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
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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
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Rotaviruses Methods and Protocols Edited by
James Gray Ulrich Desselberger Clinical Microbiology and Public Health Laboratory Addenbrooke's Hospital Cambridge, UK
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© 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:
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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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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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B. V. VENKATARAM PRASAD • Verna and Marrs Department of Biochemistry and W. M. Keck Center for Computational Biology, Baylor College of Medicine, Houston, TX ROBERT F. RAMIG • Division of Molecular Virology, Baylor College of Medicine, Houston TX MARY RAMSAY • Public Health Laboratory Service, Communicable Disease Surveillance Centre, London, UK LINDA J. SAIF • Food and Animal Health Research Program, Department of Veterinary Preventive Medicine, Ohio Agricultural Research and Development Center, The Ohio State University, Wooster, OH ZENOBIA TARAPOREWALA • Laboratory of Infectious Diseases, National Institutes of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD LUCY A. WARD • Department of Veterinary Preventive Medicine, Ohio Agricultural Research and Development Center, The Ohio State University, Wooster, OH
Rotaviruses: Basic Facts
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1 Rotaviruses: Basic Facts Ulrich Desselberger 1. Introduction Rotaviruses (RVs) are the chief etiologic agent of viral gastroenteritis in infants and young children, and in the young of a large variety of animal species. Since the discovery of RVs in man 25 yr ago, much has been learned about their genome and protein composition; their three-dimensional structure; their replication, pathogenesis and clinical pattern; the host’s immune response; and the epidemiology. Measures of individual treatment have recently been complemented by the licensure in the United States of a tetravalent (TV), live attenuated rhesus rotaviruses (RRV)-based, human reassortant vaccine which may to be universally applied. The brief introductory description mostly follows recent reviews (1–4) in which more special references can be found. 2. The RV Genome The genome of RVs consists of 11 segments of double-stranded RNA (dsRNA) with conserved 5' and 3' ends, ranging from 667 bp (segment 11) to 3302 bp (segment 1) in size (SA11 simian RV strain), and totaling 6120 kDa, or 18,555 bp. 3. Gene-Protein Assignment This is complete for several RV strains, and is shown for the SA11 strain in Table 1. With the exception of two genes (RNA 9 and 11), all genes are monocistronic, and the untranslated 5' and 3' regions are very small. 4. RV Proteins There are six structural viral proteins (VPs: termed VP1, VP2, VP3, VP4, VP6, and VP7) and five nonstructural proteins (termed NSP1–NSP5). The From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ
1
2
Size (bp)
1 2 3 4
3302 2690 2591 2362
VP1 VP2 VP3 VP4
5
1611
6 7a
1356 1104
8a
1059
9a
1062
10
751
11
667
Posttranslational modification
125.0 102.7 88.0 86.7
– Myristylation – Proteolytic cleavage (VP5* + VP8*)
VP5 (NS53) VP6 VP8 (NS35) VP9 (NS34) VP7 (1) VP7 (2)
58.6
–
44.8 36.7
Myristylation –
34.6
–
VP10 (NS29) VP11 (NS26)
20.3
Cleavage of signal sequence; glycosylation Glycosylation
21.7
Phosphorylation
37.4 33.9
Location and function Inner core protein; RNA polymerase Inner core protein; RNA binding; leucine zipper Inner core protein; guanylyl transferase; methalyse Surface protein (dimer) Hemagglutinin Neutralization antigen (serotype specific) Fusogenic protein Virulence Pathogenicity Nonstructural (?) Zinc fingers; assembly Inner capsid protein (trimer); group and subgroup antigen Nonstructural; RNA replication? Nonstructural; RNA binding Surface glycoprotein Neutralization antigen (serotype specific); Ca2+ binding site? Nonstructural; intracellular receptor; morphogenesis; enterotoxin Nonstructural
aGene protein coding assignment for SA11 RV strain; assignment different in other strains. Adapted with permission from ref. 4.
Desselberger
RNA segment
Protein product Deduced mol Designation wt (kDa)
2
Table 1 Gene-Protein Assignments, Protein Location, and Function of Group A RVs (SA11 RV/strain)
Rotaviruses: Basic Facts
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functions of all proteins are summarized in Table 1 and reviewed below (as far as known): 1. 2. 3. 4.
VP2 provides a scaffolding function of the inner core (see Subheading 5.). VP6 carries group- and subgroup-specifying determinants. VP7 and VP4 both carry neutralizable, and thus type-specifying, epitopes. VP7 is a glycoprotein of 326 amino acids (aa) (a second in-frame initiation codon lies 30 codons downstream), with nine variable regions contributing to type specificity at varying degrees (see Subheading 6.). 5. VP4 is a nonglycosylated protein of 776 aa, and has a large number of functions: It is the viral hemagglutinin; it is posttranslationally cleaved (in aa positions 241 and 247, the latter being the preferred cleavage site) into the larger VP5* and the smaller VP8* subunits, and cleavage of VP4 enhances infectivity; it is the determinant for protease-enhanced plaque formation and growth restriction; it interacts with the cellular receptor; it has a fusion domain (of still unclear function); it is a virulence determinant. 6. The NSPs have various functions in replication (see Subheading 7.). NSP4 has been found to act as a viral enterotoxin.
5. RV Particle Structure The particle is of icosahedral symmetry, measures 75 nm in diameter, and consists of three layers: 1. The core layer, formed by VP2, and containing the viral genome and the proteins VP1 (the RNA-dependent RNA polymerase) and VP3 (a guanylyltransferase and methylase) (these proteins may have other enzyme functions); 2. The inner capsid (intermediate layer), consisting of 260 VP6 trimers, which are interrupted by 132 aqueous channels of three different kinds in relation to the capsid’s symmetry; 3. The outer capsid (third layer), consisting of 260 VP7 trimers and 60 spike-like VP4 dimers. VP4 interacts with VP7 and VP6.
6. Virus Classification According to VP6 reactivities, there are at least seven different groups (groups A–E are confirmed by complete crossreactivities; groups F and G are likely to be new groups). Within group A, subgroups I, II, I + II, and non-I, non-II are distinguished (according to reactivities of VP6 with two monoclonal antibodies). Because there are two neutralizable outer capsid proteins (VP4 and VP7), a dual classification system has emerged (1), similar to the dual classification established for influenzaviruses (distinguishing different hemagglutinins and neuraminidases): 1. There are at least 14 different VP7-specific types, termed G-types (derived from glycoprotein);
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Desselberger 2. There are at least 19 different VP4-specific types, termed P-types (derived from protease-sensitive protein).
All G- and P-types can be unambiguously distinguished by sequencing of the relevant genes (genotypes). All G-genotypes have been characterized as serotypes; however, this is not the case for all P-genotypes. Therefore, the following nomenclature has been agreed upon: Each virus has a P-type, indicated by an order number for the serotype and by an order number in square brackets for the genotype, and a G-serotype, indicated by an order number (coinciding with genotype number). Thus, the human Wa strain is defined as P1A[8]G1 (P-serotype 1A, P-genotype 8; G-sero/genotype 1); the equine RV L338 is P[18]G13 (P-genotype 18, P-serotype not determined; G-sero/genotype 13); and so on. Because VP4 and VP7 are coded for by different RNA segments (RNA4, and RNA7–9, respectively), various combinations of G- and P-types can be observed in vivo and in vitro, both in man and in animals (4). 7. RV Replication The major features of RV replication are shown in Fig. 1, and can be described as a sequence of the following steps (all replication takes place in the cytoplasm): 1. Adsorption to cellular receptor(s) and receptor-mediated endocytosis, or direct penetration. 2. Messenger RNA (mRNA) production in the cytoplasm from single-shelled (bilayered) subviral particles. 3. mRNA translation to synthesize six structural and five nonstructural proteins (see Subheading 4.). 4. Assembly of single-shelled particles containing VP2, VP1, VP3, and VP6, and a full complement of 11 single-stranded RNAs (ssRNAs); involvement of NSP2 and NSP5; formation of dsRNA (replication) within particle precursors (no free dsRNA or negative ssRNA in cytoplasm); formation of aggregates of bilayered particles (pseudocrystals termed “viroplasm”); 5. Particle maturation to double-shelled (triple-layered) particles: a. Glycosylation of VP7 in rough endoplasmic reticulum (RER), NSP4 acting as an intracelluar receptor for bilayered particles. b. Transiently enveloped particles in RER containing VP4 and VP7. c. Envelope removal. 6. Liberation of double-shelled (triple-layered) infectious particles (virions) by cell lysis.
For further details, see ref. 1; for questions to be investigated further, see Chapter 12. Double infection of cells with two different RV strains leads to simultaneous replication of the genes and synthesis of proteins of both viruses, and, at
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Fig. 1. Major features of the RV replication cycle. Adapted with permission from ref. 1. the stage of assembly, the formation of reassortants of various gene segment combinations; those also include the emergence of different G/P reassortants (see Subheading 6.). 8. Pathogenesis and Animal Models RVs infect the apical cells of the villi of the small intestine, causing cell death and desquamation. At the zenith of the disease, up to 1011 virus particles/mL stool have been counted, concomitant with infection of all susceptible cells in a very short period. The necrosis of the apical villi reduces digestion, causing diarrhea because of primary maladsorption, and leads to villous atrophy (Fig. 2). This is compensated for by a reactive crypt cell hyperplasia accompanied by increased secretion, which also contributes to the diarrhea. Recovery is by replacement of villous epithelium by enteroblasts ascending from crypts. The disease process takes 5–7 d (5). The viral factors determining pathogenicity of RVs have been investigated in several animal models. The product of RNA segment 4, VP4 (Table 1), is likely to be a major determinant, but products of other structural genes (RNA3 coding for VP3; RNAs 8 or 9 coding VP7) and of some of the nonstructural genes (RNA5 coding for NSP1; RNA 8 coding for NSP2, and RNA 10 coding for NSP4) have also
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Fig. 2. Mid-small intestine of gnotobiotic calves. (A) Healthy control animal. (B) animal inoculated experimentally with bovine RV. The normally extended finger-like villi (A) have become stunted and misshapen (atrophy), and the enterocytes of the upper part of the villi are disarranged and swollen. Adapted with permission from ref. 5. been associated with pathogenicity (for review, see ref. 6). Host factors, such as age and host restriction of viral replication, are involved as well in determining pathogenicity, but are less clearly defined. 9. Clinical Symptoms and Treatment After a short incubation period of 24–48 h, the onset of illness is sudden, with watery diarrhea, vomiting, and rapid dehydration. Untreated RV infection is a major cause of infantile death in developing countries. It should be noted, though, that clinical symptoms after RV infection vary widely, and asymptomatic infections in neonates with so-called “nursery” strains have been described. Treatment is by oral or parenteral rehydration with oral rehydration solution (ORS) formulae, which have been approved by the World Health Organization for worldwide treatment in developing countries, and by some drugs (7). 10. Diagnosis Because of the high number of virus particles in feces during the acute illness, diagnosis is easy, using electron microscopy, passive particle agglutination tests, or enzyme-linked immunosorbent assays. 11. Immune Response and Correlates of Protection Acute RV infection is followed by a virus-specific, humoral immune response comprising immunoglobulin (Ig)M, IgG, and IgA antibodies, and by
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a cell-mediated immune response of RV-specific cytotoxic T-cells in the lamina propria of gut tissue. After prolonged research and discussions, it has become increasingly clear that RV-specific, local secretory IgA antibodies (copro-IgA) represent the best correlate of protection (8–10). 12. Epidemiology RVs are the main etiologic agents of serious diarrheal disease in infants and young children under 2 yr of age throughout the world. For developing countries, approx 125 million cases of RV infection occur annually in children under 5 yr of age, of which 18 million are moderately severe to severe; almost 900,000 children die annually from RV infections in these countries. For the USA it is estimated that RV infections cause an estimated 1 million cases of severe diarrhea and approx 150 deaths per annum. RVs are transmitted mostly by the fecal–oral route. A high degree of resistance to physical inactivation, the large number of virus particles shed, and the very low diarrhea dose 50% ensure that infection is also easily taken up from environmental sources, as demonstrated by tenacious nosocomial infections once a clinical ward has been contaminated. Animals infected by various RV types may act as a reservoir for human RV infections. The epidemiology of RVs in complex. Group A RVs are the major cause of human infections. Outbreaks with a strict seasonal winter pattern occur in temperate climates, in tropical regions infections are spread more evenly throughout the year. At any one time and site, there is cocirculation of RVs of different G- and P-types. Viruses of multiple different G/P-type combinations have been isolated. However, G1–G4 viruses represent over 95% of the human strains co-circulating worldwide, and G1 viruses approx 50%. The P/G combinations found are P1A[8]G1, P1B[4]G2, P1A[8]G3, and P1A[8]G4. Within one country, the relative incidence figures for the different types show regional differences, as well as changes over time, and changes in relative incidences of different types are unpredictable. 13. Prevention and Control Since August 1998, a TV, RRV based human reassortant vaccine has been licensed in the United States for universal use, and a decision of licensure for Europe is pending. The vaccine does not prevent infection to a significant degree (i.e., does not produce sterilizing immunity), but has been shown to prevent severe disease with an efficacy of 80%. The vaccine carries G1–G4 epitopes of human RV strains (G1, G2, and G4 on RRV monoreassortants, G3 on RRV). It remains to be seen to what extent the vaccine will produce heterotypic immunity, and whether, upon extensive use of the vaccine, new RV types emerge in man (for review, see refs. 11,12).
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References 1. Estes, M. K. (1996) Rotaviruses and their replication, in Fields Virology, 3rd ed., (Fields, B. N., Knipe, D. M., Howley, P. M., et al., eds.), Lippincott-Raven, Philadelphia, pp. 1625–1655. 2. Kapikian, A. Z. and Chanock, R. M. (1996) Rotaviruses, in Fields Virology, 3rd ed., (Fields, B. N., Knipe, D. M., Howley, P. M., et al., eds.), Lippincott-Raven, Philadelphia, pp. 1657–1708. 3. Iqbal, N. and Shaw, R. D. (1997) Rotaviruses. in Clinical Virology, (Richman, D. D., Whitley, R. J., and Hayden, R. G. eds.), Churchill Livingstone, New York-Edinburgh-London, pp. 765–785. 4. Desselberger, U. (1998) Reoviruses, in Topley and Wilson’s Microbiology and Microbial Infections. 9th ed. vol. 1: Virology, (Mahy, B. W. J. and Collier, L., eds.), E. Arnold, London-Sydney-Auckland, pp. 537–550. 5. Greenberg, H. B., Clark, H. F., and Offit, P. A. (1994) Rotavirus pathology and pathophysiology in Rotaviruses (Ramig, R. F., ed), Springer Verlag, Berlin-Heidelberg, pp. 256–283. 6. Burke, B. and Desselberger, U. (1996) Rotavirus pathogenicity. Virology 218, 299–305. 7. Desselberger, U. (1999) Rotavirus infection: guidelines for treatment and prevention. Drugs 58, 447–452. 8. Offit, P. A. (1994). Rotaviruses: immunological determinants of protection against infection and disease. Adv. Virus. Res. 44, 161–202. 9. Yuan, L. J., Ward, L. A., Rosen, B. I., To, T. L., and Saif, L. J. (1996) Systemic and intestinal antibody secreting cell responses and correlates of protective immunity to human rotaviruses in a gnotobiotic pig model of disease. J. Virol. 70, 3075–3083. 10. Moser, C. A., Cookinham, S., Coffin, S. E., Clark, H. F., and Offit, P. A. (1998) Relative importance of rotavirus-specific effector and memory B cells in protection against challenge. J. Virol. 72, 1108–1114. 11. Vesikari, T. (1997) Rotavirus vaccines against diarrhoeal disease. Lancet 350, 1538–1541. 12. Desselberger, U. (1998) Towards rotavirus vaccines. Rev. Med. Virol. 8, 43–52.
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2 Electron Cryomicroscopy and Computer Image Processing Techniques Use in Structure–Function Studies of Rotavirus B. V. Venkataram Prasad and Mary K. Estes 1. Introduction Rotavirus (RV), a double-stranded (ds)RNA virus in the family Reoviridae, is a complex, relatively large (diameter, including spikes = 1000 Å), nonenveloped icosahedral virus. Once RV was recognized as a major human pathogen, it was extensively studied using modern molecular genetic and biological techniques, as discussed elsewhere in this book. These studies provided basic information about gene-coding assignments, protein processing, genome expression and replication, viral morphogenesis, and pathogenesis (1). In addition, molecular epidemiological studies, coupled with the characterization of neutralizing monoclonal antibodies (MAbs) and sequencing of the genes that encode the neutralizing antigens, provided an understanding at the molecular level of the antigenic and genetic variability of the RVs. Medical relevance, intriguing structural complexity, and several unique strategies in the morphogenesis of RVs have provoked extensive structural studies on these viruses in recent years (2–10). A detailed architectural description of these complex viruses, including the topographical locations of the various structural proteins and their stoichiometric proportions, was obtained as the resolution of these techniques improved. Together with molecular biological studies, structural studies are permitting a dissection of the molecular mechanisms that underlie biological processes of the virus, such as cell entry, neutralization, transcription, gene expression, and virus assembly (8). This chapter reviews current knowledge of RV structure and the methods used in structural analysis. From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ
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2. Computer Image Processing of Electron Cryomicrographs Over the past four decades, X-ray crystallography has been the technique of choice to study three-dimensional (3-D) structures of biological macromolecules and macromolecular assemblies at atomic resolution. Structures of several animal viruses have been studied using this technique (11–12; see ref. 13 for a brief review of the technique). Until now, the largest structure that has been studied using this technique is simian vacuolating virus 40, ~500 Å in diameter (14). However, recently, the structure of bluetongue virus (BTV) cores, ~680 Å in diameter, a virus in another genus of the Reoviridae family, has been determined to near-atomic resolution, using X-ray crystallographic techniques (15). X-ray crystallographic techniques have also been used on individual viral proteins (VPs), such as the hemagglutinin and neuramindase proteins of influenza virus (12), several proteins of HIV (16,17) and, most recently, VP6 of RV (18). Attempts are being made to determine the atomic resolution structure of the entire RV and subassemblies of RV using X-ray crystallographic techniques. Prerequisite to successful structural analysis by X-ray crystallography is the ability to obtain crystals, of the specimen of interest, which diffract X-rays to atomic resolution. In the past decade, advances in electron cryomicroscopy (cryo-EM) and computer image processing techniques have provided another powerful tool for studying the 3-D structures of biological macromolecules and assemblies. This technique does not require the specimen to be in a crystalline state. Another advantage of this technique that it allows structural studies of the specimen not only in its native state, but also under various physiological conditions. For example, this technique has been used to study complexes of antibody-bound virus (3,19–22) and receptor-bound virus (23), transcribing virus (9), and specimens under varied chemical states, such as different pH and ionic strengths (24). All information obtained on structure–function relationships in RVs has come from using cryo-EM and computer image processing techniques. The following sections briefly discuss these techniques, and then describe how they have been useful in dissecting structure–function correlations in RVs.
2.1. Computer Image Processing Because of the large depth of focus of conventional electron microscopes, transmission electron micrographs, in effect, represent two-dimensional (2-D) projections of the specimen. Inference of the detailed 3-D structure by direct examination of electron micrographs is therefore often a difficult task. Over the past two decades, computerized procedures have been developed to reconstruct the 3-D structure of a specimen from such projections. These procedures offer an objective way of extracting 3-D structural information from electron
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micrographs. Similar procedures of reconstruction from projections are used in other contexts, such as diagnostic tomography. To determine the 3-D structure of any object, it is necessary to combine the information content from different views of that object. These different views are sometimes provided by the specimen lying in different orientations, as in the case of icosahedral viruses, or they may be obtained by tilting the specimen in the microscope. 3-D reconstruction from electron micrographs is based on what is known as the projection theorem, which states that the 2-D Fourier transform of a projection of a 3-D object is a central section, normal to the direction of the view, of the 3-D Fourier transform of the object. If all the different orientations of the specimen can be identified with respect to a common frame of reference, the 3-D Fourier transform of the specimen can be built from the 2-D Fourier transforms of different views. Fourier inversion of the 3-D transform thus obtained gives the 3-D structure of the specimen. For icosahedral viruses, Crowther et al. (25–27) have developed image-processing procedures to determine their 3-D structure from micrographs. The first step in the computer processing of electron micrographs is converting the image into digitized data by microdensitometry. Regions in the micrographs with a sufficient number of particles are digitized using a computer-controlled microdensitometer. The digitized regions are put into the computer, where each particle is windowed and centered inside a box. The orientation of each particle is determined in a computer from its Fourier transform using the so-called common lines procedure (26). Orientations of enough particles are determined so that they evenly represent the asymmetric unit of an icosahedron. The number of particles required for a reconstruction depends on the size of the specimen and the resolution sought. Typically, to attain a resolution of 20 Å for RV, 200 particles with unique orientations are used. For resolution closer to 10 Å, the number of particles with unique orientations have to be in the thousands. Particle images with well-determined orientation parameters are then combined in Fourier space using cylindrical expansion methods, to obtain a 3-D Fourier transform (26). An inverse Fourier transform then gives the 3-D structure. Orientation parameters of the particles are refined, either with respect to one another, using the cross-common lines, or with respect to the projections obtained from the initial 3-D map, in an iterative fashion. The visualization and interpretation of the 3-D reconstruction are carried out using computer graphics software. These procedures have been modified and refined in the past few years by several laboratories (27–31). A simplified outline of the image processing scheme is shown in Fig. 1. A detailed description of every aspect of the processing protocol is beyond the scope of this chapter but can be found in ref. 29. Computations of difference maps between closely related
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Fig. 1. A brief schematic representation of the major steps involved in the 3-D reconstruction of the RV structure.
strains (e.g., wild-type and mutant [4]), or viruses in different states (6), allow additional structural information to be obtained.
2.2. Electron Cryomicroscopy In conventional transmission EM one normally uses metal-shadowing or negative-staining techniques to enhance image contrast. These preparative techniques may potentially alter the structure of the specimen, and sometimes may destroy fragile structural features because of chemical modification, dehydration, and desiccation. As a result, these preparative techniques are not suitable, particularly when the goal is to obtain 3-D structural information. However, for diagnostic purposes, these conventional techniques are adequate, and have proven very useful. In 1975, Taylor and Glaeser (32) introduced a method of embedding the specimen in a thin layer of ice, and imaging at low temperatures using a low electron dose. Since then, several laboratories have been involved in the improvement of this technique (for review, see ref. 33). Cryo-preparative techniques not only provide good contrast but also preserve the structural integrity of the specimen. One further advantage of structural analysis using electron images of ice-embedded specimens is the ability to retrieve the details of the internal structure of particles. This contrasts with
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structure determination from images of negatively stained specimens, in which it is possible to obtain only surface features. The image contrast in the absence of stains or metal decoration is caused by the scattering density difference between protein and ice. Cryo-EM involves three principal steps: quick-freezing of the specimen, transfer of the specimen to the microscope, and examination of the specimen in the microscope at low temperature. The first step is carried out using a mechanical guillotine-type plunging device (34). The specimen grid is held by a tweezer in the quick-freezing device. After the excess liquid on the grid is blotted using filter paper, the specimen grid is rapidly plunged into liquid ethane (cryogen) at its melting point. This rapid freezing produces a thin layer of vitreous or amorphous ice in which the specimen is embedded. In the second step, the frozen specimen is initially transferred to a cryospecimen holder maintained at liquid nitrogen temperature (–173°C) in a workstation. Then the cryo-specimen holder is quickly transferred to the electron microscope. The specimen is examined in the microscope at a temperature below –160°C, using low electron doses (typically 5–10 e – /Å 2 on the specimen). Several technical problems associated with this technique have been addressed during the past few years, which have made this a successful technique (for details, see refs. 33,35).
2.3. Recent Developments in Cryo-EM In recent years, there have been exciting developments in the high-resolution electron imaging of biological specimens: for example, the use of computer-controlled spot-scan imaging with medium-high-voltage electrons to reduce beam-induced motion images and to increase the efficiency of recording high resolution (7,36–39); the use of a field emission gun with intermediate-high-voltage electrons to increase the high-resolution image contrast by improving temporal and spatial coherence (40–42), and the use of energy-filtered electrons to remove the background intensity caused by inelastic scattering, and to increase the scattering signal from the specimen (43). Advances in computer image processing schemes have also been made, taking advantage of high-speed, high-performance computers and multiprocessor computers (44). The future looks promising for attaining 10 Å resolution and higher by cryo-EM, for icosahedral particles. The structure of the hepatitis B virus has already been determined to a resolution of ~8 Å, using cryo-EM techniques (41,42).
2.4. Combined Use of X-ray Crystallography and Cryo-EM Techniques An exciting trend in recent years is the marriage of X-ray crystallography and cryo-EM techniques (45). Several studies have shown that these two
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techniques are not mutually exclusive, but can effectively complement one another, particularly in studying large macromolecular assemblies. If the structures of individual components of an assembly are available at atomic resolution, this information can be fitted into the structure of the whole assembly determined by cryo-EM techniques, to enhance understanding of the molecular interactions. The complementarity of these two techniques offers a significant step forward in structural biology, as is shown in structural studies on such viruses as adenovirus (46), flock-house virus (47), rhinovirus (48), and Sindbis virus (49). Recent structural analysis of the BTV core is an excellent example of the combined use of cryo-EM and X-ray crystallography techniques to determine the structure of a complex macromolecular assembly at atomic resolution (50). Similarly, the cryo-EM structure of Norwalk virus has been very useful for determining the structure at atomic resolution of this virus by X-ray crystallographic data (51). Such a combinatorial approach is being used not only on icosahedral viruses, but also in the structural studies of ribosomes (52), which lack any symmetry, and other nonicosahedral complex structures, such as bacteriophage φ29 (53). 3. 3-D Structure of RV 3-D structural studies using cryo-EM techniques have been carried out on two RV strains: simian RV SA11 (2–4) and rhesus RV (5,6). The structural features seen in these two strains of RV are very similar, because both have three concentric layers of protein. A surface representation of a 3-D reconstruction of triple-layered (mature) RV along the icosahedral threefold axes is shown in Fig. 2A. Some of the structural features inside the RV structure are shown in a cross-sectional slice taken from the 3-D reconstruction of RV in Fig. 2B. The structure, which is based on a left-handed T = 13 icosahedral lattice, exhibits distinct structural features, including aqueous channels and surface spikes. The overall diameter of the particle, including the spikes, is 1000 Å.
3.1. Outer Layer Biochemical experiments have shown that the outer shell of the RV contains two proteins, VP7 and VP4 (54,55). The major component is VP7, a glycoprotein. A close examination of surface features (Fig. 2A) indicates that VP7 molecules cluster into triangular-shaped trimers surrounding the aqueous channels, on the T = 13 icosahedral lattice. The VP7 trimers have a small depression at the center, and they are connected to one another on the sides. Whether VP7 molecules aggregate into trimers prior to the assembly of the shell is not known, nor whether they cluster into trimers on their assembly into the icosahedral shell. The T = 13 icosahedral symmetry of the VP7 layer dictates that each virion has 780 molecules (or 260 trimers) of VP7. The overall thickness of the VP7 layer is ~35 Å.
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Fig. 2. (A) 3-D structure of the triple-layered particle (TLP) at a resolution of 23 Å. A set of icosahedral symmetry axes and the locations of the three types of channels are indicated. The surface is a left-handed T = 13 icosahedral lattice. In such a lattice, a fivefold axis is reached from its neighboring fivefold axis by stepping over three six-coordinated positions and taking a left turn. (B) Central cross section extracted from the 3-D map of the virus. The mass density breaks up into three distinct shells between the radii of 210 and 500 Å. These three shells are between the radii 500 and 340 Å, 340 and 270 Å, and 270 and 210 Å. The VP4 spike density is also shown; notice the lower domain of VP4 inside one of the channels. (C) Surface representation of the 3-D structure with anti(VP4) Fab bound. (D) Interaction of VP4 spikes with the VP6 layer. Scale bar = 20 Å.
3.2. Aqueous Channels A distinctive feature of the RV structure is the presence of aqueous channels at all of the five- and six-coordinated positions on the T = 13 icosahedral lattice (see Fig. 2A). These channels penetrate into the outer two layers of the structure and are ~140 Å deep. They have been classified into three types based on their location with respect to the icosahedral symmetry axes. Type I channels run down the icosahedral fivefold axes, type II channels are those on the
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six-coordinated positions surrounding 2the fivefold axes, and type III channels are those on the six-coordinated positions neighboring the icosahedral threefold axes. In each virion, there are 132 channels: 12 type I, 60 type II, and 60 type III channels. Type II and III channels are about 55-Å wide at the outer surface of the virus. The type I channels, in contrast, have a narrower and more circular opening of ~40 Å in diameter. All the channels constrict before widening in the interior, and have their maximum width at the position close to the surface of the inner shell proteins, as seen in the side view of the type III channels in Fig. 2B.
3.3. Surface Spikes Another distinctive feature of the RV structure is surface spikes. From the surface of the particles, 60 spikes extend to a length of 120 Å (Fig. 2A). The RV spikes are located at the outer edge of the type II channels that surround the icosahedral fivefold axes. Each spike has a distinct bilobed structure at the distal end. Each lobe has a diameter of ~25 Å. These lobes are individually connected to rod-shaped densities that are separated from each other by a hole that is ~20 Å in diameter. These rod-shaped densities, with a left-handed helical twist, merge together as they approach the surface of the outer layer, making two points of contact with the vertices of the triangular-shaped VP7 trimers. When the spikes on RV were first discovered, Prasad et al. (2) predicted that these surface projections were made up of VP4, which was confirmed by immunolabeling studies using MAbs against VP4 (3). Two antigen-binding fragments (Fab) molecules bound to the sides of the distal bilobes of each spike (Fig. 2C). The volume of the spike indicates that each spike can accommodate two molecules of VP4. Thus, each virion contains 120 copies of VP4. The observed structural features of individual spikes, as described above, and in vivo radiolabeling studies, further support the idea that the spikes are dimers of VP4 (4). Gel filtration analyses of expressed VP4 also have provided biochemical evidence that VP4 is an oligomer, probably a dimer. Interactions between these oligomers are apparently maintained by hydrophobic interactions, because these are readily disrupted by detergents (56), which explains why dimers of VP4 have not been detected by simple analysis of VP4 in virus disrupted by sodium dodecylsulfate (SDS) and separated by SDS-polyacrylamide gel electrophoresis.
