Advances in
VIRUS RESEARCH VOLUME 46
ADVISORY BOARD DAVIDBALTIMORE
PAULKAESBERG
ROBERT M. CHANOCK
BERNARD Moss
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Advances in
VIRUS RESEARCH VOLUME 46
ADVISORY BOARD DAVIDBALTIMORE
PAULKAESBERG
ROBERT M. CHANOCK
BERNARD Moss
PETERC. DOHERTY
ERLINGNORRBY
N. FIELDS BERNARD
AKIRAOYA
H. J. GROSS
J. J. SKEHEL
B. D. HARRISON
R. H. SYMONS
M. H. V. VANREGENMORTEL
Advances in VIRUS RESEARCH Edited by
KARL MARAMOROSCH
FREDERICK A. MURPHY
Department of Entomology Rutgers University New Brunswick, New Jersey
School of Veterinary Medicine University of California, Davis Davis, California
AARON J. SHATKIN Center for Advanced Biotechnology and Medicine Piscataway, New Jersey
VOLUME 46
W ACADEMIC PRESS San Diego New York Boston London Sydney Tokyo Toronto
This book is printed on acid-free paper.
@
Copyright 0 1996 by ACADEMIC PRESS, INC. All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher.
Academic Press, Inc. A Division of Harcourt Brace 19Company 525 B Street, Suite 1900, San Diego, California 92101-4495 United Kingdom Edition published by Academic Press Limited 24-28 Oval Road, London NWI 7DX
International Standard Serial Number: 0065-3527 International Standard Book Number: 0- 12-039846-X PRINTED IN THE UNITED STATES OF AMERICA 95 96 9 7 9 8 99 0 0 B C 9 8 7 6 5
4
3 2
1
CONTENTS
Poliovirus Assembly and Encapsidation of Genomic RNA
DAVIDANSARDI.DONNAC. PORTER. MARIEJ . ANDERSON. AND CASEYD . MORROW Overview ........................................................ 2 3 Genomic Organization ............................................ Poliovirus Life Cycle ............................................. 6 14 Poliovirus Virion ................................................. 19 Morphogenesis of Poliovirus ...................................... 30 RNA Encapsidation Process ....................................... 34 New Methods to Study Poliovirus Assembly ........................
I. I1. I11. IV . V. VI . VII . VIII . Complementation System to Study Poliovirus Encapsidation ........ IX . Perspectives on Poliovirus Assembly ............................... References .......................................................
39 53 56
Genome Rearrangements of Rotaviruses
I. I1. I11.
IV . V. VI . VII . VIII . IX . X.
ULRICH DESSELBERGER Discovery of Genome Rearrangements ............................. Extent of Genome Rearrangements in Rotaviruses ................. Sequence Data of Rearranged Genes ............................... Genome Rearrangements Generated in Vitro ....................... Mechanisms of Genome Rearrangements .......................... Biophysical Data ................................................. Function of Rearranged Genes and Their Products ................. Genome Rearrangements and Evolution of Rotaviruses ............. Genome Rearrangements in Other Genera of Reouiridae ............ Outlook ......................................................... References .......................................................
71 75 75 79 82 86 86 91 92 92 93
Human ImmunodeficiencyVirus Type 1 Reverse Transcriptase and Early Events in Reverse Transcription
ERICJ . ARTSAND MARKA . WAINBERG I. Introduction .....................................................
I1. Overview of Human Immunodeficiency Virus Type 1 Replication V
....
99 101
vi
CONTENTS
I11. Human Immunodeficiency Virus Type 1 ........................... IV . Human Immunodeficiency Virus Type 1 Reverse Transcription ...... References .......................................................
107 119 146
Hepadnaviruses: Current Models of RNA Encapsidation and Reverse Transription
DOROTHY A . FALLOWS AND STEPHEN P. GOFF I. I1. I11. IV . V. VI .
Introduction ..................................................... Transcription and Translation .................................... RNA Encapsidation .............................................. The Hepadnaviral Polymerase .................................... Reverse Transcription ............................................ Concluding Remarks ............................................. References .......................................................
167 172 176 180 184 192 193
Cell Types Involved in Replication and Distribution of Human Cytomegalovirus
BODOPLACHTER. CHRISTIAN SINZGER. AND GERHARD JAHN I. I1. I11. IV . V. VI . VII .
Introduction ..................................................... Determinants of Human Cytomegalovirus ......................... Organ Tropism of Human Cytomegalovirus ........................ Cells Types Involved in Acute Human Cytomegalovirus Disease ..... Viral Spread and Pathogenesis .................................... Latent Cytomegalovirus Infection ................................. Summary ........................................................ References .......................................................
197 198 216 219 232 236 241 241
Varicella-Zoster Virus: Aspects of Pathogenesis and the Host Response to Natural Infection and Varicella Vaccine
ANN M . ARVIN.JENNIFER F . MOFFAT. AND REBECCA REDMAN I. Introduction ..................................................... 265 I1. The Virus ....................................................... 266
I11. Cell-Associated Viremia in the Pathogenesis of Varicella-Zoster Virus Infection ................................................... IV. The Cell-Mediated Immune Response to Varicella-Zoster Virus ...... V. Summary ........................................................ References .......................................................
267 280 306 307
Anatomy of Viral Persistence: Mechanisms of Persistence and Associated Disease
JUANCARLOS DE
LA
TORREAND MICHAELB . A . OLDSTONE
I . Introduction ..................................................... I1. Requirements for Establishment of Viral Persistence ...............
313 315
CONTENTS
vii
111. Virus-Induced Alterations of Host Cellular Differentiated Functions
in Absence of Cytolysis ........................................... IV . Conclusions ...................................................... References .......................................................
323 338 340
The lridoviruses
TREVOR WILLIAMS I. I1. I11. IV . V. VI . VII .
Introduction ............. ........................... Classification .................................................... Structure ................... ...................... Replication ...................................................... Molecular Biology ................................................ Ecology ...................... ..................... Future Directions for Iridoviruses ................................. References .............. .....................
347 350 366 372 386 391 399 401
Molecular Biology of Luteoviruses
I . Introduction
M . A . MAYOAND V . ZIEGLER-GRAFF .................. ..........................
Mechanisms of Gene Expr ................... Particle Structure ................................................ Location of Luteovirus Replication ....................... Phytopathology .................................................. Taxonomy .............. Concluding Remarks . . . . References .... ...................................
416 417 424 435 444 449 450 453 457 457
INDEX ...........................................................
463
I1. Genome Structure ................................................ 111. Functions of Gene Products .......................................
IV . V. VI . VII . VIII .
IX .
This Page Intentionally Left Blank
t
POLIOVIRUS ASSEMBLY AND ENCAPSIDATION OF GENOMIC RNA David C. Ansardi, Donna C. Porter, Marie J. Anderson, and Casey D. Morrow Department of Microbiology University of Alabama at Birmingham Birmingham, Alabama 35294
I. Overview 11. Genomic Organization 111. Poliovirus Life Cycle A. Virus Entry and Uncoating B. Translation of Viral RNA C. Release of Individual Proteins by Viral Proteases D. Replication of Viral RNA IV. Poliovirus Virion A. Properties of Virion B. Virus Structure C. Myristylation of Poliovirus Capsid Proteins V. Morphogenesis of Poliovirus A. 5s Protomer B. 14s Pentamer C. Empty Capsid D. Provirion VI . RNA Encapsidation Process A. RNA Requirements for Encapsidation B. Poliovirus Defective Interfering Particles C. RNA Encapsidation Signals D. Subcellular Location of Encapsidation VII. New Methods to Study Poliovirus Assembly Process A. Studies of Poliovirus Assembly Process Using Recombinant Vaccinia Viruses B. Expression of Poliovirus P1 and 3CD Using Recombinant Vaccinia Virus Vectors C. Functional Significance of Poliovirus Capsid Myristylation VIII. Complementation System to Study Poliovirus Encapsidation A. Proteolytic Cleavage of Capsid Precursor B. Capsid Mutations Affecting RNA Encapsidation C. Studies on Maturation Cleavage Using Complementation System IX. Perspectives on Poliovirus Assembly References
1 Copyright 0 1996 by Academic Press,Inc. All rights of reproduction in any form resewed.
2
DAVID C. ANSARDI et al.
I. OVERVIEW The biology of poliovirus has been a subject of intense study since the 1950’s. Poliovirus is the causative agent of the paralytic disease poliomyelitis, once a major health problem in the United States that has largely been eradicated since the development of two highly effective vaccines (Sabin and Boulger, 1973; Salk, 1960). Despite control of the disease in industrialized nations, poliomyelitis continues to be a health concern in the undeveloped world. Poliovirus is a member of a family of viruses, the Picornauiridae, that includes members responsible for several diseases of humans, including the human rhinoviruses (common cold), hepatitis type A, and the coxsackieviruses (cardiac infections) (Rueckert, 1990). Other members of the Picornauiridae are responsible for important diseases of livestock, including foot-and-mouth disease virus, bovine enterovirus, and the causative agent of swine vesicular disease. Another group of picornaviruses, the cardioviruses, primarily infect mice and includes members such as mengo virus and encephalomyocarditis virus (EMCV). Poliovirus, like all of the members of the Picornauiridae, is a spherical, single-stranded RNA virus. The viral genome is a n approximately 7500-nucleotide-long RNA molecule of positive polarity (messenger-sense) and is encapsidated within a virion particle that is approximately 30 nm in diameter (Kitamura et al., 1981; Koch and Koch, 1985). The poliovirus genome has been cloned and sequenced (Kitamura et al., 1981; Racaniello and Baltimore, 1981a1, greatly facilitating the analysis of specific proteins and cis-acting regions of the RNA genome in the life cycle of the virus. Three different antigenically distinct serological types of virulent poliovirus have been identified, designated types 1 , 2 , and 3 (Koch and Koch, 1985). The polioviruses are members of the enterovirus genus of the Picornauiridae,and as such primarily inhabit the alimentary canal of the host. Most infections with poliovirus do not result in paralytic disease. However, in some instances, poliovirus spreads from the intestine to the central nervous system; lytic replication of the virus in motor neurons results in paralysis, which can often be fatal (Koch and Koch, 1985). Control of poliovirus infection in modern nations is largely based on the success of a highly effective oral vaccine consisting of live, attenuated strains of poliovirus (Sabin and Boulger, 1973). The attenuated strains given to the vaccine recipient replicate in the intestine, where they stimulate immunity against poliovirus infection, but are incapable of causing paralytic disease. The highly effective nature of the poliovirus vaccines has led to intensive research into the application of poliovirus as a vector for delivering foreign antigens to the
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
3
immune system, opening the possibility that this human pathogen might be harnessed for helpful purposes (Almond and Burke, 1990; Ansardi et al., 1994b; Porter et al., 1993a, 1995). Several developments have made poliovirus an excellent model for studying the molecular processes of viral replication. Poliovirus can be grown in many tissue culture cell lines of human and primate origin (Koch and Koch, 1985). The viral genome has been cloned and sequenced, revealing the nucleotide sequence of the RNA genome and the predicted amino acid sequences for the viral proteins (Kitamura et al., 1981; Racaniello and Baltimore, 1981a). cDNA copies of the poliovirus RNA genome are infectious and result in a productive virus infection on transfection into suitable host cells (Racaniello and Baltimore, 1981b; Semler et al., 1984). The infectivity of poliovirus cDNA has allowed the use of techniques such as site-specific mutagenesis to alter the coding sequence of the virus (Zoller and Smith, 1983). A further advance was made with the finding that positive-sense RNA genomes transcribed in uitro from poliovirus cDNA, under the control of the promoter for bacteriophage T7 RNA polymerase, were highly infectious on transfection into host cells (Van der Werf et al., 1986). In 1985, the three-dimensional structure of poliovirus was solved, providing detailed information about the structure of poliovirus capsid proteins and insight into possible mechanisms of poliovirus morphogenesis (Hogle et al., 1985). The cell surface protein receptor used by poliovirus to gain entry into the host cell has been cloned and sequenced (Mendelsohn et al., 1989). Transgenic mice which express the poliovirus receptor have also been generated, providing an animal model in which the molecular mechanisms of poliovirus pathogenesis can be studied (Ren et al., 1990).
11. GENOMICORGANIZATION The organization of the poliovirus RNA genome and the cascade of the formation of individual viral proteins are presented in Fig. 1. The positive-sense RNA genome of poliovirus is 7441 nucleotides in length (Kitamura et al., 1981; Racaniello and Baltimore, 1981a). The 5' end of the RNA genome is not linked to a 7-methylguanosine cap, but instead is covalently linked to a virus-encoded basic peptide of 22 amino acids, known as VPg (genome-linked protein), through a phosphodiester linkage between the 0 4 hydroxyl group oxygen of a tyrosine residue in VPg and the phosphate of the 5' terminal uridine residue of the RNA genome (Ambros and Baltimore, 1978; Lee et al., 1977; Morrow et al., 1984; Nomoto et al., 1976; Rothberg et al., 1980; Wimmer, 1982).The 3'
4
DAVID C. ANSARDI et al.
