, as will be modeling of vaccine interventions. For RV infections in which superinfections are possible, and protection may only be partially crossprotective, mathematical modeling, as outlined by McLean (146,147), should be applied. This involves application of a number of equations developed to describe infections/vaccinations, after which total immunity/crossimmunity, or partial immunity allowing superinfection, is the outcome. The models will allow questions to be addressed about whether it is possible to eliminate an infection, what proportions of hosts must be vaccinated to achieve this, and at what intervals hosts should be revaccinated (148). The models will also allow predictions of the emergence of superinfecting or vaccine-resistant strains and their spread in the population (147,149,150). Other key questions relate to whether new group A serotypes will arise, and whether infections with non-group A RVs will become more widespread, if group A RV disease is controlled by the vaccine. 10. Antivirals Treatment of RV infections is primarily based on oral or parenteral rehydration (e.g., see ref. 151). Passively administered human anti-RV immunoglobulins have been found to be useful (152–155). Such treatments may be improved by including high-titer Ab to the RV enterotoxin NSP4, but that remains uncertain.

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Antiviral compounds have been found to interfere in vitro at various stages of the replication cycle: trypsin inhibitors (153,156) to reduce viral infectivity; N-acetyl neuraminic acid (NANA)-based inhibitors (157,158) or milk and other proteins (159,160), to prevent viral adsorption; interferon and other cytokines inhibiting intracellular replication and translation (161); adenosine analogs, probably preventing the formation of viral mRNA (162); and Brefeldin inhibiting viral maturation in the endoplasmic reticulum (163). Only trypsin inhibitors and NANA-based inhibitors have been found to have an antidiarrheic effect in vivo (156,164); none of these agents has so far been applied in clinical trials. Acknowledgment The authors thank Lynne Bastow for typing of the manuscript. References 1. Adam, W. K. and Kraft, M. M. (1963) Epizootic diarrhoea in infant mice: identification of the etiologic agent. Science 141, 359–360. 2. Mebus, C. A., Underdahl, N. R., Rhodes, M. B. and Twiehaus, M. (1969) Calf diarrhoea (scours): Reproduced with a virus from a field outbreak. Univ. Nebraska Res. Bull. 233, 1–16 3. Bishop, R. F., Davidson, G. P., Holmes, I. H., and Ruck, B. J. (1973) Virus particles in epithelial cells of duodenal mucosa from children with viral gastroenteritis. Lancet 2, 1281–1283. 4. Flewett, T. H., Bryden, A. S., and Davies, H. (1973) Virus particles in gastroenteritis. Lancet 2, 1497. 5. Wimmer, E. (ed.) (1994) Cellular Receptors for Animal Viruses. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. 6. Tyler, K. L. and Fields, B. N. (1996) Pathogenesis of viral infections, in Fields Virology, 3rd ed., (Fields, B. N. and Knipe, D. M., et al., eds.), Lippincott Raven, Philadelphia, pp. 173–218. 7. Bastardo, J. W. and Holmes, I. H. (1980) Attachment of SA-11 rotavirus to erythrocyte receptors. Infect. Immun. 29, 1134–1140. 8. Fukudome, K., Yoshie, O., and Konno, T. (1989) Comparison of human, simian and bovine rotaviruses for the requirement of sialic acid in hemagglutination and cell absorption. Virology 172, 196–205. 9. Ciarlet, M. and Estes, M. K. (1999) Human and most animal rotavirus strains do not require the presence of sialic acid on the cell surface for efficient infectivity. J. Gen. Virol. 80, 1943–1948. 10. Kaljot, K. T., Shaw, R. D., Rubin, D. H., and Greenberg, H. B. (1988) Infectious rotavirus enters cells by direct cell membrane penetration, not by endocytosis. J. Virol. 62, 1136–1144. 11. Ruiz, M. C., Alonso, T. S., Charpilienne, A., Vasseur, M., Michelangeli, F., Cohen, J., and Alvarado, F. (1994) Rotavirus interaction with isolated membrane vesicles. J. Virol. 68, 4009–4016.