3.4. Internal Domain of VP4 From the 3-D structure of the native virus alone, it is not possible to determine whether there is an inward extension of the VP4 spikes. That is, do the spikes terminate at the surface of the virion, or do they penetrate into the VP7 layer? Based on the volume of VP4 calculated from the 3-D reconstructions of
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the native and the Fab-bound virions, Prasad et al. (3) estimated that about 30 kDa of VP4 is buried inside the virion surface. By computing a difference map between the structures of the native strain and a reassortant strain of RV that lacks spikes, the existence of a large internal domain of VP4 beneath the VP7 surface was confirmed (4). This domain of VP4 is centered inside the type II channel, in close association with the walls of the channel made of trimers of VP6, as shown in Fig. 2D (the structure of the VP6 shell is described in more detail in Subheading 3.5.). Similar results were obtained by difference imaging of the virus structure at normal pH and alkaline pH (6). At alkaline pH, the RV spikes fall off without affecting the integrity of the VP7 layer (6,57). The finding that VP4 interacts extensively with the VP6 layer may have implications in the morphogenesis of the RV particles. Shaw et al. (4) have suggested that VP4 may play an important role, along with the nonstructural protein (NSP4), in facilitating the budding of progeny double-layered particles (DLPs) through the endoplasmic reticulum (ER) membrane, and that assembly of VP4 onto newly made particles occurs prior to VP7 assembly. An interesting aspect of RV infection is trypsin-enhanced infectivity (54,58). Trypsin, present at the natural site of infection, cleaves VP4 at a conserved arginine residue to produce VP8* (28 kDa, aa 1–247) and VP5* (~60 kDa, aa 248–776). Trypsin cleavage of VP4 is accompanied by a significant increase in RV infectivity (54,58), which is associated with enhanced cell entry (59,60). Although the overall structure of the VP4 spike is visualized in the cryo-EM reconstructions, the topographical locations of the proteolytic fragments VP5* and VP8* are not known. The MAb that was used in the structural studies by Prasad et al. (3), 2G4, is a VP5*-specific antibody, which binds to the sides of the distal lobes of the VP4-spike, suggesting that these distal globular domains of VP4 contain some portions of VP5*. The MAb 2G4 is a neutralizing antibody, and has been shown to block virus penetration, but not cell attachment. The exposed region of the distal tip of the spike is probably involved in initial attachment to cells, and the region of the spike binding to 2G4 molecules may be involved in cell penetration.
3.5. The Intermediate Layer Treatment of intact triple-layered virions with chelating agents (e.g., ethylenediamine tetra-acetic acid) removes the outer shell, reduces infectivity by several log steps, and exposes the inner shell proteins. The resulting DLPs are indistinguishable from those produced in infected cells (61). Electron micrographs of DLPs embedded in vitreous ice show that these particles are 705 Å in diameter, with a bristly surface. The 3-D structure of the DLPs (Fig. 3A) has been determined to ~19 Å, using cryo-EM techniques (7). The protein mass is mostly concentrated into 260 morphological units positioned at
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Fig. 3. (A) Surface representation of the 3-D structure of the double-layered particle (DLP) at a resolution of ~19 Å. A set of icosahedral symmetry axes and the locations of the three types of channels are indicated. (B) VP6 computationally isolated from the 3-D map of the DLP. (C) Surface representation of the VP2 layer extracted from the structure of DLP; the threshold used represents 120 molecules of VP2. (D) As viewed at a higher threshold, the boundaries of two molecules of VP2 that make the icosahedral asymmetric unit are denoted as A and B. Scale bar = 200 Å.
all the local and strict threefold axes of the T = 13 icosahedral lattice. The location and shape of the capsomeres strongly suggest a trimeric clustering of the inner capsid protein, VP6. This has been confirmed by separating VPs on nondenaturing conditions (62,63). These 260 capsomeric units are arranged in such a way that there are channels at all five- and six-coordinated centers. These channels are in register with the channels in the VP7 layer. The structure of the VP6 molecule (Fig. 3B) appears to have two domains: the globular upper domain, and a slender lower domain. The upper globular domains of individual monomers interact with one another in stabilizing the trimer. The sides of the lower domain seem to be involved in intercapsomeric interactions, and the bottom part of the lower domain interacts
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with the VP2 layer. VP6 molecules interact with VP7 through their upper domain. Flat upper domains of the VP6 trimers protrude into the VP7 layer, so that the close contact between the two shells occurs primarily around the local and strict threefold axes. The triangular-shaped VP7 trimers spread across the VP6 trimers, and portions of the VP6 trimers are visible through the outer-shell channels (Fig. 2A). It is possible that small molecules or small enzymes may be able to interact with portions of the VP6 layer, even in the presence of the outer VP7 layer. Consistent with this idea, at least one MAb to VP6 has been shown to bind to both triple-layered particles and DLPs (64). Recently, the structure of VP6 has been determined by X-ray crystallographic analysis to 2.8 Å (18). The tertiary structure of VP6 bears several similarities to the structure of VP7 of BTV (65), which is the counterpart of VP6 in RV. Both these proteins consist of two distinct domains: a distal eight-stranded β-barrel domain and a lower α-helical domain. The distal β-barrel domain interacts with VP7; the α-helical domain interacts with VP2. Fitting of the X-ray structure of VP6 into the cryo-EM structure of RV is in progress, to delineate the regions of VP6 that interact with VP7, VP4, and VP2.
3.6. Inner Layer Although early protein and EM analyses defined the composition of the outer two layers of the virion with certainty, the existence and composition of the third shell initially was based on conjecture. The total mass density in the outer shell accounts for 780 and 120 molecules of VP7 and VP4, respectively. The volume of the protein in the intermediate shell accounts for 780 molecules of VP6. On the basis of the radial density profile, and the fact that VP2 is the most abundant of the remaining three structural proteins, it was proposed that the density between the radii of 230 and 270 Å is caused by the shell formed by VP2 molecules (66). The existence of an inner shell was confirmed when single-layered (core-like) particles were produced by expression of VP2 alone (67). 3-D structural analysis of recombinant particles containing VP2 alone (2-virus-like particles [VLPs]) has indicated that the VP2 layer extends from a radius of about 230 Å to 270 Å, in agreement with interpretation based on the radial density profile computed from the 3-D structure of the virion. The structure determination of 2-VLPs to a resolution closer to ~20 Å has been hampered by extensive aggregation of 2-VLPs. However, knowing the radial extension of the VP2 layer, the structural features of the VP2 layer has been deduced from the reconstruction of native RV particles or recombinant particles (68) containing VP2 and VP6, referred to as 2/6-VLPs (10). In contrast to the VP7 and VP6 layers, the VP2 layer is a rather featureless, continuous bed of density (Fig. 3C). Examination of the structure of this layer at a slightly higher
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threshold of protein mass reveals the arrangement of 120 VP2 molecules on the T = 1 icosahedral lattice (Fig. 3D). Each asymmetric unit of the icosahedron consists of two molecules, referred to as type A and type B monomers. The type A monomer lies close to the fivefold axis, the type B monomer originates slightly away from the fivefold axis, and extends toward the threefold axis. The T = 1 icosahedral organization, with 120 molecules, poses an interesting structural question about how this layer, consisting of 120 molecules, interacts with the VP6 layer, consisting of 780 molecules. Such a T = 1 icosahedral structure, with 120 molecules, is unique, because it renders the two molecules in the asymmetric unit quasiequivalent, and has been observed in other dsRNA viruses besides RV. In the Reoviridae family, both aquareovirus (69) and BTV (15) have inner layers composed of 120 subunits on a T = 1 icosahedral lattice. A T = 1 symmetry with dimers is also observed in other dsRNA viruses outside Reoviridae: The fungal viruses L-A of Saccaromyces cerevisiae and P4 of Ustilago maydis exhibit similar organization (70). This unique T = 1 organization may be of fundamental significance in the endogenous transcription of the genome of dsRNA viruses.
3.7. Internal Organization In addition to VP7, VP4, VP6, and VP2 reviewed so far, the RV structure should account for VP1 and VP3, the remaining structural proteins, and the genomic RNA. In the 3-D structure of RV, several internal features are seen. To understand the internal organization of RV and to interpret the internal structural features in terms of minor proteins and the genomic RNA, a twofold strategy was used by Prasad et al. (7). First, difference imaging between native DLPs and various recombinant VLPs was used to identify the internal features. Second, a higher-resolution (~19 Å) structure of the DLP was carried out, to delineate the internal features more clearly.
3.7.1. Locations of VP1 and VP3 Although VP1 and VP3 are present in small amounts, they play an important role in the endogenous transcription process of RV. Several biochemical studies have indicated that VP1 is the RNA-dependent RNA polymerase (71) and VP3 is the guanylyltransferase (72). The structure of 1/2/3/6-VLPs shows flower-shaped features attached to the inside tip of the VP2 at all the fivefold positions (Fig. 4B), which is absent in 2/6-VLPs (Fig. 4A); it is also absent in 1/2/6-VLPs and 3/2/6-VLPs. These studies suggest that the flower-shaped feature becomes discernible only when VP1 and VP3 are both present, and that this structural feature represents a complex of VP1 and VP3. Although these studies strongly suggest the location of VP1 and VP3, the observed shape of the proposed VP1–VP3 is uncertain, because of the reconstruction procedures that
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Fig. 4. (A) Identical cross-section of 2/6-VLP structure. Notice the absence of the flower-shaped structural feature seen in B. (B) Cross-section (85 Å thick) of 1/2/3/6-VLP structure. Flower-shaped structures are seen attached to the inside surface of VP2 at all the icosahedral fivefold axes. (C) Cutaway from the 19-Å structure of the DLP, exposing the mass density caused by the genomic RNA. (D) Dodecahedral shell of the ordered RNA, extracted from the 19-Å structure. Scale bar = 200 Å.
implicitly use icosahedral symmetry. The shape that has been observed can only be real if each virion has 60 molecules of VP1–VP3, or this flower-shaped complex has an internal fivefold symmetry. The volume occupied by each flower-shaped structure, assuming a protein density of 1.30 g/cm3, accounts for about 25% of the expected mass of a complex of VP1 (125 kDa) and VP3 (98 kDa). Biochemical data indicate that there are 12 molecules of VP1 and 12 VP3 molecules per virion (72). The remaining portions of VP1 and VP3 may extend further inside the radius of 160 Å, and are transparent to structural analysis, either because they are disordered or because they move away from the fivefold axes, and lack any semblance of icosahedral symmetry. Thus, the flower-shaped structure may represent the structurally discernible portion of the VP1–VP3 enzyme complex.
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3.7.2. Location of Genomic RNA Comparison of the radial density profiles computed from the structures of DLPs and VLPs clearly indicate that mass density caused by the genomic RNA lies inside the radius of 230 Å (7). At this radius, the DLP structure shows strong mass density surrounding the fivefold axes, and between the neighboring threefold axes (Fig. 4C). This mass density is completely absent in the 2/6-VLP and the 1/3/2/6-VLP structures. The density attributed to ordered RNA in the 23-Å DLP is clearly resolved into parallel strands of tube-like mass density that form a dodecahedral shell in the 19 Å structure of the DLPs, determined using micrographs taken by using a 400 keV electron microscope. These tubes of mass density have an average diameter of 20 Å, typical of a dsRNA double helix (Fig. 4D). These strands encircle the VP1–VP3 complex located at the fivefold axis. Icosahedral packing of RNA does not imply 60-fold repetition in the gene sequence, but indicates that portions of the dsRNA occupy an icosahedrally equivalent volume in the structure. Icosahedrally ordered nucleic acid has been previously observed in small virus structures by X-ray crystallography (73–77). The maximum number of base pairs previously visualized in a virus structure is about 300 (76). In the RV structure, ordered RNA accounts for ~4500 of a total 18,555 bp. The 19 Å-DLP structure shows that there are several points of contact between the inwardly protruding portions of VP2 and the RNA surrounding each fivefold axis (7). These observations are consistent with biochemical results that show VP2 has dsRNA binding activity (78). Thus, VP2, which is icosahedrally assembled,
Fig. 5. (A) 3-D structure of transcribing DLP. (B) Proposed pathway for the nascent mRNA molecule exiting through a type I channel.
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appears to be responsible for inducing icosahedral ordering on closely interacting portions of RNA. Such extensive ordering of dsRNA is also seen in the recently determined X-ray structure of the BTV core (15), which is the equivalent of the RV DLP. The extensive ordering of the genome, described above, is perhaps critical for endogenous transcriptase activity. It may facilitate an orchestrated movement of the genome through the enzyme complex. This motion could be driven by the continuous exit of newly synthesized mRNA. The pores surrounding the fivefold axes, in the VP2 layer, are large enough to permit mRNA to exit, although it is also possible that VP2 may undergo conformational changes during transcription, to facilitate the exit of mRNA. Understanding of this process will be made clear by observing changes in the RNA and the VPs during active transcription, and also under conditions in which the DLP has been rendered transcriptionally incompetent. Though it has not been possible, from any of the current structural studies, to address the question of where the different segments of dsRNA are located inside the structure, it is tempting to speculate that a substantial portion of each of the 11 segments is ordered around a fivefold axis, and that each segment interacts with a specific VP1–VP3 complex.
3.8. Structure of Actively Transcribing DLPs From earlier studies, the authors had postulated that channels in the capsid layers would be used for the import of the precursors necessary for transcription and the exit of nascent mRNA molecules (2). As described in Subheading 3.5., there are 132 channels in the VP6 layer. Which of these channels are used for export of the nascent transcripts? To determine how the mRNA transcripts are translocated through the intact DLPs during transcription, the 3-D structure of actively transcribing DLPs has been determined (9). There are two chief inferences from these studies: First, the DLPs remain structurally intact during transcription; second, the nascent transcripts exit through the type I channels in the VP6 layer (Fig. 5A). The observation that the DLPs remain structurally intact is especially important, considering that the particles are capable of unlimited transcription, provided that the necessary RNA precursors are continuously supplied (79). The structural integrity of the particles is suggested not only from the images and the reconstruction, but also from the observation that the comparison of particle images with projections of the reconstruction by cross-common lines yields phase residual values that are very similar to those observed for nontranscribing DLPs. These results strongly imply that icosahedral symmetry is maintained to the same degree in both structures, at least to the resolution that these structures have been analyzed. Why is the structural integrity of DLP necessary for transcription? Biochemical studies show that no structural protein by itself is capable of carrying
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out transcription, which occurs within the confines of the intact DLPs. Structural and biochemical studies seem to indicate that the VP2 protein may play a scaffolding role in the architecture of RV. The inner surface of the VP2 layer not only provides a structural support for the RNA, but also helps to properly position the transcription complex consisting of VP1 and VP3. Recent biochemical studies have shown that the amino terminus of the VP2 molecule possesses nonspecific single-stranded RNA and dsRNA binding activity, and also that it is essential for the incorporation of VP1 and VP3 (80). Structural analysis of 2/6-VLPs, with an intact and a truncated aminoterminus of VP2, have shown that the amino-terminus is located at the inside surface of the VP2 layer, and close to the fivefold axis (10). The unique arrangement of VP2 molecules on a T = 1 icosahedral lattice, discussed previously (Fig. 3D), may be particularly useful in carrying out the dual roles of providing structural support to the RNA and transcription complex. It is likely that type A monomers of VP2, as they interact in a head-to-head manner at the fivefold axis, are exclusively involved in the interactions with VP1 and VP3, which are anchored to the inner surface of the VP2 capsid layer, along the fivefold axis; the N-terminal portions of type B monomers may provide the necessary interactions with RNA. The outer surface of the VP2 layer provides a structural platform for the assembly of VP6 trimers, preventing aggregation of the core particles (VP2, VP1, VP3, and the genome), which are known to be highly hydrophobic (81). The assembly of VP6 provides well-defined conduits for the exiting RNA molecules. From structural and biochemical studies, it can be hypothesized that the structural integrity of the DLP is necessary for the observed transcriptional efficiency and continuous reinitiation, because of the need to hold the components of the transcription machinery in their proper arrangements throughout repeated cycles of initiation–elongation, as well as to enable the efficient and continuous release of the mRNA transcripts.
3.9. Model for Exit Pathway of mRNA Although structural studies on the actively transcribing particle strongly suggest that the type I channels are used for exporting the nascent mRNA molecules, the precise pathway from the site of synthesis to the exterior of the particle remains to be elucidated. Based on the observed internal organization (7), and the structure of the transcribing particle, a model for the exit pathway in RV has been proposed (9). The RNA synthesis probably occurs within the core of the virion, very near to the fivefold axes, because this is the location of the transcriptional complexes composed of VP1 and VP3. Newly transcribed mRNA then exits the core through the channels in the VP2 layer, which are immediately adjacent to the fivefold axis, and probably in clos-
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est proximity to the site of synthesis. Once through the VP2 layer, the exiting strands of mRNA migrate out of the particle through the type I channels (at the fivefold axes) in the VP6 layer (Fig. 5B). In the electron images of the actively transcribing particles, 3–4 strands of mRNA, associated with the particles, have been observed (9). From these observations, it appears that multiple mRNA transcripts can be released simultaneously from an actively transcribing particle. Each genome segment may be transcribed by a specific polymerase complex, and the resulting transcript may exit the particle through the channel system at the fivefold axis adjacent to its site of synthesis. The mechanism of transcription and mRNA release proposed in these studies might also explain why no dsRNA virus with more than 12 genome segments has ever been found. 3-D structural studies using cryo-EM, similar to those described by Lawton et al. (9) on transcribing RV particles, have been carried out on orthoreoviruses (M. Yeager, personal communication). These studies have confirmed the finding of earlier studies using classical Kleinschmidt techniques, that the mRNA release occurs through the fivefold vertices in these viruses (82). In orthoreoviruses, the transcription complex is also suggested to be anchored to the inner surface of the innermost capsid layer at the fivefold axes (83). The release of mRNA through the fivefold vertices is probably a common characteristic of segmented dsRNA viruses, including BTV (15,84), aquareovirus (69), and the bacteriophage φ6 (85). 4. Conclusions and Future Challenges Structural studies on RVs have helped to provide a foundation for understanding the molecular mechanisms underlying some of the functions of these viruses, and are just the beginning of obtaining a detailed understanding of the structure–function relationships in this complex and large virus. Present and future studies are aimed at answering several questions. Are there specific receptors for RVs and how do these viruses initiate the cell-entry process? How does the transcription take place inside intact DLPs? How does the virus encapsidate a correct set of genome segments? What are the roles of the nonstructural proteins in virus replication and self-assembly, and what molecular interactions regulate their functions? What is the molecular mechanism of budding of the progeny DLPs into the ER membrane? The authors anticipate that a more complete picture of how these viruses replicate will emerge as structural analyses improve in resolution, either by cryo-EM or X-ray crystallography, or combinations thereof, in conjunction with advances in biochemical and molecular biological studies. Obtaining detailed molecular and structural information should also allow the development of more effective strategies to combat, prevent, or treat the clinical outcome of infections with these viruses. Increasing knowledge of the structure and
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function of the RV genes and proteins, and the ability to produce VLPs of various components, are already being exploited to develop interesting vaccine candidates (see Chapters 3 and 9). Acknowledgments The authors’ work is supported in part by grants from the NIH (AI 36040 and DK 31044), the W. M. Keck Foundation and the National Center for Research Resources (RR 02250). We thank J. Lawton and B. Pesavento for useful discussions and their help in making the figures. References 1. Estes, M. K. (1996) Rotaviruses and their replication, in Virology (Fields, B. N., Knipe, D. M., and Howley, P. M., eds.), Lippencott Raven, Philadelphia, pp. 1625–1655. 2. Prasad, B. V. V., Wang, G. J., Clerx, J. P., and Chiu, W. (1988) Three-dimensional structure of rotavirus. J. Mol. Biol. 199, 269–275. 3. Prasad, B. V. V., Burns, J. W., Marietta, E., Estes, M. K., and Chiu, W. (1990) Localization of VP4 neutralization sites in rotavirus by three-dimensional cryo-electron microscopy. Nature 343, 476–479. 4. Shaw, A. L., Rothnagel, R., Chen, D., Ramig, R. F., Chiu, W., and Prasad, B. V. V. (1993) Three-dimensional visualization of the rotavirus hemagglutinin structure. Cell 74, 693–701. 5. Yeager, M., Dryden, K. A., Olson, N. H., Greenberg, H. B., and Baker, T. S. (1990) Three-dimensional structure of rhesus rotavirus by cryoelectron microscopy and image reconstruction. J. Cell Biol. 110, 2133–2144. 6. Yeager, M., Berriman, J. A., Baker, T. S., and Bellamy, A. R. (1994) Three-dimensional structure of the rotavirus haemagglutinin VP4 by cryo- electron microscopy and difference map analysis. EMBO J. 13, 1011–1018. 7. Prasad, B. V. V., Rothnagel, R., Zeng, C. Q., Jakana, J., Lawton, J. A., Chiu, W., and Estes, M. K. (1996) Visualization of ordered genomic RNA and localization of transcriptional complexes in rotavirus. Nature 382, 471–473. 8. Prasad, B. V. V. and Estes, M. K. (1997) Molecular basis of rotavirus replication: structure-function correlations, in Structural Biology of Viruses (Chiu, W., Burnett, R., and Garcia, R., eds.), Oxford University Press, New York and Oxford, pp. 239–268. 9. Lawton, J. A., Estes, M. K., and Prasad, B. V. V. (1997) Three-dimensional visualization of mRNA release from actively transcribing rotavirus particles Nature Struct. Biol. 4, 118–121. 10. Lawton, J. A., Zeng, C. Q., Mukherjee, S. K., Cohen, J., Estes, M. K., and Prasad, B. V. V. (1997) Three-dimensional structural analysis of recombinant rotavirus-like particles with intact and amino-terminal-deleted VP2: implications for the architecture of the VP2 capsid layer. J. Virol. 71, 7353–7360. 11. Rossmann, M. G. and Johnson, J. E. (1989) Icosahedral RNA virus structure. Annu. Rev. Biochem. 58, 533–573.
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43. Frank, J., Zhu, J., Penczek, P., Li, Y., Srivastava, S., Verschoor, A., et al. (1995) Model of protein synthesis based on cryo-electron microscopy of the E. coli ribosome. Nature 376, 441–444. 44. Zhou, Z. H., Chiu, W., Haskell, K., Spears, H., Jr., Jakana, J., Rixon, F. J., and Scott, L. R. (1998) Refinement of herpesvirus B-capsid structure on parallel supercomputers. Biophys. J. 74, 576–588. 45. Baker, T. S. and Johnson, J. E. (1996) Low resolution meets high: towards a resolution continuum from cells to atoms. Curr. Opin. Struct. Biol. 6, 585–594. 46. Stewart, P. L., Fuller, S. D., and Burnett, R. M. (1993) Difference imaging of adenovirus: bridging the resolution gap between X-ray crystallography and electron microscopy. EMBO J. 12, 2589–2599. 47. Cheng, R. H., Reddy, V. S., Olson, N. H., Fisher, A. J., Baker, T. S., and Johnson, J. E. (1994) Functional implications of quasi-equivalence in a T = 3 icosahedral animal virus established by cryo-electron microscopy and X-ray crystallography. Structure 2, 271–282. 48. Smith, T. J., Chase, E. S., Schmidt, T. J., Olson, N. H., and Baker, T. S. (1996) Neutralizing antibody to human rhinovirus 14 penetrates the receptor-binding canyon. Nature 383, 350–354. 49. Cheng, R. H., Kuhn, R. J., Olson, N. H., Rossmann, M. G., Choi, H. K., Smith, T. J., and Baker, T. S. (1995) Nucleocapsid and glycoprotein organization in an enveloped virus. Cell 80, 621–630. 50. Grimes, J. M., Jakana, J., Ghosh, M., Basak, A. K., Roy, P., Chiu, W., Stuart, D. I., and Prasad, B. V. (1997) An atomic model of the outer layer of the bluetongue virus core derived from X-ray crystallography and electron cryomicroscopy. Structure 5, 885–893. 51. Prasad, B. V. V., Hardy, M. E., Dokland, Bella, J., M., Rossmann, M. G., and Estes, M. K. (1999) X-ray crystallographic structure of the Norwalk virus capsid. Science, in press. 52. Ban, N., Freeborn, B., Nissen, P., Penczek, P., Grassucci, R. A., Sweet, R., et al. (1998) A 9 Å resolution X-ray crystallographic map of the large ribosomal subunit. Cell 93, 1105–1115. 53. Tao, Y., Olson, N. M., Xu, W., Anderson, D. L., Rossmann, M. G., and Baker, T. S. (1998) Assembly of tailed bacterial virus and its genome release studied in three-dimensions. Cell 95, 431–437. 54. Estes, M. K., Graham, D. Y., and Mason, B. B. (1981) Proteolytic enhancement of rotavirus infectivity: molecular mechanisms. J. Virol. 39, 879–888. 55. Arias, C. F., Lopez, S., and Espejo, R. T. (1982) Gene protein products of SA11 simian rotavirus genome. J. Virol. 41, 42–50. 56. Zhou, Z., Crawford, S., and Estes, M. K. Personal communication. 57. Anthony, I. D., Bullivant, S., Dayal, S., Bellamy, A. R., and Berriman, J. A. (1991) Rotavirus spike structure and polypeptide composition. J. Virol. 65, 4334–4340. 58. Arias, C. F., Romero, P., Alvarez, V., and Lopez, S. (1996) Trypsin activation pathway of rotavirus infectivity. J. Virol. 70, 5832–5839.
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59. Kaljot, K. T., Shaw, R. D., Rubin, D. H., and Greenberg, H. B. (1988) Infectious rotavirus enters cells by direct cell membrane penetration, not by endocytosis. J. Virol. 62, 1136–1144. 60. Keljo, D. J., Kuhn, M., and Smith, A. (1988) Acidification of endosomes is not important for the entry of rotavirus into the cell. J. Pediatr. Gastroenterol. Nutr. 7, 257–263. 61. Cohen, J. and Dobos, P. (1979) Cell free transcription and translation of rotavirus RNA. Biochem. Biophys. Res. Commun. 88, 791–796. 62. Sabara, M., Ready, K. F. M., Frenchick, P. J., and Babiuk, L. A. (1987) Biochemical evidence for the oligomeric arrangement of bovine rotavirus nucleocapsid protein and its possible significance in the immunogenicity of this protein. J. Gen. Virol. 68, 123–133. 63. Shen, S., Burke, B., and Desselberger, U. (1994) Rearrangement of the VP6 gene of a group A rotavirus in combination with a point mutation affecting trimer stability. J. Virol. 68, 1682–1688. 64. Tosser, G., Labbé, M., Bremont, M., and Cohen, J. (1992) Expression of the major capsid protein VP6 of group C rotavirus and synthesis of chimeric single-shelled particles by using recombinant baculoviruses. J. Virol. 66, 5825–5831. 65. Grimes, J., Basak, A. K., Roy, P., and Stuart, D. (1995) The crystal structure of bluetongue virus VP7. Nature 373, 167–170. 66. Prasad, B. V. V. and Chiu, W. (1994) Structure of rotavirus. Curr. Top. Microbiol. Immunol. 185, 9–29. 67. Labbé, M., Charpilienne, A., Crawford, S. E., Estes, M. K., and Cohen, J. (1991) Expression of rotavirus VP2 produces empty corelike particles. J. Virol. 65, 2946–2952. 68. Crawford, S. E., Labbé, M., Cohen, J., Burroughs, M. H., Zhou, Y. J., and Estes, M. K. (1994) Characterization of virus-like particles produced by the expression of rotavirus capsid proteins in insect cells. J. Virol. 68, 5945–5952. 69. Shaw, A. L., Samal, S. K., Subramanian, K., and Prasad, B. V. (1996) Structure of aquareovirus shows how the different geometries of the two layers of the capsid are reconciled to provide symmetrical interactions and stabilization. Structure 4, 957–967. 70. Cheng, R. H., Caston, J. R., Wang, G. J., Gu, F., Smith, T. J., Baker, T. S., et al. (1994) Fungal virus capsids, cytoplasmic compartments for the replication of double-stranded RNA, formed as icosahedral shells of asymmetric Gag dimers. J. Mol. Biol. 244, 255–258. 71. Valenzuela, S., Pizarro, J., Sandino, A. M., Vasquez, M., Fernandez, J., Hernandez, O., Patton, J., and Spencer, E. (1991) Photoaffinity labeling of rotavirus VP1 with 8-azido-ATP: identification of the viral RNA polymerase. J. Virol. 65, 3964–3967. 72. Liu, M., Mattion, N. M., and Estes, M. K. (1992) Rotavirus VP3 expressed in insect cells possesses guanylyltransferase activity. Virology 188, 77–84. 73. Chen, Z. G., Stauffacher, C., Li, Y., Schmidt, T., Bomu, W., Kamer, G., Shanks, M., Lomonossoff, G., and Johnson, J. E. (1989) Protein-RNA interactions in an icosahedral virus at 3.0 A resolution. Science 245, 154–159. 74. Tsao, J., Chapman, M. S., Agbandje, M., Keller, W., Smith, K., Wu, H., Luo, M., Rossmann, M. G., and Compans, R. W. (1991) The three-dimensional structure of canine parvovirus and its functional implications. Science 251, 1456–1464.
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3 Virus Replication John T. Patton, Vladimir Chizhikov, Zenobia Taraporewala, and Dayue Chen 1. Introduction The effort to understand the molecular biology of rotaviruses (RVs) has led to the development of procedures that can be used to study the replication and transcription of the RV genome, the assembly and structure of the rotavirion, and the structure and function of RV proteins. Because it is not possible to provide a detailed description of all the techniques developed, this chapter stresses only those that have broad application, or which represent important new technical advances. In particular, this chapter emphasizes procedures used to prepare large amounts of purified triple- (TLP), double- (DLP), and single-layered (core) RV particles; to synthesize viral RNAs in vitro, through the transcriptase and replicase activities associated with RV particles; to evaluate the RNA-binding activity of RV proteins; and to assemble core-like and virus-like particles (CLPs and VLPs, respectively) via the expression of RV recombinant proteins.
1.1. Growth In Vitro With the discovery two decades ago that trypsin-like proteases enhance the infectivity of RVs (1), cultivation of these viruses to higher titers became possible, and the large amounts of virions necessary for the study of their structural and enzymatic properties could be purified. This Subheading describes protocols routinely used in the authors’ laboratory to propagate and titer RVs and to generate cores and DLPs from triple-layered rotavirions.