VPg-IRESy 33;86 W APSID-NON-C
5,
APSI7370~ 7 4 4 1
AA(AEnAA 3' n=dO
OPEN READING FRAME
POLYPROTEIN
A
A
-PLF 1[yp3lpiq P qm p q
?+N.A.+PO
A
4 m VP4
2BC
3AB
MlzC1ElO38
uncleaved
3C + 3D
(VPg)
A cleavage catalyzed by 2A A cleavage catalyzed by 3CD A cleavage cntdyzd by 3C A ~ ~ ~ ~ ; ~ ~
~
~
~
l
,
w
n
FIG.1. Poliovirus genomic organization and cascade of polyprotein processing. The poliovirus genome is a single-stranded messenger (plus sense) RNA molecule that is approximately 7500 bases in length. The 5' end of the RNA molecule is covalently linked to a small peptide, VPg, and the 3' end contains a genetically encoded polyadenylate tail that is approximately 60 nucleotides long. The first 742 nucleotides at the 5' end of the genome comprise the 5'-N"R, which contains the internal ribosome entry sequence (IRES). The poliovirus genome contains a single open reading frame encoding a 2209amino acid polyprotein precursor. Virus-encoded proteases 2A and 3C catalyze cis-acting cleavages of the polyprotein to initiate the cascade of formation of the individual viral proteins. Further processing of the viral proteins is primarily mediated by 3C, although the 3CD polyprotein catalyzes cleavage of the P1 capsid precursor to VPO, VP3, and VP1. Both 3C and 3CD catalyze cleavages a t glutamine-glycine dipeptides, whereas 2A catalyzes cleavages between tyrosine-glycine amino acid pairs. The final cleavage event occurs at an asparagine-serine amino acid pair on the interior of the virion, resulting in conversion of VPO to VP2 and VP4. The source of this cleavage is unknown, but it is speculated to occur intramolecularly.
end of the genome is polyadenylated, with a tail length of approximately 60 adenine residues. The poly(A) tail is genetically encoded by the virus rather than added by host cell polyadenylation enzymes (Kitamura et al., 1981; Racaniello and Baltimore, 1981a; Spector and Baltimore, 1975; Yogo and Wimmer, 1975). The majority of the viral RNA genome (6627 nucleotides) constitutes a long open reading frame that encodes a single translation product of 2209 amino acids. An unusually long nontranslated region of 742 nucleotides (5'-NTR) pre-
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
5
cedes the open reading frame upstream of the initiation codon used for translation of the genomic polyprotein. The 5'-NTR contains eight AUG triplets prior to the one which actually serves as the initiation codon for translation of viral proteins (Kitamura et al., 1981; Pelletier et al., 1988; Racaniello and Baltimore, 1981a). Translation of poliovirus as well as other picornavirus RNA genomes occurs by a capindependent method, in which ribosome binding occurs at an internal sequence known as the internal ribosome entry sequence (IRES) (Jang et al., 1988; Pelletier et al., 1988; Pelletier and Sonenberg, 1988; Sonenberg, 1990; Trono et al., 1988). The coding portion of the poliovirus genome is subdivided into three distinct regions, designated P1, P2, and P3 (Kitamura et al., 1981; Rueckert and Wimmer, 1984). The P1 region encodes the viral capsid proteins VP1, VP2, VP3, and VP4. The P2 region encodes nonstructural viral proteins including a protease, 2A, and 2B and 2C, which are believed to play roles in replication of the RNA genome. The P3 region encodes nonstructural proteins required for virus replication, including 3Dpo1, the RNA-dependent RNA polymerase, a protease, 3Cpr0, and the VPg protein (also known as 3B). Many of the viral proteins have important functions in polyprotein forms; for example, the membrane-bound 3AB protein is a component of the replication complex (Giachetti and Semler, 1991; Semler et al., 1982), and the 3CD polyprotein catalyzes proteolytic cleavages of the capsid precursor (Jore et al., 1988; Ypma-Wong et al., 1988a). All of the proteolytic cleavages required to liberate individual poliovirus proteins required for replication and encapsidation of the genomic RNA are catalyzed by virus-encoded proteases which cleave the primary translation product both in cis and in trans (Dewalt and Semler, 1989; Hanecak et al., 1982; Harris et al., 1990; Lawson and Semler, 1990; Palmenberg, 1990; Toyoda et al., 1986). The primary cleavage of the genomic polyprotein is an intramolecular event in which the 2A protease processes the peptide bond between a tyrosine-glycine dipeptide, releasing the 97-kDa polyprotein encoded by the P1 region (Toyoda et al., 1986). The P1 protein is a precursor from which the individual capsid proteins of the virus are derived. The virus-encoded protease 3Cpr0, acting in a polyprotein form, 3CD, is responsible for cleavage of the P1 precursor to VPO, VP3, and VP1 (Jore et al., 1988; Ypma-Wong et al., 1988a). Cleavage of VPO to VP2 and VP4 is catalyzed during or after RNA encapsidation and is widely believed to occur intramolecularly (Arnold et al., 1987; Jacobson et al., 1970). The viral proteins encoded in the P2 or P3 regions are released from polyprotein precursors by the protease 3Cpro (Hanecak et al., 1982). These cleavages occur'exclusively at glutamine-glycine dipeptides, although
6
DAVID C. ANSARDI et al.
not every glutamine-glycine dipeptide present in the genomic polyprotein is a substrate for 3C-mediated cleavages. An additional tyrosineglycine dipeptide substrate for 2A~rolies in the 3CD polyprotein, resulting in the production of two proteins, 3C' and 3D'. This cleavage may simply be a fortuitous event as poliovirus mutants without this cleavage site have no apparent growth defects (Lee and Wimmer, 1988).
111. POLIOVIRUS LIFECYCLE Infection of cells by poliovirus is associated with several pronounced cytopathic effects on the host cell, including shrinkage in cell size, an increase in intracellular membranous vesicles, deformation of the nucleus, and changes in the cell cytoskeleton (Koch and Koch, 1985). A schematic representation of the events which take place during a single cycle of poliovirus replication are depicted in Fig. 2. The virus initially attaches to the host cell by binding to a cell-surface glycoprotein molecule. The normal cellular function of the poliovirus receptor is unknown, but the predicted amino acid sequence derived from the cloning of the receptor gene indicates that the molecule belongs to the immunoglobulin-like superfamily of proteins (Mendelsohn et al., 1989). On attachment to the receptor, the virus undergoes conformational changes, and the internal capsid protein VP4 is expelled from the virion (De La Torre et al., 1992; DeSena and Mandel, 1976, 1977; Guttman and Baltimore, 1977a; Rueckert, 1990). The virus is believed to be internalized into the cytoplasm by receptor-mediated endocytosis (Madshus et al., 1984, 1985). The mechanism by which the virus releases its RNA genome across the endosomal membrane and into the cytoplasm is not understood. Once present in the cytoplasm, the messenger-sense viral RNA genome is translated on host ribosomes to yield viral proteins. Translation of the poliovirus genome is an obligatory first step because the virus does not package any of the proteins required to initiate replication of the viral RNA genome. An important consequence of poliovirus infection is the shutoff of translation of host cell mRNA, which occurs primarily as a result of cleavage of the large subunit ( ~ 2 2 0of ) the cap binding complex (eIF-4F) (Etchison et al., 1982). This cleavage is indirectly mediated by the viral protease 2Apro (Bernstein et al., 1985; Krausslich et al., 1987; Lloyd et al., 1988; Wycoff et al., 1990). Once viral proteins are synthesized, RNA synthesis occurs exponentially from approximately 30 min postinfection to 3 hr postinfection, then occurs in a linear fashion until approximately 4.5 hr postinfection followed by a rapid decline in the rate of synthesis (Rueckert, 1990).
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
7
Replication of the viral genome requires that the plus-strand RNA molecule is transcribed first to yield a complementary minus-strand RNA, which is also linked at its 5' end to VPg (Kuhn and Wimmer, 1987; Paul et al., 1987b; Richards and Ehrenfeld, 1990). This minusstrand RNA then serves as the template for synthesis of new plus strands of RNA. Synthesis of plus- and minus-strand RNA molecules is an asymmetric process, with plus strands produced in excess of minus strands by at least 10-fold. Replication of the poliovirus RNA genome occurs in association with smooth membrane vesicles which proliferate on infection, and the combination of these membranes with the viral proteins and RNA template molecules required for RNA replication is referred to as the replication complex (Caliguiri and Tamm, 1970; Ehrenfeld et al., 1970; Kuhn and Wimmer, 1987; Paul et al., 1987b; Richards and Ehrenfeld, 1990). Progeny plus-strand RNA serves as both mRNA for synthesis of additional viral proteins and as the RNA molecule encapsidated in progeny virions. Encapsidated virion RNA is linked to VPg, whereas the VPg protein is removed from the 5' end of plus-strand RNA molecules destined for translation (Hewlett et al., 1976; Nomoto et al., 1977; Petterson et al., 1977). The final aspect of the poliovirus life cycle is the formation of progeny virions. The capsid proteins assemble subviral oligomeric particles, probably prior to interaction with the RNA genome, although the precise pathway of assembly has not been deduced (Putnak and Phillips, 1981a; Rueckert, 1990).Encapsidation of plus-strand VPg-linked RNA may occur by condensation of 12 pentamers of VPO, VP3, and VP1 [(VPO-3-1),] around the RNA molecule or by insertion of VPg-linked RNA into a preformed empty capsid or procapsid consisting of 60 copies of VPO-VP3-VP1 [(VP0-3-1),,1 (Jacobson and Baltimore, 1968; Rueckert, 1990). The encapsidation process is specific for both VPglinked RNA and plus strands as packaging of minus strands does not occur, despite the presence of VPg (Nomoto et al., 1977; Novak and Kirkegaard, 1991; Petterson et al., 1978). At the end of infection, lysis of the cell occurs and virions exit, although mechanisms for active release of virus prior to lysis may exist (Tucker et al., 1993).
A . Virus Entry and Uncoating The mechanism by which poliovirus enters the host cell is poorly understood (Rueckert, 1990). Progress in this field will likely proceed at a faster pace with the identification, cloning, and sequencing of the poliovirus receptor (Mendelsohn et al., 1989). On attachment of poliovirus virions t o the glycoprotein receptor, the virus undergoes conformational changes that are marked by a conver-
8
DAVID C. ANSARDI et al.
FIG.2. Events in a single cycle of poliovirus infection. Poliovirus virions initiate infection by attaching to a glycoprotein receptor on the cell surface. The virus is believed to be internalized into the cell by receptor-mediated endocytosis. On attachment to the receptor and entry into the cell, the capsid undergoes conformational changes, and the messenger-sense RNA genome is released into the cytoplasm in a n unknown manner. The viral RNA genome is translated on host ribosomes to generate proteins required for RNA replication and encapsidation of progeny genomes. RNA replication occurs in virus-induced complexes of viral and host protein(s) that are associated with smooth membrane vesicles. RNA replication proceeds by synthesis of minus-sense RNA followed by synthesis of nascent plus strands, which occurs in excess over minus-strand formation. RNA structures in which several nascent plus strands are simultaneously being synthesized on the same minus-strand template are known as RI or replicative
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
9
sion of the sedimentation coefficient of the virion from 155s to 135s (DeSena and Mandel, 1976, 1977; Everaert et al., 1989; Fricks and Hogle, 1990; Guttman and Baltimore, 1977a;Kaplan et al., 1990).This process is associated with the expulsion of the small, myristylated internal capsid protein, VP4, from the virion. In addition to release of VP4, the conformational changes associated with attachment to the receptor also lead t o exposure of the amino terminus of the viral capsid protein VP1 on the surface of the virion, a location which is far removed from its normal location on the capsid interior (Fricks and Hogle, 1990).Exposure of the amino terminus of VP1 on the surface of the virion increases its hydrophobicity, giving the structurally altered virions the ability to bind to liposomes (Fricks and Hogle, 1990).The amino terminus of VP1 has been modeled as an amphipathic helix (this region of VP1 was unresolved in the X-ray structure), and the hypothetical formation of this structure has led to the proposal that amino-terminal residues of VP1 may be involved in forming a pore through endosomal membranes through which the viral RNA genome can be released into the cytoplasm (Fricks and Hogle, 1990). A role for the amino terminus of VP1 in virus uncoating has been supported by the phenotypes of two temperature-sensitive poliovirus mutants which contain small deletions in the VP1 amino terminus and which are defective in virus uncoating at the nonpermissive temperature (Kirkegaard, 1990; Kirkegaard and Nelson, 1990). Attachment of virus to the cellular receptor is not a guarantee of successful entry into the cell, as this process appears to be largely abortive and is associated with sloughing of a large percentage of attached, altered particles (Mandel, 1965; Rueckert, 1990).Once bound to the receptor, the virus is believed to be internalized through receptor-mediated endocytosis (Madshus et al., 1984, 1985). The process by which RNA is released from the endosomes and into the cytoplasm is not well understood. Acidification of the endosomes might be responsible for conformational changes required for capsid protein fusion with the membrane and release of RNA (Madshus et al., 1984, 1985). A study conducted on a mutant of human rhinovirus, another member of the Picornauiridae, suggested that the conformational changes associated with receptor attachment were not sufficient for RNA release into the cytoplasm, and led to the proposal that the uncoating capsid must form a membrane-associated structure, termed an
intermediate RNA. Plus strands produced in the replication complexes are either encapsidated or translated (following removal of VPg) to generate additional viral proteins. Poliovirus infection results in lysis of the host cell, allowing progeny virions to exit.
10
DAVID C. ANSARDI et al.
infectosome, responsible for injecting the RNA genome into the cytoplasm (Lee et al., 1993).
B . Translation of Viral RNA Poliovirus has evolved a cap-independent method of translation which allows it t o shut off host cell cap-dependent translation by inactivating a component of a translation initiation factor (eIF-4F) which recognizes the capped 5’ ends of host mRNA molecules (Etchison et al., 1982; Pelletier et al., 1988; Pelletier and Sonenberg, 1988; Sonenberg, 1987, 1990; Trono et al., 1988). A host cellular enzyme is believed to unlink the VPg protein from the 5’ end of poliovirus virion RNA prior to translation (Ambros and Baltimore, 1978; Hewlett et al., 1976; Lee et al., 1977; Morrow et al., 1984; Nomoto et al., 1977; Rothberg et al., 1980; Wimmer, 1982). Initiation of translation of poliovirus mRNA does not proceed by the scanning model proposed by Kozak (1989). Ribosome binding to poliovirus RNA occurs in the 5’-NTR, upstream of the initiator AUG codon, and is mediated by an internal sequence of several hundred nucleotides, which has been designated the internal ribosome entry sequence (IRES) (Jang et al., 1988; Pelletier et al., 1988; Pelletier and Sonenberg, 1988; Sonenberg, 1990; Trono et al., 1988). The determinants for recognition of the IRES by host translational machinery have not been elucidated, but secondary RNA structures present in the 5’-NTR between nucleotides 240 and 620 may mediate the internal binding of ribosomes (Sonenberg, 1990).How the IRES operates is not yet clear; possibly the ribosome binds the IRES region and scans the RNA genome until it encounters the initiator AUG codon at position 743 and begins translation. The 5‘-NTR of poliovirus type 1 contains eight AUG codons upstream of the initiating AUG that are not used as initiator codons (Kitamura et al., 1981; Racaniello and Baltimore, 1981a). However, the 100 nucleotides between the 3‘ end of the IRES and the initiator methionine at nucleotide 743 contain no AUG codons. Translation of the single long open reading frame results in the synthesis of a long polyprotein. The actual existence of this translation product in uiuo is doubtful, however, as 2Apro cleaves the P1 portion out of the growing polyprotein intramolecularly and probably cotranslationally (Toyoda et al., 1986). The shutoff of host-cell mRNA translation in poliovirus-infected cells is largely associated with cleavage of the p220 component of the cap-binding complex (eIF-4F)(Etchison et al., 1982).Inactivation of the cap-binding complex prevents binding of the translation initiation factors eIF-4A and eIF-4B to the 5’ end of mRNA molecules, which are
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
11
believed to be required for melting of RNA secondary structure in the 5' ends of mRNA molecules to allow binding of the ribosome (Sonenberg, 1990). Cleavage of the p220 protein is indirectly mediated by the viral protease 2Ap*o in some way associated with eIF-3, although the exact mechanism by which 2Apro induces p220 cleavage is not certain (Bernstein et al., 1985; Krausslich et al., 1987; Lloyd et al., 1988; Wycoff et al., 1990). Generally, 2Apro is not believed to catalyze the cleavage of p220 directly but may somehow activate a latent cellular protease which cleaves p220.
