Rotaviruses: Methods and Protocols

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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)


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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 cryoelectron 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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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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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 (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 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 (

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