1.1.1. Virus Propagation Although many cell lines will support the growth of group A RVs, MA104 cells (fetal monkey kidney cells) are the most commonly used for their From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ
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propagation. MA104 cells can be grown to confluency in either minimum essential medium or Medium 199 containing 5% fetal bovine serum (FBS). However, in this laboratory, higher virus yields are obtained with the latter.
1.1.2. Preparation of Virions Containing Uncleaved Virion Protein VP4 The presence of trypsin in the medium during RV propagation will result in the cleavage of the virion protein VP4 into VP5* (60kDa) and VP8* (27kDa) (2) and, thus, virus purified from trypsin-containing media will contain little if any intact VP4. To obtain purified virus that contains only intact VP4, cells are infected at a high multiplicity of infection (MOI) (3–10 plaque-forming units [PFU]/cell) with trysin-activated virus, washed several times with serumfree medium following absorption, and maintained in trypsin-free medium containing protease inhibitors (see Subheading 3.2.2.).
1.1.3. Purification of Triple-Layered and Double-Layered Virions To generate the large quantities of RV particles that are often necessary for structural and biochemical analyses, the authors’ usually purify virus from 50 150-cm2 flasks of infected cell lysate. The virions are recovered from the cell lysate after treatment with trichlorotrifluoroethane, followed by high-speed centrifugation in cesium chloride (CsCl) to separate triple- and double-layered virions (see Subheading 3.2.3.). Triple-layered virions can be easily converted to DLPs through the removal of the VP4–VP7 outer shell by chelating agents such as ethylenediaminetetraacetic acid (EDTA) or ethylene glycoltetraacetic acid (EGTA) (3,4; see Subheading 3.2.4.).
1.1.4. Preparation of Cores DLPs can be converted to core particles by chaotropic agents such as calcium chloride (CaCl2) or sodium thiocyanate (5,6; see Subheading 3.2.5.).
1.2. Cell-Free Synthesis of Viral RNA In the infected cell, the RV RNA-dependent RNA polymerase, VP1, serves dual functions: as the viral transcriptase with which it catalyzes the synthesis of capped, but nonpolyadenylated messenger (m)RNAs (7); and as the viral replicase with which it catalyzes the synthesis of double- stranded (ds)RNA (8). During transcription, the RNA polymerase uses genomic dsRNA as the template to synthesize mRNA; during RNA replication, the RNA polymerase uses mRNA as the template to synthesize minus-strand RNA, resulting in the formation of dsRNA. The RNA polymerase activity, associated with DLPs and open core particles, allows the in vitro synthesis of viral mRNA and dsRNA, respectively (9,10).
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1.2.1. Transcription by DLPs In the presence of ribonucleotides, the transcriptase of DLPs can use the endogenous dsRNA genome as the template for synthesis of capped and methylated mRNA (9,11). The ability of DLPs to efficiently transcribe the genome is dependent on the presence of the VP6 shell, because removal of this protein from such particles yields core particles that are transcriptionally inactive (5,12). Several lines of evidence suggest that VP3 may be the viral guanylyltransferase, including the observation that, when incubated with guanosine 5'-triphosphate (GTP), the protein will form a covalently linked VP3–guanosine 5'-monophosphate (GMP) complex (13,14). Less is known about the identity of the methyltransferase of the DLP. Because a cell-free system has not been developed that supports the synthesis of mRNA from exogenous genomic dsRNA, many questions regarding the molecular details of the transcription process remain unanswered. The preparation of RV mRNA is described in Subheading 3.3.1.
1.2.2. Preparation of Plus-Sense Template RNA from cDNAs Full-length complementary (c)DNAs, corresponding to all the genome segments of SA11 and bovine RV and to some segments of many other RVs have been synthesized. Sequencing of the cDNAs has shown that the mRNAs of the group A RVs generally begin with the sequence 5'-GGC-3' and end with the sequence 5'-ACC-3' (15). To prepare viral mRNAs that have authentic 5' and 3' ends, the full-length cDNAs can be placed into a vector immediately downstream of the promoter for T7 RNA polymerase and upstream of a SacII site. The sequence of the joined T7 promoter (lower case) and the 5'-end of the viral cDNA (upper case) should be 5'-taatacgactcactataGGC-3' and the sequence of the joined 3'-end of the cDNA (upper case) and SacII site (underlined) should be 5'-ACCgcgg-3'. As illustrated below, digestion of the vector with SacII leaves a 3'-overhang that can be removed by the 3' to 5' exonuclease activity of T4 DNA polymerase. Run-off transcription of the treated vector (see Subheading 3.2.2.) will produce viral mRNAs that have authentic 5'- and 3'- termini. 5'-ACCgcgg-3' 3'-TGGcgcc-5'
SacII
→
5'-ACCgc 3'-TGG
gg-3' cgcc-5'
T4 DNA pol
→
5'-ACC
gg-3'
3'-TGG
cc-5'
Polymerase chain reaction (PCR) amplification is an alternate approach for preparing DNA templates for the synthesis of RV mRNAs with authentic 5' and 3'-termini (16). In this case, the plus-sense primer will contain the T7 promoter sequence, followed directly by the sequence formed by the first 18 nucleotides of one of the viral mRNAs, and the minus-strand primer will be the inverse complement of the sequence formed by the last 18 nucleotides of the
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homologous mRNA. The template used for PCR amplification can be either the cDNA product of a reverse transcription reaction of viral mRNA or a cDNA that has been cloned into a DNA plasmid. If Taq DNA polymerase is used to generate the template for transcription, then the amplified product should be treated with T4 DNA polymerase, to remove the nontemplated A residues that may have been added to the 3'-ends of the cDNAs. If pfu or vent DNA polymerase is used instead, then the nontemplated addition of an A residue will not occur, and therefore treatment of the amplified DNA with T4 DNA polymerase is not required. The cDNA templates are transcribed with T7 RNA polymerase under the condition provided by the supplier of the enzyme. With a T7 Megascript Kit (Ambion, Austin, TX), transcription of the cDNA templates will yield 50–100 µg viral mRNA. The quantity of the RNA is determined by spectrophotometry, and the quality of the RNA is assessed by electrophoresis on a 5% polyacrylamide gel containing 7 M urea (see Subheading 3.7.2.). RNA in the gel can be detected with ethidium bromide or by silver staining.
1.2.3. RNA Replication by Open Cores The replicase activity associated with open cores can use viral transcripts made from DLPs or from cDNAs as templates for the sythesis of dsRNA (10). Coupled with baculovirus (BV)-expressed core proteins (VP1, VP2, and VP3), the open core system has proved to be an important tool for understanding the mechanism of RV RNA replication. Recent studies have shown that a conserved stretch of seven nucleotides at the 3'-end of the viral mRNAs forms an essential cis-acting signal for minus-strand synthesis (16,17); the 3'-end of the viral mRNA must be single stranded for efficient minus-strand synthesis (18); and both VP1 and VP2 are required for RV replicase activity (19,20,21). A step-by-step protocol for the in vitro synthesis of minus-strand RNA by open cores is described in Subheadings 3.4.1. and 3.4.2.
1.2.4. Capping-Related Activities of Open Cores VP3 and other guanylyltransferases function in mRNA capping by transferring GMP to the phosphate end of the nascent RNAs (22). During this process, the guanylyltransferase first interacts with GTP and generates a guanylyltransferase–GMP intermediate complex, or, for RVs, a reversible and covalently-linked VP3–GMP intermediate complex (13,14). The guanylyltransferase activity then transfers GMP from the intermediate complex to the pyrophosphate end of the nascent RNA, producing a GpppN cap structure in which N is a purine residue. Protocols for generating the VP3–GMP intermediate complex, and for capping the nascent RNA by the guanylyltransferase activity associated with open cores, are shown in Subheadings 3.4.3. and 3.4.4.
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1.3. Detecting RNA-Binding Activity of RV Proteins Of the 12 proteins encoded by the RV genome, at least one-half possess RNA-binding activity (23). Although the role of many of these proteins in RV replication is unclear, their function may be related to RNA synthesis and packaging, mRNA transport to the site of genome replication, and mRNA translation and the regulation of gene expression. Many methods have been used to analyze the RNA-binding activity of RV proteins although the most important has been the gel-shift assay (for VP1, VP3, nonstructural proteins NSP1, NSP2, NSP3, see refs. 19,21,24,25,26, respectively). Other methods that have also been useful for analysis of binding proteins include Northwestern blot assays (VP2) (26,27), RNA-capture assays with immunoadsorbed protein (NSP1, NSP2) (28,29) and ultraviolet (UV)-crosslinking assays (VP2, NSP2, NSP3) (20,25,26,30). RV proteins that have been used in RNA-binding assays have been obtained from infected cell lysates (25,27) and purified virions (19,21) and have been expressed in vitro with rabbit reticulocyte lysates (25) and in vivo with BV (18,26) and vaccinia expression vectors (28).
1.3.1. Gel-Shift Assays The electrophoretic mobility shift assay, or gel-shift assay, is the simplest and most rapid of the RNA-binding assays. Its usefulness for the study of RVs is perhaps best illustrated by the fact that, of the six known viral RNA-binding proteins, five have been characterized by the gel-shift assay. 1.3.1.1. PREPARATION OF PROBES
Choosing a suitable probe to use in the gel-shift assay often requires making assumptions about the location of the recognition signals in an RNA. In detecting and analyzing the RNA-binding activities of the RV proteins, the authors have assumed that the specific recognition signals for these proteins are more likely to reside in the 5' and 3'-untranslated regions of the viral mRNAs than in the open reading frame (ORF). Indeed, this idea is consistent with data showing that cis-acting signals in viral mRNAs, which function in RNA replication and in gene expression, reside at the ends of the mRNA (16,17). In designing probes, one must consider that many RNA-binding proteins may recognize defined structural motifs in an RNA, e.g., stem-loops, bulges, panhandles, rather than, or in addition to, the primary sequence of the RNA. Therefore, it may be worthwhile to determine whether the probe to be used in the gel-shift assay assumes a fold pattern that is similar to its predicted secondary structure in the full-length RNA from which it is derived. The secondary structures of RNAs are easily computed with the mfold program made available on the home page of Michael Zuker of Washington University (http://www.ibc.wustl.edu/~zuker).
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Methods for preparing 32P-labeled RNA probes and performing the gel-shift assay are described in Subheadings 3.5.1. and 3.5.2. 1.3.1.2. ASSESSING STABILITY OF PROBE–PROTEIN INTERACTIONS
The strength of the interaction between an RNA and a protein can be evaluated by monitoring the effect of including different concentrations of salt in the gel-shift assay. 1.3.1.3. DISTINGUISHING BETWEEN SPECIFIC AND NONSPECIFIC RNA-BINDING ACTIVITIES
Because RNA-binding proteins generally display some degree of nonspecific affinity for single-stranded RNA, competitive gel-shift assays should be performed to determine whether the RNA-binding activity observed for any protein is specific or nonspecific (21). In the competitive gel assay, cold competitor RNA and 32P-labeled probe are usually included in the reaction mixture at molar ratios of 1:1, 10:1, and 100:1. Particularly for those proteins that appear to have specific affinity for the probe, the competitor RNA should be of the same approximate size as the probe, so that the relative concentration of the probe to competitor RNA used in the assay will be about the same at the molar and mass level. By using dsRNA as the competitor RNA, the comparative gel-shift assay can also be used to evaluate the ability of RV proteins to bind dsRNA. The effect of the competitor RNA on the formation of protein–probe complexes is evaluated by electrophoresis of the reaction mixture on a nondenaturing 8% polyacrylamide gel, and by quantitation of the intensity of bands formed by the shifted probe with a phosphorimager. If the cold competitor RNA causes a loss of the complex formed between the 32P-labeled probe and the viral protein, this indicates that the complex represents a nonspecific interaction between the probe and protein. On the other hand, if the presence of cold competitor RNA does not affect the formation of the probe–protein complex, or causes a decrease in the amount of probe–protein complex formed, which is less than the molar ratio of competitor RNA:probe used in the assay, then the interaction between the viral protein and the probe probably includes a specific component. 1.3.1.4. IDENTIFYING PROTEINS WITH RNA-BINDING ACTIVITY
Often, an extract prepared from infected cells or virus particles is found to contain an RNA-binding activity, but the identity of the protein in the extract responsible for the activity is unknown. Four alternative procedures that can be used to identify the protein component of a probe–protein complex detected by the gel-shift assay are described in Subheading 3.5.3.1.
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1.3.2. Drawbacks of Gel-Shift Assay There are three significant limitations associated with using the gel-shift assay to study RNA-binding proteins. First, the assay does not work well for proteins that are poorly soluble, or that form multimeric structures of large or varying size. For example, analysis of the major core protein VP2 is difficult, if not impossible, by the gel-shift assay, because this protein self-assembles into poorly soluble structures that cannot migrate into nondenaturing gels (10). Second, the assay is not suitable for the analysis of proteins with weak or short-lived affinity for RNA, or for resolving RNA–protein complexes that are not stable in a Tris-glycine (pH 8.8) electrophoresis buffer system. Third, the gel-shift assay restricts the size of the RNA probe that can be used as the bait for detecting and analyzing the RNA-binding activity of a protein. This is particularly a problem when the location of a recognition signal on the RNAs is unknown. In such cases, a longer stretch of RNA, possibly a full-length mRNA in the case of RV, might serve as a more effective probe, compared to short RNA probes that represent fragments of the RNA.
1.4. RNA-Capture Assay with Immunoadsorbed Protein Most of the limitations associated with the gel-shift assay can be avoided by using the RNA-capture assay, in which a protein is immunoadsorbed onto an affinity matrix, such as protein-A Sepharose beads, and the ability of the immobilized protein to bind RNA is then evaluated by incubation with radiolabeled probes. This technique has been described in detail elsewhere (29), and therefore is only summarized here. 1. An extract containing the protein of interest is prepared under nondenaturing conditions from RV-infected cells or cells programmed to produce the protein with a recombinant expression vector. The lysate is then incubated with a monospecific monoclonal or polyclonal antiserum that recognizes the protein. Because some antibodies (Abs) might block or inactivate the RNA-binding activity of a protein by altering its conformation, it is advisable to test as many different monospecific antisera as possible for the ability to bind to the protein without affecting its activity. 2. The Ab-protein complex is incubated with protein-A or -G Sepharose beads in a buffer that is anticipated to allow the protein to retain its physiological conformation. Often, this precludes the presence of strong detergents, e.g., sodium dodecyl sulfate (SDS). The beads are then thoroughly washed with the same buffer to remove unbound protein. To verify that the protein of interest has been immunoadsorbed onto the beads, and to determine which, if any, other proteins are bound to the beads, a portion of the material is analyzed by SDS-12% polyacrylamide gel electrophoresis (PAGE). (Instead of using an Ab and protein-A or -G to immobilize the protein onto an affinity matrix, it is also
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possible to use iminodiacetic acid, coupled to Sepharose, to immobilize his-tagged RV recombinant proteins, and then to use this protein-loaded matrix in the RNA-binding assay). 3. The RNA-binding activity of the immunoadsorbed protein is assessed by incubating with radiolabeled RNA probes in the appropriate RNA-binding buffer for 1 h on a rotating wheel. Following several washes to remove the unbound probe, the bound probe is eluted under high-salt conditions, typically in the range of 0.8–1 M NaCl. Electrophoresis of a fraction of the eluted radiolabeled probe on a 5 or 8% polyacrylamide gel containing urea (see Subheading 3.7.2.) allows visualization and quantitation of the bound RNA by phosphorimaging. Alternatively, the RNA-binding activity of the protein can be analyzed by quantitating the radioactivity that is eluted from the beads with a scintillation counter.
For detecting and defining interactions between RNAs and proteins, the RNA-capture assay offers some technical advantages over other systems: 1. The RNA-capture assay provides a mechanism to study the RNA-binding properties of proteins that normally exist as oligomers, be they homo- or hetero-oligomeric; this is particularly important, because the strength and specificity of the RNA-binding activity of such a complex may differ significantly from that of any individual component of the complex. 2. The RNA-capture assay can also be used to evaluate the RNA-binding activity of a protein, without resorting to the use of strong denaturants and without a priori purification of the protein. The former is an important advantage over the Northwestern blot assay, in which the proteins must be treated with SDS, and electophoretically resolved before assaying their RNA-binding activity. 3. Compared to the gel-shift assay, the RNA-capture assay allows the use of probes that range in size from small to large, and that may even be full-length. Perhaps better than any other approach, because of the flexibility in probe size, this assay system can be used in a systematic manner, to map the precise site of a recognition signal for an RNA-binding protein, beginning with a large RNA. 4. Finally, a strong advantage of the RNA-capture assay is that the system can be used easily and rapidly to define the optimal conditions that promote the interaction between a protein and an RNA. For example, metal ions (zinc), nucleotides (adenosine triphosphate [ATP] or GTP), or Na or K salts, can be added to the RNA-binding buffer to determine whether they enhance the RNA-binding activity of a protein. Similarly, the effect of temperature, pH, buffering system, stabilizing agents (glycerol and bovine serum albumin [BSA]), and the presence of protease and RNase inhibitors on the formation of RNA–protein complexes can also be rapidly evaluated with this system.
1.5. Assembly of CLPs and VLPs from Recombinant Protein The expression of RV proteins with recombinant baculoviruses (rBVs) has been useful not only for analyzing the properties of the RV proteins, but,
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perhaps even more so, for defining the structure of the rotavirion. In particular, expression of VP2 alone, with VP6, or with VP6, VP7, and VP4, in Spodoptera frugiperda (Sf9) insect cells, infected with appropriate rBVs, has been used to generate single-layered CLPs, double-layered VLPs, and triple-layered VLPs, respectively (31–33). Analysis of the recombinant particles by cryoelectron microscopy and image reconstruction (see Chapter 2) has helped to establish the number, location, and arrangement (e.g., triangulation number) of structural proteins in the virion (34). More recently, the co-expression of recombinant rVP1 and/or rVP3 with VP2 and VP6 has also provided insight into the structure and location of the viral RNA polymerase and guanylyltransferase in the core of the virion (35; see Chapter 2). From an applied point of view, the fact that purified recombinant TLPs, when administered orally to animals, can induce a neutralizing Ab response, raises the possibility that rVLPs may be used to generate RV vaccines (see Chapter 9). Although BV vectors are noted for expressing high concentrations of recombinant protein, the production of protein by this system is a relatively slow process, because it requires several steps, including the cloning of a cDNA into a specialized transfer vector, the generation of a rBV with the transfer vector, and the selection and plaque purification of the rBV. Therefore, other expression systems may be more useful in situations in which many different recombinant proteins need to be generated (e.g., in mutagenesis studies), and when the recovery of large amounts of recombinant protein is not critical. As an alternative to the BV system, recombinant proteins can be produced by transfecting T7 transcription vectors containing cDNAs of RV genes into CV1 or MA104 cells infected with vTF7-3, a recombinant vaccinia virus that constitutively expresses T7 RNA polymerase (36). With respect to the study of RV replication, the vTF7-3 expression system has several benefits: The expression of RV proteins by the vTF7-3 system can reach levels that are as high as those found in RV-infected cells; the vTF7-3 system can express RV proteins in cells which are permissive for RV replication, and therefore is likely to support authentic modification of recombinant proteins; and structural proteins expressed by the vTF7-3 system can also assemble into CLPs and VLPs.
1.5.1. Expression and Purification of VLPs Made by BV Vectors The co-expression of various combinations of RV structural proteins with rBV vectors in insect cells has been use to produce large amounts of highly purified single-layered CLPs, and double- and triple-layered VLPs. Detailed protocols for producing and purifying the recombinant particles are given elsewhere in the literature (8,31–33,35,37), and therefore will not be repeated here (see Chapter 9). However, there are several features of these protocols that are worth noting.
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1. Double- and triple-layered VLPs are released into the media of infected Sf9 cells; CLPs will remain cell-associated (31,32). 2. The detergent sodium deoxycholate (DOC) is commonly used in the purification of CLPs, because it enhances their solubility and aids in their release from cellular material (33). Sonication has also been use to dissociate CLPs from host debris, and to break up CLP aggregates. 3. CLPs and double- and triple-layered VLPs have been successfully purified by centrifugation on isopycnic CsCl gradients (8,31,32). However, because CLPs can be somewhat unstable in CsCl, procedures have also been developed for the purification of CLPs by sedimentation on sucrose gradients (33,35,37). 4. Double- and triple-layered VLPs are stable upon extraction in the lipid solvent trichlorotrifluoroethane (32). 5. Concentration of CLPs may result in their conversion into sheet-like, helical, or elongated bristly structures (37). 6. VP2 expressed by BV vectors is extremely susceptible to proteolytic cleavage (37), and therefore protease inhibitors (e.g., leupeptin and aprotinin) must be added to all solutions used to produce and purify the protein.
1.5.2. Expression of CLPs with the Vaccinia-Virus vTF7-3 System Infection of mammalian cells with the recombinant vaccinia virus vTF7-3 leads to the production of T7 RNA polymerase, which in turn can direct the synthesis of mRNAs from T7 transcription vectors contained in the same cell. Because of the production of virus-encoded capping enzymes in vTF7-3-infected cells, a significant percentage of the T7 transcripts will gain 5'-cap structures that will enhance their ability to direct protein synthesis. By modification of transcription vectors, so that they produce T7 transcripts that initiate with sequences specifying an independent ribosome entry site, translation of the transcripts becomes cap-independent, which often will result in even higher levels of recombinant protein expression in the cell. The vTF7-3 expression system has been used successfully to express nearly all RV proteins, including those that are structural components of the virus and those that have RNA-binding activity. Subheadings 3.6.1. and 3.6.2. describe methods used to express and purify VP2 CLPs from vTF7-3-infected cells. Derivative protocols may be used to study the assembly of double- and triple-layered VLPs, and to identify, by mutagenesis, domains in the structural proteins that are important for virion assembly.
1.6. SDS-12% PAGE for Separation of dsRNAs and Proteins Viral dsRNAs and proteins can be electrophoretically resolved on discontinuous gels (100 × 100 × 1.5 mm) containing a resolving portion of 12% polyacrylamide and a stacking portion of 4% polyacrylamide (see Subheading 3.7.).
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2. Materials 2.1. Virus Propagation 1. 2. 3. 4. 5. 6. 7. 8. 9.
MA104 cell cultures. Medium 199. Phosphate buffered saline (PBS), pH 7.2. Trypsin (cryst., 5 µg/mL, 1 µg/mL), or 5 µg/mL pancreatin. EDTA. FBS. Cell culture flasks (150 cm2). Six-well tissue culture plates. 1% Nutrient agar: prepared by mixing an equal volume of a 2% sterile solution of Bacto agar (Difco, all locations) or cell culture grade agarose and 2X Medium 199 containing 25 µg/mL of diethylaminoethyl dextran, 200 U/mL penicillin, and 200 µg/mL streptomycin. 10. 1% Neutral red agar: prepared by mixing an equal volume of a 2% sterile solution of cell-culture grade agarose and 2X Medium 199 containing 50 µg/mL neutral red. 11. Centrifuge. 12. Incubator.
2.2. Virus Preparation 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.
12. 13. 14. 15. 16. 17.
Methionine-free cell culture medium. 35S-labeled amino acids. Medium 199. Aprotinin. Leupeptin. Trichlorotrifluoroethane. Sorvall (DuPont Co., Wilmington, DE) Omni-mixer. Centrifuge. Ultracentrifuge. Ultracentrifuge tubes. Tris-buffered saline (TBS): 8 g NaCl, 0.38 g KCl, 0.1 g Na2HPO4, 1 g dextrose, 3 g Tris-base, 0.1 g MgCl2, 0.1 g CaCl 2. Dissolve in distilled H2O, adjust to pH 7.4 with HCl, and make up to 1 L. CsCl. 21-gage needle. Sodium azide (NaN3). 0.5 M EDTA, pH 8.0. Spectrophotometer. SDS-12% PAGE (see Subheading 2.7.).
2.3. RV RNA Preparation 1. Transcription cocktail: 100 mM Tris-HCl, pH 7.8, 50 mM sodium acetate, 10 mM magnesium acetate, 1.0 mM dithiothreitol, 2.5 mM each nucleoside
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2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.
triphosphate, 8.0 mM phospho(enol)pyruvate, 50 µg/mL pyruvate kinase, 1.0 mM S-adenosyl-L-methionine, 160 U/mL RNasin (Promega, Madison, WI). 10% SDS. Phenol:chloroform (1:1). Chloroform:isoamyl alcohol (24:1). 3 M sodium acetate. Ethanol. Centrifuge. RNase-free distilled water. Spectrophotometer. SacII restriction enzyme. T4 DNA polymerase. 10X T4 DNA polymerase buffer: 0.5 M NaCl, 0.1 M Tris-HCl, 0.1 M MgCl2, 10 mM dithiothreitol, pH 7.9 (New England BioLabs, Beverly, MA). 10 mg/mL BSA.
13.
2.4. Preparation and Use of Open Cores 1. Low-salt buffer (LSB): 2 mL 1 M Tris-HCl, pH 7.5, 1 mL 0.5 M Na 2EDTA, pH 7.5, 77 mg dithiothreitol. Make up to 1 L with distilled H 2O. 2. Digestion buffer: 340 U micrococcal nuclease, 10 mM Tris-HCl, pH 8.0, 10 mM NaCl, 1 mM CaCl2. 3. 1% agarose. 4. 50X TAE buffer: 242 g Tris-base, 136.1 g sodium acetate.3H2O, 19g Na2EDTA.2H2O Dissolve in distilled H2O, adjust to pH 7.2, and make up to 1 L. 5. EGTA. 6. 1 M Tris-HCl, pH 7.1. 7. 0.2 M magnesium acetate. 8. RNasin (40 U/µL). 9. 30% polyethylene glycol. 10. 10 mM nucleoside triphosphates. 11. Dithiothreitol. 12. [α-32P]-uridine triphosphate (UTP) (10 mCi/mL, 800 Ci/mmol). 13. RNase-free distilled water. 14. SDS-PAGE sample buffer (see Subheading 2.7.). 15. [α-32P]-GTP (10 mCi/mL, 800 Ci/mmol). 16. 5 mM MgCl2. 17. NaCl. 18. Phenol:chloroform (1:1). 19. Chloroform. 20. Sephadex G50. 21. SDS-12% polyacrylamide gel (see Subheading 2.7.). 22. 6 M urea-8% polyacrylamide gel (see Subheading 2.7.). 23. X-ray film. 24. Autoradiography cassette.
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2.5. Gel-Shift Assay 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18.
Ambion T7 MEGAshort transcription system. [α-32P]-UTP (10 mCi/mL, 800 Ci/mmol). 1.9 mM cold UTP. RNase-free DNase. Phenol:chloroform (1:1). Chloroform. 4 M NaCl. Ethanol. 6 M urea–8% polyacrylamide gel (see Subheading 2.7.). LSB. 8% polyacrylamide gel in Tris-glycine buffer (see Subheading 2.7.). Whatman 3M paper. X-ray film. Autoradiography cassette. 8% polyacrylamide gel containing 50 mM Tris-HCl, pH 8.8 (see Subheading 2.7.). SDS-PAGE sample buffer (see Subheading 2.7.). SDS-PAGE 4% stacking and 12% resolving gel (see Subheading 2.7.). Fluorographic enhancer (Amplify [Amersham, Piscataway, NJ] or Enhance [New Life, Boston, MA]). 19. 254 nm 25 W germicidal UV lamp. 20. RNase A.
2.6. Expression and Purification of VP2 CLPs from vTF7-3-Infected Cells 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15.
MA104 cell cultures. Medium 199. FBS. Penicillin (100 U/mL). Streptomycin (100 µg/mL). CO2 incubator. Cytosine β-D-arabinofuranoside (AraC). Vaccinia virus recombinant - vTF7-3. Qiagen Maxi-prep DNA purification kit. LipofectAMINE reagent (Gibco). Aprotinin. Leupeptin. PBS, pH 7.2. Centrifuge. Lysis buffer: 10 mM Tris-HCl, pH 7.4, 0.1 M EDTA, 1% DOC, 1 µg/mL aprotinin, 1 µg/mL leupeptin. 16. 20% sucrose (w/v) in 10 mM Tris-HCl, pH 7.4, 0.1 M EDTA, 0.2% deoxycholate (DOC). 17. Ultracentrifuge. 18. TE buffer: 10 mM Tris-HCl, pH 8.0, 0.1 mM EDTA.
46 19. 20. 21. 22.
Patton et al. Amicon Centricon microconcentrator. 3% phosphotungstic acid-NaOH, pH 7.2. Formvar-carbon coated grid. Transmission electron microscope.
2.7. Electrophoresis Systems 1. Electrophoresis power supply. 2. Electrophoresis tanks, vertical and horizontal. 3. Electrophoresis buffers: a. 10X SDS-PAGE running buffer: 105 g Tris-base, 504 g glycine, 35 g SDS, 3 L distilled H2O. Adjust to pH 9.1 with NaOH, and bring to a final volume of 3.5 L. b. SDS-PAGE sample buffer: 10 mL 50% glycerol (v/v), 1.25 mL 1 M Tris-HCl, pH 6.8, 5.0 mL 10% SDS (w/v), 8.0 mL distilled H2O, 150 µL 0.2% bromophenol blue in ethanol, 38.5 mg dithiothreitol. Bring to a final volume of 25 mL with distilled H2O. c. 10X TBE running buffer: 121.1 g Tris-base, 55 g boric acid, 7.5 g Na2EDTA. Bring to a final volume of 1 L with distilled H2O; pH should be 8.3. d. 2X Formamide sample buffer: 200 µL 0.5 M EDTA, pH 8.0, 200 µL 0.2% bromophenol blue in ethanol, 9.6 mL deionized formamide. 4. Electrophoresis gels: a. 12% Resolving gel: 15.5 mL 30% acrylamide (w/v), 6.4 mL 2% bis-acrylamide (w/v), 10.0 mL 1.5 M Tris-HCl, pH 8.8, 0.4 mL 10% SDS, 7.45 mL distilled H 2O, 20 µL TEMED, 0.2 mL 10% ammonium persulfate. b. 4% Stacking gel: 2.6 mL 30% acrylamide, 1.6 mL 2% bis-acrylamide, 2.5 mL 1 M Tris-HCl, pH 6.8, 0.2 mL 10% SDS, 13.7 mL distilled H2 O, 20 µL TEMED, 0.1 mL 10% ammonium persulfate. c. 6 M Urea-8% polyacrylamide gel: Dissolved 18.75 g of urea in 11.25 mL distilled H2O, then add 3.75 mL 10X TBE, 7.5 mL 40% acrylamide solution (38 g, acrylamide + 2 g bis-acrylamide in 100 mL distilled H 2 O). Add 7.5 µL TEMED and 0.38 mL 10% ammonium persulfate. d. 7 M Urea-5% polyacrylamide gel: Dissolve 16.8 g of urea in 17.5 mL distilled H 2O, then add 4 mL 10X TBE; 4 mL 50% acrylamide solution (50 g acrylamide + 1 g bis-acrylamide in 100 mL distilled H2O). Add 0.1 mL TEMED and 0.4 mL 10% ammonium persulfate.