C . Release of Individual Proteins by Viral Proteases The polyprotein organization of the poliovirus RNA genome translation product dictates that proteases required to liberate individual proteins play a critical role in the life cycle of the virus. All cleavages of poliovirus proteins, except for the maturation cleavage of VPO to VP2 and VP4, have been shown to be mediated by virus-encoded proteases (Hanecak et al., 1982; Toyoda et al., 1986). Although not formally proven, the maturation cleavage of VPO to VP2 and VP4 is likely to occur through an intramolecular mechanism subsequent to encapsidation of the genomic RNA (Arnold et al., 1987; Jacobson et al., 1970). A description of the two poliovirus proteases, 2A~r0and 3Cpr0,is given in the following sections. 1 . Protease 2 A p r o
The viral protease 2Apr0 is responsible for two cleavages of poliovirus polyproteins, one which occurs in cis and the other which occurs in trans, and is indirectly involved in the inactivation of the p220 component of eIF-4F, as described in the previous section (Toyoda et al., 1986). The 2Apro protein is speculated to be a member of the serine protease family, but instead of serine the enzyme may use a cysteine residue as the nucleophile in the catalytic active site (Bazan and Fletterick, 1988). These predictions have been substantiated by data showing that mutations of putative members of the catalytic triad of residues inhibit 2A~roactivity (Yuand Lloyd, 1991). The 2Apro protease is responsible for the cotranslational primary cleavage of the poliovirus translation product which occurs in cis at a tyrosine-glycine bond, releasing the P1 capsid precursor protein (Toyoda et al., 1986).The only other confirmed cleavage of poliovirus proteins by ~ A Poccurs ~ o in trans at a tyrosine-glycine dipeptide in the 3CD polyprotein, releasing two proteins designated 3C' and 3D'. These proteins are not required for viral replication, and their formation may simply be the result of a fortuitous processing site (Lee and Wimmer, 1988).
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DAVID C. ANSARDI et al.
The determinants for substrate recognition by 2Apro have not entirely been identified but clearly involve aspects other than primary sequence, because cleavage occurs at only 2 of 10 tyrosine-glycine dipeptides in the poliovirus polyprotein (Kitamura et al., 1981; Racaniello and Baltimore, 1981a). A study demonstrated that the P2 and P1’ residues relative to the cleavage site (P2 refers to the second residue amino terminal to the scissile bond, and P1’ is the position of the residue immediately carboxyl terminal to the site of cleavage) were important determinants of cleavage site recognition, and that the primary sequence requirements for cleavage site recognition in trans were more stringent than for the cis cleavage at the site between the VP1 protein and 2A protein (Hellen et al., 1992). The requirement for the proteolytic cleavage activity of the enzyme 2A~rohas been shown to be dispensable for replication of a poliovirus replicon containing foreign gene sequences substituted for the capsid gene in vaccinia virusinfected cells (Ansardi and Morrow, 1995; Ghosh and Morrow, 1993). 2 . Protease 3Cpro
The enzyme 3Cpr0 is the viral protease responsible for the majority of poliovirus protein cleavages (Hanecak et al., 1982). The 3Cpr0 enzyme has been predicted to share structural homology with the serine family of proteases, but a cysteine residue is believed to function as the nucleophile in the catalytic triad (Bazan and Fletterick, 1988; Gorbalenya et al., 1989; Ivanoff et al., 1986; Lawson and Semler, 1991). The 3Cpro protease catalyzes proteolytic cleavages at glutamineglycine (QG) dipeptide sites in the poliovirus polyprotein (Hanecak et al., 1982). As with the 2Apro protease, 3Cpro activity is responsible for both cis and trans cleavages of the polyprotein, although the precise pathways of generation of each of the individual polypeptides is still under investigation (Dewalt and Semler, 1989; Hanecak et al., 1984; Harris et al., 1990; Lawson and Semler, 1990,1992; Palmenberg, 1990). Poliovirus is unique among picornaviruses in that all of the cleavages catalyzed by 3Cpro occur at QG bonds. In other picornaviruses, more flexibility in primary sequence at the 3Cpro cleavage sites is evident, primarily at the P1’ residue (Palmenberg, 1990).Information about the tolerance of different substituents at 3Cpro cleavage sites by the poliovirus enzyme is limited, although one study indicated that an alanine substitution for the glycine residue at the QG site between proteins 3C and 3D was compatible with cleavage, whereas more drastic substitutions inhibited cleavage (Kean et al., 1990). Determinants for 3Cpro-mediated cleavages other than a QG primary sequence must exist, because only 8 of the 13 QG dipeptides present in the poliovirus translation product are actually used as cleavage sites (Kitamura et
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
13
al., 1981; Racaniello and Baltimore, 1981a). The selection of QG cleavage sites is at least partially determined by accessibility; for example, unused QG sites present in the P1 capsid precursor protein are located within buried core regions of the capsid proteins and are likely to be inaccessible to the protease (Ypma-Wong et al., 1988b). In addition, it has been reported that cleavage site sequences must be presented in a region of flexible structure, and an alanine at the -4 amino acid position (four residues upstream of the scissile bond) has been shown to be a determinant for efficient cleavage (Blair and Semler, 1991; Mirzayan et al., 1991; Pallai et al., 1989). In addition to cleavage of viral polyproteins, poliovirus 3Cpro has been shown to cleave the transcriptional activator protein, TATA binding factor, at a QG dipeptide, and this cleavage event may be a mediator of the shutoff of host cell transcription which occurs in poliovirus-infected cells (Das and Dasgupta, 1993). An important aspect of 3Cpro-mediated cleavages of the P1 capsid precursor protein is a requirement for the sequences of the 3DpoI protein in addition t o those of 3Cpro in the form of an uncleaved 3CD polyprotein (Jore et al., 1988; Ypma-Wong et al., 1988b). Studies have indicated that 3CD is a stable viral protein and not the precursor to 3Cpro and 3Dpo1; instead, a longer polyprotein, SABCD, is the likely precursor from which 3C and 3D are liberated (Lawson and Semler, 1992; Porter et al., 1993b). The nature of the requirement of the 3D portions of the 3CD protein for cleavage of the P1 precursor have not been defined but are speculated to potentially involve interaction of hydrophobic portions of the 3D domain with hydrophobic regions of the P1 precursor (Harris et al., 1992; Krausslich et al., 1990; Nicklin et al., 1988). A more recent study suggests that a host-cell factor may be involved in a 3CD-P1 processing complex required for efficient P1 precursor cleavage (Blair et al., 1993). The 3D sequences of 3CD are apparently most important for cleavage of P1 between VPO and VP3, as cleavage of P1 between VP3 and VP1 can be catalyzed in uitro by 3Cpro at enzyme concentrations much lower than those required for cleavage between VPO and VP3 (Krausslich et al., 1990; Nicklin et al., 1988).
D . Replication of Viral RNA The poliovirus virion does not contain any of the viral proteins required for replication of the viral genome, making translation of the genome a prerequisite to RNA replication. Proteins encoded in the P2 and P3 regions of the genome are required for the replication process, whereas proteins encoded in the P1 capsid region of the genome are
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DAVID C. ANSARDI et al.
dispensable for RNA replication (W.-S. Choi et al., 1991; HadzopoulouCladaras et al., 1989; Hagino-Yamagishi and Nomoto, 1989; Kaplan and Racaniello, 1988). Mutants of many of the proteins encoded in the P2 and P3 regions are not complementable in trans,perhaps reflecting the requirement for some components of the replication machinery to remain associated with the template from which they were translated (Bernstein et al., 1986; Dewalt and Semler, 1987; Hagino-Yamagishi and Nomoto, 1989; Johnson and Sarnow, 1991). Despite years of study, the process of poliovirus RNA replication is not completely understood, although some insights have been gained from both in uitro and in viuo analyses (Kuhn and Wimmer, 1987; Paul et al., 1987b; Richards and Ehrenfeld, 1990). In addition to virally encoded proteins, there are cis-acting features of the poliovirus RNA genome required for replication, likely including the formation of secondary structures at the terminal regions of the RNA genome (Andino et al., 1990; Jacobson et al., 1970). The general strategy for poliovirus RNA replication is to first synthesize a full-length complementary strand (minus-strand RNA) to serve as template RNA molecules for the synthesis of progeny plus-strand RNA genomes. Initiation of synthesis of plus- and minus-strand RNA molecules requires recognition by the replication machinery of different 3' template ends, as the 3' ends of plus strands but not minus strands are polyadenylated (Larsen et al., 1980; Richards and Ehrenfeld, 1980). Replication of poliovirus RNA is associated with the formation of two types of fully double-stranded or partially double-stranded RNA. Completely double-stranded RNA in which a full-length plus strand is hybridized to a full-length minus strand is known as the replicative form (RF). A minus-strand RNA genome partially hybridized to a series of nascent plus strands concurrently being synthesized by different polymerase proteins on a single minus-strand template is a structure known as the replicative intermediate (RI) (see Fig. 2). IV. POLIOVIRUS VIRION A productive poliovirus infection must include synthesis of progeny virions and release of these virions from the host cell. The steps in these processes require formation of a capsid shell with icosahedral symmetry beginning with monomeric subunits and encapsidation of a single copy of a VPg-linked plus-strand RNA genome. The mature poliovirus virion serves many functions: it must protect the RNA genome from nucleases in the environment, it binds to a receptor protein on the surface of the host cell to initiate the infection process, and it
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
15
provides a mechanism for uncoating of the RNA genome on entry into the host cell.
A. Properties of Virion The poliovirus virion is a spherical particle with a diameter of approximately 30 nm (Putnak and Phillips, 1981b). The virion is composed of an icosahedral capsid shell formed by 60 copies of each of the four mature viral capsid proteins, VPl(306 amino acids, 33 kDa), VP2 (272 amino acids, 30 kDa), VP3 (238 amino acids, 26 kDa), and VP4 (69 amino acids, 7.5 kDa), and a single copy of the plus-strand RNA genome (Hogle et al., 1985). One or two copies of VP2 and VP4 may be present in the mature virion in the uncleaved precursor form, VPO (Jacobson et al., 1970), but the biological significance of the uncleaved VPO proteins in the mature virion has not been determined. The interior of the virion contains a single copy of the viral RNA genome linked to VPg (Wimmer, 1982). The poliovirus capsid does not contain sufficient basic amino acid residues to neutralize the negative charges of the RNA backbone and therefore packages numerous cations in addition to the RNA (Koch and Koch, 1985). The cations packaged include approximately 4900 K+ ions, 900 Na+ ions, 110 Mg2+ ions, and a few molecules of the polyamines putrescine and spermidine. Two types of lipid substituents are also present in the mature virion. The amino termini of the VP4 proteins are linked to a single molecule of the fatty acid myristate by an amide linkage (Caliguiri and Tamm, 1968; Chow et al., 1987; Page et al., 1988; Paul et al., 1987a). A second type of lipid, possibly sphingosine, occupies a hydrophobic pocket within the VP1 P-barrel core (Filman et al., 1989). The mature virion is a very stable structure and is resistant to concentrations of sodium dodecyl sulfate (SDS) as high as 1%,high salt concentrations, and exposure to acidic pH (Koch and Koch, 1985). The poliovirus capsid is less permeable than those of most other picornavirus members, a property which is reflected by the lower buoyant density (1.34 g/cm3) of the poliovirus virion in CsCl gradients. Other picornaviruses, such as rhinoviruses and cardioviruses, have higher densities in CsCl (1.4 g/cm3), reflecting their ability to uptake Cs+ ions, whereas the poliovirus virion is impermeable to Cs+ (Burness and Clothier, 1970; Mapoles et al., 1978; Medappa and Rueckert, 1974). The poliovirus virion has a sedimentation coefficient ( s ~ ~of, 155S, ~ ) a value typical for most picornavirus members (Putnak and Phillips, 1981a; Rueckert, 1990). The poliovirus virion also exists in a second type of conformation which is induced on attachment of the viral particle to the host cell glycoprotein receptor (Kaplan et al., 1990; Rueckert,
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DAVID C . ANSARDI et a2.
1990). The sedimentation coefficient of the virus drops from 155s to approximately 135S, and the VP4 protein is released from the particle. This altered virion is more lipophilic than the 155s virion, possibly as a result of the externalization of the VP1 amino termini (DeSena and Mandel, 1976, 1977; Fricks and Hogle, 1990; Putnak and Philips, 1981b). The conformational changes of the altered virus are also marked by a conversion from a neutral pl in the native particle to a more acidic pl and by a change in the antigenic determinants displayed on the capsid to those resembling heated or denatured virus (Koch and Koch, 1985; Putnak and Philips, 1981a). The precise function of the altered virion in virus entry has not yet been determined.
B . Virus Structure The three-dimensional structure of the type 1poliovirus virion was solved in 1985 at a resolution of 2.9 A (Hogle et al., 1985).Structures of several other picornaviruses have also been solved, including human rhinovirus type 14, mengo virus, foot-and-mouth disease virus, and poliovirus type 3/Sabin (Acharya et al., 1989; Filman et al., 1989; Luo et al., 1987; Rossman et al., 1985). The poliovirus capsid exhibits the symmetrical qualities of a T = 3 icosahedron, with 180 major subunits comprising a complete shell which has five-, three-, and twofold axes of symmetry (Hogle et al., 1985; Rossman and Johnson, 1989). Because the three major proteins which make up the asymmetric unit of the capsid are nonidentical in sequence, the poliovirus virion is said to be a pseudo T = 3 capsid, or a P = 3 capsid. The three major capsid proteins, VP1, VP2, and VP3, have a high degree of structural similarity despite major differences in amino acid sequence. Five copies of VP1 surround each of the twelve fivefold axes of symmetry of the capsid, whereas VP2 and VP3 alternate around each of the twenty threefold axes of symmetry. Each of these proteins forms a P-barrel core structural domain characteristic of the structural proteins of most spherical viruses whose structures have been solved to date (Rossman and Johnson, 1989). This p-barrel structure is formed by eight strands of P-sheet structure arranged in an antiparallel fashion. The P-sheets are named alphabetically from B to I as they occur from the amino to carboxyl termini of the proteins and form a wedge or trapezoidal-shaped structure. The B, I, D, and G P strands are contained within a twisted p-sheet structure that forms the floor and one wall of the P-barrel. The C, H, E, and F strands form a smaller, flatter wall on the opposite side of the barrel from the B-I-D-G wall. The P strands are connected by loop structures that are designated by the two P strands they connect (e.g., the B-C or D-E loops). The P-barrel cores are flanked by two a
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
17
helices. In some cases, these loops extend outward from the core structure of the capsid proteins and are responsible for the surface features of the virus exterior. Whereas the basic core structure of the P barrel is similar for each of the major capsid proteins VP1, VP2, and VP3, the connecting loops differ markedly between the different proteins and account for their unique structures. The VP4 protein is much smaller than the other three capsid proteins and lies entirely on the interior of the viral capsid. The VP4 protein is essentially a continuance of the N-terminal arm of the VP2 protein, and it is released from VP2 either during or after RNA encapsidation. The VP4 protein is involved in the networking of capsid protein termini on the capsid interior and may also be involved in interacting with the RNA genome. The amino and carboxyl termini of capsid proteins VP1, VP2, and VP3 extend outward from the core structure. On the interior of the viral capsid, terminal portions of the capsid proteins form extensive networks responsible for linking the capsid proteins together. The most striking example of this networking occurs on the capsid interior at the fivefold axes of symmetry, where the amino termini from each of the fivefold related copies of VP3 intertwine and form an unusual P-sheet structure, the P annulus, which resembles a twisted tube and represents a conserved structure among the picornaviruses of known structure (Acharya et al., 1989; Hogle et al., 1985; Luo et al., 1987; Rossman et al., 1985; Rossman and Johnson, 1989). The P annulus is flanked by five copies of a short, two-stranded antiparallel P sheet formed by residues 3-8 and 25-29 of VP4. The amino-terminal glycine residue of VP4 is covalently linked to a myristate moiety by an amide linkage (Chow et al., 1987; Paul et al., 1987a). The myristate moieties from the fivefold related VP4 molecules form a hydrophobic cluster which mediates the interaction between the amino termini of VP3 and VP4 (Chow et al., 1987; Filman et al., 1989). A third P strand is formed from portions of the amino-terminal segment of VP1, extending the p-sheet structure toward the capsid interior (Filman et al., 1989). These networking structures likely play key roles in capsid integrity. The virus capsid contains determinants necessary for interacting with the host-cell receptor glycoprotein. The three-dimensional structure of the poliovirus virion revealed a depression or canyon on the surface encircling the fivefold axes of the capsid. This canyon structure is similar t o that seen for human rhinovirus type 14 and is analogous to a pit on the surface of mengo virus (Luo et al., 1987; Rossman et al., 1985). This inaccessibility of the surface of the floor of the canyon to antibody molecules led to the canyon hypothesis which suggested that the canyon floor contained the receptor binding sites for these viruses
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DAVID C. ANSARDI et al.