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Index Adjuvants, 150, see also Vaccine Agarose gel electrophoresis, 205 Animal models, 5, 101, 120 gnotobiotic pig, 101, 133 challenge, 110 derivation, 108 immune response, 104 inoculation, 110 maintenance, 110 sample collection, 110 mouse, 101, 133, 159, see also Vaccine adult mouse model, 154 ascites fluid production, 139 cell depletion, 139 cell transfer, 137 enrichment of CD8+ cells, 141 husbandry, 167 immunization, 171 immunodeficient strains, 134 immunodepletion, 136, 139 isolation of intraepithelial lymphocytes, 139 isolation of lamina propria lymphocytes, 140 propagation of virus stocks in vivo, 167 sample collection, 172 viral clearance, 134 rabbit, 101, 159, 163, see also Vaccine husbandry, 167 immunization, 169 propagation of virus stocks in vivo, 167 sample collection, 170 Antibody secreting cells, 102, see also Correlates of protection Ascites fluid production, 139 Antivirals, 246 Baculovirus expression system, 41, 69, 149 insect cell growth, 73 plaque purification, 73, 166 virus growth, 73 virus titration, 73 VLP production, 40, 73, 150, 166 CLPs, 59, 150 DLPs, 150 TLPs, 150

Cell depletion, 139 Cell transfer, 137 Classification, 3, see also Genotyping; Serotyping; Diagnosis group, 3 G-type, 3 P-type, 4 subgroup, 3 CD8+ cell enrichment, 141 Clinical symptoms, 6 scoring for severity, 226 Cloning of PCR products, 194, 211 CLPs, 59, 150, see also Baculovirus expression systems; Virus assembly Colostrum/milk, 104, see also Immunity Complementation tests, 80 calculation of results, 94 significance of results, 97 Computer image processing, 10, see also Structure Correlates of protection, 6, 119, 243 antibody secreting cells, 102 copro-IgA, 7, 102, 243 Cross linking RNA-protein, 56 Cryo-electron microscopy, 9, 12, see also Structure Cytokine depletion, 139 3-D structure, 14, see also Structure DD50, 102 DEPC treatment of water, 198 Diagnosis, 6 electron microscopy189, 202 ELISA, 190, 203 immune electron microscopy, 190, 202 PAGE, 189, 204 passive particle agglutination test, 191 RT-PCR, 191 agarose gel electrophoresis, 205 group A, 204 group B, 206 group C, 206 nucleic acid extraction, 203 Disease burden, 227 Disease severity score, 226

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260 DLPs, 23, 41, 48, see also Baculovirus expression system; Virus assembly DNA sequencing, 193, 212 cloning of PCR products, 194, 211 PAGE, 195 purification of extension products, 212 purification of PCR products, 194, 209, 210 quantitation, 195 sequence analysis196, 213 Electron cryomicroscopy, 9, 12, see also Structure Electron microscopy, 189, 202 Electrophoresis, 59–61, 204–205, see also Diagnosis; PAGE ELISA, 126, 173, 190, 203, 206, see also Diagnosis; Immunity; Serotyping; Vaccine ELISPOT assay, 128, see also Immunity Endogenous peroxidase, 113 Enterotoxin, 102, 103, 241, see also Virus proteins; NSP4 Entry into cells, 67, 239, see also Virus entry Epidemiology, 7, 217, 244 age, 217 clinical reporting schemes, 221 death certificates, 224 dissemination of data, 227 laboratory reporting schemes, 222 medical care records, 224 sentinel physician reporting schemes, 222 special surveys, 225 diversity of virus strains, 220 geographical distribution, 219 morbidity, 218 population dynamics, 245 risk factors, 219 seroprevalence data, 224 sex, 217 surveillance, 221 analysis and interpretation of data, 225 disease burden, 226, 227 dissemination of data, 227 evolution of surveillance systems, 226 temporal distribution, 219 transmission, 220 vaccination strategy, 227 Expression systems, 41, 42, 69, 149, see also Baculovirus; Vaccinia Fluorescent focus assay, 174 Fluorescent focus neutralization assay, 175, see also Immunity

Index Formalin inactivation, 165 Gel electrophoresis, 59–61 Gel-shift assay, 53, see also RNA-binding activity Gene-protein assignment, 1, 2 Genome, 1, 22 Genotyping, 192 G-type multiplex RT-PCR, 209 G-type semi-nested RT-PCR, 192, 207 nucleic acid extraction, 203, see also Diagnosis P-type multiplex RT-PCR, 207 P-type semi-nested RT-PCR, 192, 208 random priming, 192, 204 Gnotobiotic pig, 101, see also Animal models Growth in vitro, 33, see also Virus propagation Guanylyltransferase, 1–3, 20, 41, 57, 104, 240, see also Virus proteins; VP3 Histopathology, 106, 111, 112, see Pathogenesis ID50, 102, see also Median infective dose Immune electron microscopy190, 202 Immune response, 6, 119, 121 Immunity, 104, 133 active, 105 ELISA, 105, 176 fecal antibody, 127, 176 serum antibody, 126, 176 ELISPOT assay, 105, 128 fluorescent focus neutralisation assay, 175 lymphoproliferative assay (LPA), 105 mononuclear cell isolation, 128 neutralising antibodies, 119 passive, 104 colostrum/milk, 104 maternal serum antibodies, 104 plaque reduction neutralisation assay, 122, 124, 175 Immunodeficiency mouse, 134, 135 man, 243 Immunodepletion studies, 134, 136, see also Animal models Immunofluorescence cell culture, 110 intestinal epithelial cells, 111 Inactivation, 165 formalin, 165 psoralen-UV, 165 Infectivity titration, 110, 167 Interference tests, 81 Lymphoproliferative assay, 105, see also Immunity