3. Methods 3.1. Virus Propagation
3.1.1. Subculture of MA104 Cells 1. Disrupt a confluent cell monolayer of MA104 cells, by incubating in a solution of PBS containing 0.005% trypsin and 0.1% EDTA. 2. Resuspend the detached cells in serum-containing medium, (see Note 1). 3. Dilute the MA104 cell suspension from 1 in 5 to 1 in 10 in medium, and seed new cell culture flasks. The cells will produce a confluent monolayer in 4–7 d.
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3.1.2. Virus Propagation 1. Wash confluent MA104 cells maintained in 150-cm2 flasks, 3× with serum-free medium, or incubate several hours to overnight in one change of serum-free medium. 2. Prepare the viral inoculum by incubating virus stocks of known titer with 5 µg/mL trypsin for 30 min at 37°C, and diluting the activated virus stock to a volume of 5 mL/flask with serum-free medium, (see Note 2). 3. Remove the serum-free medium from the cell monolayer, and add 5 mL inoculum to each flask. 4. Incubate the flasks for 1 h at 37°C, with gentle swirling of the inoculum over the monolayer every 15 min. 5. Remove the inoculum and add 20 mL serum-free medium containing 1 µg/mL trypsin to the flasks, maintain the cells at 37°C until the cytopathic effect (CPE) is complete (see Note 3). 6. Freeze and thaw the infected cells 3× and remove large debris from the lysate by low-speed centrifugation (see Note 4). 7. The lysate is titered by plaque assay (see Subheading 3.1.3.), stored frozen at –20°C, and used as stock for subsequent propagation of the virus.
3.1.3. Titering Virus by Plaque Assay 1. Seed MA104 cells into six-well plates at a density that will allow the monolayers to reach confluency in 2–3 d (see Note 5). 2. Incubate the confluent monolayers in serum-free medium for 16–18 h, and inoculate with serial 10-fold dilutions of trypsin-activated virus (0.1 mL/well). 3. Incubate the plates at 37°C for 1 h, with occasional rocking to allow for virus adsorption. 4. Gently overlay the monolayers with 3.0 mL/well 1% nutrient agar or agarose containing pancreatin (3–5 µg/mL) or trypsin (1 µg/mL) cooled to 45°C. 5. Immediately after adding the overlay, swirl slowly, allowing the inoculum and overlay to mix. 6. Once solidified, incubate the plates for 3–4 d at 37°C. 7. Add 1 mL of a second overlay, consisting of 1% neutral red agar, cooled to 45°C, to each well, to aid plaque counting. 8. Count the plaques 3–5 d following the addition of the second overlay. Typically, the titer of rhesus rotavirus (RRV), simian rotavirus (SA11)-4F, and prototypic RVs will reach 1 × 108 PFU/mL.
3.2. Virus Preparation 3.2.1. Preparation of 35S-Labeled Virions When it is necessary to obtain 35S-labeled virions, monolayers are infected with virus and maintained in 85% methionine-free medium containing 5–10 µCi of 35S-labeled amino acids (Specific radioactivity: 1175 Ci/mmol) per mL. The infection is allowed to go until CPE is complete. Afterwards, the virus is purified from the lysate, as described in Subheading 3.2.3.
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3.2.2. Preparation of Virions Containing Uncleaved VP4 1. Infect MA104 cells at a high MOI (3–10 PFU/cell) with trypsin-activated virus. 2. Wash the cell sheet several times with serum-free medium following adsorption. 3. Maintain the cells in trypsin-free medium containing 1 µg/mL aprotinin or 0.5 µg/mL leupeptin (see Note 6). 4. Harvest the cells once the CPE has reached 70–80% (~24 h post infection).
3.2.3. Purification of Triple-Layered Virions 1. Combine the lysate with an equal volume of trichlorotrifluoroethane, and homogenize the mixture with a Sorvall Omni-mixer for three 1 min cycles on ice. 2. Centrifuge the homogenates at low speed to separate the organic and aqueous phases (4100g for 10 min in a Sorvall GSA rotor at 4°C). 3. Pellet the virus particles from the aqueous phase by centrifugation at 90,000g in a Beckman (Palo Alto, CA) SW28 rotor or at 45,000g in a Beckman Type 19 rotor for 2 h at 4°C. 4. Resuspend the pellet in TBS, and add CsCl to the virus suspension to achieve a density of 1.370 g/cm3, as determined by refractometry (see Note 7). 5. Centrifuge the mixture at 110,000g in a Beckman SW50.1 rotor for 20 h at 4°C. 6. Inspect the gradients in a darkened room with an inverted light source, to reveal two bands (see Note 8). 7. Recover the virus from the gradient by puncturing the bottom of the centrifuge tube with a 21-gage needle, and collect drops containing TLPs and DLPs, respectively, into separate tubes. 8. Remove the CsCl by dialyzing the virus extensively against TBS at 4°C. 9. Store the dialyzed samples at 4°C in the presence of 0.01% NaN3 (see Note 9).
3.2.4. Preparation of DLP Virus 1. Add 0.5 M EDTA, pH 8.0, to purified virus TLPs in TBS buffer, to give a final concentration of 10 mM EDTA. 2. Incubate for 1 h at 37°C with gentle periodic mixing. 3. Centrifuge the virus sample at 200,000g in a Beckman SW50.1 rotor for 1 h at 4°C, to pellet the DLPs. 4. Resuspend the pellet in TBS containing 10 mM EDTA. 5. Purify the DLPs by banding on a CsCl gradient (see Subheading 3.2.3 and Note 10).
3.2.5. Preparation of Cores 1. 2. 3. 4. 5. 6.
Dilute DLPs with TBS to an optical density OD 260 of 2 or less. Mix the sample with an equal volume of 2.0 M CaCl2 (see Note 11). Incubate for 1 h or more at 37°C, with frequent agitation (see Note 12). Centrifuge the sample at 16,000g for 2 min in an Eppendorf centrifuge at 4°C. Wash the pellet once with 0.5 mL 1 M CaCl2. Resuspend the cores in 2.5 µL TBS/mL of starting infected cell lysate.
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Fig. 1. Protein composition of purified TLPs (lane 1), DLPs (lane 2), and single-layered (core) particles (lane 3), as evaluated by SDS-12% PAGE. Virus was propagated in the presence of trypsin, causing the loss of VP4 and the appearance of VP5* and VP8* in the TLPs. Proteins in the gel were detected by staining with Coomassie blue R-250. 7. Dialyze extensively against TBS, to remove the residual CaCl2. 8. Purify the cores by centrifugation for 60 min at 200,000g in a Beckman SW50.1 rotor at 4°C through a 0.5-mL cushion of 40% sucrose in TBS (w/v). 9. Resuspend the pellets in TBS, and briefly sonicate to disrupt the core aggregates. 10. Adjust the suspension to a density of 1.45 g/cm3 with CsCl (by adding 3 g CsCl for every 4 mL core suspension). 11. Centrifuge the sample at 150,000g in a Beckman SW50.1 rotor for 20 h at 4°C. 12. Collect the cores by bottom puncture (see Note 13). 13. Dialize extensively against TBS to remove CsCl. 14. Determine the purity and quality of cores, DLPs and TLPs by SDS-12% PAGE (see Subheading 3.7., Fig. 1 and Note 14).
3.3. RV RNA Preparation 3.3.1. Preparation of mRNA from RV Particles 1. Pellet at least 100 µg CsCl gradient-purified DLPs in TBS by centrifugation at 200,000g in a Beckman SW50.1, or at 150,000g in a Beckman TLS55 rotor for 1 h at 4°C. 2. Resuspend in 1 mL transcription cocktail (see Note 15).
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3. Incubate for 2 h at 40°C (see Note 16). 4. Add 50 µL 10% SDS. 5. Extract the sample twice with an equal volume of phenol:chloroform (1:1, pH 4.5), and once with an equal volume of chloroform:isoamyl alcohol (24:1). 6. Add a 1/10 vol of 3 M sodium acetate and 3 vol of ethanol, mix, and incubate for 2 h at –70°C to precipitate the RNA transcripts. 7. Collect the RNA precipitate by centrifugation at 12,000g in a Sorvall SS34 rotor for 20 min at 4°C. 8. Resuspend the RNA pellet in 0.5 mL RNase-free water, and determine the concentration by spectrophotometry. The yield should be approx 200–300 µg RNA (see Note 17).
3.3.2. Production of Viral mRNA from Transcription Vector 1. Incubate 10 µg purified plasmid DNA with 30 U SacII restriction enzyme in a final volume of 50 µL for 1 h at 37°C. 2. Add 10 U T4 DNA polymerase, 10 µL 10X T4 DNA polymerase buffer, 1 µL 10 mg/mL BSA, 1 µL each of 2.5 mM dATP, dCTP, dGTP, and dTTP, and water to a final volume of 100 µL. 3. Incubate the reaction mixture for 20 min at room temperature. 4. Recover the plasmid by phenol:chloroform extraction and ethanol precipitation (see Subheading 3.3.1.).
3.4. Preparation and Use of RV Open Cores 3.4.1. Preparation of Open Cores 1. Prepare open cores by extensive dialysis of purified core particles (see Subheading 3.2.5.) against LSB at 4°C (see Note 18). 2. Incubate 100 µg open cores (see Note 19) in 1 mL digestion buffer, containing 340 U micrococcal nuclease, 10 mM Tris-HCl, pH 8.0, 10 mM NaCl, and 1 mM CaCl2 for 60 min at 30°C, to remove the dsRNA genome from open cores. 3. Confirm digestion of the dsRNA by electrophoresis of a 15-µL aliquot of the digest on a 1% agarose gel in TAE buffer. 4. Inactivate the micrococcal nuclease activity by adding EGTA to a final concentration of 3 mM to the digest.
3.4.2. Replicase Assay 1. Mix the following components: 1 µL 1 M Tris-HCl, pH 7.1; 1 µL 0.2 M magnesium acetate; 1 µL RNasin (40 U/µL); 1 µL 30% polyethylene glycol; 2.5 µL 10 mM mixture of nucleoside triphosphates (2.5 mM of each); 0.4 µL dithiothreitol; 1.5 µL [α-32P]-UTP (10 mCi/ml, 800 Ci/mmol); 2.5 µL open cores (100–300 µg/mL); 0.1–1.0 µg viral mRNA; RNase-free distilled water, to bring the reaction mix to a total volume 20 µL. 2. Incubate the mixture at 32°C for 2–3 h.
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Fig. 2. Replicase activity associated with untreated and micrococcal nuclease-treated open cores. The exogenous RNA added to some reaction mixtures (lanes 2 and 4) was produced in vitro by T7 transcription of a linearized plasmid containing a cDNA of the SA11 gene encoding NSP2 (gene 8). 32P-labeled dsRNA products recovered from the reaction mixtures were detected by SDS-12% PAGE and autoradiography. Note the appearance of radiolabeled background bands corresponding to the 11 genome segments made in the reaction mixture containing untreated open cores and no exogenous mRNA (lane 1). Micrococcal nuclease-treated open cores without exogenous RNA act as a negative control (lane 3). 3. Combine the reaction mixture with 50 µL SDS-PAGE sample buffer, and incubate for 15 min at 37°C. 4. Analyze the samples by SDS-12% PAGE (see Figs. 2 and 3, and Notes 20–23).
3.4.3. Formation and Detection of VP3–GMP Intermediate Complexes 1. Incubate 1–10 µg open cores with 100 µCi [α-32P]-GTP (800 Ci/mmol), 5 mM MgCl 2, and 50 mM NaCl in a final volume of 30 µL for 30 min at 37°C. (see Fig. 3 and Note 24). 2. Detect the 32P-labeled VP3 by SDS-12% PAGE and autoradiography.
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Fig. 3. The various biological activities of open cores and the influence of NaCl. (Top panel) 32P-labeled VP3–GMP intermediate complexes were formed by incubating 32P-GTP with open cores, and were detected by SDS-12% PAGE and autoradiography. Not only were 32P-labeled VP3–GMP complexes generated in reaction mixtures lacking NaCl, but, through an unknown mechanism, 32P-labeled VP1 was generated as well. In the absence of MgCl2, neither 32P-labeled VP3–GMP nor 32P-labeled VP1 were produced. The addition of NaCl to reaction mixtures prevented the formation of 32P-labeled VP1, but had little (<2-fold) effect on the formation of the 32 P-labeled VP3–GMP complex. (Second panel) RNA capping assays contained 32P-GTP, open cores, cold SP72-v3'40 RNA, and either no MgCl or MgCl and the 2 2 indicated level of NaCl. The products were analyzed by electrophoresis of reaction mixtures on a 6 M urea–8% polyacrylamide gel, and detected by autoradiography. As the results show, capping was dependent on MgCl2, and was nearly blocked at high
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3.4.4. Assay for 5'-Capping of RNA 1. Incubate 20 µg open cores in LSB, 5 mM MgCl2, 400 µCi [α-32P]-GTP (800 Ci/mmol), and 2.5 µg cold RNA in 160 µL for 3 h at 37°C, to produce capped and 32 P-GMP-labeled RNA (see Fig. 3 and Note 25). 2. Deproteinize the capped RNA by extraction with phenol:chloroform and chloroform; if necessary, unincorporated 32 P-GTP can be removed by passing the sample through a Sephadex G50 column. 3. Depending on the length of the RNA, the capped RNA can be detected by electrophoresis on either an SDS-12% polyacrylamide gel or a 6 M urea-8% polyacrylamide gel, followed by autoradiography or phosphorimaging.
3.5. Gel-Shift Assay 3.5.1. Preparation of 32P-Labeled RNA Probes 1. Prepare the 32P-labeled RNA probes for use in the gel-shift assay by run-off transcription with an Ambion T7 MEGAshort kit transcription (Austin, TX). The reaction conditions are as described by the manufacturer, except that 2.5 mCi/mL [32P]-UTP (800 Ci/mmol) and 1.9 mM cold UTP are included in the reaction mixture. 2. The templates used for transcription are generated either by PCR amplification of a viral cDNA or of a nonviral DNA using a plus-sense primer that also contains the promoter sequence for T7 RNA polymerase, or by linearizing a plasmid containing a promoter for T7 RNA polymerase at an appropriate site with a restriction enzyme. If necessary, T4 DNA polymerase can be used to produce blunt-ends the template before transcription (see Subheading 3.3.2.). 3. Incubate the transcription mixture for 8–18 h at 37°C. 4. Treat with RNase-free DNase to remove the DNA template. 5. Deproteinize by extracting sequentially with phenol:chloroform and chloroform. 6. Precipitate the RNA from the sample by adding 1/20 vol of 4 M NaCl and 2.5 vol of ethanol. Fig. 3. (continued) concentrations of NaCl (100–200 mM). (Third panel) Replicase assays contained 32P-UTP, open cores, no mRNA, or gene 8 mRNA and the indicated level of NaCl. The synthesis of 32P-labeled dsRNA in the reaction mixtures was quantitated by SDS-12% PAGE, followed by phosphorimaging. As the concentration of NaCl increased in the reaction mixtures, the level of dsRNA synthesis decreased. (Bottom panel) Gel-shift assays were performed to evaluate the RNA-binding activity of the core proteins. Reaction mixtures containing 32P-labeled SP72-v3'40 probe, open cores, no MgCl2, and the indicated amount of NaCl. The probe–protein complexes were detected by electrophoresis of the reaction mixtures on a nondenaturing 8% polyacrylamide gel. The presence of 50–150 mM NaCl had either no effect or an enhancing effect on the formation of VP1– and VP3–probe complexes. At 200 mM NaCl, the level of VP1–probe complexes formed in the reaction mixture was the same as that formed in reaction mixtures containing no added NaCl, but the level of VP3–probe complexes was reduced.
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7. Dissolve the probe in 20 µL RNase-free water, and gel-purify by electrophoresis on an 8% polyacrylamide gel containing 6 M urea, according to the protocol described in Subheading 3.7.
3.5.2. Assay Conditions (see Note 26) 1. Combine 1 pmol gel-purified 32P-labeled RNA probe with 2.5–5 µg open cores. 2. Adjust the mixture to a final volume of 10 µL with LSB. Include a control reaction mixture that contains the labeled probe, but lacks open cores. 3. Incubate the mixture for 60 min at room temperature (see Note 27). 4. Add 10 µL gel-shift sample buffer to the reaction mixture, and electrophorese the samples at 175 V on a nondenaturing 8% polyacrylamide gel in Tris-glycine buffer for 3–6 h (see Subheading 3.7. and Note 28). 5. Dry the gel onto Whatman 3M paper, and determine the location of the RNA probe in the gel by autoradiography (see Fig. 4 and Note 29).
Fig. 4 shows an example of the gel-shift pattern obtained with open cores and 32P-SP72-v3'40, an RNA probe that is 72 nucleotides long. The last 40 nucleotides of SP72-v3'40 are identical in sequence to the last 40 nucleotides of the SA11 gene 8 mRNA. The positions of the VP1–probe and VP3–probe complexes in the nondenaturing gel are indicated (lane 2, lower and upper bands, respectively). Despite containing a smaller protein, the VP3–probe complex migrates much slower in the gel than the VP1–probe complex. One explanation for this observation is that the VP3–probe complex may represent the interaction of a VP3 multimer with the probe. However, it is possible that the charge or Stokes radius of the protein is causing the slow rate of migration of the VP3–probe complex relative to the VP1–probe complex. Fig. 4 also exemplifies the usefulness of cold competitor RNAs as a tool to analyze the specificity of the interaction between viral proteins and probe in the gel-shift assay. The presence of the unlabeled luciferase RNA, Xenopus elongation factor-1 RNA, and brome mosaic virus RNA prevented the formation of a complex with VP3 and 32P-labeled SP72-v3'40, but not with VP1 and the same probe (21). Because the probe contains sequences present at the 3'-end of the RV gene 8 mRNA, the results of the gel-shift experiment suggest that VP3 does not specifically recognize the 3'-end of the mRNA, but VP1 does. Therefore, this experiment provides evidence that the 3'-terminus of the gene 8 mRNA may contain a specific recognition signal for VP1. As shown in Fig. 3, the RNA-binding activity of VP1 and VP3 was not inhibited by the presence of NaCl, except at 200 mM, at which the binding activity of VP3 was reduced by approx 30%. Indeed, in the presence of low concentrations of NaCl (50 mM), the RNA-binding activity of VP1 and VP3 was enhanced. Given that the physiological concentration of salt in the cell is
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Fig. 4. RNA-binding activities of open cores, as evaluated by the competitive gelshift assay. Open cores and 1 pmol 32P-labeled probe, SP72-v3'40, were incubated alone or with 5 pmol luciferase RNA, 1 pmol xenopus elongation factor-1 RNA, or 2 pmol brome mosaic virus RNA. Probe–protein complexes in the reaction mixtures were resolved by electrophoresis on a nondenaturing 8% polyacrylamide gel, and were detected by autoradiography. When open cores were not added to the reaction mixture, the upper and lower complexes were not detected (lane 1). Protein analysis has indicated that VP3 and VP1 are responsible for the formation of the upper and lower bands, respectively (lane 2). Note that in the presence of cold competitor RNA, the upper band is lost (lanes 3–5), indicating that the interaction between VP3 and the probe is nonspecific.
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taken as 100–150 mM, these data would indicate that VP1 and VP3 can form stable complexes with RNA in the infected cell. 3.5.3. Identifying Proteins with RNA-Binding Activity 3.5.3.1. DETECTION OF 35S-LABELED PROTEIN IN PROBE-PROTEIN COMPLEXES
This procedure requires the use of 35S-labeled viral protein in the gel-shift assay, and was used to establish that VP1, VP3, and NSP2 have RNA-binding activity (19,21,24). 1. Combine 35S-labeled open cores, or an extract prepared from 35S-labeled infected cells, with an appropriate 32P-labeled RNA probe. 2. Incubate for 30–60 min at room temperature (see Subheading 3.5.2.). 3. Resolve the probe–protein complexes in the mixture by electrophoresis on a nondenaturing 8% polyacrylamide gel containing 50 mM Tris-HCl, pH 8.8 (see Subheading 3.7.). 4. Dry the gel onto Whatman 3M paper and detect the positions of the probe–protein complexes by autoradiography. 5. With the autoradiograph as a guide, cut out portions of the gel containing probe-protein complexes. 6. Rehydrate the pieces for 5 min in 200 µL SDS-PAGE sample buffer at room temperature. 7. Separate the gel pieces from the Whatman paper with tweezers. 8. Incubate the gel pieces for 30 min more at 37°C in 200 µL of fresh sample buffer. 9. Insert the rehydrated pieces into the well of an SDS-PAGE 4% stacking gel layered over an SDS-PAGE 12% resolving gel (see Subheading 3.7.). Parallel wells are loaded, with appropriate viral protein markers and mol wt standards. 10. Following electrophoresis, soak the gel in a fluorographic enhancer agent such as Amplify or Enhance. 11. Detect the 35S-labeled protein in the gel by autoradiography.
3.5.3.2. CROSSLINKING OF RNA–BINDING PROTEIN TO 32P-LABELED PROBE
The basis for this procedure is that exposure of RNA–protein complexes to UV light can induce the formation of covalent linkages between the RNA and protein. 1. Incubate unlabeled open cores or extracts of infected cells with a high specific activity 32P-labeled probe, under the same conditions used for the gel-shift assay (see Subheading 3.5.2.). 2. Expose the reaction mixtures to UV light for various lengths of time (e.g., 0, 5, 10, 15, 30, 60 min) using a 254-nm 25-W germicidal lamp placed 3–4 cm above the samples. 3. Treat the samples with 10 µg/mL RNase A at 37°C, to remove all but the few nucleotides of the RNA probe that are covalently linked to the protein via exposure to UV light.
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4. Identify the 32P-labeled protein by co-electrophoresis on an SDS-12% polyacrylamide gel, with appropriate viral protein markers and mol wt standards (see Note 30).
3.5.3.3. RNA-BINDING ACTIVITY OF RECOMBINANT PROTEIN
Large amounts of nearly all the RV proteins have been produced with both recombinant eukaryotic (BV and vaccinia virus) and prokaryotic (bacterial) expression vectors. Some of the recombinant proteins have also been purified to homogeneity, or nearly so, and used in gel-shift assays to assess the potential RNA-binding activity of the protein. The obvious advantage of using purified recombinant proteins in the gel-shift assay is that detection of any RNA–protein complexes in the gel leads to the conclusion that the protein has RNA-binding activity. For example, experiments that showed that BV-expressed VP1 interacted with the 3'-specific viral probe SP72-v3'40 were useful for confirming results of earlier gel-shift assays, which suggested that VP1 has RNA-binding activity (18). Similarly, NSP1 produced in bacteria and NSP2 produced in CV1 cells, with the vaccinia virus vTF7-3 expression system, have also contributed to verifying and defining the RNA-binding activities of these proteins (28,29). The limitation of using recombinant protein to evaluate the RNA-binding activity is that the recombinant protein may behave differently than the naturally occurring protein. As an example, VP3 contained in open core preparations interacts readily with probes to form RNA–protein complexes that can be detected by the gel-shift assay. In contrast, recombinant VP3 produced with a BV expression vector is largely insoluble, and, as yet, has not been shown to have RNA-binding activity. 3.5.3.4. SUPERSHIFTING PROBE–RNA COMPLEXES WITH SPECIFIC ABS
Monospecific polyclonal and monoclonal antisera can be used together with the gel-shift assay as another approach for identifying proteins that interact with RNA probes to form complexes. The basis for this approach is that the specific interaction of an Ab molecule with the protein component of the probe–protein complex will dramatically increase the overall mass of the complex, which in turn will decrease its rate of migration during nondenaturing electrophoresis. Regarding application, binding of the Ab will cause the probe–protein complex to produce a band that is shifted upwards (supershifted) in the gel, relative to the same probe–protein complex that is Ab-free. Occasionally, binding of the Ab will generate a complex sufficiently large that it will not migrate into an 8% polyacrylamide gel (see Subheading 3.7.). Because monospecific and monoclonal antisera have been generated to most RV proteins, supershift assays are a reasonable approach for studying RNA–protein complexes formed with RV proteins. The quantity of Ab used in the assay must be empirically determined, because it will depend on the concentration of
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both the Ab and the protein, and on the strength of the affinity of the Ab for the protein. It is also critical that the antiserum be free of RNase activity, and that the binding activity of the Ab does not destabilize the probe–protein complex. Finally, it is important to realize that, although supershift assays can be used to identify a protein component of a probe–protein complex, it does not rule out the possibility that the complex may be heteromultimeric, and therefore may also contain other protein components.
3.6. Expression and Purification of VP2 CLPs from vTF7-3-Infected Cells 3.6.1. Infection and Transfection of Cells 1. The day prior to transfection, the appropriate number of 75-cm2 cell culture flasks are each seeded with 2 × 106 MA104 cells in 20 mL Medium 199 supplemented with 7% FBS, 100 U/mL penicillin, and 100 µg/mL streptomycin. The cells are incubated at 37°C in an atmosphere of 5% CO2-air, and should reach 80% confluency the following day. 2. On the day of transfection, the cell monolayers are washed with serum-free and antibiotic-free Medium 199. Afterwards, 20 mL Medium 199 containing 40 µg/mL AraC (Medium 199-AraC) is placed in each flask, and the cells are incubated for 1 h at 37°C in a CO2 incubator. AraC is an inhibitor of vaccinia virus DNA replication (38) and, in the authors’ experience, enhances the expression of recombinant proteins in vTF7-3-infected cells. 3. To prepare the virus inoculum, a vTF7-3 stock of known titer is vortexed vigorously, to disperse any clumps of material contained in the preparation (see Note 31). If this treatment is not adequate, then the stock should be sonicated for 15–30 s on ice. For each flask to be infected, 3 mL inoculum is prepared by diluting the vTF7-3 stock in Medium 199-AraC, to yield a MOI of 10 PFU/cell. Following removal of the medium from each flask, the inoculum is added, and the cells maintained for 1 h at 37°C in a CO2 incubator. During this period, the flasks are gently rocked every 10–15 min to allow the inoculum to swirl over the monolayer. 4. The production of VP2 CLPs requires a plasmid that contains a cDNA of the RV gene 2 ORF, positioned downstream from the promoter for T7 RNA polymerase. For the authors, plasmids that give the highest level of expression when transfected into cells are either purified by banding on CsCl-ethidium bromide gradients or with a Qiagen Maxi-prep DNA purification kit. About 30 min before the end of the vTF7-3 infection period, two solutions, A and B, are prepared, following the protocol provided by Life Technologies (Gaithersburg, MD) for transfecting cells with LipofectAMINE reagent. Solution A is made by adding 20 µg VP2 transcription vector to 700 µL Medium 199 containing 40 µg/mL AraC. Solution B is made by adding 100 µL LipofectAMINE reagent (Gibco) to 700 µL Medium 199, also containing AraC. Solutions A and B are combined,
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mixed gently, and then incubated for 30 min at room temperature. Afterwards, the combined solutions A and B are increased to a final volume of 6 mL by the addition of Medium 199-AraC. 5. Immediately before transfection, the vTF7-3 inoculum is removed from the flasks, and the cells are washed once with 10 mL Medium 199-AraC. The plasmid–LipofectAMINE mixture is gently layered over the cell monolayers and the flasks are incubated for 5 h at 37°C in a CO2 incubator. 6. Five h posttransfection, 6.0 mL Medium 199-AraC, containing 20% FBS, 100 U/mL penicillin, and 100 µg/mL streptomycin, are added to each flask. To inhibit the degradation of the expressed VP2, 1 µg/mL each of aprotinin and leupeptin are also added to the medium. The cells are then maintained at 37°C in a CO2 incubator. Overt cytopathic effects caused by infection of the cells with vTF7-3 virus will become apparent as early as 3–4 h postinfection. Although the infected cells will round up, the monolayer mostly will remain attached to the surface of the flask until time of harvest.
3.6.2. Purification of VP2 CLPs 1. At 24–36 h postinfection, the medium is removed from the flasks, and the cell monolayer is carefully washed with cold PBS. A rubber policeman or scraper is then used to scrape the cells into a small volume of cold PBS. 2. The cells are pelleted from the suspension by centrifugation at 1000g for 5 min at 4°C. The supernatant is removed, and the cell pellet is resuspended in 0.5 mL lysis buffer (10 mM Tris-HCl, pH 7.4, 0.1 mM EDTA, 1% DOC, 1 µg/mL each of aprotinin and leupeptin) per 75-cm2 cell culture flask harvested. The cell lysate is then incubated for 5 min at 15°C. 3. The lysate is transferred to a 1.5-mL microcentrifuge tube, and then clarified by centrifugation at 16,000g for 10 min at 4°C in a microcentrifuge. 4. The clarified lysate is layered onto 10-mL linear gradients of 5–20% sucrose (w/v) in 10 mM Tris-HCl, pH 7.4, 0.1 mM EDTA, and 0.2% DOC, and the gradients are centrifuged at 110,000g for 2 h in a Beckman SW 40.1 rotor at 4°C. 5. One-mL fractions are collected from the gradient, and the distribution of VP2 in the gradient is determined by SDS-PAGE of the fractions. Usually, VP2 forms a peak in the middle of the sucrose gradient. 6. Fractions containing VP2 are adjusted to 1 µg/mL each of aprotinin and leupeptin to inhibit protease activity, and are dialyzed against 10 mM Tris-HCl, pH 7.4, containing 0.1 mM EDTA, to remove sucrose. If necessary, VP2 in samples can be concentrated by centrifugation in an Amicon Centricon 30 microconcentrator. 7. To evaluate VP2 samples for the presence of CLPs by EM, a drop of the sample is placed on a Formvar-carbon-coated grid (EM Science, Gibbstown, NJ) and left for 1 min. The sample fluid is drawn off the grid with a piece of filter paper, and then a drop of 3% phosphotungstic acid-NaOH, pH 7.2, is added to the grid. After 1 min, the stain is removed with a piece of filter paper, and the grid is inspected for CLPs at a magnification of 50,000× (Fig. 5).