(Luo et al., 1987; Rossman et al., 1985). The predominant idea behind this hypothesis was that the canyon was too narrow for the Fab portion of an antibody molecule to bind to residues lining the floor of the canyon, whereas the viral receptor might be a structure of lesser width able t o interact with the floor residues. This strategy would provide a mechanism by which the receptor binding domain of the virus could be maintained and escape immune surveillance. This hypothesis has been confirmed for human rhinovirus, and the structure of human rhinovirus type 16 complexed with extracellular domains of intracellular adhesion molecule (ICAM-11, the host receptor used by rhinovirus, has been determined (Olson et al., 1993). In addition to the receptor binding site, the exterior of the virus capsid contains the major antigenic determinants of the virus (Minor, 1990).Four major antigenic epitopes have been mapped on the surface of the virus by use of escape mutants. Three of the epitopes are present both on the intact virus and on subviral 14s pentamers (Page et al., 19881, whereas a fourth site is formed by the interaction between two pentamers and is present only in the completed, natively antigenic shell (Rombaut et al., 1990a). One of the antigenic sites, site 1, is composed of amino acid sequences in the loop connecting the B and C p strands of the VP1 protein core and has been the site of substitution of foreign antigenic determinants into the capsid to produce antigenic chimeras of poliovirus (Almond and Burke, 1990; Evans et al., 1989; Jenkins et al., 1990; Kitson et al., 1991).
C . Myristylation of Poliovirus Capsid Proteins N-Myristylated proteins are linked cotranslationally to a single molecule of the 14-carbon fatty acid myristate (n-tetradecanoic acid) by the enzyme N-myristoyltransferase (Towler et al., 1987; Wilcox et al., 1987). The myristylation reaction requires a glycine amino terminus, which is generated on the P1 capsid precursor on cleavage of the initiator methionine residue from the polyprotein (Dorner et al., 1982). The myristate moiety is linked to the glycine residue via an amide bond between the a-amino group of the glycine residue and the carbonyl carbon of the myristate molecule. The N-myristylated terminus becomes the amino terminus of the P1 capsid precursor on cotranslationa1 cleavage of the capsid precursor from the genomic polyprotein (Toyoda et al., 1986). Subsequent to cleavage of P1, the N-myristylated glycine is the amino terminus of capsid protein VPO, and finally becomes the amino terminus of VP4 on virion maturation. Electron density associated with the N-linked myristate chains was apparent in structural refinements of the poliovirus capsid (Chow et al., 1987; Filman et al., 1989).The five myristate moieties from within a common
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
19
pentameric subunit were found to cluster together, underneath the p annulus formed by the VP3 N-termini at the fivefold vertices of the capsid. The myristate cluster cradles the p annulus, implicating a role for the myristate moieties in capsid stability. The cotranslational N-myristylation of numerous other cellular and viral proteins has been reported (Schultz et al., 1988; Towler et al., 1988). The determinants for addition of myristate to a nascent peptide include a myristylation signal at the N terminus of the protein. This signal has an absolute requirement for glycine at the amino terminus of the protein and a preference for serine, alanine, or threonine at position 5 relative to the glycine acceptor (Towler et al., 1988). Substitution of the glycine residue with alanine completely abolishes myristate addition to the poliovirus P1 precursor (Krausslich et al., 1990; Marc et al., 1991). In the cases of many other viral and cellular N-myristyl proteins, the myristate moiety has been demonstrated to play a n important role in subcellular localization of the protein by contributing to a targeting signal which directs the protein to the plasmid membrane or to an intracellular membrane (Bryant and Ratner, 1990; Buss et al., 1989; Heuckeroth and Gordon, 1989; Johnson et al., 1990; Rhee and Hunter, 1987; Schult et al., 1988; Schultz and Rein, 1989; Towler et al., 1988). The myristate moiety may also participate in anchoring proteins within a lipid bilayer. The myristate moiety alone, however, is not sufficient to direct intracellular targeting to membranes (Rhee and Hunter, 1990), and several N-myristylated proteins are located in the cytosol (Schultz et al., 1988; Towler et al., 1988). By analogy to the properties of other N-myristylated proteins, the myristic acid moiety of the poliovirus capsid might participate in a targeting signal for directing capsid proteins to intracellular sites of assembly, or it may function as a membrane anchor for the capsid proteins (Chow et al., 1987; Paul et al., 1987a). A final possibility for a role for myristate in the poliovirus life cycle is at the point of uncoating. The hydrophobic myristate moieties might participate in interactions with endosomal membranes required for expulsion of the RNA genome across the membrane and into the cytoplasm. Direct analyses of a role for the myristate moieties of poliovirus in virus entry have been hampered by the complication that poliovirus mutants which do not encode a functional myristylation signal are nonviable (Krausslich et al., 1990; Marc et al., 1989, 1990).
V. MORPHOGENESIS OF POLIOVIRUS The morphogenesis of poliovirus has been a topic of intense study since the 1960’s. Much of the information gathered to date has relied
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DAVID C. ANSARDI et al.
on studies conducted in uitro and has been largely confined to assembly of empty shells rather than RNA-containing virions because reconstitution of poliovirus virions in uitro from purified components has not been achieved (Putnak and Philips, 1981a; Rombaut et al., 1984,1991). The cumulative information from poliovirus morphogenetic studies has resulted in a hypothetical pathway for assembly, which is depicted in Fig. 3. The proposed steps of assembly include ordered proteolytic cleavages and formation of capsid protein subviral particles prior to RNA encapsidation and maturation of the virion (Putnak and Philips, 1981a; Rueckert, 1990). Briefly, the hypothesized order of these events is as follows: (i)cotranslational release of the 97-kDa P1 capsid precursor from the genomic polyprotein by an intramolecular proteolytic cleavage catalyzed by the 2A protease; (ii) cleavage of the P1 precursor to the individual capsid proteins VPO, VP3, and VP1, catalyzed by the 3CD polyprotein, the form of 3Cpr0 active on the P1 precursor; (iii) assembly of five 5s promoters [(VPO-3-1),] to form a 14s pentamer intermediate [(VP0-3-1),1; (iv) assembly of a 70-80s empty capsid or procapsid consisting of 60 copies of VPO, VP3, and VP1; (v) encapsidation of VPg-plus-strand RNA genome, proceeding either from a 1 4 s intermediate or from an empty capsid, to form a provirion [(VP0-3-1),,1 or immature virion; (vi) maturation of the virion by cleavage of VPO to VP2 and VP4, an event which is probably catalyzed intramolecularly. Each of these steps and pathway intermediates are discussed in the following sections.
A. 5s Protomer The 5s protomer, or (VP0-3-1),, is the smallest identical subunit from which the complete poliovirus capsid is built (VPO is uncleaved VP4 plus VP2). The protomer is derived by proteolytic cleavage of the P1 capsid precursor polyprotein after release from the genomic polyprotein by the 2Apm protease (Toyoda et al., 1986). Cleavage of the P1 precursor occurs at two glutamine-glycine dipeptides in the precursor to generate three proteins, VPO, VP1, and VP3, which have molecular masses of 37.4,33, and 26 kDa, respectively (Koch and Koch, 1985).As reviewed in Section II1,C,1, the glutamine-glycine cleavage sites are substrates for cleavages catalyzed by the virus-encoded enzyme 3Cpro (Hanecak et al., 1982). These cleavages have been demonstrated in uitro to be catalyzed more efficiently by the polyprotein, 3CD, which consists of uncleaved 3Cpm and 3Dpo1(Jore et al., 1988; Ypma-Wong et al., 1988a). Cleavage of P1 by purified 3Cpr0 can still occur in uitro,but cleavage between VPO and VP3 requires high enzyme concentrations (Krausslich et al., 1990; Nicklin et al., 1988).
POLIOVIRUS ASSEMBLY AND RNA ENCAF'SIDATION
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FIG.3. Proposed pathways of poliovirus assembly. The poliovirus capsid proteins are initially translated as part of the genomic polyprotein and are released by a n autocatalytic cleavage by viral protease 2A as a 97-kDa precursor designated P1. The viral polyprotein 3CD catalyzes cleavage of the P1 precursor at two glutamine-glycine amino acid pairs to generate capsid proteins VPO, VP3, and VP1. The three proteins derived from a common precursor are believed to remain associated, comprising a 5s protomer subunit. Five protomers assemble 14s pentamer subviral particles [(VPO-3-1),1, which are believed to be virion precursors. Twelve pentamers assemble 755 empty capsid (procapsid) particles [(VPO-3-1),,], which some studies suggest are the direct virion precursor, with virion formation proceeding by condensation of twelve pentamers around a nucleating RNA genome. On RNA encapsidation, VPO is cleaved at a n asparagine-serine amino acid pair, releasing VP2 and VP4, a 69-amino acid protein located on the interior of the virion. The mature virion may be directly preceded by a provirion intermediate (not shown) in which the RNA genome has been encapsidated in a complete VPO-3-2 capsid.
The less efficient cleavage reaction at the VPO-VP3 bond has in part been attributed to the primary sequence near the cleavage sites. An alanine residue is present in the -4 position relative t o the scissile
22
DAVID C. ANSARDI et al.
bond at the site between VP3 and VP1, and the presence of alanine at the -4 position has been demonstrated to be a determinant for site recognition by 3Cpro and 3CD (Blair and Semler, 1991; Pallai et al., 1989). In contrast, the -4 position relative to the VPO-VP3 bond is a proline residue. The unfavorable -4 position residue also affects 3CDcatalyzed cleavage at the VPO-VP3 site (Blair et al., 1993). Substitution of this proline residue with an alanine improves cleavage by 3CD in uitro, perhaps by alleviating the requirement for a cellular cofactor to facilitate cleavage at that site (Blair et al., 1993); however, the substitution is lethal for virus growth when introduced into a poliovirus mutant RNA genome. The molecular nature of the requirement of 3D sequences to catalyze efficient cleavage of P1 to VPO, VP3, and VP1 is not understood, but speculations have been made that hydrophobic regions of 3D interact with hydrophobic regions of P1 to promote enzyme-substrate interaction (Harris et al., 1992; Krausslich et al., 1990; Nicklin et al., 1988). This hypothesis is based on the observation that 3CD activity on P1 in uitro is reduced in the presence of nonionic detergent. Speculation that the myristate molecule linked to the amino terminus of VPO might be involved in these hydrophobic interactions arose after separate studies found that 3CD did not efficiently cleave nonmyristylated P1 in uitro (Krausslich et al., 1990; Marc et al., 1989). This question has been addressed by our laboratory using an intracellular system to study proteolytic processing of P1 precursors by the 3CD enzyme (discussed in Section VI1,B). After cleavage of P1 to VPO, VP3, and VP1, the three individual proteins generated from a single precursor most likely remain associated as a 5s protomer subunit (Bruneau et al., 1983). The individual capsid proteins are always found to sediment in sucrose gradients at a 5s position, and free forms of these proteins have not been detected. Formation of the 5s protomer from the uncleaved P1 precursor is likely associated with significant conformational changes in the protomer (Hogle et al., 1985). The amino and carboxyl termini freed from one another by proteolytic cleavage are located on opposite sides of the promoter in the mature virion, indicating that structural rearrangements occur following cleavage. These structural rearrangements may be required to activate the domain responsible for the next step in assembly: formation of the 14s pentamer (Hogle et al., 1985; Rueckert, 1990).
B . 14s Pentamer On formation of the 5s protomer, the capsid subunits rapidly assemble into 14s pentamer structures consisting of five copies of each of the
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
23
individual proteins VPO, VP3, and VP1 (Putnak and Phillips, 1981a; Watanabe et al., 1962). In poliovirus-infected cells incubated with radiolabeled amino acids, incorporation of label into 14s pentamers has been reported to occur as rapidly as within 7-10 min (Putnak and Phillips, 1981b). A point of controversy in picornaviral assembly has been whether 14s pentamer formation occurs prior or subsequent to P1 precursor cleavage (Putnak and Phillips, 1981b).Evidence has been presented for some picornavirus members, including encephalomyocarditis virus (EMCV)and rhinovirus, that P1 precursors assemble to form a 13.4s pentamer precursor prior to proteolytic cleavage (McGreggoret al., 1975; McGreggor and Rueckert, 1977). The 13.4s pentamer might then be converted t o a 14s pentamer structure on cleavage of the pentamerized precursors. It has also been proposed that formation of P1 pentamers precedes proteolytic processing of the P1 precursor for hepatitis A virus (Borovec and Anderson, 1993). Information from the threedimensional structure of various picornaviruses, however, casts doubt on the likelihood of this pathway (Acharya et al., 1989; Hogle et al., 1985; Luo et al., 1987; Rossman et al., 1985). The amino termini of five VP3 proteins within a common pentamer subunit of the capsid interact with one another, forming the @-annulus structure near the 6-fold axes of symmetry. The @-annulusstructure likely provides stabilizing interactions required for pentamer formation. Proteolytic cleavage of the P1 precursor at the site between VPO and VP3 is required to free the VP3 amino termini from the carboxyl end of VPO. Unless the interprotomer interactions that occur within a pentameric subunit of the mature virion are different from those in the subviral pentamer, cleavage of the precursor would appear to be a prerequisite for assembly. In studies of cell-free assembly of in vitro translated EMCV capsid proteins, complete cleavage of the precursor was required for pentamer formation (Palmenberg, 1990; Parks and Palmenberg, 1987). The assembly of a pentamer as a capsid precursor is compatible with the notion that construction of an icosahedral capsid from monomeric subunits requires a stepwise assembly process, with formation of one building block required to activate the domains necessary for assembling the next intermediate structure (Caspar and Klug, 1962; Rossman and Johnson, 1989).A controversy exists, however, about whether the pentamer is the direct precursor to the poliovirus virion (Putnak and Philips, 1981a; Rueckert, 1990). A few lines of evidence suggest that 1 4 s pentamers are the immediate precursor to the virion. In pulse-chase metabolic radiolabeling experiments using poliovirusinfected cells, radiolabel flows from 5s protomers into 14s pentamers and into both empty capsids and virions (Jacobson and Baltimore,
24
DAVID C. ANSARDI et al.