Index MA104 cells, 33, 69 sub-culture, 46, 87, 164 Maternal antibodies, 104, see also Immunity Median diarrheal dose, 102 Median infective dose, 102 Median shedding dose, 154 Mixed infections, 79 asynchronous co-infection, 86, 95 nonselective, 82 selective, 82 Mononuclear cell isolation, 128 Morphology, 3, 9, 190, 240, see also Structure Mouse, 101, 133, 159, see also Animal models mRNA, 49, see also Transcription Mutants, 79, 83, 89 Neutralising antibodies, 119, see also Immunity Nomenclature, 4 NSP4, 102, 103, 241, see also Enterotoxin Nucleic acid extraction, 203 Open cores, 36, 50 capping, 36 RNA binding activity, 57 RNA replication, 36 ORS, 6, see also Treatment PAGE, 42, 59, 60, 61, 189, 195, 204, see also Diagnosis; DNA sequencing Passive particle agglutination test, 191 Pathogenesis, 5, 101, 241 gnotobiotic pig, 103 histopathology, 106, 111, 112 immunohistochemistry, 111, 112 pathophysiology, 107 Phylogenetic analysis, 196, 213, see also DNA sequencing Plaque assay, 47, 90, 174 Plaque reduction neutralization assay, 124, see also Immunity Polymerase chain reaction, 192, 204-209 Population dynamics, 245 Prevention and control, 7, see also Vaccine Primers of PCR, 199-201 Protease inhibitors, 61 aprotinin, 48 leupeptin, 48 Proteins, 1, 2, see also Virus proteins Proteolytic cleavage, 33, 34, 96, 164 Psoralen-UV inactivation, 165 Rabbit, 101, see also Animal models

261 Reassortment, 79, 81, see also Mixed infections calculation of results, 91 kinetics, 83, 86 significance of results, 97 Recombination, 79 pseudorevertants, 91 recombinants, 91 recombination tests, 80 revertants, 91 Replicase assay, 50 Replication, 4, 33, 36, 240 RNA binding activity, 37–40, 56, 57, see also Open cores; Virus proteins gel-shift assay, 37, 53 RNA-capture assay, 39 RNA capping, 36 RNA capture assay, 39 RNA dependant RNA polymerase, 1–3, 20, 34, 57, 240, see also Virus proteins; VP1 RNase-free distilled water, 198 RNA synthesis, 34 RNA transcription, 23, 35 Reverse genetics, 242 Reverse transcriptase, 192, 204, see also Genotyping, RT-PCR subentries Rotavirus, see specific entries α-Sarcin assay, 69, 72, see also Virus entry Sequence analysis, 195, 196 Sequencing, 193, 209, 212 Serotyping, 191 G-type ELISA, 191, 207 P-type ELISA, 191, 207 sub-group ELISA, 191, 206 Staining hematoxylin and eosin, 111, 112 immunofluorescence, 111 immunohistochemistry, 111, 112 silver, 191, 204 Statistics, 169 Structure, 3, 190, 240 3-D structure, 14 aqueous channels, 15 computer image processing, 10 electron cryomicroscopy, 9, 12, 240 genomic RNA, 22 inner layer, 19 intermediate layer, 17 internal organization, 20 mRNA exit pathway, 24 outer layer, 14 surface spikes, 16 transcription, 2

262 Structure (cont.), 3-D structure (cont.), VP1, 20 VP2, 24 VP3, 20 VP4, 16 VP6, 17 VP7, 14 X-ray crystallography, 13, 240 Supershifting assay, 57 Syncytia assay, 67, see also Virus entry T-cell response, 105, see also Immunity Temperature sensitive mutants, 79 TLPs, 41, 48, see also Virus assembly Transcription, 33–35 Treatment, 6 oral rehydration solution, 6 drugs, 6, 246 Trypsin, 33, 34, 96, 164 Vaccine, 7, 147, 221, 244 adjuvants, 148–151 parenteral, 150 mucosal, 151 delivery systems, 150, 152 edible vaccines, 245 evaluation, 157, 229 experimental design, 168 live attenuated vaccine, 7, 147, 156, 245 mouse model, 147, 154 antibody detection, 156, 173, 175, 176 collection of samples, 155, 172 immunization, 155, 171 virus detection, 155 rabbit model, 147, 153 antibody detection, 156, 171, 175, 176 collection of samples, 154, 170 immunization1, 154, 169 virus detection, 155 strategy, 227 virus shedding, 173 antigen ELISA, 173 fluorescent focus assay, 174 plaque forming assay, 174 VLPs, 148, 245, see also Virus assembly