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Fig. 5. Production of VP2 CLPs and VP2-VP6 VLPs with the vTF7-3 expression system. MA104 cells were infected with vaccinia virus vTF7-3 and transfected with a VP2 transcription vector (A) alone, or in combination with a VP6 transcription vector (B and C). Particles were purified from the cell lysates, stained with phosphotungstic acid, and examined by electron microscopy. The bar is equivalent to 100 nm.
3.7. Electrophoresis Systems 3.7.1. SDS-12% Polyacrylamide Gel Electrophoresis 1. Prior to electrophoresis, protein samples are mixed with sample buffer, and are usually heated to 95°C for 3 min. (see Notes 32 and 33). 2. Electrophoresis is performed in 1X running buffer for 15 h at 30 mA for dsRNAs and 3–4 h at 60 mA for proteins. 3. Gels can be stained with ethidium bromide or silver, to detect dsRNAs, and with Coomassie blue R250 or silver, to detect proteins.
3.7.2. Urea–Polyacrylamide Gels for Purification of RNA Probes and Resolving Viral mRNAs 1. Immediately prior to electrophoresis, 32P-labeled probes in a volume of 20 µL distilled H2O are mixed with an equal volume of 2X formamide sample buffer, and are heated to 75°C for 10 min (see Note 34).
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2. The probes are electrophoresed on an 8% polyacrylamide gel containing 6 M urea (100 × 100 × 1.5 mm) in 1X TBE at 250 V, until the bromophenol dye front migrates to the bottom of the gel (2–3 h). 3. Positions of probes in wet gels are identified by autoradiography, and portions of the gel containing probes are cut out and soaked overnight at 37°C in 0.5 mL distilled H2O. 4. The eluted probes are extracted once with phenol:chloroform, and recovered by ethanol precipitation.
3.7.3. Nondenaturing Polyacrylamide Gels Used in the Gel-Shift Assays RNA–protein complexes are resolved by electrophoresis on a nondenaturing 8% polyacrylamide gel (100 × 100 × 1.5 mm) for 3–4 h at 175 V. Gels are dried and analyzed by autoradiography.
3.7.4. RNA Strand Separation Gel The plus-sense and minus-sense RNAs of RV dsRNAs can be resolved by electrophoresis on 1.75% agarose gels containing 6 M urea and 25 mM citrate buffer, pH 3.0. The details for this procedure have been published elsewhere (39). 4. Notes 1. Antibiotics (100 U/mL penicillin and 100 µg/mL streptomycin) can be included in the medium to reduce the possibility of bacterial contamination. 2. The concentration of virus in the inoculum should be sufficient to produce a MOI of 0.01–0.5 PFU per cell, depending on the growth characteristics of the virus. In general, slow-growing RVs require a higher MOI than fast-growing viruses, to obtain similar virus yields. 3. Under the conditions described in Subheading 3.1.2., CPEs are usually complete within 24–48 h of infection, especially for the well cell culture-adapted, fast-growing strains of RVs such as RRV and SA11. 4. Low-speed centrifugation is not necessarily required, if the lysate will be used for preparing CsCl-purified virus. 5. This can be accomplished by resuspending a confluent monolayer of MA104 cells (1–1.5 × 107 cells) from a 150-cm2 flask into 120 mL serum-containing medium and placing 3 mL cell suspension in each well of a six-well plate. 6. The protease inhibitors are also included in all buffers used in the purification of the virus. 7. A density of approx 1.37 g/mL can be produced by adding 2 g CsCl to 4 mL virus. 8. The upper band consists of triple-layered virions, and will have a density of 1.36 g/cm3; the lower band consists of DLPs and will have a density of 1.38 g/cm3. 9. The samples can be stored for at least 6 mo without noticeable degradation.
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10. If recovery of purified TLPs is not required, DLPs can be obtained directly from infected cell lysates as follows: Add 0.5 M EDTA (pH 8.0) to the lysates, to give a final concentration of 50 mM; extract the cell lysates with trichlorotrifluoroethane, and pellet the virus particles from the lysates, as described in Subheading 3.2.4. The pellets are resuspended in TBS, and the DLPs are purified by centrifugation in a CsCl gradient. To ensure high purity of the DLPs, an additional round of CsCl centrifugation is strongly recommended. 11. The efficiency of disruption of the VP6 layer of protein by CaCl2 decreases as the concentration of DLPs increases, and may differ, depending on the strain of virus used. Even in preparations in which CaCl2-treatment fails to remove as much as one-half of the VP6 layer of protein, the open core preparation will have a high level of replicase activity, and will lack transcriptase activity. 12. The treatment will cause the sample to become cloudy, and will eventually result in aggregation of the core particles. 13. The cores may appear as a flocculent band in the gradient, and will have a density of approx 1.45–1.50 g/cm3, depending on the amount of residual VP6 associated with the cores. 14. As shown in Fig. 1, purified DLPs should not contain detectable VP7 and VP4, or the trypsin-cleavage products of VP4, VP5*, and VP8*, and core particles should contain virtually no VP6. Although purified cores are not as stable as DLPs and TLPs, all three types of particles can be stored for several months at 4°C without loss of structural integrity. Frequently, as cores age, a specific degradation of VP2 generates a set of fragments that are slightly smaller than the wild-type protein, and which, upon SDS-PAGE, can obscure detection of VP3. Degradation of VP2 can be slowed by maintaining cores in the presence of 1 µg/mL leupeptin and aprotinin. 15. S-adenosyl-L-methionine is added to the reaction mixture, because it provides the methyl group for methylation of the cap structures added to the 5'-end of the mRNAs by guanylyltransferase. 32P-UTP can also be included in the reaction mixture, to label mRNA products. 16. If it is necessary to remove the endogenous dsRNA, the reaction mixture is centrifuged at 200,000g in a Beckman SW50.1 or TLS55 rotor for 1 h at 4°C. If necessary, the tubes can be filled by overlaying the reaction mixture with mineral oil. 17. Because of disruption of some DLPs during the transcription reaction, it is common to find some residual dsRNA contamination of the mRNA preparation. The residual dsRNA can be removed by lithium chloride precipitation (40). The quality of the mRNA preparation is assessed by electrophoresis of 1–2 µg on a 5% polyacrylamide gel containing 7 M urea (see Subheading 3.7.). A high quality preparation should contain all 11 transcripts, although the larger transcripts (of RNA segments 1–4) usually will be less abundant than the smaller transcripts. 18. Dialysis should continue until the sample has clarified (>24 h). The open cores can be used directly in replicase assays, or can be treated with micrococcal nuclease (see Subheading 3.4.1.2.) to remove endogenous dsRNA before being used in replicase assays.
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19. The concentration of protein can be determined with a Bio Rad (Hercules, CA) Protein Assay kit, or by co-electrophoresis with known amounts of BSA. The concentration of VP2 in the open core preparations is usually 100–300 µg/mL. 20. The 32P-labeled dsRNA products are detected by autoradiography, and can be quantified with a phosphorimager. Shown in Fig. 2 are the 32P-labeled dsRNA products made in reaction mixtures that contained gene 8 mRNA and untreated open cores (lane 2) or micrococcal nuclease-treated open cores (lane 4). 21. The product of assays performed with open cores that have not been treated with micrococcal nuclease, and that lack exogenous ssRNA will include low levels of 32P-labeled dsRNA (see Fig. 2, lane 1). 22. Synthesis of dsRNA will occur in the replicase assay if the open cores are replaced with purified rVP1 and rVP2 or CLPs containing rVP1 and rVP2. Synthesis of dsRNA with recombinant proteins is optimal when the molar ratio of rVP1 and rVP2 in the assay is approx 1:10 (8). The synthesis of dsRNA by rVP1 and rVP2 is much less efficient than the synthesis of dsRNA by open cores. 23. Synthesis of dsRNA by the replicase assay decreases as the level of salt in the reaction mixture increases (Fig. 3). At 200 mM NaCl, the amount of dsRNA produced in the assay will be at least 10-fold less than the amount of dsRNA produced in the control reaction mixture. 24. Formation of the VP3–GMP adduct is not affected by NaCl concentrations of 1–200 mM, but the presence of NaCl in the reaction mixture prevents the radiolabeling of VP1 with 32P-GTP (Fig. 3). 25. The 5'-terminal base of the nascent RNA can be either A or G. Like the formation of VP3–GMP intermediate complexes, the capping of RNA by open cores is a MgCl2-dependent process (Fig. 3). However, unlike the formation of VP3-GMP intermediate complexes, the capping of RNA by open cores is inhibited by the addition of NaCl to the assay. 26. Subheading 3.5.2. can be used to detect the RNA-binding activity of the RNA polymerase, VP1, and the guanylyltransferase, VP3, in open core preparations. Because both VP1 and VP3 have nonspecific affinity for RNA, any probe of 50–100 nucleotides can be used in the gel-shift assay. However, because there is evidence that VP1 also binds specifically to the 3'-end of viral mRNA, the authors generally use a probe that contains the 3'-terminal sequence of one of the viral mRNAs (19). 27. The temperature and length of incubation of the mixtures are not critical for the formation of RNA–protein complexes containing VP1 and VP3, but the authors normally incubate the mixtures for 60 min at room temperature. 28. The time of electrophoresis will vary from 3–6 h, depending on the length of the probe, and should be performed until free (unbound) probe, migrates near, but not off, the bottom of the gel. 29. Bands of probe that migrate in the gel more slowly than free probe indicate the presence of protein–probe complexes, and suggest that a protein with RNA-binding activity is contained in the reaction mixture. To verify that novel bands detected by the gel-shift assay do indeed represent complexes containing RNA and
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31.
32.
33.
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Patton et al. protein, reaction mixtures can be treated with 10 µg/mL RNase A and proteinase K, respectively, for 30 min at room temperature prior to electrophoresis. Crosslinking of radiolabeled nucleotides of the probe to the protein generates a 32P-labeled protein. The 32P-labeled protein may migrate slower in the gel, because the cross-linked nucleotides will add to the total mass of the protein (usually <1 kDa). The identity of the 32P-labeled protein can be verified by Western blot or immunoprecipitation assay using monospecific antiserum. UV-crosslinking has been used successfully to show that VP1, NSP2, and NSP3 have RNA-binding activity (24,26,41). BL2 experimentation: CAUTION: All procedures with vaccinia virus must be carried out using NIH-Biosafety Level 2 Guidelines. Smallpox vaccination is recommended for personnel who potentially may be exposed to vaccinia virus. VP4 and VP7 contain intramolecular disulfide bonds, which cause these proteins to migrate more rapidly when reducing agents are not added to the SDS-PAGE sample buffer (42). There is also some evidence that VP6 may also contain intrachain disulfide bonds (42). The intermediate shell of the RV triple-layered virion is formed by 260 trimers of VP6. The trimers are quite stable and are also formed by VP6 that is expressed by BV vectors containing the gene 6 cDNA (43), and by translation of the gene 6 mRNA in rabbit reticulocyte lysates (44). To detect trimers, VP6-containing preparations are incubated in SDS-PAGE sample buffer at 37°C for 5 min, and are then electrophoresed on an SDS-12% polyacrylamide gel. The trimer form of VP6 will migrate with a mol wt of 135–150 kDa. If, instead, the sample is heated to 100°C for 5 min, the VP6 trimers will be disrupted, and the VP6 will then migrate as a 45 kDa monomer upon SDS-12% PAGE. Viral mRNAs are denatured by heating in 2X formamide sample buffer. The samples are then electrophoresed on 5% polyacrylamide gels containing 7 M urea (100 × 100 × 0.75 mm) in 1X TBE running buffer at 200 V for 5 h. The mRNAs can be detected in the gel by staining with ethidium bromide or by silver staining.
References 1. Graham, D. Y. and Estes, M. K. (1980) Proteolytic enhancement of rotavirus infectivity: biological mechanisms. Virology 101, 432–439. 2. Espejo, R. T., Lopez, S., and Arias, C. (1981) Structural polypeptides of simian rotavirus SA11 and the effect of trypsin, J. Virol. 37, 156–160. 3. Cohen, J., Laporte, J., Charpilienne, A., and Scherrer, R. (1979) Activation of rotavirus RNA polymerase by calcium chelation. Arch. Virol. 60, 177–186. 4. Shirley, J. A., Beards, G. M., Thouless, M. E., and Flewett, T. H. (1981) The influence of divalent cations on the stability of human rotavirus. Arch. Virol. 67, 1–9. 5. Bican, P., Cohen, J., Charpilienne A., and Scherrer, R. (1982) Purification and characterization of bovine rotavirus cores. J. Virol. 43, 1113–1117. 6. Almeida, J. D., Bradburne, A. F., and Wreghitt, T. G. (1979) The effect of sodium thiocyanate on virus structure. J. Med. Virol. 4, 269–277. 7. Valenzuela, S., Pizarro, J., Sandino, A. M., Vasquez, M., Fernandez, J., Hernandez, O., Patton, J., and Spencer, E. (1991) Photoaffinity labeling of
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rotavirus VP1 with 8-azido-ATP: Identification of the viral RNA polymerase. J. Virol. 65, 3964–3967. Patton, J. T., Jones, M. T., Kalbach, A. N., He, Y.-W., and Xiaobo, J. (1997) Rotavirus RNA polymerase requires the core shell protein to synthesise the double-stranded RNA genome. J. Virol. 71, 9618–9626. Cohen, J. (1977) Ribonucleic acid polymerase activity associated with purified calf rotavirus. J. Gen. Virol. 36, 395–402. Chen, D. Y., Zeng, C. Q.-Y., Wentz, M. J., Gorziglia, M., Estes, M. K., and Ramig, R. F. (1994) Template dependent, in vitro replication of rotavirus RNA. J. Virol. 68, 7030–7039. Mason, B. B., Graham, D. Y., and Estes, M. K. (1980) In vitro transcription and translation of rotavirus SA11 gene products. J. Virol. 33, 1111–1121. Sandini, A. M., Jashes, M., Faundez, G., and Spencer, E. (1986) Role of the inner protein capsid on in vitro human rotavirus transcription. J. Virol. 60, 797–802. Pizarro, J. L., Sandino, A. M., Pizarro, J. M., Fernandez, J., and Spencer, E. (1991) Characterization of rotavirus guanylyltransferase activity associated with polypeptide VP3. J. Gen. Virol. 72, 325–332. Liu, M., Mattion, N. M., and Estes, M. K. (1992) Rotavirus VP3 expressed in insect cells possesses guanylyltransferase activity. Virology 188, 77–84. Desselberger, U. and McCrae, M. A. (1994) The rotavirus genome. Curr. Top. Microbiol. Immunol. 185, 31–66. Patton, J. T., Wentz, M., Xiaobo, J., and Ramig, R. F. (1996) Cis-acting signals that promote genome replication in rotavirus mRNAs. J. Virol. 70, 3961–3971. Wentz, M. J., Patton, J. T., and Ramig, R. F. (1996) The 3'-terminal consensus sequence of rotavirus mRNA is the minimal promoter of negative-strand RNA synthesis. J. Virol. 70, 7833–7841. Chen, D. and Patton, J. T. (1998) Rotavirus RNA replication requires a single-stranded 3'-terminus for efficient minus strand synthesis. J. Virol. 72, 7387–7396. Patton, J. T. (1996) Rotavirus VP1 alone specifically binds to the 3'-end of viral mRNA but the interaction is not sufficient to initiate minus-strand synthesis. J. Virol. 70, 7940–7947. Labbé, M., Baudoux, P., Charpilienne, A., Poncet, D., and Cohen, J. (1994) Identification of the nucleic acid binding domain of the rotavirus VP2 protein. J. Gen. Virol. 75, 3423–3430. Patton, J. T. and Chen, D. (1999) RNA binding and capping activities of proteins in rotavirus open cores. J. Virol. 73, 1382–1391. Shuman, S. (1995) Capping enzyme in eukariotic mRNA synthesis. Prog. Nucleic Acids Res. Mol. Biol. 50, 101–129. Patton, J. T. (1996) Structure and function of the rotavirus RNA-binding proteins. J. Gen. Virol. 76, 2633–2644. Brottier, P., Nandi, P., Bremont, M., and Cohen, J. (1992) Bovine rotavirus segment 5 protein expressed in the baculovirus system interacts with zinc and RNA. J. Gen. Virol. 73, 1931–1938. Kattoura, M., Clapp, L. L., and Patton, J. T. (1992) The rotavirus nonstructural protein, NS35, is a nonspecific RNA-binding protein. Virology 191, 698–708. Poncet, D., Aponte, C., and Cohen, J. (1994) Four nucleotides are the minimal requirement for RNA recognition by rotavirus non-structural protein NSP3. EMBO J. 13, 4165–4173. Boyle, J. F. and Holmes, K. V. (1986) RNA-binding proteins of bovine rotavirus. J. Virol. 58, 561–568.
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28. Hua, J., Chen, X., and Patton, J. T. (1994) Deletion mapping of the rotavirus metalloprotein NS53 (NSP1): The conserved cysteine-rich region is essential for virus-specific RNA binding. J. Virol. 68, 3990–4000. 29. Patton, J. T. and Hua, J. (1995) Using the RNA-capture assay to assess the RNA-binding activity of viral proteins, in Methods in Molecular Genetics, vol. 7: Molecular Virology Techniques (Adolph, A. W., ed.), Academic Press, New York, p. 373–387. 30. Poncet, D., Aponte, C., and Cohen, J. (1996) Structure and function of rotavirus nonstructural protein NSP3. Arch. Virol. 12, Suppl. S29–S35. 31. Zeng, Q.-Y., Wentz, M. J., Estes, M. K., and Ramig, R. F. (1996) Characterization and replicase activity of double-layered and single-layered rotavirus-like particles expressed from baculovirus recombinants. J. Virol. 70, 2736–2742. 32. Crawford, S. E., Labbé, M., Cohen, J., Burroughs, M. H., Zhou, Y. J., and Estes, M. K. (1994) Characterisation of virus-like particles produced by the expression of rotavirus capsid proteins in insect cells. J. Virol. 68, 5945–5952. 33. Labbé, M., Charpilienne, A., Crawford, S. E., Estes, M. K., and Cohen, J. (1991) Expression of rotavirus VP2 produces empty corelike particles. J. Virol. 65, 2946–2952. 34. Lawton, J. A., Zeng, C. Q., Mukherjee, S. K., Cohen, J., Estes, M. K., and Prasad, B. V. (1997) Three-dimentional structural analysis of recombinant rotavirus-like particles with intact and amino-terminal-deleted VP2: implications for the architecture of the VP2 capsid layer. J. Virol. 71, 7353–7360. 35. Zeng, C. Q.-Y., Estes, M. K., Charpilienne, A., and Cohen, J. (1998) The N terminus of rotavirus VP2 is necessary for encapsidation of VP1 and VP3. J. Virol. 72, 201–208. 36. Fuerst, T. R., Niles, E. G., Studier, F. W., and Moss, B. (1986) Eukaryotic transient-expression system based on recombinant vaccinia virus that synthesizes bacteriophage T7 RNA polymerase. Proc. Natl. Acad. Sci. USA 83, 8122–8126. 37. Zeng, C.Q.-Y., Labbé, M., Cohen, J., Prasad, B. V. V., Chen, D., Ramig, R. F., and Estes, M. K. (1994) Characterization of rotavirus VP2 particles. Virology 201, 55–65. 38. Cochran, M. A., Puckett, C., and Moss, B. (1985) In vitro mutagenesis of the promoter region for a vaccinia virus gene: evidence for tandem early and later regulatory signals. J. Virol. 54, 30–37. 39. Patton, J. T. and Stacey-Phipps, S. (1986) Electrophoretic separation of the plus and minus-strands of rotavirus SA11 double-stranded RNAs, J. Virol. Methods 13, 185–190. 40. Chen, D., Gombold, J. L., and Ramig, R. F. (1990) Intracellular RNA synthesis directed by temperature-sensitive mutants of simian rotavirus SA11. Virology 178, 143–151. 41. Poncet, D., Aponte, C., and Cohen, J. (1993) Rotavirus protein NSP3 (NS34) is bound to the 3’ end consensus sequence of viral mRNAs in infected cells. J. Virol. 67, 3159-3165. 42. Patton, J. T., Hua, J., and Mansell, E. A. (1993) Location of interchain disulfide bonds in VP5* and VP8* trypsin cleavage fragments of the rhesus rotavirus spike protein VP4. J. Virol. 67, 4848–4855. 43. Estes, M. K., Crawford, S. E., Penaranda, M. E., Petrie, B. L., Burns, J. W., Chan, W.-K., et al. (1987) Synthesis and immunogenicity of the rotavirus major capsid antigen using a baculovirus expression system. J. Virol. 61, 1488–1494. 44. Clapp, L. L. and Patton, J. T. (1991) Rotavirus morphogenesis: domains in the major inner capsid protein essential for binding to single-shelled particles and for trimerization. Virology 180, 697–708.
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4 Rotavirus Entry into Tissue Culture Cells Joanna M. Gilbert and Harry B. Greenberg 1. Introduction Rotavirus (RV) is a triple-protein-layered icosahedral virus, for which studies have established that the two outer-layer proteins, viral protein 4 (VP4) and viral protein 7 (VP7), are required for viral infectivity (1,2). VP7, a glycoprotein, is the major component of the outer-layer, but its role in viral entry is unclear. VP4 forms dimers extending out from the VP7-coated viral surface (3,4) and have been shown to be a determinant of host range and virulence, and is directly involved in cell attachment and RV entry into cells (5–8). Proteolytic cleavage of VP4 into two noncovalently associated subunits, VP8* and VP5* (2,9,10), significantly enhances viral infectivity (11–13). The mechanism of cell penetration during infection is not fully understood, but it probably consists of multiple steps, including binding, protease cleavage, and entry, in which binding is independent of the other two steps, and can occur without infection. VP4 is intimately involved in each of these steps of RV entry. Experiments have demonstrated that RV enters cells in a pH-independent fashion, and most, but not all, of the evidence indicates that RV penetrates cells directly through the plasma membrane rather than through the early endosomal membrane after receptor-mediated endocytosis (6,14–16). This chapter describes two different assays that measure the entry of RV into tissue culture cells: a cell-cell fusion (syncytia) assay (Fig. 1;17) and a toxin co-entry assay (Fig. 2;18–20). It is hypothesized that the ability of RV to induce both cell-to-cell fusion and toxin co-entry is related to its ability to penetrate the plasma membrane during an infection. The former assay, examining RV-induced cell fusion-from-without, allows direct visualization of the results of virus–cell interaction: RV-induced fusion of cellular plasma membranes. The toxin co-entry assay examines viral entry by determining the extent of α-sarcin From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ
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Fig. 1. Syncytia assay.
ingress into cells, concomitant with RV penetration. This toxin disrupts protein translation, and therefore allows quantitation of viral entry by measuring the decrease of protein synthesis (21,22). Both assays require cleavage of VP4 by the protease trypsin, and also depend on the presence of VP4 and VP7; particles depleted of both outer layer proteins are nonfunctioning in these assays (15–17,19,20,23). The assays can differentiate between cells that are permissive for RV infection from cells that are not permissive (19,20,23). Monoclonal antibodies, which neutralize RV infectivity, can be demonstrated to block both assays (17,19,20). Therefore, these assays provide us with methods to identify factors involved in RV entry into cells.
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Fig. 2. α-Sarcin coentry assay.
One major difficulty impeding the understanding of RV entry into cells is that a method to alter a specific RV gene product, and recover it in infectious virus, is not yet available. For molecular analysis, the authors have therefore employed VLPs (24) as an alternative to intact RV particles. VLPs are formed by co-infecting Spodoptera frugiperda 9 (Sf9) cells with four different recombinant baculoviruses (rBVs), each expressing one of the four main structural proteins of RV (VP2, VP4, VP6, or VP7). The co-expressed RV proteins assemble into particles that are similar to intact RV particles by electron microscopic, biochemical, and immunological criteria (24). The VLPs have also been shown to bind specifically to MA104 cells (24) and to function identically to intact RV particles in these entry assays (19,20,23).
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Ultimately, there are several major questions concerning the mechanism of RV entry into cells. Using recombinant VLPs, containing either VP4 or VP7 proteins engineered with specific mutations, allows the examination of the role of these proteins in RV entry at a molecular level (23). For example, recent work (25) has demonstrated that arginine residue 247 is the critical site for the proteolytic activation of VP4 required for penetration. VP4 is the primary protein involved in entry and contains two provocative sequence motifs that are hypothesized to be involved in virus penetration of host cells (26). One of these motifs is a region that has sequence similarity to the putative internal fusion peptides of Sindbis virus and Semliki Forest virus. This domain is proposed to form an amphipathic α-helix, with the hydrophobic face interacting directly to disrupt the cell plasma membrane. Using site-directed mutagenesis to reduce the hydrophobicity of this helix would allow one to examine the role of this region in RV entry. Fusion peptides have only been demonstrated to function in enveloped viruses. The second motif is a putative α-helical, coiled-coil domain that is conserved among viruses as diverse as retroviruses, alphaviruses, and paramyxoviruses (27). Using specifically mutated VLPs, whether this region is required for VP4 dimerization, or has a role in viral entry, can now be determined. Finally, it appears that there is a functional interaction between VP4 and VP7 (23,28), and they may cooperate in order for RV to penetrate host cells. The contributions of these individual proteins could be assessed using site-directed mutagenesis and one of the above entry assays. 2. Materials 1. The fetal African green monkey kidney cell line, MA104 (BioWhittaker, Walkersville, MD). 2. Dulbecco’s modified Eagle’s medium (DMEM; BioWhittaker), with 4.5 g/L glucose containing 100 IU/mL penicillin, 100 IU/mL streptomycin, 0.29 mg/mL L-glutamine (Irvine Scientific, Santa Ana, CA), 0.25 mg/mL amphotericin B (Gibco-BRL, Gaithersburg, MD), and 15, 10, 7%, or no heat-inactivated fetal bovine serum (hiFBS; Hyclone, Logan, UT), as indicated. 3. DMEM without cysteine or methionine (DMEM cys– met–; Gibco-BRL), with 4.5 g/L glucose containing 100 IU/mL penicillin, 100 IU/mL streptomycin, and 0.29 mg/mL L-glutamine. 4. Dulbecco’s phosphate buffered saline (DPBS) and DPBS calcium- and magnesiumfree (CMF; Gibco-BRL). 5. 0.5% trypsin in Versene (sodium ethylenediamine tetraacetic acid [NaEDTA], Gibco-BRL). 6. Trypsin (10 mg/mL in media without serum; Sigma, St. Louis, MO.; Type XIII, N-tosyl-L-phenylalanine chloromethyl ketone [TPCK]-treated). 7. N-α-p-tosyl-L-lysine chloromethyl ketone: 10 mM in media without serum (TLCK; Sigma). 8. 10 mM cholesterol (Sigma), freshly made in 100% ethanol.
RV Entry into Tissue Culture Cells 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25.
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α-sarcin (1 mg/mL in DMEM cys– met–, stored at –20°C; Sigma). Translabel (35S cysteine and methionine; ICN, Irvine, CA). 10% trichloroacetic acid (TCA). Absolute ethanol. 0.5% sodium dodecylsulfate (SDS) in 0.1 M sodium hydroxide (NaOH). 1 M hydrochloric acid (HCl). Aqueous scintillation fluor. The insect cell line derived from Sf9 (Invitrogen; San Diego, CA). Sf-900 II media (Gibco-BRL) supplemented with 100 IU/mL penicillin, 100 IU/mL streptomycin, 0.29 mg/mL L-glutamine, and containing either 2.5% or no hiFBS. 2X Graces media (Gibco-BRL). 2% low-melt agarose in double-distilled H2O (Gibco-BRL). The protease inhibitors pepstatin, aprotinin, and leupeptin (0.5 µg/mL; Sigma). Genentron (trichlorotrifluoroethane) (Dupont, Wilmington, DE. 10 mM Tris-HCL (Tris [hydroxymethyl] aminomethane), pH 7.4, (Dupont, Wilmington, DE) 100 mM NaCl, 5 mM CaCl 2 (TNC). 35% sucrose (w/v) in TNC. Cesium chloride (CsCl) (1.36 g/mL) in TNC. The BAC to BAC system rBV system (Gibco-BRL) containing the pFASTBAC subcloning and transposition vector and the DH10BAC Escherichia coli bacteria containing the BV genome.
3. Methods 3.1. Syncytia Assay 1. Plate 5 × 10 5 MA104 cells in 25-cm 2 flasks in 5 mL DMEM with 10% hiFBS, in an atmosphere of 5% CO 2-air at 37°C. 2. After 24 h, replace medium with 5 mL DMEM with 7% hiFBS containing 100 µM cholesterol, from the 10 mM cholesterol in ethanol stock, and incubate overnight (14–16 h) at 37°C in a CO 2 incubator (see Note 1). 3. Treat the RV or RV VLPs (see Subheading 3.3.) with 10 mg/mL trypsin at a final concentration of 20 µg/mL for 30 min at 37°C. Inactivate trypsin by adding TLCK to a final concentration of 100 µM. 4. Remove the cholesterol-supplemented cells from the flasks, first by washing with DPBS CMF, and then by treating with 1% trypsin in versene. 5. Centrifuge the cells at 1000g at room temperature. 6. Resuspend the cells in DMEM at 1 × 10 6 cells/mL and add 100 µL, containing 1 × 10 5 cells, to each 1.5-mL Eppendorf tube on ice. 7. Centrifuge samples for 30 s at 400g at 4°C, then carefully aspirate the supernatant without disturbing the cell pellet (see Note 2). 8. Resuspend the cell pellet in 100 µL virus in DMEM, on ice, at a dilution of 7.5 × 107 plaque forming units (PFU)/mL to 1 × 108 PFU/mL (approx 75–100 particles/cell), or the protein equivalent of VLPs (e.g., for a CsCl purified, titered stock of wild-type rhesus RV with a determined protein concentration
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Gilbert and Greenberg of 2.0 × 10 9 PFU/mg protein, 3.75 µg VLP protein is equivalent to approx 75 PFU for 1 × 10 5 cells). Incubate the samples on ice for 15 min, to allow the virus to bind to the cells. Centrifuge the tubes at 400g for 15 s, and then incubate at 37°C for 15 min. Gently resuspend the cell and virus pellet after addition of 400 µL DMEM. Plate the cells into six-well tissue culture dishes containing 1 mL/well DMEM with 15% hiFBS (10% hiFBS final). Incubate the plates for 2 h at 37°C in an atmosphere of 5% CO2 -air (see Note 3). Examine the wells for fusion, with an inverted phase-contrast microscope. Cells containing two or more nuclei are considered to have formed syncytia: Cells with two nuclei have undergone one fusion event; cells with three nuclei have undergone two fusion events, and so on. (see Note 4). Samples are tested in duplicate, and a minimum of 500 nuclei are counted per sample (five sets of approx 100 nuclei each) and the number of nuclei in polykaryons noted. Data are averaged and presented as number of nuclei in syncytia per 100 total nuclei counted.