1968). Experiments in which the drug guanidine was used to inhibit poliovirus RNA replication in poliovirus-infected Mi0 cells provided evidence that 14s pentamers are direct precursors to the virion (Ghendon et al., 1972). When RNA replication is halted by guanidine treatment, virion formation also abruptly halts. Under guanidine treatment conditions, capsid protein radioactivity in Mi0 cells was found to accumulate in 14s pentamers. On removal of the guanidine, the 14s pentamer radioactivity was rapidly converted to virions without formation of detectable empty capsids. An additional line of experimental evidence has supported the hypothesis that the 14s pentamer is the direct precursor to the virion (Rombaut et al., 1990b). In cells infected with poliovirus at 30"C, radioactivity in radiolabeled capsid proteins was found to accumulate in 14s pentamers without formation of empty capsids or RNA-containing virions. On shift of temperature to 37"C, a temperature permissive for virion and empty capsid assembly, radioactivity in the 14s pentamer fractions was rapidly chased into mature virions without significant accumulation of an empty capsid intermediate. The investigators could not, however, rule out the possibility that RNA encapsidation was occurring so rapidly that an empty capsid intermediate was obscured. This explanation might also account for the lack of detection of an empty capsid intermediate in the guanidineinhibition studies in Mi0 cells (Ghendon et al., 1972). Electron microscope immunocytochemistry studies have offered further evidence that the 14s pentamer is the direct precursor to the virion (Pfister et al., 1992). These studies were conducted with subcellular fractions containing virus-induced smooth membrane vesicles associated with replication complexes. By using monoclonal antibodies specific for subsets of capsid protein structures, 14s pentamers were detected around the peripheries of the replication complexes in association with the membrane vesicles. In contrast, empty capsids could not be detected in association with the complexes by these methods. The hypothesis was made that 14s pentamers associate with the replication complexes and interact with pools of nascent RNA chains being released from the replication complexes (Troxler et al., 1992). Interestingly, solubilization of membrane-associated replication complexes with nonionic detergents resulted in conversion of 14s pentamers to natively antigenic empty capsids. The investigators suggested that linkage to a membrane support prevents 14s pentamers from coalescing into a capsid until interaction with RNA takes place. When the membranous support is dissolved with nonionic detergents, the 14s pentamers may rapidly assemble empty capsids, opening the possibility that empty capsids previously reported to be associated with replication complexes may actually be artifacts produced on lysis of the host cell and solubilization of membrane-associated pentamers.
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
25
C. Empty Capsid The empty capsid or procapsid has probably been the most controversial intermediate in the proposed assembly pathway. Empty capsids are composed of sixty copies of each of the individual capsid proteins VPO, VP3, and VP1 (Putnak and Philips, 1981a; Rueckert, 1990). In sucrose density gradients, the empty capsid sediments at a rate approximately one-half that of mature virions, and it has a sedimentation coefficient reported t o be between 65s and 80s (Rueckert, 1990). In addition, the empty capsid can exist in two different conformations that are distinguishable by the antigenic epitopes they display and their relative stabilities. One type of empty capsid is very labile and displays the same antigenic epitopes as native virus (N-antigenicity) and is also referred to as the natural empty capsid (Gauntt eta,?.,1981; Maronginu et al., 1981; Putnak and Phillips, 1982; Rombaut et aZ., 1982, 1984; Rueckert, 1990). The other type of empty capsid is much more stable and has antigenicity consistent with heated poliovirus virions, which display a completely different subset of antigenic epitopes (H-antigenic) (Maize1et al., 1967).The labile empty capsid can be dissociated under mild alkaline conditions and is rapidly converted to the H-antigenic form if heated even briefly after extraction from the infected cell (Maronginu et al., 1981; Onodera et al., 1986). An additional difference in the two types of empty capsids is their reported sedimentation velocities. Native antigenic empty capsids are reported to sediment in sucrose gradients at a 65-708 position, whereas the more stable H-antigenic empty capsids sediment at a position of 8 0 s (Putnak and Phillips, 1982; Rombaut et al., 1982). These properties may reflect a more condensed capsid structure in the stable H-antigenic particle (Koch and Koch, 1985). In the course of studies of poliovirus assembly, the empty capsid has been proposed in conflicting hypotheses to be the direct precursor to the virion (Jacobson and Baltimore, 1968),a by-product of assembly in which excess pentamers assemble empty capsids (Koch and Koch, 19851, and an artifact of solubilization methods used to analyze subviral particles by their sedimentation properties (Pfister et al., 1992).A major problem with acceptance of the empty capsid as the direct precursor to the lririon is a conceptual one because envisioning how a 7450-base RNA genome can be tightly wound and threaded into a preformed empty shell is difficult (Putnak and Phillips, 1981a; Rueckert, 1990). Nevertheless, numerous experiments have been presented which suggest that the empty capsid, or procapsid, is the direct precursor to the virion. Poliovirus 14s subunits can self-assemble empty capsids in vztro in the absence of full-length poliovirus RNA (Onodera and Phillips, 1987;
26
DAVID C. ANSARDI et al.
Phillips, 1971; Phillips and Wiemert, 1978; Rombaut et al., 1991).This inherent ability of 14s pentamers to form empty capsids was taken as evidence for their precursor role in virion morphogenesis (Putnak and Phillips, 1981b). In some experiments, the radiolabel in poliovirusinfected cells appeared in empty capsid particles (15-20 min) before appearing in mature virions (20-30 min), and this observation was taken as evidence for a precursor role for empty capsids in virion morphogenesis (Putnak and Phillips, 1981b).In addition, pulse-chase radiolabeling experiments conducted using cells infected with footand-mouth disease virus (FMDV) in the presence of protein synthesis inhibitors indicated a flow of radioactivity from 5s to 14s to empty capsids to virions (Yafal and Palma, 1979).Radiolabeling experiments conducted in the presence of guanidine in poliovirus-infected HeLa cells showed that radiolabel accumulated in empty capsids quickly on inhibition of RNA replication, and the empty capsid-associated radiolabel was rapidly chased into virions on removal of the drug (Fiszman et al., 1972; Jacobson and Baltimore, 1968). This experiment was similar to that conducted in Mi0 cells in which radioactivity accumulated in 14s pentamers in the presence of guanidine was rapidly chased into virions on removal of the inhibitor (Ghendon et al., 1972). Two early studies conclude that poliovirus empty capsids were associated with the viral RNA replication complexes (Caliguiri and Compans, 1973; Yin, 1977);however, those findings have been challenged more recently as being an artifact of solubilization methods used to extract capsid particles from the membranous complexes (Pfister et al., 1992). The inherent ability of 14s pentamers to assemble empty capsids has been studied extensively in uitro (Phillips, 1969, 1971; Phillips et al., 1968, 1980; Phillips and Wiemert, 1978; Putnak and Phillips, 1981a; Rombaut and Boeye, 1991; Rombaut et al., 1984, 1991). On incubation at 37"C, purified 14s pentamers isolated from poliovirusinfected cells assemble empty capsids (Phillips, 1971; Phillips and Wiemert, 1978; Rombaut et al., 1991). Attempts to reconstitute virions in uitro from purified virion RNA and 14s pentamers or empty capsids isolated from poliovirus-infected cells have failed (Putnak and Phillips, 1981b; Rombaut and Boeye, 1991). In in uitro assembly experiments, purified 14s pentamers at sufficient concentrations assemble empty capsids in the absence of additional poliovirus-specific factors, but the empty capsids which form are H-antigenic (Putnak and Phillips, 1982). When 14s pentamers were incubated in the presence of a poliovirus infected-cell extract, however, empty capsids were assembled much more rapidly, and the resulting empty capsids displayed native antigenic epitopes (Phillips, 1969; Putnak and Phillips, 1981b, 1982; Rombaut et al., 1984). These findings led to the search for the morphopoietic factor present
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
27
in poliovirus-infected cells which facilitated assembly of empty capsids from 14s pentamers and which conferred native antigenicity on the in uitro assembled shells. Although the source of the assembly-promoting activity originally was not believed to be simply the endogenous supply of 1 4 s pentamers provided by the infected cell extracts (Phillips, 1969; Putnak and Phillips, 1981b), extracts of cells infected with poliovirus defective interfering particles, which do not encode functional capsid proteins (discussed more extensively in Section VI,B), lacked the assembly-promoting activity (Phillips et al., 1980). The source of the assembly-promoting activity has been identified by Rombaut et al. (19911, who demonstrated a threshold concentration (-1.6 nM) above which purified 14s pentamers rapidly assembled empty capsids in uitro when incubated at 37°C.The ability of infected cell extracts to facilitate the assembly of 14s pentamers at concentrations below the assembly threshold was directly correlated with the supply of 1 4 s pentamers provided in the infected cell extract which brought the final concentration of 14s subunits above the assembly threshold. These observations demonstrated that the assembly-promoting activity was simply the additional 14s pentamers provided by the extract. However, the factor contributing to native antigenicity of empty capsids assembled in the presence of an infected cell extract appears to be different. The VP1 core p barrel contains a hydrophobic pocket normally occupied by an unidentified lipid molecule, probably sphingosine (Filman et al., 1989). This pocket is analogous to the pocket in human rhinovirus VP1 which binds a series of candidate antiviral drugs known as WIN compounds, which act at multiple levels in preventing infection of cells by drug-complexed virions (Badger et al., 1988; Fox et al., 1986; Smith et al., 1986). The WIN compounds have also been demonstrated t o inhibit poliovirus uncoating (Fox et al., 1986) and to protect poliovirus N-antigenic empty capsids from thermal denaturation (Rombaut and Boeye, 1991). Rombaut et al. (1991) found that purified 14s pentamers assembled empty capsids with H-antigenicity in uitro. However, if the drug molecule disoxaril, a WIN compound, was provided in the assembly reactions, the empty capsids which assembled displayed native antigenic epitopes (Rombaut and Boeye, 1991). The authors speculated that the drug mimics a lipid compound provided by the infected cell extracts by binding in the VP1 pocket and promoting native antigenicity of the assembled empty capsid particles. The assembly-enhancing features of infected cell extracts were thus twofold: supply of 1 4 s pentamers to bring concentrations above the threshold required for assembly and provision of some compound, possibly a lipid molecule which was mimicked by the drug disoxaril, to maintain native antigenicity. The existence of different forms of the empty capsid (Nand H-anti-
28
DAVID C. ANSARDI et al.
genic) has led to different interpretations about the role the empty capsid plays in morphogenesis. When extracted from infected cells, empty capsids were originally reported to be very stable, H-antigenic structures (Maize1et al., 1967).Because of this property, empty capsids were thought to not be capable of equilibrating with 14s pentamers. Thus, Jacobson and Baltimore (1968) proposed the procapsid hypothesis in which the viral genome is directly inserted into a procapsid. Subsequent studies have shown, however, that empty capsids in uiuo likely exist in a natively antigenic, dissociable state (Maronginu et al., 1981). The observance of H-antigenic empty capsids probably reflects handling methods, as natively antigenic empty capsids are rapidly thermally denatured (Maronginu et al., 1981). Rapid thermal denaturation of 14s pentamers also occurs in uitro on incubation at 37°C (Rombaut and Boeye, 1991). The lability of the N-antigenic empty capsid suggests that interconversion between 14s and empty capsid forms may occur in uiuo (Rueckert, 1990). This property is consistent with a model in which empty capsids are a storage depot for excess pentamers which can readily dissociate back into 14s pentamers (Maronginu et al., 1981; Rueckert, 1990). The lability of the empty capsid might also reflect, however, a more flexible structure which can uptake RNA (Koch and Koch, 1985). Several methods for how the RNA genome could be inserted into an empty capsid have been proposed. One model suggests that the energy released during synthesis of the RNA genome provides the driving force for inserting the RNA into the capsid (Rueckert, 1990). Such a strategy implies a very close link between RNA synthesis and encapsidation, which is supported by studies which have noted that guanidine inhibition of RNA synthesis is associated with a concurrent inhibition of RNA encapsidation (Caliguiri and Tamm, 1968; Fiszman et al., 1972). Another hypothesis suggests that empty capsids may not have a full complement of capsid subunits and may contain holes available for inserting an RNA genome, a hypothesis primarily based on studies of mengo virus assembly in which empty capsid structures were believed to contain 10 rather than 12 pentameric subunits (Lee and Colter, 1979). Other hypotheses have from time to time been based on more elusive intermediates that may exist in poliovirus-infected cells. A few reports have offered evidence for the transient formation of half-shells, with sedimentation coefficients of approximately 50s (Corrias et al., 1987; Koch and Koch, 1985; Lee et al., 1978;Rombaut et al., 1985). A half-shell might possibly serve as a direct virion precursor, with the RNA being enclosed within the two halves. Interestingly, electron microscopy studies of subcellular fractions containing poliovirus replication complexes have identified structures attached to the
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
29
membrane vesicles which had a half-shell appearance (Pfister et al., 1992).
D . Provirion Whatever the capsid precursor serving as the RNA-binding particle may be, the RNA encapsidation step appears to lead to production of a provirion particle in which the RNA genome has been encapsidated in a completed shell composed of 60 copies of VPO, VP3, and VP1 (Putnak and Phillips, 1981a; Rueckert, 1990). The original evidence for the existence of this intermediate was based on sedimentation studies in which a 125s shoulder was observed on the 155s virion peak on sucrose density gradients (Fernandez-Thomas and Baltimore, 1973; Fernandez-Thomas et al., 1973). The capsid protein composition of this shoulder was found to be enriched for VPO over the 155s peak, in which most if not all VPO had been cleaved to VP2 and VP4. Subsequent studies reported the sedimentation coefficient of these provirions or immature virions to be 150s (Guttman and Baltimore, 197713). These findings led to the speculation that cleavage of VPO to VP2 and VP4 occurred subsequent to RNA encapsidation and possibly by an intramolecular mechanism. l ' b o studies have provided further evidence for the existence of the provirion intermediate. Compton et al. (1990) isolated a temperaturesensitive mutant of poliovirus with a glutamine substitution for arginine at residue 76 of VP2 which accumulated provirions at the nonpermissive temperature. These studies showed that RNA encapsidation and VPO cleavage to VP2 and VP4 could be unlinked. The resulting provirion particles, however, were not infectious. A subsequent study of site-directed mutants of human rhinovirus type 14 has more thoroughly characterized the provirion particle (Lee et al., 1993). Rhinovirus mutants with a threonine substitution for asparagine at the carboxyl terminus of VP4 (at the cleavage site between VP4 and VP2) accumulated provirion particles in cells transfected with an in uitro transcribed RNA genome encoding the substitution. The provision particles were shown to be noninfectious, and the lack of infectivity was traced to a step in the uncoating process of rhinovirus. Provirion particles attached to host receptors normally and underwent the associated conformational changes (155s to 125s conversion). The block with the provirion mutant appeared to occur at the level of RNA release, leading to the hypothesis of an infectosome intermediate in the uncoating pathway in which a membrane-associated virus particle expels its RNA across an endosomal membrane and into the cytosol. Formation of this structure is apparently dependent on cleavage of VPO t o VP2
30
DAVID C. ANSARDI et al.
and VP4. At the other end of the spectrum, why the virus has evolved to delay cleavage of VP4 from VP2 until after RNA encapsidation takes place is not known. Intact VPO may be needed to maintain the required conformations of the 5s and 14s capsid subunits for subsequent assembly events (Koch and Koch, 1985). The maturation cleavage of VPO to VP2 and VP4 is the final proteolytic cleavage in the maturation of poliovirus capsid proteins (Arnold et al., 1987; Hellen and Wimmer, 1992a,b).Following the solution of the three-dimensional structures of several picornaviruses, a potential mechanism for how maturation cleavage might occur was proposed (Arnold et al., 1987). In this autocatalytic model, a serine residue in VP2 (amino acid number 10 in VP2), which forms a hydrogen bond with the carboxyl terminus of VP4 in the mature virion, was believed to be the residue responsible for nucleophilic attack on the peptide bond. Because a nearby histidine residue, which would serve as the proton-abstracting base for the nucleophilic attack, was not present, a nitrogenous base from the RNA molecule was speculated to activate the serine residue for nucleophilic attack. This model provided a convenient explanation for how the maturation cleavage event was dependent on RNA encapsidation since RNA would be required to complete the catalytic triad of the protease. This theory was disproved, however, when Harber et al. (1991) demonstrated that the putative catalytic serine residue could be substituted with other amino acids without affecting the maturation cleavage event. Despite the collapse of this model, a role for the RNA genome in contributing to the catalytic site of the intramolecular protease has not been ruled out.