Index Vaccinia virus expression system, 42 expression of VP2 CLPs, 58 purification of VP2 CLPs, 59 vTF7-3 strain, 42 Vector pFASTBAC, 74 Virion proteins, 1, see also Virus proteins Virus assembly, 40 core-like particles, 41, 48, 58, 59 double-layered particles, 23, 41, 48 triple-layered particles, 41, 48 Virus-cell interactions, 241 Virus core, 34, 48 Virus detection, 173, 189, 190 Virus entry, 67, 239 α-sarcin assay, 69, 72 syncytia assay, 68, 71 Virus infectivity, 110, 167, see also Infectivity Virus propagation, 33, 46–47, 164, 167 35S-labelled virus, 47 Virus proteins, 1 NSP1, 5, 37, 57, 241 NSP2, 4, 5, 37, 57, 241 NSP3, 27, 241 NSP4, 4, 5, 102, 103, 241 NSP5, 4, 241 RNA binding activity, 37 VP1, 1–3, 20, 34, 57, 240 VP2, 1-3, 19, 41, 150, 240 VP3, 1-3, 20, 41, 57, 104, 240 VP4, 1-3, 14, 16, 34, 41, 48, 67, 70, 102, 150, 191, 220 VP5*, 1-3 VP6, 1-3, 17, 19, 41, 150, 190, 240 VP7, 1-3, 14, 41, 67, 70, 102, 150, 191, 220, 241 VP8*, 1-3 Virus purification, 34, 48-49, 164 Virus receptors, 239 Virus replication, 4, 33, 36, 240, see also Replication Virus structure, 3, 9, 190, 240, see also Structure VLP, 40, 73, 148, 150, 166, 245, see also Baculovirus expression system; Virus assembly; Vaccine X-ray crystallography, 13, 240, see also Structure

M E T H O D S I N M O L E C U L A R M E D I C I N E TM Series Editor: John M. Walker

Rotaviruses Methods and Protocols Edited by

James Gray and Ulrich Desselberger Clinical Microbiology and Public Health Laboratory Addenbrooke’s Hospital, Cambridge, UK

In Rotaviruses: Methods and Protocols, James Gray and Ulrich Desselberger have assembled a comprehensive collection of established and cutting-edge methods for studying and illuminating the structure, molecular biology, pathogenesis, epidemiology, and prevention in animal models of infection with rotaviruses, an important cause of infant morbidity and mortality. Presented by experts in the fields of animal and human rotavirus infections and rotavirus vaccine research, these readily reproducible methods detail molecular and other modern techniques, and include relevant background information and various notes to ensure reproducible and robust results. Authoritative and up-to-date, Rotaviruses: Methods and Protocols offers researchers today’s benchmark compendium of experimental methods for the investigation of this medically significant virus. Features • Comprehensive and up-to-date collection of established and experimental methods • Step-by-step instructions by hands-on experts of each method • Animal models for understanding pathogenesis, immune response, and correlates of protection

• Techniques applicable to the study of other viruses and virus–host systems • Electron cryomicroscopy and complex computer imaging techniques • Comprehensive guide for detection and typing methods

Contents Rotaviruses: Basic Facts. Electron Cryomicroscopy and Computer Image Processing Techniques: Use in Structure–Function Studies of Rotavirus. Virus Replication. Rotavirus Entry into Tissue Culture Cells. Mixed Infections with Rotaviruses: Protocols for Reassortment, Complementation, and Other Assays. Pathogenesis and Animal Models. Immunologic Methods and Correlates of Protection.

In Vivo Study of Immunity to Rotaviruses: Selected Methods in Mice. Evaluation of Rotavirus Vaccines in Small Animal Models. Methods of Rotavirus Detection, Sero- and Genotyping, Sequencing, and Phylogenetic Analysis. Epidemiology of Group A Rotaviruses: Surveillance and Burden of Disease Studies. Future Rotavirus Research. Index.

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Methods in Molecular Medicine™ Rotaviruses: Methods and Protocols ISBN: 0-89603-736-3

9 780896 037366