3.2. α -Sarcin Assay 1. Plate 5 × 10 3 MA104 cells/well into 96-well dishes in DMEM with 10% hiFBS, and incubate at 37°C in an atmosphere of 5% CO2-air for 24–48 h (to confluency at approx 2.5 × 10 4 cells/well). 2. Aspirate the media and add DMEM cys – met –, and incubate at 37°C in an atmosphere of 5% CO2 for 1 h. 3. Treat the RVs or RV VLPs (see Subheading 3.3.) with 10 mg/mL trypsin at a final concentration of 10 µg/mL for 30 min at 37°C. Inactivate the trypsin by adding TLCK to a final concentration of 100 µM. 4. Aspirate the media, and incubate the cells with trypsin-activated virus or VLPs (at the amount or protein equivalent between 50 and 100 PFU/cell; J. Angel, personal communication) in DMEM cys – met –, in a volume of 50 µL, and add 50 µL α-sarcin in DMEM cys– met– at a final concentration of 100 µg/mL. 5. Incubate the cells at 37°C for 1 h in an atmosphere of 5% CO 2-air. 6. Remove the virus and α-sarcin, and wash the plates twice with DMEM cys – met – . 7. Overlay the cells with 100 µL DMEM cys – met – containing 0.1 µCi 35S Translabel/well, and incubate at 37°C for 1 h in an atmosphere of 5% CO 2. 8. Remove the media, and wash the plates twice with DMEM cys– met –. 9. Add 100 µL/well 10% TCA, and incubate at 4°C for 1 h (see Note 5). 10. Remove the TCA, and wash the wells twice with ethanol. Let the plates air-dry. 11. Lyse the cell monolayers with 50 µL/well 0.5% SDS in 0.1 M NaOH, and incubate at room temperature for 1 h (see Note 6). 12. Neutralize the NaOH with 5 µL 1 M HCl/well. Add the lysates to the scintillation fluor, and count samples. Data are expressed as a percent inhibition of protein synthesis, compared with control, untreated cells (see Notes 7 and 8).
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3.3. VLP Production 3.3.1. Cell Growth Sf9 cells are grown in monolayer culture in Sf-900 II media with 2.5% hiFBS in a humidified incubator at 28°C. Cells are subcultured by passaging at a 1 in 4 dilution every 3–4 d.
3.3.2. Virus Titration 1. BV stocks are grown in Sf9 cells in Sf-900 media without serum, to titers between 1 × 10 8 and 1 × 10 9 PFU/mL. To titer the virus, plate Sf9 cells into 6-well dishes at 2 × 10 6 cells/well for 1 h at 28°C in Sf-900 II media without serum (see Note 9). 2. Dilute the virus to be titered in 1.5 mL media without serum from 1 in 10–1 to 1 in 10–9 in 12 mm × 75 mm polystyrene fluorescence-activated cell sorter tubes. 3. Infect the Sf9 cells with 600 µL diluted virus mixture for 90 min at 28°C, rocking gently. 4. Aspirate media, and overlay with agar mix (1:1 2% low-melt agarose: 2X Graces’ media with 10% hiFBS), and allow the agar to set at room temperature for 20 min. Incubate at 28°C for 7–10 d, or until plaques are clearly visible (see Note 10).
3.3.3. Plaque Purification 1. To plaque-purify the virus, pick well-spaced plaques, and add them to microcentrifuge tubes containing 300 µL Sf-900 II media without serum. 2. Gently vortex the microcentrifuge tubes, and infect 0.5 × 106 Sf9 cells/well grown in 12-well dishes. Incubate the dishes at 28°C for 90 min, and then overlay with 1 mL Sf-900 II media without serum. 3. Incubate the virus and the cells until infection is complete (7–10 d). Harvest the supernatants containing the purified BV aseptically, and analyze the remaining cells for expression of the desired protein.
3.3.4. Virus Growth 1. To grow up virus, add 100 µL positive viral supernatant to a 25-cm2 flask containing 2.5 × 106 Sf9 cells in 750 µL Sf-900 II media without serum. Incubate the flask for 90 min at 28°C, rocking occasionally. 2. After incubation, overlay the cells with a final volume of 6 mL Sf-900 II media without serum. 3. Harvest the supernatant after all the cells are infected, approx 6–10 d at 28°C. Spin down the cell debris at 2000g, and store the supernatant at 4°C in the dark. For long-term storage, add 2% hiFBS, and store at –70°C. Virus stocks can be amplified from this initial stock.
3.3.5. VLP Preparation 1. Infect monolayers of Sf9 cells with the desired combination of recombinant rBVs (for the triple-layered particles required for entry studies, VP2, VP6,
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4. 5.
6.
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8. 9. 10.
Gilbert and Greenberg VP4, and VP7), at a multiplicity of infection (MOI) of 6, in Sf-900 II media in the absence of serum. Incubate the cells for 24–36 h at 28°C, and then remove the inoculum by aspiration. Overlay the cells with Sf-900 II media containing 0.5% hiFBS and the protease inhibitors pepstatin, aprotinin, and leupeptin (0.5 µg/mL final concentration of each). Incubate the cells at 28°C, with the addition of protease inhibitors daily until the cells are harvested at 5–6 d postinfection. Remove the cells from the flasks by shaking, and rinse the flasks with DPBS. Pellet the cells by centrifugation at 3300g for 15 min at 4°C, and save the supernatant on ice. Extract the cell pellet with Genentron, and add the aqueous layer to the supernatant on ice. Centrifuge the samples, first at 12,000g, in a JA-14 rotor (Beckman, Fullerton, CA) for 30 min at 4°C, and then at 200,000g in a Ti 45 rotor (Beckman) for 2 h at 4°C. Resuspend the pellet in TNC containing (0.5 µg/mL) aprotinin, (0.5 µg/mL) leupeptin, and (0.5 µg/mL) pepstatin, and centrifuge through a 35% (w/v) sucrose cushion in TNC at 300,000g for 90 min at 4°C. Resuspend the VLP pellet in TNC containing CsCl (1.36 g/mL), and centrifuge at 250,000g for 24–36 h at 4°C in an SW-40 rotor (Beckman). The VLP-containing bands are collected, and recentrifuged through CsCl. Collect the VLP band. The particles can be dialyzed extensively against TNC, and stored at 4°C for use in the short term. For longer-term storage, the particles should be stored in aliquots at –70°C. The VLPs are stable in CsCl at 4°C.
3.4.6. rBV Production rBVs can be easily prepared using the BAC to BAC system available from Gibco-BRL. cDNAs for VP4 and/or VP7, containing the desired changes, can be subcloned into the pFASTBAC vector. The mutant cDNAs can be transposed into the BV genome, resulting in the desired recombinant viruses. The mutant viruses can then be used in place of either wild-type VP4 or wild-type VP7 to create mutant VLPs, in order to examine the function of these mutants in RV entry. 4. Notes 1. To solubilize cholesterol in ethanol, warm the tube in a 37°C water bath, vortexing occasionally. When the cholesterol is added to the DMEM with 7% hiFBS, the media will become slightly cloudy. 2. If cells have been treated with too much cholesterol, they will not pellet at this g force, nor will they plate well after syncytia formation, making quantitation difficult. 3. If cells are left too long after plating (longer than 3.5 h), they will begin to die. 4. It is vital to include, as controls, cells alone, and cells incubated with untrypsinized virus, to establish the background levels of syncytia formation. Cells alone will usually have an average of three nuclei in syncytia/100 nuclei
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counted. An average of 4.5 nuclei in syncytia/100 nuclei counted can be expected with cells treated with untrypsinized virus. For trypsin-treated virus at an MOI of 75, the level of syncytia seen is approx 18 nuclei in syncytia/100 nuclei counted. Samples in TCA can be left overnight at 4°C. Samples in 0.5% SDS/0.1 M NaOH can be left overnight at 4°C. It is vital to include as controls, cells alone, cells incubated with α-sarcin alone, cells with trypsinized virus without α-sarcin, and cells incubated with untrypsinized virus with α-sarcin, to establish the background levels. Translation is expected to be diminished by 90% using virus at an MOI between 75 and 100. An alternative procedure is to examine the loss of translation using SDS polyacrylamide gel electrophoresis (PAGE). The assay should be scaled up to 24-well dishes. After the cells are labeled and washed, they are lysed in buffer containing 130 mM NaCl, 20 mM Tris (pH 7.4), 1 mM EDTA, 1.0 % Nonidet P-40, and 1 mM phenylmethyl sulfonylfluoride, and cleared of cellular debris by centrifugation at 13,000g for 15 min. The lysates are boiled in an equal volume of sample buffer (0.25 M Tris-HCl, pH 7.4, 30% sucrose, 8% SDS, and 50 mM dithiothreitol) for 5 min. These samples are run on SDS-PAGE gels, and examined by autoradiaography (F. Liprandi, J. Angel, and N. Shulman, personal communications). It is vital that cells are at the appropriate density, or they will overgrow and make detection of plaques impossible. It is crucial that the final concentration of agarose is not greater than 1%. The temperature of the agarose overlay should also not be greater than 28°C, because Sf9 cells are temperature-sensitive.
References 1. Bridger, J. C. and Woode, G. N. (1976) Characterization of two particle types of calf rotavirus. J. Gen. Virol. 31, 245–250. 2. Estes, M. K., Graham, D. Y., Smith, E. M., and Gerba, C. P. (1979) Rotavirus stability and inactivation. J. Gen. Virol. 43, 403–409. 3. Prasad, B. V. V., Burns, J. W., Marietta, E. Estes, M. K., and Chiu, W. (1990) Localization of VP4 neutralization sites in rotavirus by three-dimensional cryo-electron microscopy. Nature 343, 476–479. 4. Shaw, A. L., Rothnagel, R., Chen, D., Ramig, R. F., Chiu, W., and Prasad, B. V. V. (1993) Three-dimensional visualization of the rotavirus hemagglutinin structure. Cell 74, 693–701. 5. Greenberg, H. B., Flores, J., Kalica, A. R., Wyatt, R. G., and Jones, R. (1983) Gene coding assignments for growth restriction, neutralization and subgroup specificities of the W and DS-1 strains of human rotavirus. J. Gen. Virol. 64, 313–320. 6. Kaljot, K. T., Shaw, R. D., Rubin, D. H., and Greenberg, H. B. (1988) Infectious rotavirus enters cells by direct cell membrane penetration, not by endocytosis. J. Virol. 62, 1136–1144. 7. Offit, P. A., Blavat, G., Greenberg, H. B., and Clark, H. F. (1986) Molecular basis of rotavirus virulence: role of gene segment 4. J. Virol. 57, 46–49.
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8. Ruggeri, F. M. and Greenberg, H. B. (1991) Antibodies to the trypsin cleavage peptide VP8 neutralize rotavirus by inhibiting binding of virions to target cells in culture. J. Virol. 65, 2211–2219. 9. Espejo, R. T., Lopez, S., and Arias, C. (1981) Structural polypeptides of simian rotavirus SA11 and the effect of trypsin. J. Virol. 37, 156–160. 10. Lopez, S., Arias, C. F., Bell, J. R., Strauss, J. H., and Espejo, R. T. (1985) Primary structure of the cleavage site associated with trypsin enhancement of rotavirus SA11 infectivity. Virology 144, 11–19. 11. Babiuk, L. A., Mohammed, K., Spence, L., Fauvel, M., and Petro, R. (1977) Rotavirus isolation and cultivation in the presence of trypsin. J. Clin. Microbiol. 6, 610–617. 12. Barnett, B. B., Spendlove, R. S., and Clark, M. L. (1979) Effect of enzymes on rotavirus infectivity. J. Clin. Microbiol. 10, 111–113. 13. Clark, S. M., Roth, J. R., Clark, M. L., Barnett, B. B., and Spendlove, R. S. (1981) Trypsin enhancement of rotavirus infectivity: mechanism of enhancement. J. Virol. 39, 816–822. 14. Fukuhara, N., Yoshie, O., Kitaoka, S., and Konno, T. (1988) Role of VP3 in human rotavirus internalization after target cell attachment via VP7. J. Virol. 62, 2209–2218. 15. Nandi, P., Charpilienne, A., and Cohen, J. (1992) Interaction of rotavirus particles with liposomes. J. Virol. 66, 3363–3367. 16. Ruiz, M. C., Alonso, T. S., Charpilienne, A., Vasseur, M., Michelangeli, F., Cohen, J., and Alvarado, F. (1994) Rotavirus interaction with isolated membrane vesicles. J. Virol. 68, 4009–4016. 17. Falconer, M. M., Gilbert, J. M., Roper, A. M., Greenberg, H. B., and Gavora, J. S. (1995) Rotavirus-induced fusion-from-without in tissue culture cells. J. Virol. 69, 5582–5591. 18. Carrasco, L. (1994) Entry of animal viruses and macromolecules into cells. FEBS Lett. 350, 151–154. 19. Cuadras, M. A., Arias, C. F., and Lopez, S. (1997) Rotaviruses induce an early membrane permabilization of MA104 cells and do not require a low intracellular Ca2+ concentration to initiate their replication cycle. J. Virol. 71, 9065–9074. 20. Liprandi, F., Moros, Z., Gerder, M., Ludert, J. E., Pujol, F. H., Ruiz, M. C., Michelangeli, F., Charpilienne, A., and Cohen, J. (1997) Productive penetration of rotavirus in cultured cells induces coentry of the translation inhibitor α-sarcin. Virology 237, 430–438. 21. Brigotti, M., Rambelli, F., Zamboni, M., Montanaro, L., and Sperti, L. (1989) Effect of α-sarcin and ribosome-inactivating proteins on the interaction of elongation factos with ribosomes. Biochem. J. 257, 723–727. 22. Endo, Y. and Wool, I. G. (1982) The site of action of α-sarcin on eukaryotic ribosome. J. Biol. Chem. 257, 9054–9060. 23. Gilbert, J. M. and Greenberg, H. B. (1997) Virus-like particle induced fusion-from-without in tissue culture cells; role of outer-layer proteins VP4 and VP7. J. Virol. 71, 4555–4563.
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24. Crawford, S. E., Labbé, M., Cohen, J., Burroughs, M. H., Zhou, Y. J., and Estes, M. K. (1994) Characterization of virus-like particles produced by the expression of rotavirus capsid proteins in insect cells. J. Virol. 68, 5915–5922. 25. Gilbert, J. M. and Greenberg, H. B. (1998) Cleavage of rhesus rotavirus VP4 after arginine 247 is essential for rotavirus-like particle-induced fusion-from-without. J. Virol. 72, 5323–5327. 26. Mackow, E. R., Shaw, R. D., Matsui, S. M., Vo, P. T., Dang, M.-N., and Greenberg, H. B. (1988) The rhesus rotavirus gene encoding protein VP3: Location of amino acids involved in homologous and heterologous rotavirus neutralization and identification of a putative fusion region. Proc. Natl. Acad. Sci. USA 85, 645–649. 27. Buckland, R. and Wild, F. (1989) Leucine zipper motif extends. Nature 338, 547–548. 28. Mendez, E., Arias, C. F., and Lopez, S. (1996) Interactions between the two surface proteins of rotavirus may alter the receptor-binding specificity of the virus. J. Virol. 70, 1218–1222.
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5 Mixed Infections with Rotaviruses Protocols for Reassortment, Complementation, and Other Assays Robert F. Ramig 1. Introduction Mixed infection of tissue culture cells is the primary means of studying genetic and nongenetic interactions between viral mutants. The purpose of the mixed infection is to place two different viral genomes into the same cell, where interactions between the genomes and their encoded gene products can take place. In some cases, interactions are observed in the context of the mixed infected cell, but generally the yield of progeny virus from the mixed infected cells is assayed to reveal the result of the interactions. Here, methods for mixed infections between conditional-lethal mutants of the temperature-sensitive (ts) phenotype will be presented and considered. (Mutants with ts phenotype are designated according to the following nomenclature: tsA [778], in which ts indicates the phenotype, A indicates the mutant belongs to recombination group A, and [778] indicates the number of the specific mutant in group A.) Temperature-sensitive mutants are particularly useful for genetic studies, because they can be propagated at the permissive temperature (PT; 31°C in the case of rotavirus [RV] ts mutants), but the mutant phenotype is expressed at the nonpermissive temperature (NPT; 39°C in the case of RV ts mutants). The conditional-lethality of the ts mutations allows the use of selective conditions in the analysis of the progeny of the mixed infections, so that the results of genetic and nongenetic interactions can be easily revealed.
1.1. Mixed Infections for Recombination (Reassortment) Tests Along with other members of the Reoviridae, RVs have a segmented genome (1), which results in a high frequency of recombination by the mechanism of From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ
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reassortment of genome segments. Thus, mutations that reside on noncognate segments are expected to reassort at a high frequency; mutations that reside on cognate segments are expected to be unable to reassort. The all-or-none nature of recombination in the RVs confirms reassortment as the recombination mechanism (2,3). In recombination tests, one determines if two different mutant genomes can interact to form viable genomes with new combinations of genetic material. To allow the genomes of mutants tsA and tsB to interact, the mixed infection is performed at PT, so that the mutants can replicate and increase the size of the intracellular genome pools which, in turn, increases the probability of recombinational interaction. The formation of wild-type (WT, also called ts+; RV WT grows at both 31 and 39°C) recombinant genomes from the two ts mutant genomes is generally assayed, because this progeny class is easily detected among the yield (progeny virus that results from the replication of the parental viruses during a mixed or single infection) of the cross, which also contains the parental ts mutants (used to initiate mixed infections) and the tsA–tsB double mutant. The WT recombinant class is detected at NPT at which neither the ts parental viruses nor the double mutant recombinant can grow. The recombination test is used to divide the ts mutants of RV into groups. Mutations within a group fail to generate WT reassortants in mixed infection, but mutations in different groups generate WT reassortants at high frequency (4–8). Because genome segments of the Reoviridae contain single genes, all mutations on a single genome segment (reassortment group) reside in the same gene or in its cis-acting regulatory sequences.
1.2. Mixed Infections for Complementation Tests Complementation tests are performed to determine if two mutant viruses contain lesions in the same function. With ts mutants, the mixed infection of tsA with tsB is performed at NPT, so that the mutant phenotype is expressed. At NPT in the mixed infection, neither the tsA or tsB parental virus can replicate, unless tsB can provide the function that is mutant in tsA and tsA can provide the function that is mutant in tsB. This requires that the mutant function in tsA can be supplied in trans by the nonmutant function of tsB (and vice versa), and results both in growth of mutants and the production of a yield that is primarily the two mutants. Transcomplementation is detected by assaying the yield at PT, and documenting an increase in yield over the infection of each parent alone. Failure of the mixed infection to increase the yield indicates that both mutants contained lesions in the same function. The segmented nature of the RV genome, with each segment encoding a single gene product, suggests that the gene products function in trans, and that complementation should be efficient. However, complementation is not useful
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for assignment of RV mutants to functional groups (3), because results of complementation analyses are generally negative and irreproducible (5). The failure of RV mutants to complement is ascribed to the interference phenotype associated with all of the mutants (6,7).
1.3. Interference Tests Mixed infections between a ts mutant and WT at either PT or NPT are used to determine if the ts mutant has an interference phenotype (6,7). The fact that all ts mutants tested interfered with the growth of WT suggests that ts mutants also interfere with each other in mixed infections, which is thought to account for failure to complement (see Subheading 1.2.). Interference tests are performed to determine if a virus mutant interferes with growth of another virus. Interference tests are done between a ts mutant and WT virus, because interference can be easily detected as a decrease in yield of WT from the mixed infection, compared to a control infection of WT alone. It is much more difficult to determine the interference of one ts mutant, with another ts mutant, because neither parental virus can be easily scored. To determine the interference phenotype of a ts mutant, mixed infections are performed between the ts mutant and WT, together with control infections of the ts mutant and WT alone (6,7). The mixed infections are incubated at either 31 or 39°C, depending on the condition at which the interference phenotype is of interest. The yield is determined at 39°C, at which only WT (ts+) virus will grow. An interfering mutant reduces the growth of the WT virus in the mixed infection, compared to the control single infection of WT. Interference tests are often performed at equal and unequal multiplicity of infection (MOI), and an interfering mutant generally shows a stronger interference phenotype when it dominates the input mixture (6,7). (MOI is the number of plaque-forming units [PFU]/cell used to initiate an infection). PFUs are the number of infectious units present in a virus stock, measured by their ability to form plaques on cell monolayers; generally expressed as PFU/mL. The number of PFU/mL in a virus stock is the titer of that stock, and is nearly always determined at PT.)
1.4. Generation of Reassortant Viruses Mixed infections between WT RV strains are used to generate reassortant viruses that contain defined constellations of genome segments derived from the two parental strains. The resulting reassortants are useful in studies to map expression of various phenotypes to specific genome segments or constellations of genome segments (see refs. 2 and 3 for examples). The methods for generation and selection of reassortant viruses are described in detail elsewhere (9).
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1.4.1. Nonselective Mixed Infection for Random Isolation of Reassortant Progeny Viruses Reassortant viruses, with carefully defined constellations of genome segments derived from two different virus strains, are usually generated by mixed infection of cultured cells. For these infections, it is best to use WT, nonmutagenized viruses as parents, because viral mutants selected following mutagenesis often contain unknown mutations that can affect the phenotype of reassortants in unpredictable ways (10). Because the parental viruses are WT, both the mixed infections and subsequent plaque assays for isolation of reassortant progeny are performed at the nonselective temperature of 37°C. The absence of selective markers in the mixed infections, and the absence of selective pressures during isolation of progeny reassortant, means that a large number of randomly chosen progeny plaques must be screened to identify the reassortant with the segment constellation of interest. When screening progeny from a mixed infection for reassortants, the host cell used can affect the spectrum of reassortant constellations isolated. Evidence has been presented that the host cell used for plaque formation can affect the spectrum of reassortant constellations isolated (11,12). Thus, a failure to isolate the desired reassortant constellation from a large number of screened plaques may be remedied by changing the cell line from which the plaques for screening are picked. Alternatively, a genome segment that is absent or underrepresented may be increased in representation in progeny by the use of unequal MOI for the mixed infections (11).
1.4.2. Mixed Infection with Selection for Reassortants with Defined Segment Constellation Often, mixed infections are performed with the intention of isolating reassortants with a specific constellation of parental genome segments. This often occurs when generating RV reassortants with viral protein VP4 and VP7 neutralization proteins in heterologous combination, relative to either parental virus. In such a case, a standard mixed infection is performed using the nonselective temperature of 37°C. The plaque assay from which plaques will be picked is overlaid with agar containing a neutralizing monoclonal antibody (MAb) directed at the VP4 or VP7 that one does not want in the reassortants. This antibody (Ab) selection during plaque formation selects against one of the parental viruses, and against reassortants that contain the protein being selected against. This significantly reduces the number of progeny plaques that must be screened to identify the reassortant with the
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segment constellation of interest. However, caution must be exercised in the use of such selective techniques, because reassortant viruses may express phenotypes that are not consistent with parental origins of the proteins (3). In one study (13), in a cross between RV strains SA11-4F and B223, a MAb directed against VP4 of SA11-4F was used to select reassortants with VP4 from the B223 parent. However, all of the reassortants isolated from this selection contained VP4 from SA11-4F (13). When the reassortant of interest was isolated using nonselective conditions (e.g., without selecting Ab), it was found to be neutralized by the Ab directed against SA11-4F VP4, although the reassortant contained VP4 from B223. Conversely, the reassortant containing SA11-4F VP4 was not neutralized by the Ab as expected (13). This unexpected presentation of neutralizing epitopes was demonstrated to result from interactions between VP4 and VP7 outer capsid proteins of heterologous parental origin (14). This example emphasizes the problems that may be encountered in the use of selective conditions when making reassortants with specific constellations of genome segments.
1.5. Mapping RV Mutants Mixed infections between RV ts mutants (15), or between a ts mutant and WT virus of a different strain (16), are used to map the ts mutations to genome segments. The ts phenotype of the ts parent(s) is counterselected to simplify identification of the ts+ reassortant progeny useful for mapping (15,16).
1.6. Mixed and Single Infections for Kinetic Analyses If mixed or single infections are desired for a kinetic analysis, the protocol outlined here is easily adapted. The preparation of vials for infection and preparation of activated virus stocks for infection are scaled-up, and multiple vials are infected with each single virus or virus mix from a single inoculum preparation. Vials are then incubated at the appropriate temperature, and individual vials of each single or mixed infection are harvested at the appropriate time postinfection. Single infections harvested in kinetic fashion have been used to generate highly reproducible one-step growth curves, and mixed infections have been used to examine the kinetics of reassortment (5–7). 2. Materials The sources and catalog numbers in the following list indicate items and sources the author has used with consistent success over the past 20 yr. Only with the 2-dram vials is the specific source and catalog number important. BioWhittaker (Walkersville, MD) is the only commercial source for MA104 cells.
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2.1. Cells and Viruses 1. African Green monkey kidney MA104 cells; (BioWhittaker, cat. no. 75-104F). For optimal results, MA104 cells should be used at passages lower than 100–120. 2. High-titer stocks of RV ts mutants and WT.
2.2. Equipment 1. Tissue culture incubators (Forma [Marietta, OH], Model 3110), humidified and flushed with 5% CO2-air. One each at 39, 37, and 31°C. 2. Water bath at 37°C. 3. Probe-type sonication device equipped with a microprobe for small volumes (Braun-Sonic [Melsungen, Germany] Model U, equipped with 3 mm diameter microprobe). Probe is sterilized between uses by immersion in virus killer (0.5 gm KI, 1.0 gm I 2 , dissolved in 1000 mL 95% ethanol) for 10 s; excess virus killer wiped off with tissue, immersion in sterile H 2O to briefly rinse. 4. Vacuum aspirator with trap. 5. Inverted microscope. 6. Hemocytometer.
2.3. Reagents 1. Medium 199 + fetal bovine serum (FBS). (Medium 199 with Earle’s salts, Irvine Scientific [Santa Ana, CA], cat. no. 9466) Resuspended and sterilized according to manufacturer’s instructions. The medium is then completed by making the following additions: 0.3 g/L glutamine; 1.22 g/L NaHCO3; 0.16 g/L penicillin; 0.25 g/L streptomycin; and 5% [v/v] FBS. 2. Medium 199 without FBS. Medium 199 completed as above, except FBS is omitted. 3. 2× Medium 199. Medium 199 with Earle’s salts resuspended at twice the concentration recommended by the manufacturer. For use, 2× 199 is supplemented with 0.6 g/L glutamine; 2.0 g/L NaHCO3; 0.16 g/L penicillin; 0.25 g/L streptomycin; 0.025 g/L gentamicin; 5000 U/L nystatin; and pancreatin (135 trypsin-activity U/L). 2× 199 should be supplemented with NaHCO 3 immediately prior to use, because precipitates rapidly form as complete medium is stored. 4. 2% Bacto-agar (Difco, Detroit, MI, cat. no. 0140-01): 20 g/L in distilled H2O and sterilized by autoclaving. 5. Gelatin-saline: 8.0 g/L NaCl, 0.03 g/L CaCl2, 0.17 g/L MgCl2, 1.2 g/L H3BO3, 0.05 g/L Na2B4O7, 5.0 g/L gelatin (pH 7.2). 6. 1 mg/mL trypsin, (Worthington, Freehold, NJ) dissolved in 0.001 N HCl, and stored in single-use 1.0-mL aliquots at –80°C. 7. Pancreatin (Oxoid, London, UK) One tablet dissolved in 50 mL distilled H2O, filter sterilized. Trypsin activity determined and frozen in 1.0-mL aliquots until use. Added to 2× 199 at the rate of 135 trypsin activity U/L. 8. Gentamicin (Sigma) at a final concentration of 0.025 g/L in 2× 199. 9. Nystatin (Gibco-BRL, Gaithersburg, MD) at a final concentration of 5000 U/L in 2× 199.
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10. Two-dram vials (8 mL): flat bottom, borosilicate glass, with rubber-lined screw cap. (Wheaton, Millville, NJ, cat. no. 224884). Unwashed vials directly from manufacturer are loosely capped, wrapped in butcher paper, and sterilized by autoclaving for 20 min on a drying cycle. 11. Sterile 0.1 and 1.0 mL pipets or automatic pipeters with sterile tips. 12. Sterile Pasteur pipets. 13. Six-well sterile tissue culture cluster plates (Costar, Costar, NY, cat. no. 3506).
3. Methods The methods and procedures outlined here are specifically for reassortment tests using ts mutants. Variations in procedure for complementation tests appear as notes throughout Subheadings 3.3.–3.9., as appropriate. Variations in procedure for other tests using mixed infection are covered in Subheadings 3.10.1.–3.10.4.