VI. RNA ENCAPSIDATION PROCESS As discussed in the preceding sections, the precise pathway leading to the formation of poliovirus virions is a major unresolved question in poliovirus morphogenesis. Not only is the identity of the direct capsid precursor to the virion not known, but the mechanisms involved in capsid protein-RNA genome interaction are also not well understood. The two components of this interaction, the capsid protein determinants involved in RNA binding and the regions of RNA specifically recognized by the capsid proteins, have not been identified. The threedimensional structure of poliovirus provided few clues about this interaction, as the encapsidated RNA molecule does not adopt the icosahedral symmetry of the capsid shell (Hogle et al., 1985).Because the VPg-linked RNA molecules exist in multiple conformations within a
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
31
crystal lattice of poliovirus virions, structural determinations for the RNA genome and VPg could not be made.
A . RNA Requirements for Encapsidation Poliovirus virions encapsidate only plus-strand VPg-linked genomic RNA (Lee et al., 1977; Novak and Kirkegaard, 1991; Wimmer, 1982). Beyond these characteristics of encapsidated RNA, few other determinants required for packaging of a poliovirus RNA molecule have been recognized. Most information regarding the RNA requirements for encapsidation has come from studies of defective interfering (DI) particles of poliovirus (Cole, 1975). Studies of the naturally occurring DI genomes, which contain in-frame deletions within the P1 coding portion of the genome, indicate that shorter RNA genomes can be encapsidated and have suggested a minimal size constraint for encapsidation of 80-87% the length of the wild-type genome (Cole et al., 1971; Kuge et al., 1986; Lundquist et al., 1979). A poliovirus RNA genome 108% of the length of the wild-type genome, namely, a genetically engineered dicistronic RNA genome containing an IRES element of EMCV inserted between the P1 and P2 genes, has been demonstrated t o be compatible with virion formation, indicating that the virus can accommodate a lengthier RNA molecule (Molla et al., 1992). Beyond the ability of the poliovirus capsid to accommodate genomes of different sizes, and the requirement for VPg linkage for encapsidation, few other properties of poliovirus RNA necessary for encapsidation have been uncovered.
B . Poliovirus Defective Interfering Particles Cole et al. (1971) were the first to describe the appearance of DI particles within populations of poliovirus passaged at very high multiplicities of infection [>200 pfu (plague-forming units)/celll. The DI particles were first identified by their slower sedimentation properties in sucrose density gradients and were then shown to have lower buoyant densities in CsCl density gradients relative to wild-type virus (1.31-1.325 g/cm3 versus 1.34 g/cm3 for wild-type). The DI particles were found t o exhibit properties of interference with wild-type poliovirus production in mixed infections and were shown to enrich in proportion to wild-type virions on multiple passages (Cole and Baltimore, 1973b,c). Deletions within the DI genomes were mapped to the 5’ region of the genome and were believed to have limitations in minimal size permissible for propagation, with the smallest naturally oc-
32
DAVID C. ANSARDI et al.
curring DI genomes identified having approximately 80%of the length of wild-type genomes (Cole and Baltimore, 1973a; Cole et al., 1971). Subsequently, other investigators reported separate generation of DI particles in populations of poliovirus passaged at high multiplicities of infection (Kajigaya et al., 1985; Lundquist et al., 1979). Electron microscopy studies provided additional evidence that these genomes contained deletions in the 5’ region of the RNA genome in the region believed to encode the capsid proteins (Lundquist et al., 1979). Determination of the nucleic acid sequences of several DI genomes of poliovirus type 1 Sabin confirmed that DI genomes contained deletions in the P1 capsid region which maintained the translational reading frame for the P2 and P3 regions of the genome (Kuge et al., 1986). Poliovirus DI genomes containing deletions in the P2 or P3 regions have never been identified, reflecting the property that replication of genomes encoding mutations of P2 and P3 region proteins is not readily complementable by viral proteins provided in trans (Kuhn and Wimmer, 1987; Page et al., 1988; Paul et al., 1987b; Richards and Ehrenfeld, 1990). The P1 deletions characterized by sequence analysis appeared to have specific boundaries within the P1 gene at both the 5‘ and 3’ ends, as the naturally occurring in-frame deletions were all contained within an internal segment of the P1 gene between nucleotides 1226 and 2705, encompassing much of the VP2 and VP3 genes (Kuge et al., 1986).
C . RNA Encapsidation Signals The finding that portions of the P1 gene were maintained in every isolate of naturally occurring DI genomes of poliovirus type 1 Sabin led to the speculation that portions of the P1 coding region might contain cis elements required for RNA replication and/or encapsidation of the genome (Kuge et al., 1986). Kaplan and Racaniello (1988) generated in uitro transcribed poliovirus RNA genomes which contained genetically engineered deletions in the P1 gene, encompassing all but the final 320 nucleotides of the P1 gene. On transfection into HeLa cells, the deletion-containing genomes replicated normally, indicating that most, if not all, of the P1 gene is dispensable for replication of the RNA genome. The investigators did not report on whether the deletion-containing RNA genomes could be encapsidated if transfected into cells infected with wild-type helper poliovirus. Another report has demonstrated that sequences at the 5’ end of the P1 coding region are dispensable for both RNA replication and encapsidation. These studies demonstrated replication and encapsidation of a poliovirus RNA replicon containing a reporter chloramphenicol
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
33
acetyltransferase (CAT) gene inserted for part of the P1 gene, beginning with the AUG codon for translation initiation (Percy et al., 1992). The CAT gene replaced P1 gene sequences from nucleotides 756 to 1805, indicating that an encapsidation signal does not exist in the 5' portion of the P1 gene. Further evidence that P1 regions are dispensable for RNA replication was provided by W.-S. Choi et al. (1991), who showed that internal regions of the P1 gene could be substituted with foreign gene segments encoding human immunodeficiency virus type 1 (HIV-1) proteins in the same translational reading frame as the poliovirus polyprotein. Experiments have demonstrated that RNA genomes with foreign genes substituted for the complete P1 gene can be encapsidated (Ansardi et al., 199413; Porter et al., 1995). Interestingly, the presence of nucleotides 743-959, which encompass the VP4 gene, appeared to facilitate encapsidation of the replicon RNA, pointing to the possibility that this region of the poliovirus genome might be involved in encapsidation after all (Porter et al., 1995).
D . Subcellular Location
of
Encapsidation
Successful encapsidation of poliovirus RNA might require interaction of the capsid proteins with the RNA at a specific subcellular location. Poliovirus RNA replication occurs in replication complexes associated with smooth intracellular vesicles, and capsid proteins of poliovirus have also been found in association with smooth vesicles as discussed in Section I11 (Caliguiri and Compans, 1973; Caliguiri and Mosser, 1971; Ehrenfeld et al., 1970; Girard et al., 1967; Hewlett et al., 1976). In RNA-labeling experiments conducted using short pulses of incubation with PHluridine, virions associated with the smooth membrane fractions were found to have higher specific activity than those found in other subcellular fractions (Caliguiri and Compans, 1973), implying that the most recently made virions were associated with the smooth membrane complexes. Immunoelectron microscopy studies have demonstrated the presence of capsid-related particles, probably 1 4 s pentamers, associated with the peripheral membrane vesicles of replication complexes isolated from poliovirus-infected cells (Hewlett et al., 1976). Poliovirus capsid proteins, in a precursor form to virions, may possibly be directed to and associate with intracellular membrane vesicles in a location required for interaction with newly synthesized RNA genomes (Hewlett et al., 1976; Koch and Koch, 1985).The mechanisms by which capsid proteins associate with the membranes is not understood. One hypothesis suggests that the myristate molecule linked to the amino terminus of VPO mediates association with intracellular membranes (Chow et al., 1987; Paul et al., 1987a). The lipo-
34
DAVID C. ANSARDI et al.
philic amino terminus of capsid protein VP1 might represent another candidate determinant for capsid protein association with intracellular membranes (Filman et al., 1989).
VII. NEWMETHODS TO STUDY POLIOVIRUS ASSEMBLYPROCESS Until recently, much of the information about picornavirus assembly was gathered from attempts to reconstitute the assembly process in uitro (Putnak and Phillips, 1981a). Although empty capsids assemble from 14s pentamer subunits in uitro, the formation of virions from purified components has not been achieved. Molla et al. (1991)reported on the de nouo synthesis of poliovirus in uitro. This system relied on in uitro translation of poliovirus proteins from full-length genomic RNA in the presence of intracellular membranes which in turn resulted in replication of the RNA genome and encapsidation of RNA to form infectious poliovirions. Other methods of studying the poliovirus assembly process have relied on isolation of temperature-sensitive mutant polioviruses or on the recovery of mutant viruses on transfection of in uitro transcribed RNA genomes containing site-directed mutations (Comptonet al., 1990; Kirkegaard, 1990; Kirkegaard and Nelson, 1990; Marc et al., 1990; Moscufo and Chow, 1992; Moscufo et al., 1991; Reynolds et al., 1992). An inherent problem exists in characterizing these types of poliovirus mutants because the poliovirus replicase, with no known editing capabilities, is prone to error, and reversions of mutations arise with great frequency (De La Torre et al., 1992). RNA genomes encoding capsid mutations replicate normally since capsid proteins are dispensable for replication (W.-S. Choi et al., 1991; Hagino-Yamagishi and Nomoto, 1989; Kaplan and Racaniello, 19881, so opportunity for reversion of mutations in the capsid gene is great. Thus by transfecting in uitro transcribed RNA genomes into cells and recovering mutant viruses, it becomes difficult to assess definitively whether intermediate phenotypes observed are a reflection of populations of revertants (Marc et al., 1990, 1991). In addition, it was also difficult to recover enough material from the transfected cells to characterize the physical features of the subviral particles thoroughly.
A . Studies of Poliouirus Assembly Process Using Recombinant Vaccinia Viruses To understand further the molecular details of poliovirus assembly, it was critical to develop an intracellular system in which the early
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
35
events in the assembly of poliovirus could be studied without having to depend on recovery of mutant polioviruses or on expression of capsid proteins from a replicating template, which had the potential to revert a capsid gene mutation. Vaccinia virus vectors have several features which make them attractive for the expression of poliovirus proteins (Mackett et al., 1985). Among these are the following: the cytoplasmic site of vaccinia virus replication ensures that messenger RNA molecules encoding poliovirus proteins are not exposed to nuclear splicing machinery; the vaccinia virus genome is capable of accepting large amounts of foreign DNA; and generation of recombinant vaccinia viruses is greatly facilitated by recombination plasmids that direct homologous recombination of foreign genes into the thymidine kinase gene of the vaccinia virus genome, thereby providing a mode of selection because the resulting recombinants do not synthesize thymidine kinase. Finally, the recombination plasmid coexpresses P-galactosidase, providing another selection marker for recombinant viruses (Chakrabarti et al., 1985).
B . Expression of Poliovirus PI and 3CD Using Recombinant Vaccinia Virus Vectors Previous studies demonstrated that stable recombinant vaccinia viruses could not be isolated which contained the poliovirus 2A gene (Jewel1 et al., 1990; Turner et al., 1989). The lethal effect of 2A~r0was probably associated with its role in shutting off translation of capped mRNA molecules (Etchison et al., 1982; Krausslich et al., 1987; Lloyd et al., 1988; Wycoff et al., 1990). This observation was important because the carboxyl terminus of the P1 precursor is generated by a cisacting proteolytic cleavage by 2Apr0 (Toyoda et al., 1986). To overcome the need for 2A-mediated cleavage to generate an authentic P1 carboxyl terminus, termination codons were engineered into a recombinant P1 gene downstream of the codon for the authentic tyrosine carobxyl terminal residue (Ansardi et al., 1991). Infection of cells with recombinant vaccinia virus that contains the P1 gene (VVP1) resulted in expression of a 97-kDa protein. Coinfection of cells with VVPl and a second recombinant vaccinia virus, VVP3, which expressed the 3CD protein (Porter et al., 1993b),resulted in expression of both P1 and 3CD in coinfected cells. The P1 precursor was rapidly cleaved to VPO, VP3, and VP1 by the 3CD protease (Fig. 4). In addition, these cleavage products assembled both 14s pentamers and empty capsid particles. The rapidity with which P1 precursors were cleaved to VPO, VP3, and VP1 and assembled subviral particles in VVPl/VVP3-~oinfected cells demonstrates that all of the virally encoded information required
36
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FIG.4. Analysis of poliovirus assembly using recombinant vaccinia viruses which express P1 and 3CD. In previous studies, we described construction and characterization of recombinant vaccinia viruses that express the poliovirus capsid precursor protein P1 and the viral protease 3CD (Ansardi et al., 1991). In this system, cells are coinfected with vaccinia viruses VV-P1 and W-P3. The infection of cells with VV-P1 results in the expression of the poliovirus P1 protein. Expression of 3CD from VV-P3 results in the proteolytic processing of P1 to give the capsid proteins VPO, VP3, and VP1. Once proteolytic processing occurs, the capsid proteins assemble into poliovirus subviral intermediates: 55 protomers, 14s pentamers, and 75s empty capsids. Because no poliovirus RNA is present in this system, the final end point of the assembly is the 75s empty capsid in which VPO is not cleaved to VP4 and VP2.
for these stages in poliovirus assembly is present in the P1 and 3CD proteins and can occur in the complete absence of replicating poliovirus RNA. Poliovirus replication occurs in association with intracellular membranes (Kuhn and Wimmer, 1987; Paul et al., 1987b; Richards and Ehrenfeld, 19901, and more recent studies suggest that poliovirus P2 and P3 proteins required for replication are localized on mem-
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branes via common precursor polyprotein (Lawson and Semler, 1992). In contrast, the P1 capsid precursor is cotranslationally cleaved away from the P2 and P3 proteins by the autocatalytic activity of 2Apr0 (Toyoda et al., 1986) and presumably can diffuse away from the other viral proteins. The results of our studies demonstrate that the proteolytic cleavage and subviral particle assembly steps can occur independently of the replication complexes. On formation of assemblycompetent capsid subunits, the capsid proteins may then be targeted through some unknown mechanism to sites of RNA encapsidation. Such a strategy of partitioning capsid assembly away from sites of RNA replication may ensure that immature or assembly-incompetent capsid subunits are restricted from entering sites of encapsidation. Alternatively, RNA released from the replication complexes may diffuse into the soluble sites of capsid assembly. This mechanism would ensure that capsid proteins, with affinity for viral RNA, would not enter the replication complexes and potentially interfere with RNA synthesis. The rapid assembly of VPO, VP3, and VP1 proteins generated in VVPl/VVPS-coinfected cells suggests that P1 precursors and 3CD proteins might form a processing/assembly complex in which P1 precursors are brought together and can rapidly form 14s pentamers on proteolytic processing. The vast majority of radiolabeled VPO, VP3, and VP1 recovered from VVPl/VVP3-~oinfectedcells was present in 14s pentamer or 75s empty capsid fractions. Although no evidence for assembly of specific oligomeric structures from P1 capsid precursors has been found, it was possible that such structures might be very labile and subject to disruption on lysis of the cells. Formation of labile precursor oligomers might account for the “P1 pentamers” reported in early studies of rhinovirus and EMCV assembly (McGreggor et al., 1975; McGreggor and Rueckert, 1977). Although the threedimensional structure of the P1 precursor is not known, the precursors may have enough affinity for one another to associate prior to cleavage. Formation of stable pentamers is almost certainly dependent on cleavage of P1 to free the amino terminus of VP3, five copies of which intertwine at the fivefold axes of symmetry t o form the p annulus (Acharya et al., 1989; Hogle et al., 1985; Luo et al., 1987; Rossman et al., 1985). The 3CD protein may also help in some unknown way to nucleate the P1 protomers.
C . Functional Significance of Poliovirus Capsid Myristylation To define further the role of myristylation in poliovirus assembly, a recombinant vaccinia virus was constructed that expressed a P1 pre-
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cursor with a glycine to alanine substitution at the amino terminus of P1 to prevent myristic acid addition (Ansardi et al., 1992). Using the VVPl/VVP3 coexpression system, the importance for myristylation of the capsid precursor in cleavage by 3CD and assembly of subviral particles was investigated. Previous reports in the literature had indicated that nonmyristylated P1 was not processed as efficiently as myristylated P1 in uitro by 3CD (Krausslich et al., 1990; Marc et al., 1989). Fkports describing intracellular studies of assembly of nonmyristylated capsid proteins expressed on transfection of in uitro transcribed RNA genomes encoding amino acid substitutions that prevented myristylation did not address rates of cleavage of the nonmyristylated precursors, but completely processed nonmyristylated P1 cleavage products were detected (Marc et al., 1990, 1991). Another study had been reported in which mutant polioviruses encoding altered P1 myristylation signals expressed mixed populations of myristylated and nonmyristylated P1 precursors (Moscufo et al., 1991). Interpretation of results from these previous approaches had been difficult, and one author cited reversion of the mutations back to wild type as a complication in data interpretation (Marc et al., 1990, 1991; Moscufo et al., 1991). To circumvent this problem, a nonmyristylated P1 precursor and 3CD protease were coexpressed in the same cell from separate recombinant vaccinia viruses. Surprisingly, the nonmyristylated P1 was completely cleaved to VPO, VP3, and VP1 (Ansardi et al., 1992). The reasons for this contrast with the in uitro cleavage studies is not clear. Enzyme/substrate ratios in the W P l m y r - / VVP3-coinfected cells may have been more favorable for complete proteolytic cleavage than those in the in uitro studies. Alternatively, additional host-cell components may contribute to the cleavage reaction. Evidence has been presented that a host cellular protein factor facilitates cleavage of the P1 precursor by 3CD protease (Blair et al., 1993). Several studies had suggested that myristylation of the P1 precursor was required for assembly of stable poliovirus. By using the P113CD coexpression system, it became clear that a block in assembly of nonmyristylated promoters occurred at the level of 1 4 s pentamer formation. The results of the studies confirmed the speculation based on structural information that the myristate moieties are required to assemble a stable capsid (Chow et al., 1987; Paul et al., 1987a). The use of VVPllVVP3 coexpression system, then, demonstrated that this requirement for myristylation occurs prior to cleavage of VPO to VP2 and VP4, as nonmyristylated subviral particles did not assemble (Ansardi et al., 1992).
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VIII. COMPLEMENTATION SYSTEM TO STUDY POLIOVIRUS ENCAPSIDATION Development of the recombinant vaccinia virus system for intracellular coexpression of the P1 precursor and 3CD protease provided the opportunity to analyze poliovirus capsid mutants for defects in proteolytic cleavage and assembly of subviral particles. The utility of this system was confirmed by analyzing the nonmyristylated P1 precursor expressed by a recombinant vaccinia virus. However, the assembly system at this point did not permit analysis of the RNA encapsidation step of morphogenesis. Information about this stage of assembly is particularly lacking, owing to the inability thus far t o reconstitute poliovirus virions in vitro from purified components (Putnak and Phillips, 1981b; Rombaut and Boeye, 1991),and because of the difficulty in obtaining poliovirus mutants with encapsidation defects. Previous studies by Jewell et al., (1990) had suggested that P1 precursors expressed by recombinant vaccinia viruses could not serve as a supply of capsid proteins in a mixed infection with poliovirus. Because of the large degree of intracellular localization of the processes of poliovirus RNA replication and possibly encapsidation, it was possible that the recombinant P1 precursors expressed by VVPl would be excluded from intracellular poliovirus compartments involved in RNA replication and encapsidation. The first indication that this was not the case came from experiments which repeated the recombinant vaccinia virus/ poliovirus coinfection experiments of Jewell et al. (1990). In contrast to these previous studies, in cells coinfected with VVPl and type 1poliovirus, the P1 precursor expressed by VVPl was proteolytically cleaved in trans by poliovirus protease 3CD, resulting in production of VPO, VP3, and VP1. In addition, mature virion protein VP2 derived from recombinant precursors was observed, strongly suggesting that vacciniaexpressed capsid proteins were incorporated into the poliovirus encapsidation pathway. Analyzing the incorporation of recombinant vaccinia-expressed capsid proteins into poliovirus virions in mixed infections with wild-type poliovirus provided one method for determining whether a mutant P1 precursor had defects at the encapsidation stage of assembly (Ansardi et al., 1992). However, this system suffered from an inherent complication because mutant capsid proteins expressed by the recombinant vaccinia virus were synthesized in the presence of wild-type capsid proteins expressed by the coinfecting poliovirus. Assembly defects of the mutant capsid subunits might then be overshadowed if they were incorporated into mixed wild-type and mutant particles. This appeared
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to be the case with the nonmyristylated capsid proteins expressed by a recombinant vaccinia virus which did not assemble in VVPlmyr-/ VVP3 coinfected cells, but which were incorporated into poliovirus virions to some degree in cells coinfected with VVPlmyr- and type 1 poliovirus (Ansardi et al., 1992). To establish a trans-complementation system in which the only source of functional capsid proteins was the recombinant vaccinia virus, a poliovirus defective interfering (DI) genome was used as the source of a poliovirus replicon which did not express functional capsid proteins (Cole, 1975; Hagino-Yamagishi and Nomoto, 1989; Kuge et al., 1986). The DI genome we used for these studies had been previously described and could be generated in uitro by transcription of a cDNA copy of the genome contained in plasmid pSMl(T7)l under the control of a promoter for bacteriophage T7 RNA polymerase (HaginoYamagishi and Nomoto, 1989). The DI genome contained a deletion of 816 nucleotides of the P1 gene (-31% of the P1 gene) and had been constructed in vitro by ligating a cDNA copy of a segment of the deletion-containing P1 gene from a naturally occurring DI genome of poliovirus type 1Sabin into a type 1Mahoney cDNA background. The in-frame deletion encompasses sequences from portions of the VP2 and VP3 genes. In their studies, Hagino-Yamagishi and Nomoto (1989) demonstrated that the in uitro transcribed DI genome replicated on transfection into poliovirus-infected cells and was encapsidated by capsid proteins provided in trans by helper wild-type poliovirus. Furthermore, the genetically engineered DI genome was maintained in serial passage in a mixed stock of wild-type and DI viruses. To establish a complementation system, Ansardi et al. (1993) transfected the RNA derived from the in uitro transcription of the DI cDNA into cells previously infected with VVP1. The DI RNA replicated and was encapsidated by the P1 provided in trans by VVP1; the encapsidated defective genome was referred to as PVdefSM (Fig. 5). This was the first demonstration of trans-complementation of a defective poliovirus genome and generation of a homogeneous population of defective poliovirus particles free of contaminating wild-type helper virus. By serially passaging PVdefSM in the presence of VVP1, stocks of PVdefSM were generated that could be used subsequently as a means of delivering the capsid gene-deficient replicon to every cell in a monolayer, overcoming limitations of transfection efficiencies. These studies established that vaccinia virus vectors expressing P1 capsid precursors could be used as the exclusive source of capsid proteins for a capsid gene-deficient poliovirus replicon, providing the opportunity to analyze the poliovirus assembly process in all of its stages, including
FIG.5. Complementation system in which to study poliovirus assembly (Ansardi et al., 1993). We utilize a cDNA clone of a poliovirus defective interfering genome (DI) (Hagino-Yamagishi and Nomoto, 1989). Cells are first infected with VV-P1, followed by transfection with the DI RNA obtained from in uitro transcription. The genome is defective because i t lacks a complete coding region for the poliovirus P1 protein. Transfection of this RNA into cells results in the complete replication cycle of poliovirus because the DI genome encodes all of the necessary proteins for RNA replication. The first step following transfection is translation of the DI RNA, which results in the production of poliovirus proteins required for replication including 3CD, which processes the poliovirus P1 protein expressed from VV-P1, resulting i n cleavage and assembly of subviral intermediates. In parallel, the viral proteins replicate the defective viral genome, resulting in the synthesis of multiple copies of the plus-stranded RNA. The plus-stranded RNA genome interacts with subviral intermediates, resulting in encapsidation. The encapsidated RNA is then released from the cells by a n as yet undetermined mechanism. Serial passage of the encapsidated RNA (referred to as a replicon) in the presence of VVP1 results in amplification; following extended serial passage of greater than 20 or more, stocks of the encapsidated replicons can be obtained. Removal of residual VV-P1 from the stocks can be achieved using centrifugation in combination with anti-vaccinia antibodies. The resulting stock of encapsidated replicons is devoid of any VV-P1. The stock of encapsidated replicons can be used in combination with VV-P1 variants containing defined mutations in the poliovirus capsid genes to assess the effects of the mutations on poliovirus assembly and encapsidation (Ansardi and Morrow, 1993, 1995; Ansardi et al., 1994a).
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encapsidation, without having to rely on expression of mutant capsid proteins from a replicating poliovirus genome. To characterize the complementation system, the role myristylation of P1 has on processing, assembly, and encapsidation of RNA was reexamined. In assembly experiments conducted by using cells coinfected with VVPlmyr- and PVdefSM, results markedly different than those obtained with the VVPlmyr- /VVP3-coinfected cells were observed (Ansardi et al., 1993). Low levels of the VPO, VP3, and VP1 proteins recovered from cells coinfected with VVPlmyr- and PVdefSM had sedimentation properties on sucrose density gradients consistent with empty capsids. Importantly, this difference in assembly phenotypes of mutant capsid proteins between the two systems was not a general property of all of the assembly-defective mutants because in most cases in which assembly of mutant capsid proteins was analyzed in both systems, those that did not assemble in cells coinfected with the mutant P1-expressing recombinant vaccinia virus and VVP3 also did not assemble in cells coinfected with PVdefSM and the mutant P1expressing recombinants. The difference in assembly phenotypes indicated that some factor associated with VVPlmyr-/PVdefSM-coinfectedcells, but not with VVPlmyr- /VVP3-coinfected cells, facilitated assembly of nonmyristylated protomers. One explanation might be that cleavage of the nonmyristylated P1 precursor occurred much more rapidly in cells coinfected with PVdefSM, increasing pools of nonmyristylated 5s protomers available for assembly. This would seem to be a plausible explanation since the processed proteins generated in VVPlmyr-/VVP3-~oinfected cells were unstable; more efficient cleavage of nonmyristylated P1 might then allow greater concentrations of nonmyristylated protomers to build prior to their degradation. However, no delays were observed in complete cleavage of the nonmyristylated precursor in comparison to the wild-type precursor in cells coinfected with VVP3, and expression levels of the 3CD protease in cells infected with 20 pfdcell of VVP3 appear to be comparable to those expressed by the defective poliovirus genomes when introduced at levels sufficient to infect every cell in a monolayer. A second difference associated with the VVPlmyr- /PVdefSMcoinfected cells was the presence of the nonfunctional capsid precursor with an internal deletion of 272 amino acids expressed by the defective poliovirus genome. This precursor retains the myristylation signal at the amino terminus and is presumably cotranslationally modified by myristate addition. We have observed that this deletion-containing P1 protein is unstable. The possibility that the myristylated deletioncontaining P1 protein is facilitating assembly of nonmyristylated cap-
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sid proteins seems unlikely. The deletion-containing P1 precursor would also not be able to supply a myristylated VPO protein through phenotypic mixing with the individual proteins derived from the recombinant nonmyristylated precursor, as the deleted region of the precursor encompasses portions of VPO. Other components present during a poliovirus infection may be contributing to the assembly of nonmyristylated protomers. Although clearly poliovirus P1 and 3CD proteins are the only poliovirus proteins required for assembly of subviral particles, this does not rule out the possibility that another poliovirus protein plays a facilitory role in capsid assembly. A more intriguing possibility is that the viral RNA genome plays a nucleating role in virus assembly, and capsid protein interactions with the RNA genome might play a stabilizing role in formation of the capsid. If the viral RNA plays some active role in nucleating the capsid protomers at an early stage, then interaction of nonmyristylated subunits with the viral RNA might facilitate their assembly into capsid particles. Clearly, a pronounced block exists in forming nonmyristylated RNA-containing virions; therefore, this type of assembly model for poliovirus suggests that at least some populations of subviral particles may be derived from the dissociation of unstable ribonucleoprotein complexes on extraction from the infected cell. Some evidence in the literature suggests that empty capsids may be a degradation product of an unstable ribonucleoprotein complex that easily dissociates into empty capsids and RNA on extraction from the cell (Koch and Koch, 1985; Maronginu et al., 1981).The absence of myristate form V P O may prevent completion of the RNA encapsidation event and condensation of a mature virion, resulting in a byproduct of empty capsids. In Section VII, a hypothesis was presented that the events of poliovirus assembly and encapsidation may be sequestered from the replication complexes. If nascent RNA chains diffuse away from the replication complexes to be encapsidated, this might allow genomic RNA to participate in the assembly process and facilitate formation of nonmyristylated subviral particles. Alternatively, poliovirus capsid proteins may be targeted to membranous sites of RNA encapsidation which are adjacent to replication complexes (Caliguiri and Compans, 1973; Pfister et al., 1992).The subcellular localization of nonmyristylated and myristylated P1 precursors expressed by recombinant vaccinia viruses (cells infected with VVPl or VVPlmyr- alone) was analyzed by performing crude separations of cytosol from intracellular membranes (D. A. Ansardi and C. D. Morrow, unpublished results, 1991). The vast majority of myristylated and nonmyristylated P1 precursors partitioned in the soluble fractions. The finding that myristylated P1 is a cytosolic protein strongly
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suggests that cleavage of this protein by 3CD is not associated with intracellular membranes. The rapid assembly of cleavage products of myristylated P1 suggested that any targeting to membranous locations of RNA encapsidation occurs subsequent to pentamer formation.