3.1. General Considerations Several general considerations, taken into account in the development of this protocol, are outlined below. 1. Optimal mixed infections require mixed infection of all the cells in the population, so that the viral genomes and gene products have maximal opportunity for interaction. This requires knowledge of the number of cells present in the monolayer. The protocol below is designed to produce MA104 cell monolayers for infection, with, on average, 1 × 105 cells/monolayer. However, the number of cells in the average monolayer, under the specific conditions of the laboratory, should be confirmed by trypsinizing several monolayers, and counting the cells in a hemocytometer. The cell monolayers are formed on the flat bottoms of 2-dram vials, so that they contain a relatively small number of cells. The small number of cells minimizes the use of virus stocks, and allows the use of virus stocks with titers as low as 2 × 107 PFU/mL. 2. The prerequisite for efficient mixed infections is infection of each cell with both parental viruses. The percentage of cells mixed infected with two different parental viruses is given by the expression: (1 – e–mA)(1 – e–mB) × 100
(1)
where mA and mB are the MOI of parental viruses A and B. Theoretically, an MOI of 5 PFU/cell of each parental virus will infect 98.7% of the cells, and, as a practical matter, an MOI of 10 PFU/cell of each parental virus will mixed infect virtually 100% of the cells in the monolayer. An equal, high MOI of each parental virus should always be used for mixed infections. The use of lower MOI, so that not all cells are mixed and infected, reduces the opportunity for interaction between the parental viruses, and progressively reduces the frequency of reassortment as the MOI is decreased (7). The use of unequal MOI of the two parental viruses also reduces reassortment frequency (11; and R. F. Ramig,
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unpublished observations). This protocol is based on infection with an MOI of 10 PFU/cell of each parental virus in mixed infections. In control, single infections, the MOI is increased to 20, so that the total MOI remains constant, negating the multiplicity effects seen with some mutants. 3. The high MOI used for mixed infections should limit the infections to a single cycle. One-step growth determinations for WT and ts mutants indicate that the growth cycle is complete within 32 h at 31°C, and more rapidly at 37 or 39°C (5,6). Studies of reassortment kinetics (7) indicate that reassortant viruses can be detected as early as 12 h postinfection, and that maximal frequencies of reassortment are rapidly attained within 16–24 h postinfection. However, slight increases in reassortment frequency are observed at times later than 24 h postinfection. Studies with asynchronous mixed infections demonstrate that infection with the second virus can be delayed as much as 24 h, compared to infection with the first virus, and reassortants are still obtained (17). Asynchronous co-infection has even been shown to increase the recombination frequency in cases of one parent replicating to higher titers and forming larger plaques than the other parent (18). These observations emphasize the need to incubate the infections to completion, to allow optimal and complete interaction of the parental viruses. This protocol utilizes a 48-h period of incubation of mixed and control infections, to ensure that all viruses have completed their infectious cycle. 4. Components contained in FBS inhibit the growth of RVs. As a result, MA104 cell monolayers are formed in the presence of FBS, and the FBS must be washed out of the system before initiating the infections. This protocol minimizes manipulations by simple removal of the serum-containing medium and replacement with serum-free medium. The residual inhibitory serum components are metabolized by the cells during an 18–24 h incubation after the medium change.
3.2. Experimental Time-Scale Mixed infections require planning, so that MA104 cell monolayers in the proper medium, and at the proper cell density, are available at the appropriate time. The following is the schedule for cell preparation and performance of the mixed infections: 1. D – 2. MA104 cells in Medium 199 + FBS are seeded into 2-dram vials and incubated at 37°C to form monolayers. 2. D – 1. The medium on the MA104 cells is changed to Medium 199 without FBS, and the cells are incubated at 37°C. 3. D – 0. The monolayers are inoculated with virus mixtures to initiate the infections, and are incubated at the appropriate temperature (31°C for reassortment tests). 4. D + 2. The infected monolayers are harvested. Infected cells are harvested by tightly capping the vials, and freezing them at –20°C until they can be analyzed.
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3.3. Preparation of MA104 Cell Monolayers for Infection Monolayers of MA104 cells must be prepared in advance of infection. This protocol yields monolayers that contain, on average, 1 × 105 cells in medium without serum, which are ready for infection. 1. A confluent monolayer of MA104 cells in a 150-cm2 tissue culture flask is trypsinized and suspended in 100 mL Medium 199 + FBS, using standard tissue culture techniques (see Note 1). 2. Sterile 2-dram glass vials are seeded with 1.0 mL MA104 cell suspension, and loosely capped. Always seed 3–5 more vials than are needed for the number of crosses and controls to be performed. 3. The loosely capped vials are incubated at 37°C in an incubator with an atmosphere of 5% CO2-air, for approx 24 h, to allow the cells to form a confluent monolayer. Monitor the growth of the cells using an inverted microscope at low magnification. Removing the cap of the vial may facilitate viewing the monolayer (see Note 2). 4. When the cell monolayers are confluent (generally 24 h), the medium is removed from each vial by aspiration, using a sterile Pasteur pipet (see Note 3). 5. Each vial is re-fed with 1.0 mL Medium 199 without FBS, loosely capped, and returned to the 37°C, 5% CO2 incubator. 6. After 18–24 h incubation at 37°C, to allow serum components to be metabolized, the monolayers are inspected using an inverted microscope. If the monolayers are intact and confluent, they are ready to use for infections. 7. Before use, the monolayers in 2–3 vials should be trypsinized and counted in a hemocytometer, to establish the average number of cells/monolayer. Once the procedure is established, one assumes a constant number of cells/vial (see Note 4).
3.4. Preparation of Virus Stocks at Titers Necessary for Protocol This protocol assumes that: MA104 cell monolayers to be infected contain 1 × 105 cells; the desired MOI is 10 PFU/cell of each parental virus for mixed infections and 20 PFU/cell for single-infection controls; that virus mixes for mixed infection will be made with 0.1 mL of each parental virus, and 0.2 mL virus for single-infection controls; that virus mixes and single viruses will be activated with an equal volume (0.2 mL) of trypsin prior to infection; and that infections will be initiated with 0.2 mL activated virus or virus mix. 1. The number of PFU required for each parental virus to obtain the desired MOI: (No. cells)(PFU/cell) = (1 × 105 cells)(10 PFU/cell) = 1 × 106 PFU required (2) (see Notes 5 and 6). 2. Because the number of PFU desired must be in 0.1 mL, the titer of virus stock to contain that number of PFU in PFU/mL is calculated: (PFU/0.1 mL)(10) = (1 × 106 PFU/0.1 mL)(10) = 1 × 107 PFU/mL needed
(3)
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Table 1 Preparation of Virus Stocks at Proper Titer for Use in Mixed Infections For final titer of 2 × 10 7 PFU/mL Virus mutant
Mutant titer at PT (PFU/mL)
Volume mutant virus stock (mL)
Volume 199 w/o FBS (mL)
1.20 × 108 1.55 × 108 1.80 × 108 2.95 × 108 1.05 × 108 3.00 × 108
0.167 0.129 0.111 0.068 0.190 0.067
0.833 0.871 0.899 0.932 0.810 0.933
tsA(778) tsB(339) tsC(606) tsD(975) tsA(2289) WT
3. To adjust for dilution of the virus or virus mix with an equal volume of trypsin for virus activation, the required titer of stock virus must be multiplied by two: (PFU/mL)(2) = (1 × 107 PFU/mL)(2) = 2 × 107 PFU/mL required virus titer (4) (see Note 7). 4. RV stocks generally have titers higher than 2 × 10 7 PFU/mL and must be diluted to that concentration for use in this protocol. The amount of high-titer virus stock diluted to a total volume of 1.0 mL is calculated as follows (a sample virus dilution sheet is shown in Table 1): PFU/mL needed PFU/mL stock
=
2.0 × 10 7 PFU/mL needed 5.5 × 10 8 PFU/mL stock
=
0.037 mL diluted (5) to 1.0 mL total
3.5. Preparation and Activation of Virus Mixes and Single Viruses for Infections After virus stocks are prepared at the proper titer for use in the experiment, virus mixes and single viruses are prepared and activated as described below (a sample preparation sheet is shown in Table 2): 1. Small sterile tubes are numbered to match the control and mixed infections (Table 2), and chilled on ice. 2. Control viruses and virus mixes are prepared by adding the amounts of virus stock (prepared at a titer of 2.0 × 10 7 PFU/mL, as in Subheading 3.4.) to the individual tubes, as indicated in Table 2. Use a sterile pipet for each addition, to prevent cross-contamination of tubes. 3. Ten µg/mL trypsin is prepared by diluting 0.1 mL trypsin stock (1 mg/mL) into 9.9 mL Medium 199 without FBS. 0.2 mL 10 µg/mL trypsin is added to each of the control or virus mix tubes (Table 2) (see Note 8). Use a sterile pipet for each addition to prevent cross-contamination of tubes.
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Table 2 Preparation of Virus Mixes, Activation, and Inoculation Volumes for Performing Mixed and Control Infections Volume parental virus (2 × 10 7 PFU/mL) to make virus mixes
Tube 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17
Cross (Parent 1 ⊗ Parent 2)a
Parent 1 (mL)
Parent 2 (mL)
10 µg/mL trypsinb (mL)
Volume to inoculate (mL)
tsA(778) alone tsB(339) alone tsC(606) alone tsD(975) alone tsA(2289) alone WT alone tsA(778) ⊗ tsB(339) tsA(778) ⊗ tsC(606) tsA(778) ⊗ tsD(975) tsA(778) ⊗ tsA(2289) tsB(339) ⊗ tsC(606) tsB(339) ⊗ tsD(975) tsB(339) ⊗ tsA(2289) tsC(606) ⊗ tsD(975) tsC(606) ⊗ tsA(2289) tsD(975) ⊗ tsA(2289) Uninfected
0.2 0.2 0.2 0.2 0.2 0.2 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.2 c
– – – – – – 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.1 –
0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2
0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2
a
For viruses alone for control infections, the virus is parent 1; for virus mixes for mixed infections, the left-hand virus is parent 1 and the right-hand virus is parent 2. b After addition of trypsin, the virus mixes and single-control viruses are incubated at 37°C for 30 min, to activate infectivity. c For the uninfected cell monolayer, the mock inoculum consists of 0.2 mL 199 w/o FBS.
4. The tubes with virus and trypsin are incubated for 30 min in a water bath at 37°C, for activation of viral infectivity. 5. After activation, the tubes are chilled on ice until use for inoculation of MA104 cell monolayers. Activated viruses should be used for infection within 1–2 h after activation.
3.6. Control and Mixed Infection of Cell Monolayers Prior to initiating the infections, cell monolayers have been prepared and average cell number determined, as described in Subheading 3.3. The day the infections are to be performed, virus stock dilutions are prepared as described
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in Subheading 3.4. Finally, the virus mixtures are prepared and trypsin activated, as described in Subheading 3.5. When these steps have been completed, the infections can be initiated, as described below. 1. Vials containing monolayers of MA104 cells are numbered to match the numbers of the control and mixed infections (Table 2). The authors find it convenient to use 0.5-inch round self-adhesive labels (Avery, Diamond Bar, CA, cat. no. 05406 or equivalent) placed on the cap of each vial. 2. At room temperature, the medium is removed from a group of vials (approx 10 vials per group) by aspiration with a sterile Pasteur pipet (see Note 9). 3. A volume of 0.2 mL from the appropriate virus mix tube is added to the matched vial containing the MA104 cell monolayer, and the vial is loosely capped. Moderate care is taken to ensure that the bottom of the vial is level, so that the inoculum covers the monolayer. 4. Steps 2 and 3 are repeated until all the vials have been inoculated. 5. The inoculated vials are placed in a 31°C incubator with an atmosphere of 5% CO2-air, and incubated for 1 h to allow virus to adsorb to the cells. Again, care should be taken that the bottoms of the vials are relatively level (see Notes 10 and 11). 6. After the adsorption period, move the vials to the work space at room temperature. Again working with small groups of approx 10 vials, remove the inoculum from each vial by aspiration, using a sterile Pasteur pipet. Use a separate, sterile Pasteur pipet for each vial, to prevent cross-contamination (see Note 12). 7. To each individual vial, add 1.0 mL Medium 199 without FBS, using a separate, sterile pipet for each vial, and loosely cap the vial. 8. Steps 6 and 7 are repeated until all the vials have been fed with fresh medium (see Note 13). 9. Place the vials in an incubator with an atmosphere of 5% CO2-air at the proper temperature. With ts mutants as used in this example, the vials are incubated at 31°C for reassortment tests (see Note 14). 10. Incubate the vials with infected monolayers for 48 h. Cytopathic effects will generally be complete by 48 h, using this protocol. 11. Harvest the infected cells by tightening the cap of each vial, and place them in a freezer at –20°C. The frozen vials can be stored until it is convenient to titer the resulting infected cell lysates.
3.7. Analysis of Yields of Progeny Virus from Mixed and Control Infections The yields of progeny virus from mixed infections for reassortment tests with ts mutants are assayed for infectivity using plaque assays at 31 and 39°C. The protocols for plaque assays have been described several times (5,9,19,20), and will be described only briefly here. 1. MA104 cells (1 × 105 cells/mL) in Medium 199 + FBS are seeded into the wells of six-well cluster plates, 2 mL/well. The plates are incubated at 37°C in 5% CO2-air, until the cells form confluent monolayers.
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2. When the cells have formed confluent monolayers, the medium is removed from the wells and replaced with Medium 199 without FBS (2 mL/well), and incubated 12–18 additional hours, to allow metabolism of the residual serum components inhibitory to RV plaque formation. 3. Immediately before performing the plaque assay, the frozen yields of the mixed infections are thawed and briefly sonicated, to disrupt intact cells and virus aggregates. Serial 10-fold dilutions of the infected cell lysates are prepared using gelatin-saline as diluent (see Note 15). 4. The medium is removed from the wells of the serum-deprived confluent MA104 cell monolayers, and the wells are inoculated (0.1 mL/well) with appropriate dilutions of the mixed infection yield. 5. The plates are incubated at 31°C for 1 h, to allow virus adsorption. 6. After adsorption, the infected monolayers are overlaid with 3 mL/well of nutrient medium made by mixing equal volumes of 2% water agar and 2× 199. 7. The cluster plates are incubated at the appropriate temperature for the appropriate time in 5% CO2 incubators (see Note 16). 8. At the appropriate time, each well of the monolayer is overlaid a second time with 1.0 mL/well neutral red medium made by mixing 2% water agar with 2× 199, and adding 25 µg/mL neutral red. The plates are returned to the incubator for 12–18 h further incubation, to allow viable cells to take up the neutral red, revealing the plaques. 9. The plaques present on the plates are counted, and the counts recorded (see Table 3 for sample data).
3.8. Calculation of Results from Mixed Infections for Reassortment Assays Sample data from mixed infections between RV ts mutants, generated using the protocol described here, are shown in Table 3. This assay was performed to determine reassortment frequencies between the mutants shown in Tables 1 and 2. The crosses were performed at 31°C, so that the viruses could replicate and their genomes could interact. The progeny were assayed at 31 and 39°C, and plaque counts for the duplicate wells of each dilution plated are shown. The titer of the yield determined at 39°C for the mixed infections represents the sum of WT (ts+) reassortants, ts+ revertants, and ts+ pseudorevertants, for the single infection controls, the titer at 39°C represents ts+ revertants and ts+ pseudorevertants. Subtracting the 39°C titers of the two relevant single infection controls from the 39°C titer of the mixed infection provides a good estimate of the number of WT (ts+) reassortants in the yield of the mixed infection. The titer of the mixed infections at 31°C represents the total yield of the infection, and includes ts parentals, ts double-mutants, WT (ts+) reassortants, ts+ revertants, and ts+ pseudorevertants. The frequency of WT (ts+) reassortants is calculated from the following formula, in which the superscript indicates the temperature of the assay, to determine the titer of the progeny (see Note 17).
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Table 3 Titration Data and Recombination Frequencies
Plate
Cross (Parent 1 ⊗ Parent 2)
Dilution plated Temp (°C)
10–2
10–4
10–5
10–6
Titer (PFU/mL)
EOPd or % ts+ Recomb.e
92
1
tsA(778) alone
39 31
17/15a –
0/0 C/Cb
0/0 51/54
– 9/8
1.60 × 104 5.25 × 107
3.05 × 10–4
2
tsB(339) alone
39 31
0/4 –
0/0 C/C
0/0 32/37
– 2/3
2.00 × 103 3.45 × 107
5.80 × 10–5
3
tsC(606) alone
39 31
39/44 –
0/0 C/C
0/0 55/64
– 10/8
4.15 × 104 5.95 × 107
6.97 × 10–4
4
tsD(975) alone tsA(2289) alone
6
WT alone
7
tsA(778) ⊗ tsB(339)
8
tsA(778) ⊗ tsC(606)
9
tsA(778) ⊗ tsD(975)
C/C – C/C – C/C – C/C – C/C – C/C –
7/5 C/C 2/1 C/C C/C C/C 57/62 C/C 57/59 C/C 43/53 C/C
0/0 T/Tc 0/0 T/T 68/84 C/C 6/6 T/T 6/7 T/T 4/3 T/T
– 8/12 – 8/8 – 16/13 – 11/17 – 10/10 – 14/8
6.00 × 105 1.00 × 108 1.50 × 105 8.00 × 107 7.60 × 107 1.45 × 108 5.95 × 106 1.40 × 108 5.80 × 106 1.00 × 108 4.80 × 106 1.10 × 108
6.00 × 10–3
5
39 31 39 31 39 31 39 31 39 31 39 31
1.88 × 10–3 0.52 4.25 5.74
Ramig
3.80
tsA(778) ⊗ tsA(2289)
11
tsB(339) ⊗ tsC(606)
12
93
39 31 39 31
C/C – C/C –
3/2 C/C 33/28 C/C
0/0 T/T 2/3 72/54
– 14/9 – 6/10
2.50 × 105 1.15 × 108 3.05 × 106 6.30 × 107
0.07
tsB(339) ⊗ tsD(975)
39 31
C/C –
38/43 C/C
6/6 73/74
– 7/10
4.05 × 106 7.35 × 107
4.69
13
tsB(339) ⊗ tsA(2289)
39 31
C/C –
16/11 C/C
1/0 51/59
– 5/12
1.35 × 106 5.50 × 107
2.18
14
tsC(606) ⊗ tsD(975) tsC(606) ⊗ tsA(2289)
16
tsD(975) ⊗ tsA(2289)
17
Uninfected
C/C – C/C – C/C – 0/0 –
48/54 C/C 28/23 C/C 44/43 C/C 0/0 0/0
5/6 T/T 2/2 T/T 4/6 T/T 0/0 0/0
– 14/18 – 9/9 – 8/6 – 0/0
5.10 × 106 1.60 × 108 2.55 × 106 9.00 × 107 4.35 × 106 7.00 × 107 – –
2.79
15
39 31 39 31 39 31 39 31
a
No./no. indicates numbers of plaques in duplicate wells of six-well cluster plate.
b
C/C indicates so many plaques that monolayer was totally destroyed.
c
T/T indicates plaques too numerous to count because of overlap.
d
EOP is titer at 39°C/ titer at 31°C; shown in italics (Plates 1–6).
e%
4.77
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2.62 5.14 –
Recomb. indicates recombination frequency calculated as described in Subheading 3.8.; shown in bold (Plates 7–16).
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Ramig 39 39 39 % ts+ Reassortants = (Titer A ⊗ B) – (Titer A 31+ Titer B ) × 100. (Titer A ⊗ B)
(6)
The following points should be noted from this reassortment data (Table 3): 1. Each of the mutants retains a ts efficiency of plating EOP < 0.1 during growth at 31°C for the control infections. 2. The WT control virus retains a ts+ EOP > 0.1 during growth at 31°C for the control infection. 3. The mixed-infection tsA(778) ⊗ tsA(2289) failed to yield a significant proportion of ts+ reassortant progeny virus. This indicates that tsA(778) and tsA(2289) reside on cognate genome segments of the parental viruses, so that reassortment cannot generate ts+ progeny. Indeed, data of this type is used to place both mutants in reassortment group A. All other crosses yield significant proportions of ts+ reassortant progeny, indicating that each mutant pair contains lesions on different genome segments, so that ts+ progeny can be generated by reassortment. Indeed, these mutants are assigned to reassortment groups A, B, C, and D, based on these results.
3.9. Calculation of Results from Mixed Infections for Complementation Assays If the crosses shown in Table 3 had been performed for complementation analysis, the mixed infections would have been incubated at 39°C, at which the mutant gene products of the mutants would have been expressed (see Note 18). Expression of mutant phenotype at 39°C would prevent the mutant viruses from growing, unless they were mutant in different transacting functions. If the mutant pair is mutant in different transacting functions, each could provide the function missing in the other, resulting in a yield of mutant virus. The progeny yields of the mixed infections are assayed at 39°C, at which ts+ reassortants, ts+ revertants, and ts+ pseudorevertants are the only progeny classes that would grow. Subtracting this background of wild phenotype from the total yield of the mixed infection determined at 31°C, provides a good estimate of the titer of mutant virus yielded by the mixed infection. This number is compared to the ability of each of the parental mutants to grow alone at 31°C. The complementation index is calculated as follows (where superscripts indicate the temperature of the titration): Complementation Index (CI)
=
(Titer A ⊗ B)31 – (Titer A ⊗ B)39 (Titer A)31 + (Titer B)31
(7)
Complementation analysis is of limited use with RV mutants (see Note 19). When pairs of mutants are subjected to complementation tests, the CI is quite low and irreproducible. The complementation data cannot be used to group RV
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mutants into self-consistent groups, regardless of the level of complementation assumed to be significant (3). 4. Notes 1. This monolayer preparation protocol is designed to allow convenient intervals between steps. If monolayers are not confluent within 24 h, a larger volume of cell suspension can be seeded, or the monolayers can be allowed more time for growth to confluence. 2. 24 h is the minimal time for formation of the monolayers. If shorter times are allowed for monolayer formation, the cells often release from the bottom of the vial when the inoculum-containing trypsin is added. 3. The monolayers can be washed free of serum by several changes of medium, and immediately used. However, this increases handling and the chances of contamination, compared to the protocol as outlined. 4. Cell monolayer preparation is identical, whether the cells will be used in reassortment, complementation, or other types of tests. 5. If a different MOI is desired, appropriate changes in calculations must be made. 6. A high MOI of 10 PFU/cell is used for reassortment, complementation, and other types of tests, unless MOI is an experimental variable. Occasionally, one encounters RV mutants that cannot be grown to the minimum titer of 2 × 107 PFU/mL required for the protocol as outlined here. In such cases, two alternative approaches can be taken. The first is preferable, because it allows crosses involving the mutant in question to be performed under conditions identical to other crosses, allowing direct comparison of the results. In this case, large 100–250 mL lysates of infected cells are prepared using standard techniques for virus propagation. The lysate is extracted with Genetron (Fisher, Pittsburgh, PA) to remove lipids, and the resulting supernatant is pelleted through a sucrose cushion. The pellet is then resuspended in a minimal volume (0.5–1.0 mL), titered, and, if the partially purified virus has a titer of 2 × 107 or greater, it is used to perform the cross, using the standard technique. If the low-titer mutant virus cannot be concentrated to sufficient titer, the second approach can be used. In this approach, asynchronous mixed infections are performed, infecting first with the low-titer mutant, and allowing several hours (1–4 h) for amplification of the virus before infecting with the second virus. The asynchronous infections are performed as described (17). The disadvantage of this alternative is that the low-titer virus is often used at an MOI significantly less than that required to infect all of the cells. The second virus is used at an MOI of 10, so that all cells infected with the first virus will be infected with the second virus. The result is that all cells infected with the low-titer virus are also infected with the high-titer virus, but, in addition, a significant number of cells are infected with the high-titer parental virus alone. The results obtained are, at best, qualitative, and should not be compared to infections performed at MOI 10 for each parent where quantitative differences may be meaningful. Control mixed infections can be performed (using mutants of sufficient titer to also be used in
96
7.
8.
9. 10.
11.
12.
13.
Ramig the standard protocol) at unequal MOI, using asynchronous infection of the two mutants, to determine the effects of the unusual format of the assay. This approach has been used in the closely related bluetongue virus (11). For the protocol presented here, with trypsin activation of viruses prior to infection, each virus stock is required at a titer of 2 × 107 PFU/mL. Stocks with higher titer must be diluted to a concentration of 2 × 10 7 PFU/mL prior to preparation of virus mixes. This protocol results in a final concentration of trypsin during activation of 5 µg/mL. This amount of trypsin is consistent with activation of the viruses, and does not cause the cells to release from the monolayer during the absorption period, as often occurs if higher trypsin concentrations are used for activation. With some virus strains, lower concentrations of trypsin can be used, and full activation of infectivity will still be obtained. Preparation and activation of virus mixes and controls is identical for reassortment, complementation, and other types of tests. Small groups of vials are infected at a time, to prevent drying of the monolayer during prolonged periods with no medium. Occasionally, the monolayer releases from the substratum and rounds up into a small ball of cells during the adsorption period. If this occurs, the experiment can be salvaged if the inoculum is removed very carefully, so that the ball of cells remains in the vial. The vial is then fed normally with medium, and cytopathic effects are observed to proceed as usual. The ts mutants in this laboratory collection were selected using a 1 h adsorption period at 31°C, followed by growth or plaque formation at 39°C, so that mutants with lesions in cell attachment would not be selected (5,6). Thus, 31°C should always be used for adsorption of virus to host cells, because any mutants with lesions in binding would fail to infect the cells. When removing the inoculum from the infected monolayer after adsorption (step 6), it is critical that a new sterile pipet be used to remove the inoculum from each vial to prevent cross-contamination of vials. For the same reason, a new sterile pipet should be used to feed each vial (step 7). In this protocol, unadsorbed virus is not removed by washing the monolayer or adding neutralizing Ab. The majority of the unadsorbed virus is removed when the inoculum is aspirated from the vial, and the residual unadsorbed parental virus does not introduce an unacceptable background to the experiment. If high backgrounds of unadsorbed virus cause unacceptable background, one can wash the monolayers prior to feeding and incubating for growth. The use of neutralizing Ab to remove unadsorbed virus is not recommended, because residual Ab may neutralize a significant proportion of the progeny virus from the infected cells. The medium used to feed the infected cell monolayers does not contain trypsin. Because the cells are infected at high MOI, cell-to-cell spread of virus is not required for the infection to proceed to completion, so addition of trypsin is not required. In practice, the authors find that some residual trypsin remains in the vial, so that the progeny virus yield is mostly activated without the addition of exogenous trypsin. This facilitates titration of the yields in subsequent plaque assays.
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14. For reassortment tests, the mixed infections are incubated at 31°C, to allow the mutants to replicate and increase the size of the genome pool available for interaction. For complementation tests, the mixed infections should be incubated at 39°C, so that gene products with mutant phenotype are expressed. 15. For mixed infections performed as described here, titration of the following dilutions has been found to yield wells with countable numbers of plaques. At 39°C, dilutions 10–2, 10–4, and 10–5 are plated. At 31°C, dilutions 10–4, 10–5, and 10–6 are plated. 16. The titrations at 39°C are incubated for 5 d, and then overlaid with neutral red late on d 5. They are counted on d 6 following inoculation. The titrations at 31°C are incubated for 9 d, and then overlaid with neutral red late on d 9. They are counted on d 10 following inoculation. 17. As with many quantitative assays in biology, reassortment tests are complicated by the issue of determining the level of significance. The reassortment behavior expected of viruses with segmented genomes is all-or-none. If reassortment is random, one would expect 25% of the progeny of a mixed infection to be ts+ and, as can be seen from the data in Table 3, this high frequency is not attained with RV mutants that reassort. The lower-than-expected frequency of reassortment in RVs has been ascribed to the interference property of the mutants with WT virus (3,6,7), so that less-than-expected amounts of ts+ progeny are generated. On the other hand, as can be seen in Table 3, when reassortment is not expected, the frequency is seldom zero, but some value greater than zero. In practice, reassortment tests yielding frequencies of ts+ reassortants between 0.2 and 1.0% are repeated several times. Nearly always, one obtains an average frequency of reassortment that is either less than 0.2% or greater than 1.0% when the mixed infections are repeated (5), and this has been used as the level of significance for negative (<0.2%) and positive (>1.0%) tests. Using this criterion, self-consistent grouping of RV mutants by reassortment has been possible (4–8). 18. For complementation tests, each of the ts mutants should be used in a mixed infection with WT, as well as alone and in mixed infection with other ts mutants. The mixed infection of ts mutant (WT serves as a control, to demonstrate that the ts mutant can be complemented in trans by the cognate gene product of WT. 19. Complementation tests are not particularly useful with RV ts mutants. The complementation indices obtained with RV SA11 ts mutants are low, highly irreproducible, and cannot be used to place mutants into self-consistent complementation groups (3,5,6). Interference tests performed by mixed infections of ts mutant and WT viruses showed that the ts mutants interfered strongly with the growth of WT virus (5,6). It is assumed that ts mutants also interfere with the growth of other ts mutants, so that complementation is not observed, although this has not been directly tested. If RV ts mutants lacking an interference phenotype were available, they would probably complement as expected. This was shown to be the case in the closely related reovirus, in which ts mutants that do not interfere show normal complementation, but those that do interfere fail to complement (21).
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Acknowledgments The work in the author’s laboratory, which led to development and refinement of this protocol, was supported by research grants from the National Science Foundation (PCM80002359), and from the Institute of Allergy and Infectious Diseases, National Institutes of Health (AI16687, AI21494, AI36385). References 1. Fields, B. N. (1996) Reoviridae, in Field’s Virology, 3rd ed., (Fields, B. N., Knipe, D. M., Howley, P. M., et al., eds.), Lippincott-Raven, Philadelphia, pp. 1553–1555. 2. Ramig, R. F. and Ward, R. L. (1991) Genomic segment reassortment in rotaviruses and other reoviridae. Adv.Virus Res. 39, 163–207. 3. Ramig, R. F. (1997) Genetics of the rotaviruses. Annu. Rev. Microbiol. 51, 225–255. 4. Greenberg, H. B., Kalica, A. R., Wyatt, R. W., Jones, R. W., Kapikian, A. Z., and Chanock, R. M. (1981) Rescue of noncultivable human rotavirus by gene reassortment during mixed infection with ts mutants of a cultivable bovine rotavirus. Proc. Natl. Acad. Sci. USA 78, 420–424. 5. Ramig, R. F. (1982) Isolation and genetic characterization of temperature-sensitive mutants of simian rotavirus SA11. Virology 120, 93–105. 6. Ramig, R. F. (1983) Isolation and genetic characterization of temperature-sensitive mutants that define five additional recombination groups in simian rotavirus SA11. Virology 130, 464–473. 7. Ramig, R. F. (1983) Factors that affect genetic interaction during mixed infections with temperature-sensitive mutants of simian rotavirus SA11. Virology 127, 91–99. 8. Faulker-Valle, G. P., Clayton, A. V., and McCrae, M. A. (1982) Molecular biology of rotaviruses. III. Isolation and characterization of temperature-sensitive mutants of bovine rotavirus. J. Virol. 42, 669–677. 9. Chen, D. and Ramig, R. F. (1994) Construction and characterization of rotavirus reassortants. Methods Mol. Genet. 4, 183–194. 10. Rubin, D. H. and Fields, B. N. (1980) Molecular basis of reovirus virulence: Role of the M2 gene. J. Exp. Med. 152, 853–868. 11. Ramig, R. F., Garrison, C., Chen, D., and Bell-Robinson, D. (1989) Analysis of reassortment and superinfection during mixed infection of Vero cells with bluetongue virus serotypes 10 and 17. J. Gen. Virol. 70, 2595–2603. 12. Graham, A., Kudesia, G., Allen, A. M., and Desselberger, U. (1987) Reassortment of human rotavirus possessing genome rearrangements with bovine rotavirus: nonrandomness and evidence for host cell selection. J. Gen. Virol. 68, 115–122. 13. Chen, D., Burns, J. W., Estes, M. K., and Ramig, R. F. (1989) The phenotypes of rotavirus reassortants depend upon the recipient background. Proc. Natl. Acad. Sci. USA 86, 3743–3747. 14. Chen, D., Estes, M. K., and Ramig, R. F. (1992) Specific interactions between rotavirus outer capsid proteins VP4 and VP7 determine expression of a crossreactive, neutralizing VP4-specific epitope. J. Virol. 66, 432–439.
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15. Gombold, J. L. and Ramig, R. F. (1987) Assignment of simian rotavirus SA11 temperature-sensitive mutant groups A, C, F, and G to genome segments. Virology 161, 463–473. 16. Gombold, J. L., Estes, M. K., and Ramig, R. F. (1985) Assignment of simian rotavirus SA11 temperature-sensitive mutant groups B and E to genome segments. Virology 143, 309–320. 17. Ramig, R. F. (1990) Superinfecting rotaviruses are not excluded from genetic interactions during asynchronous mixed infections in vitro. Virology 176, 308–310. 18. Tauscher, G. I. and Desselberger, U. (1997) Viral determinants of rotavirus pathogenicity in pigs: Production of reassortants by asynchronous coinfection. J. Virol. 71, 853–857. 19. Matsuno, S., Inouye, S., and Kono, R. (1977) Plaque assay of neonatal calf diarrhea virus and the neutralizing antibody in human sera. J. Clin. Microbiol. 5, 1–4. 20. Smith, E. M., Estes, M. K., Graham, D. Y., and Gerba, C. P. (1979) A plaque assay for the simian rotavirus SA11. J. Gen. Virol. 43, 513–519. 21. Chakraborty, P. R., Ahmed, R., and Fields, B. N. (1979) Genetics of reovirus: the relationship of interference to complementation and reassortment of temperature-sensitive mutants at nonpermissive temperature. Virology 94, 119–127.
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6 Pathogenesis and Animal Models Linda J. Saif and Lucy A. Ward 1. Introduction 1.1. Rationale for Use of Gnotobiotic Pigs in Rotavirus Research Because of the limitations in studying human rotavirus (HRV) pathogenesis and mucosal immunity in the natural host (infants and children), various animal models have been utilized to investigate rotavirus (RV) disease pathogenesis and immunity. Mice and rabbits serve as useful models to evaluate and dissect immune responses to RV (see Chapter 9). However, because older mice (>14 d) and rabbits are not susceptible to diarrhea after inoculation with either homologous (murine or lapine, respectively) or heterologous (human) RVs, assessment of protective immunity is restricted to prevention of virus shedding only (1–3). Gnotobiotic pigs are susceptible to infection and disease induced by several HRVs, until at least 6 wk of age (4,5). Moreover, a virulent strain of HRV (Wa) induced intestinal lesions (villous atrophy and crypt hyperplasia) in gnotobiotic pigs (6), resembling those seen in children with natural symptomatic RV infections (7). Further advantages of gnotobiotic pigs for HRV research include the following: Colostrum-deprived pigs are devoid of maternal antibodies (Abs) at birth (the placental type in swine acts as a barrier to the transfer of maternal Abs), but are immunocompetent, thus permitting an assessment of true primary immune responses to RV (4,5,8,9), and then determination of the impact of defined titers or specificities of maternal Abs on immunity to RV (10–12); the gnotobiotic environment ensures that exposure to extraneous RVs or other enteric pathogens is not a confounding variable; gnotobiotic piglets closely resemble human infants in gastrointestinal physiology (monogastrics), mucosal immune development, size, and milk diet. From: Methods in Molecular Medicine, Vol. 34: Rotaviruses: Methods and Protocols Edited by: J. Gray and U. Desselberger © Humana Press Inc., Totowa, NJ
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1.2. Accomplishments in RV Research Resulting from Gnotobiotic Pig Studies The gnotobiotic pig model has contributed to several important advances in RV research, including adaptation of the first HRV (Wa strain) to cell culture after passage and amplification in piglets (13); delineation of the independent roles of the RV outer capsid viral proteins VP4 and VP7 in induction of neutralizing Abs and cross-protection (14); recognition of the potential role for a nonstructural protein (NSP4), besides VP4 and VP7, in RV virulence (15); and comparative pathogenesis of virulent and attenuated strains of HRV in the intestine, leading to the recognition that factors other than villous atrophy may contribute to the early diarrhea induced by virulent HRV in pigs (6) or homologous virulent RVs in gnotobiotic pigs (16) and calves (17). These latter findings, including a recently proposed role for RV NSP4 as a viral enterotoxin (18), may reveal new pathogenic mechanisms of viral diarrhea. In order to design and improve RV vaccination strategies, correlates of protective immunity against HRV-induced diarrhea have been analyzed in gnotobiotic pigs inoculated orally with virulent HRV (mimic natural infection), or live attenuated or inactivated HRV vaccines (4,5,8,9). Complete protection against RV diarrhea and virus shedding was induced by virulent RV and significant correlations were observed between the numbers of intestinal immunoglobulin A (IgA) antibody-secreting cells (ASC), intestinal IgA Ab titers and intestinal lymphoproliferative responses to RV, and the degree of protection. Such results agree with findings by others that natural symptomatic and asymptomatic RV infections in infants in the first year of life conferred a high rate of protection against subsequent diarrhea (19,20) and that pre-exposure fecal IgA Abs to RV in children remaining uninfected during a RV season were significantly higher than in symptomatically infected children (21). Thus, the gnotobiotic pig studies have established key parameters and correlates related to protective immunity, several of which were also observed in children, and have confirmed the applicability of this model to examine new strategies for the improvement of HRV vaccines for infants.
1.3. Pathogenesis of Group A HRV in Gnotobiotic Pigs Studies on RV pathogenesis need to be performed using well-defined inoculation doses expressed as the median infective dose (ID50) or median diarrheal dose (DD50) for a given virus strain within a given host species. The ID50 and DD50 of a virulent (infant stool passaged in pigs) HRV (Wa strain, P1A[8],G1) in gnotobiotic pigs was found to be extremely low (1 focus-forming unit [FFU]), indicating high susceptibility of the gnotobiotic pig to this virus (6). Inoculation doses of 105 ID50 (or 105 DD50) were found to elicit clinical disease in 100% of 3–5-d-old gnotobiotic pigs by post-inoculation h (PIH) 24,
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and were subsequently used for pathogenesis studies. Morphologic evaluation of intestinal and selected lymphoid tissues were done at the onset of diarrhea or PIH 13, during virus shedding (PIH 24–96), and after recovery from diarrhea or postinoculation d (PID) 7. Virulent Wa HRV induced diarrhea and morphologic lesions typical of homologous (porcine) RV infections (16,22,23). Macroscopic changes are characterized by mild thinning of intestinal walls, reduced chyle within mesenteric lymphatics, mesenteric lymph node enlargement, and reduction in large (spiral) colon mass. Microscopic changes include loss of normal mature absorptive epithelial cells, detachment of absorptive cells from their basement membranes (especially over the tips of the villi of the small intestine), and mild lymphoreticular hyperplasia within villous tip stroma, followed by villous atrophy and crypt hyperplasia. Lymphoreticular hyperplasia becomes widespread and more pronounced after PIH 72, with a concurrent, although transient, increase in the number of intestinal intraepithelial lymphocytes. By PID 7, diarrhea subsides and villous morphology is within normal histologic limits, with only lymphoid hyperplasia persisting. Virus antigen (Ag) distribution in intestinal tissues, like lesion distribution, is nonuniform. Few infected cells (<1–8%) are detected in the duodenum, but they are numerous (30–50%) within the jejunum and ileum. Small amounts of virus Ag are transiently present extraintestinally, within mesenteric lymph nodes at PIH 24, presumably as a result of Ag uptake by resident phagocytic cells at the peak of virus replication. The large amount of virus Ag and infectious virus within the intestines from PIH 24 to 72 results from lysis of the infected villous epithelial cells (leading to villous atrophy), which forms the basis of the proposed pathogenesis for virulent Wa HRV in gnotobiotic pigs (6). The number of cells that are infected, and presumably functionally impaired, is large and these cells are rapidly lysed. Thus, the mucosa is unable to efficiently compensate or recover sufficiently to maintain minimal capacity for normal digestion and absorption. Some absorption probably takes place in the large bowel, but it is not able to compensate for the functional loss incurred by the small intestine of the young gnotobiotic pig. During the initial phase of lesion progression, rapid cellular loss may also permit leakage of plasma proteins and electrolytes from the mucosa, which further contributes to the movement of fluid to the lumen, and to the subsequent diarrhea. The absence of significant morphologic changes at the onset of diarrhea, which also has been observed in pigs infected with porcine RVs (16,22,23), suggests that other factors (in addition to cell lysis and mucosal disruption) are contributing to the early disease expression following virulent Wa HRV infection in gnotobiotic pigs. A recent report suggests a possible role for NSP4 as a viral enterotoxin (18), and studies of RV reassortants in gnotobiotic pigs have shown that replacement of the NSP4 gene of a virulent porcine RV strain with
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the corresponding gene of an attenuated HRV strain yielded a reassortant that failed to induce diarrhea (15), thus implicating specific viral factors in disease expression. The role of host factors, such as activated T-cells and cytokines, in disease expression in gnotobiotic pigs is under active investigation by the authors. The pathogenesis of attenuated (serially passaged in cell culture) Wa strain HRV was also examined in gnotobiotic pigs (6). Because inoculation doses of >107 FFU resulted in shedding in only a few pigs (6%), and no diarrhea in any pigs, but seroconversion in 100% of pigs, the ID50 of this virus was calculated as the median seroconversion dose (SD50). Subsequently, morphologic changes associated with the attenuated virus were assessed following inoculation with 107 FFU (102 SD50). No macroscopic lesions were observed, and microscopic changes were minimal and transient, with a loss of normal absorptive cell vacuolation throughout the duodenum and jejunum from PIH 13 to 24. Lymphoid hyperplasia was also evident in mesenteric lymph nodes and Peyer’s patches from PIH 72 to PID 7. The authors’ findings suggest that infection takes place following oral inoculation with the attenuated virus strain; however, the inability to detect intestinal viral Ag, and the absence of significant histologic lesions, suggests that very few intestinal cells become infected initially, or that replication of the virus is inefficient or possibly even abortive. The attenuation process (which consisted of serial propagation in cell culture) probably leads to changes in the virus genome that ultimately influence virulence and pathogenicity. Several RV gene segments and their products have been implicated in the pathogenicity of RV in pigs, including the structural gene products, VP4, VP3, and VP7, and the nonstructural gene product, NSP4 (15,24–26). Clearly, more studies are needed to elucidate the mechanisms that account for the pathogenic differences seen between attenuated and virulent HRVs.
1.4. Studies of Immunity to HRVs in Gnotobiotic Pigs 1.4.1. Passive Immunity Gnotobiotic pigs were used to evaluate the efficacy of passively administered circulating (serum) and/or local colostrum/milk maternal (swine) or heterologous (bovine) Abs for preventing HRV-induced diarrhea (10–12,27). Cows were immunized with HRV serotypes P1A,G1 and P1B,G2 and simian RV SA11 (P[2],G3), and the immune colostrum was collected (27). Feeding of Ab concentrates from the immune bovine colostrum effectively reduced RV shedding and diarrhea, in a dose-dependent manner. Pigs protected against HRV-induced disease developed active immune responses in the presence of the heterologous, passive colostral Abs. In recent experiments, the effects of colostrum/milk (mimic breast-fed infants) and/or circulating serum maternal Abs (mimic non-breast-fed infants)
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from sows immunized with Wa HRV (immune, high Ab titer) or nonimmunized seropositive sows (control, low Ab titer), on passive protection and development of active immunity to RV were assessed in neonatal gnotobiotic pigs inoculated and challenged with HRV (10–12). At birth, pigs received immune or control circulating serum maternal Abs (via intraperitoneal injection), with or without daily immune or control colostrum/milk dietary supplements. The effect of maternal Abs on protection was assessed by analysis of diarrhea and virus shedding after inoculation and challenge, and on B-cell responses by quantitation of ASC by enzyme-linked immunospot (ELISPOT) and Abs by enzyme-linked immunosorbent assay (ELISA), and on T-cell responses by determination of lymphoproliferative assay (LPA) responses. Immune responses were examined in intestinal and systemic lymphoid tissues or secretions at selected PID and post-challenge days (PCD), as described previously (5,8). Pigs that received immune maternal Abs were at least partially protected from diarrhea and virus shedding after inoculation, compared to pigs receiving control or no maternal Abs, but had higher rates of diarrhea and virus shedding after challenge. Higher susceptibility to diarrhea after challenge coincided with lower numbers of IgG and IgA ASC in intestinal tissues, lower serum and intestinal IgA Ab titers, and lower LPA intestinal responses at challenge, compared to pigs receiving control or no maternal Abs. Thus, high titers of maternal Abs in the circulation or diet of neonatal pigs conferred at least partial passive protection against virulent HRV, but led to decreased active immune responses. Such decreased immune responses in the presence of high titers of maternal Abs could impact oral HRV vaccine efficacy, especially in developing countries.
1.4.2. Active Immunity Studies of immunity to HRV in gnotobiotic pigs have focused on the identification of correlates of homotypic protection (4,5,8,9). The viral inocula selected were designed to mimic immunity induced by natural infection (virulent Wa RV), or to assess immunity induced to live attenuated, or inactivated Wa RV vaccines. Neonatal piglets were orally inoculated with virulent, attenuated or inactivated (with 10% binary ethylenimine) RV and challenged at PID 21 with homologous virus. Pigs were examined daily for diarrhea and fecal RV shedding, by ELISA and cell culture immunofluorescence assays after inoculation and challenge. Correlates of protective immunity were evaluated by ELISPOT (B-cell responses) and LPA (T-cell responses) on systemic (blood, spleen) and intestinal (gut lamina propria, mesenteric lymph node) lymphoid tissues collected at selected PIDs and PCDs. After inoculation, almost all pigs exposed to virulent RV developed diarrhea, and all shed virus, and seroconverted with neutralizing Abs to Wa RV. None of the pigs given attenuated or inactivated RV developed diarrhea (moderate to severe) and only
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6% of pigs given attenuated RV shed virus, but 96–100% of pigs in both groups seroconverted. After challenge, most virulent virus-inoculated pigs were protected from diarrhea (87% protection rate) and virus shedding (100% protection), compared to protection rates against diarrhea of 31% for attenuated virus-inoculated pigs and 0 and 6% for inactivated virus-inoculated pigs (administered orally or parenterally, respectively). At challenge, the mean numbers of IgA ASC in the intestinal lamina propria were significantly greater in virulent virus-inoculated pigs (53 ASC/5 × 105 mononuclear cells [MNC]) than in attenuated (6 ASC/5 × 105 MNC) or inactivated (3.5 ASC/5 × 105 MNC) virus-inoculated pigs, and were significantly correlated with the degree of protection. Furthermore, the transient appearance of IgA ASC in the blood of virulent virus-inoculated pigs mirrored the IgA responses in the gut, suggesting that IgA ASC in the blood of children could serve as a surrogate marker for IgA ASC responses in the intestine after RV infection. High numbers of IgG ASC or memory IgG ASC, induced in the systemic lymphoid tissues of the inactivated virus-inoculated pigs at challenge, did not correlate with protection. Thus, inactivated RV administered orally or parenterally, was significantly less effective in inducing intestinal IgA ASC responses and conferring protective immunity than the live oral HRVs (virulent or attenuated). Based on these studies, gnotobiotic pigs should provide a useful model for examining more efficient mucosal delivery systems, mucosal adjuvants, and vaccination strategies to enhance induction of intestinal IgA ASC by candidate RV vaccines.
1.5. Derivation and Maintenance of Gnotobiotic Pigs An early description of the use of gnotobiotic pigs, in an isolator system to study disease pathogenesis, appeared in 1962, in a study of hog cholera virus (28). Since then, both gnotobiotic pigs and calves have been used as animal models to study the pathogenesis of HRV infections and immunity (4–6,8–15,27,29,30). Their prolonged susceptibility to diarrhea induced by HRV, which was not reproducible in other animal models, permitted challenge exposure to evaluate protection against disease.
1.6. Histopathology Studies The rate of autolytic change in the digestive tract is related to the microflora present within the intestine: Significant autolytic change is present in all sections of the conventional pig small intestine and colon within 24 min of death, whereas significant autolytic changes in these same tissues from gnotobiotic pigs are not observed until 1 hour after death (31). The small intestines should be among the first samples taken, because of the more rapid rate of intestinal (especially duodenal) autolysis, compared to other tissues, when microscopic
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examination is required (31). Exsanguination at euthanasia helps minimize hemorrhage artifacts, and enhances visualization of some lymphoid tissues (especially spleen). Agonal death, or application of prolonged electrical current (as during electrocution), may cause nonspecific lesions, such as petechiation in some tissues. Tissue collection should be performed rapidly and immediately following euthanasia in a cool environment. Placement of intestines on ice-filled trays, covered by plastic (cellophane wrap) or aluminum foil, helps to slow autolysis during collection of segments. Flushing the lumen of the intestinal segments collected with fixative, prior to immersion in the fixative, also reduces autolytic change and aids in fixation. A minimum of a 10:1 fixative:tissue volume ratio should be used for fixation of tissues, and tissue thickness should not exceed 0.5 cm for any specimen collected. Because lesions induced by RV may be scattered, multiple segments of intestine from each region of the intestine (duodenum, jejunum, ileum) should be collected for thorough histologic evaluation. Collection of nine 6-cm-long small intestinal segments from gnotobiotic pigs at equally spaced sites, beginning 5 cm distal to the pylorus and ending 5 cm proximal to the ileal–cecal junction, will give two to four representative segments from each region.
1.7. Pathophysiology Studies To the authors’ knowledge, no published studies exist of intestinal permeability during RV infection of gnotobiotic pigs. However, several reports evaluate the effect of RV infections on the absorption and transport of macromolecules within the intestine in species other than pigs. Readers are referred to these studies for applicable methodologies (32–37). It would appear from these studies that RV infection has diverse affects on intestinal absorption and transport of macromolecules. The uptake of polyethylene glycols (mol wt 282–1250), lactulose, and D -xylose are reduced during RV infection, but uptake of macromolecules, such as horseradish peroxidase (HRP) and L -rhamnose, are increased. This diversity may best be explained by differences between species in terms of the intestinal region involved in absorption and transport of a given macromolecule and the region most affected by the infecting RV strain. 2. Materials 2.1. Diluent and Growth Media 1. Minimal Essential Medium (Gibco-BRL, Grand Island, NY); 1% antibiotics: penicillin G (Pfizer, New York); dihydrostreptomycin sulfate (NB Biochemicals, Cleveland, OH); mycostatin (Squibb, Princeton, NJ); 1% sodium bicarbonate (Na HCO 3) (7% stock); 1% HEPES (Gibco-BRL).
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2.2. Fixatives 1. 10% neutral buffered formalin: 37–40% 100 mL formaldehyde; 900 mL distilled water; 4 g monobasic sodium phosphate; 6.5 g dibasic sodium phosphate. 2. 10% zinc formalin (Z-fix® Concentrate, Anatech, Battle Creek, MI): 4 gal distilled or deionized water; 1 gal Z-fix® concentrate. 3. Prefer (Prefer® Concentrate, Anatech): 11.6 L distilled or deionized water; 3.5 L ethanol or reagent buffer; 1.0 gal Prefer® Concentrate. Shake to mix thoroughly.
2.3. Clearants 1. Xylene. 2. Xylene substitutes: Propar ® clearant (Anatech); Clearium® (Surgipath, Richmond, IL).
2.4. Stains 1. Mayer’s hematoxylin (Sigma, St. Louis, MO). 2. Harris hematoxylin (Lerner, Pittsburgh, PA). 3. 1% eosin alcoholic stock: 1 g water soluble eosin Y; 20 mL distilled water; dissolve eosin in water, then add 80 mL 95% ethanol. 4. Eosin Y (working solution): 1 part 1% alcoholic eosin stock; 3 parts 95% ethanol. Prior to use, add 0.5 mL glacial acetic acid to each 100 mL stain, and stir.
2.5. Solutions 1. Tris-HCl buffer 0.05 M, pH 7.6: 3.03 g Trizma-HCl (Sigma), 0.69 g Trizma base (Sigma). Dissolve in 500 mL deionized water. 2. Tris - Triton-X buffer (TTBS-HS/Tx): 15.15 g Trizma-HCl (Sigma), 3.47 g Trizma base (Sigma), 1000 mL deionized water, 20.29 g NaCl, 5 mL 10 Triton-X (Sigma). 3. Pronase, 0.01%: 0.15 g Protease XIV (Sigma); 150 mL 0.05 M Tris-HCl buffer, pH 7.6. 4. Bluing agent: 0.2% NaH2CO3 and 0.04% lithium carbonate (LiCO3) in water.
2.6. Immunohistochemistry Reagents 1. 2. 3. 4.
Streptavidin-HRP (Dako, Carpinteria, CA). Biotinylated secondary Abs (Dako; Sigma). 3',3'-Diaminobenzidine (DAB) solution (Sigma). 3-Amino-9-ethylcarbazole (AEC) (Zymed, San Fransisco, CA).
3. Methods 3.1. Derivation of Gnotobiotic Pigs by Hysterectomy (38) 1. Healthy pregnant sows are selected for surgery between the d 110 and d 114 of gestation. 2. Ketamine HCl (11 mg/kg) is administered i.m., to sedate the sow.
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Fig. 1. Transfer isolator used to procure gnotobiotic piglets (see Subheading 3.1.). 3. The sow is elevated using a soft cotton rope around each posterior leg, and the abdomen is scrubbed with a germicidal solution (Prodine, Phoenix Pharmaceuticals, St. Joseph, MO). 4. The sow is rendered unconscious using CO 2 delivered into an inhalation bag, and at the end of the surgery the sow is euthanized. 5. An incision is made down the midline of the abdomen of the unconscious sow, and the entire uterus is removed and passed through a chlorinated liquid trap entry into a sterilized surgery isolator. 6. Using rubber gloves in the sides of the isolator for manipulations, the pigs are rapidly removed from the uterus and passed into a sterile transfer isolator, where the umbilical cords are tied. 7. The pigs are transferred into sterile rearing isolators maintained in a room at 35–37°C for the first few days, and thereafter maintained at room temperature. 8. The isolators are constructed of stainless steel rectangular tubs with flexible plastic bubble tops, fitted with two pairs of rubber gloves on the sides. Each tub contains a removable false bottom, metal partitions for 2–4 pigs (each pig in a separate compartment), feeders, and retaining grid. The air blower and filters are similar to ones described previously (38,39; see Fig. 1).
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3.2. Maintenance of Gnotobiotic Pigs 1. All pigs are fed 2–3× daily with a total of 240 mL liquid diet of pig milk replacer (SPF Lac, Borden, Hampshire, IL). 2. Microbiological monitoring is initiated at derivation, then at weekly intervals. Rectal swabs are cultured and examined microscopically for bacteria, yeast, and fungi. Blood agar and fresh thioglycolate broth are used for culture media, and held at least 2 wk before being judged negative.
3.3. Inoculation, Challenge, and Sample Collection 1. Pigs are immunized with various RV candidate vaccines, and challenged at 2–4 wk after the final booster immunization (5). Pigs are challenged orally with 10 6 ID 50 (the DD 50 was approx equal to the ID 50 ) of virulent Wa HRV (P1A[8],G1) or M strain HRV (P1A[8], G3), to consistently reproduce diarrhea and lesions in 100% of inoculated pigs (6). 2. Control pigs are given equal volumes of diluent alone, to monitor fecal consistency, and to collect control tissues and samples. 3. Pigs are examined daily for clinical signs, including anorexia and diarrhea (scored as fecal consistency of 0 [normal] to 3 [liquid]). 4. Blood (2–5 mL) is drawn weekly from the anterior vena cava by using a 20–22 gage, 1.5-in. needle, and serum is obtained by standard procedures. 5. Daily duplicate rectal swabs are collected at 0–10 PID and PCD into 8 mL diluent (see Subheading 2.1.).
3.4. Cell Culture Immunofluorescence (CCIF) for Titration of Infectious RV (40) 1. Monkey kidney (MA104) cells are grown for 6–9 d in flat-bottomed 96-well microtiter plates (Costar, Cambridge, MA) in growth media, but containing 10% fetal calf serum. 2. Wells are rinsed with diluent about 3 h prior to viral inoculation. Rectal swabs or fecal samples to be tested for virus are diluted 1 in 25 (two swabs in 8 mL), then 1 in 100, and serially thereafter in 10-fold dilutions in diluent, and 0.1 mL diluted viral specimens are added (in duplicate) to each well. Thirty microliters of a 1 in 200 dilution of stock pancreatin (1 in 10, 4XNF, Gibco-BRL) are added to each well. Virus positive and negative specimens are added to each plate. 3. The plates are placed in a centrifuge plate carrier and centrifuged at 1300g for 60 min. 4. After centrifugation, the plates are incubated at 37°C for 18 h in a 5% CO2-air atmosphere, and then the medium is aspirated and the wells rinsed with phosphate-buffered saline (PBS) (pH 7.2). 5. The cells are fixed with 0.2 mL of a solution containing 80% acetone and 20% distilled H 2 O for 10 min at room temperature. The fixative is aspirated, and plates are rinsed with PBS. 6. To stain, 25 µL diluted (1 in 100 in PBS) fluorescein isothiocyanate (FITC)conjugated hyperimmune anti-RV serum, prepared in gnotobiotic pigs, is added, and plates are incubated for 1 h at 37°C.
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7. Plates are rinsed with PBS, mounting media is added, and plates are examined, using a fluorescent microscope. The number of individual fluorescing cells/well are enumerated at the highest dilution with fluorescing cells, and the virus infectivity titer is calculated as FFU/mL.
3.5. Immunofluorescent (IF) Staining of RV-Infected Intestinal Epithelial Cells (41) 1. Remove small intestine from gnotobiotic pig immediately after euthanasia. 2. Cut approx 6-cm-long segments from the duodenum, jejunum, and ileum. Slit open each segment lengthwise, to expose the mucosal surface. 3. Gently rinse the mucosal surface with 0.85% saline, then use a scalpel to scrape the mucosal surface, and put the scrapings on an alcohol-cleaned microscope slide. Using a second alcohol-cleaned slide, press the two slides together, to make a mucosal impression smear. 4. Air-dry the smears, and fix in 100% acetone for 10 min. 5. Rinse in PBS (pH 7.2) for 5 min, then distilled H2O for 30 s. Air-dry and IF-stain, or store at –20°C until IF staining is done. 6. Apply 0.03 mL diluted (1 in 50–1 in 100 in PBS) FITC-conjugated anti-RV serum as described for the CCIF procedure (Subheading 3.4., step 6), and incubate for 30 min at 37°C. 7. Rinse slides using PBS, and leave in PBS for 10 min. Rinse with distilled H2O for 30 s and air-dry. 8. Add mounting medium and cover-slip, and examine the smears for fluorescing cells, using a fluorescent microscope. Calculate the approximate percentage of fluorescing cells in randomly selected fields. 9. Paraffin-embedded tissue sections may also be IF-stained (as described in Subheading 3.6.).
3.6. Processing of Tissues and Preparation of Paraffin Sections for IF Staining (40) 1. Flush the lumen of freshly collected small intestinal segments with 100% acetone, and immerse in two changes of acetone for 30 min to 1 h each. 2. Place acetone-fixed tissue directly in melted paraffin in a vacuum oven at 60°C and 25 in. (63.5 cm) Hg for 10 min. Embed and section as described in Subheading 3.7., steps 3–5. 3. Deparaffinize slides in two changes of Propar (or xylene) for 2 min each. 4. Rinse in 100% acetone for 5 min and allow to air-dry. 5. Wash in PBS for 10 min, followed by distilled H2O for 30 s. 6. Air-dry and resume IF-staining at Subheading 3.5., step 6.
3.7. Processing Paraffin-Embedded Tissues for Hematoxylin and Eosin (H&E) or Immunohistochemistry (IHC) Staining (6,42–44) 1. Process tissues through two changes of fixative, 60 min each; one change of 80%, two changes of 95%, and two changes of 100% ethanol, 30 min each;
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6.
Saif and Ward three changes of Propar, 60 min each, and two changes of melted paraffin, at least 60 min each. Paraffin-embed formalin-fixed tissues within 24–48 h of collection. Prefer-fixed tissues may stay in a fresh change of fixative for up to 2 wk before paraffinembedding. Section paraffin blocks at 4–5 µm, and float on a water bath containing only deionized or distilled water (no adhesives). Gelatin may be added to water bath for H&E sections. Pick up flattened sections on ProbeOn® Plus (Fisher Scientific, Pittsburgh, PA) slides (poly-L-lysine or chrome alum-coated slides are also acceptable), and bake in a 60°C oven for 1 h prior to deparaffinization (see Note 1). Label all slides using Secureline® (Precision Dynamics Corp., San Fernando, CA) pen (most other pens are soluble in alcohol and/or xylene, and will wash off during deparaffinization, dehydration, and clearing). Proceed with desired staining procedure.
3.7.1. H&E Staining Procedure (42) 1. Deparaffinize slides in three changes of Propar (or xylene) for 5 min each (see Note 2). 2. Transfer slides to 100% ethanol, two changes, 3 min each, followed by 95 and 70% ethanol for 3 min each. Rinse under running tap water for 1 min. 3. Transfer to Harris’ hematoxylin for 5 min (filter Harris’ just prior to use), and place under running tap water for 1 min. 4. Dip slide once in 0.25% citric acid, and rinse under running tap water for 1 min. 5. Dip slide 4× in bluing agent, and rinse under running tap water for 1 min. 6. Place in 95% and 70% ethanol for 1 min each. 7. Place in eosin Y (working solution) for 2 min. 8. Dip slide once each in 95% and 100% isopropanol. 9. Place in two changes of 100% isopropanol for 1 min each. 10. Remove s