A . Proteolytic Cleavage of Capsid Precursor Although the favored pathway for poliovirus morphogenesis indicates that complete cleavage of the capsid precursor is required prior to assembly of protomers into subviral particles or virions, this question had not been addressed previously by using intracellular assembly systems (Rueckert, 1990). In vitro assembly studies of EMCV capsid precursors with cleavage site defects indicated that complete proteolytic cleavage of the capsid precursor was required for assembly of EMCV subviral particles (Palmenberg, 1982; Parks and Palmenberg, 1987). Previous assembly studies of different picornaviruses, however, had suggested that even uncleaved P1 precursors from rhinovirus, EMCV, and poliovirus might assemble pentamer precursors (McGreggor et al., 1975; McGreggor and Rueckert, 1977). The most convincing arguments that cleavage of the P1 precursor is required for poliovirus capsid assembly were made from studies of the three-dimensional structure of picornavirus virions: formation of the p-annulus structure at the fivefold vertices of the virion requires cleavage between VPO and VP3 to free the amino termini of VP3 that form this structure (Acharya et al., 1989; Hogle et al., 1985; Luo et al., 1987; Rossman et al., 1985). Discrepancies between the early assembly studies and assembly pathways deduced from structural information suggested that an investigation of whether cleavage intermediates might assemble precursor subviral particles was warranted. As a first step in determining whether complete cleavage of P1 was required for assembly of subviral particles, P1 precursors were generated with valine substitutions for glycine at the P1’ position of the QG cleavage sites in the precursor with the hope that these substitutions would prevent cleavage at the altered sites (Ansardi and Morrow, 1993). The valine substitutions were chosen so that cleavage would be prevented by making the most conserved substitution possible; previous studies of the cleavage of the QG bond between the 3C and 3D proteins of poliovirus found that an alanine substitution for glycine still permitted cleavage, whereas a valine substitution inhibited cleavage at the site (Kean et al., 1990). However, studies by Kirkegaard suggested that glutaminemethionine could serve as a functional cleavage site between VP3 and VP1, although the use of this cleavage site had not been confirmed
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(Kirkegaard, 1990). Parks and Palmenberg had determined that a QV dipeptide was not a functional cleavage site for proteolytic processing of the EMCV capsid precursor in uitro a t the site between VP3 and VP1; the wild-type cleavage site at that location in the EMCV P1 precursor is QG as in the case of poliovirus (Parks and Palmenberg, 1987). Finally, all of the cleavage sites in poliovirus polyproteins processed by poliovirus 3Cpr0 or 3CD are QG (Kitamura et d,1981; Racaniello and Baltimore, 1981a). This strict conservation of cleavage site primary sequence is unique among picornaviruses, suggesting that the poliovirus enzyme had a particularly stringent requirement for a QG cleavage site sequence (Palmenberg, 1990). Analysis of P1 precursors which had QV cleavage sites at either the VPO-VP3 cleavage site or the VP3-VP1 cleavage site revealed that cleavage at the altered sites occurred, although less rapidly than at the QG sites (Ansardi and Morrow, 1993). Complete cleavage of these precursors prevented a thorough analysis of assembly phenotypes of incompletely cleaved precursors. However, in unpublished experiments conducted using cells coinfected with recombinant vaccinia viruses that expressed a mutant precursor and VVP3, we were able to determine the sedimentation properties of the P1 cleavage intermediates on sucrose density gradients (Ansardi and Morrow, unpublished observations, 1992). Unlike the completely cleaved proteins, the cleavage intermediates displayed nonspecific sedimentation properties without accumulating in distinct peak fractions and were more or less evenly distributed among fractions of the gradient above the 14s pentamer peak. The reasons for the nonspecific sedimentation pattern are unclear but might reflect an aggregation of cleavage intermediates into nonfunctional oligomers. Because the metabolically radiolabeled, partially cleaved precursors were presumably present in the same cell with unlabeled completely cleaved precursors, drawing definitive conclusions about the oligomeric state of P1 cleavage intermediates is difficult. Evidence from pulse-chase radiolabeling experiments indicated that P1 cleavage intermediates were degraded in the infected cells, further suggesting that they did not assemble stable subviral particles. Instability of the cleavage intermediates is compatible with the observation that assembly-defective protomers are rapidly degraded in cells. On the basis of pulse-chase experiments it was clear that not all of the partially cleaved intermediates were chased into completely cleaved proteins, and at least a portion of the partially cleaved proteins were degraded. Successful complete cleavage of the capsid precursor might require that the events occur in a single interaction event with the enzyme, with partially cleaved proteins not serving as substrates for the protease.
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A question that has not been resolved in the proteolytic cleavage step of capsid formation is whether the 3CD-mediated cleavages of the P1 precursor occur in a defined order. Evidence from previous studies suggested that the P1 precursor was cleaved first at the site between VP3 and VP1, generating an uncleaved VPO-VP3 protein, and second at the site between VPO and VP3 (Reynolds et al., 1992). This order of cleavage had been suggested because a VPO-VP3 cleavage intermediate is typically detected in lysates of poliovirus-infected cells, whereas little VP3-VP1 is detected. The P1 precursor with the QV cleavage site between VP3 and VP1 was delayed in processing at the altered site, allowing confirmation that cleavage at the site between VPO and VP3 could occur without prior cleavage between VP3 and VP1. Thus, cleavage between VP3 and VP1 is not a prerequisite for cleavage between VPO and VP3. A second component of the ordered cleavage hypothesis was that a defined processing order series of cleavages might be required to generate functional, assembly-competent capsid proteins. Interestingly, capsid protomers derived from the precursor with the altered cleavage site between VPO and VP3 (VP3-G001V, which gave rise to increased amounts of VPO-VP3 and VP1) failed to assemble subviral particles. In contrast, capsid protomers generated from the precursor with the altered cleavage site between VP3 and VP1 (VP1-G001V)were capable of assembly. Clearly an argument can be made based on these results that the previously predicted order of P1 cleavage, in which VP1 is released followed by VPO and VP3, is not required to generate assembly-competent capsid proteins. The results may even suggest that the order of cleavage required to generate assembly-competent protomers is cleavage at the VPO-VP3 site followed by cleavage at the site between VP3 and VP1. Further studies will be required to substantiate this claim, because the capsid proteins derived from the P1 precursor with the QV cleavage site between VPO and VP3 have a valine substitution at the amino terminus of VP3, and the amino termini of VP3 form the p-annulus structure responsible for interlocking common protomers within a pentamer (Acharya et al., 1989; Hogle et al., 1985; Luo et al., 1987; Rossman etal., 1985). To distinguish between these two possibilities, new mutants can be constructed with substitutions at the Q position of the QG bond, thus altering the carboxyl terminus of VPO rather than the amino terminus of VP3. If Q-substituted mutants with slower cleavage kinetics at the cleavage site between VPO and VP3 can be isolated, this might allow a resolution of these two possibilities. Secondary consequences of the valine substitutions introduced at the amino termini of both VP3 and VP1 were manifested in defects of the
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completely cleaved capsid protomers in assembly and RNA encapsidation (Ansardi and Morrow, 1993). The secondary defects of these mutants indicate that maintenance of the QG cleavage site primary sequence in the poliovirus P1 capsid precursor is required for proper function of the capsid proteins. This property is likely to be one reason why poliovirus 3Cpro (and 3CD) cleavage sites appear to have less flexibility in primary sequence than the 3Cpro cleavage sites of other picornaviruses (Palmenberg, 19901, a t least in the case of the capsid precursor.
B . Capsid Mutations Affecting R N A Encapsidation Encapsidation of a viral RNA genome requires specific recognition of the genome by the virus capsid to ensure packaging of viral RNA without nonspecific packaging of host cellular mRNA molecules. The protein-RNA interactions required for genome encapsidation consist of two elements: capsid protein determinants that specifically interact with the viral RNA, and cis-acting elements of the viral RNA genome that are recognized by the virus capsid and distinguish it from nongenomic RNA molecules. The virus capsid may also contain interior features that interact with RNA in a less specific manner not dependent on nucleotide sequence. In studies using the recombinant vaccinia virus expression systems, mutants with changes at the amino terminus of VP1 were shown to have defects in the RNA encapsidation step of assembly (Ansardi and Morrow, 1993).Two mutants were analyzed: one with a valine substitution for the glycine residue at the amino terminus of VP1 (VP1G001V) and a second in which the first four amino acids of VP1 were deleted (VPlAl-4). The VP1 deletion mutant had been previously described in the literature and was found t o have a delayed kinetics of encapsidation at 39.5"C (Kirkegaard, 1990). Surprisingly, the encapsidation defect was more pronounced in the mutant with the valine substitution for the amino-terminal glycine than in the deletion mutant. The results of these experiments raise the possibility that the amino-terminal portion of VP1 is one of the poliovirus capsid determinants involved in capsid protein-RNA interaction. The involvement of an amino-terminal arm from a viral capsid protein in interaction with nucleic acid has precedent in the literature (Geigenmuller-Gnirke et al., 1993; Rossman et al., 1985). In the cases of some icosahedral RNA viruses, including several plant viruses, the terminal extensions of capsid proteins not associated with the @-barrelcore are disordered in the three-dimensional structures and point toward the interior (Rossman et al., 1985). These amino-terminal structures are likely in con-
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tact with RNA and may play important roles in capsid-RNA interaction. Often these terminal extensions contain a large proportion of basic residues, but this is not the case with the amino terminus of poliovirus VP1. The disordered amino-terminal segment of the Sindbis capsid protein, the core of which does not follow the classic p-barrel fold of most icosahedral viruses (H. K. Choi et al., 19911, has been demonstrated to contain a segment of 32 amino acids critical for interaction with the RNA genome (Geigenmuller-Gnirke et al., 1993). This segment contains a highly conserved stretch of 10 amino acids containing three lysine and two arginine residues. In addition to the role for the VP1 amino terminus in RNA encapsidation, this region has been implicated from both genetic and biochemical studies to be involved in the processes of virus entry and release (Fricks and Hogle, 1990; Kirkegaard, 1990). Although the first 20 amino acids of VP1 were disordered in the three-dimensional structure determined for type 1 poliovirus (Hogle et al., 1988, this region was proposed by Fricks and Hogle (1990) to form an amphipathic helix structure. Even though the amino acid sequence homology is limited at the amino terminus of VP1 among different picornavirus members, Fricks and Hogle demonstrated that the first 18-23 residues of VP1 amino termini from several enteroviruses and from human rhinovirus type 14 could be modeled on an amphipathic helical wheel. This structural feature is not likely to be shared among the aphthovirus and cardiovirus members of the Picornauiridm, however, which have shorter VP1 amino termini lacking the potential structural homology of the VP1 amino termini of enteroviruses and rhinoviruses (Acharya et al., 1989; Luo et al., 1987). Further refinements of the structure of poliovirus type 3 Sabin found a partially ordered stretch of five amino acids from an unidentified portion of the VP1 amino terminus that formed a short segment of p-sheet structure along with portions of VP4. In the studies of Fricks and Hogle (19901, externalization of the amino terminus of VP1 after binding of the virus to the receptor was shown to be responsible for binding of the altered virus to liposomes, and they proposed that the amino-terminal portion of VP1 in concert with VP4, which is also extruded from the virus after receptor attachment (Everaert et al., 1989), played a role in disrupting the endosomal membrane to allow RNA to be released into the cytosol. The linkage of VP4 to the short segment of VP1 was suggested to provide a way for externalization of these two capsid features together. If the amino terminus of VP1 is also involved in RNA interaction, a mechanism might be envisioned in which extrusion of VP4 and the VP1 amino terminus triggers the release of the RNA genome from the interior of the capsid. So far, however, biochemical evidence that this region of
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VP1 interacts specifically with the RNA genome has not been reported, either with purified VP1 or with synthetic peptides corresponding to the amino-terminal portions of the VP1 protein. Future experiments might be aimed at addressing these questions. Although the VP1 amino terminus may directly interact with the poliovirus RNA genome, other regions of the virus capsid are likely to interact with the RNA genome as well. The three-dimensional structure of poliovirus type 1 did not reveal electron density that could be interpreted as RNA, but in refined structures of poliovirus type 3, some electron density was attributed to RNA base ring structures stacking with the aromatic side chains of the tryptophan-38 and phenylalanine-41 residues of VP2 (Filman et al., 1989). Although significant regions of encapsidated picornaviral RNA molecules do not adopt the regular pattern of capsid binding required for nucleic acid visualization in the crystal structures, some insight into viral capsid-RNA interactions has been gained from RNA viruses and single-stranded DNA viruses in which segments of the viral genomes do associate in a sequence-independent manner with regions of the capsid interior (Chen et al., 1989; Fisher and Johnson, 1993; Larson et al., 1993; McKenna et al., 1992; Tsao et al., 1991). In the case of bean-pod mottle virus (a comovirus), a single-stranded RNA virus with a bipartite RNA genome encapsidated in separate particles, nearly 20% of the viral RNA genome binds to the capsid interior in a symmetric fashion (Chen et al., 1989). The ordered RNA is single-stranded and associates with a hydrophilic pocket around the 3-fold symmetry axes of the capsid. The binding of the RNA around the threefold axes results in the formation of a trefoil-shaped cluster at each of the twenty three-fold axes of the capsid, with each cluster consisting of 33 ribonucleotides. The binding of RNA in this manner indicates that the protein-RNA interactions are not sequence specific as an exactly repeating set of bases in the RNA genome is not present. The interactions between the RNA segment and residues lining the binding pocket are primarily van der Waals and electrostatic interactions. RNA-protein interactions have been characterized for a number of viral and nonviral proteins (reviewed by Frankel et al., 1991; Mattaj, 1993). An example of one extensively studied RNA-protein interaction is the binding of the HIV-1 Tat protein to a segment of RNA designated TAR (Calnan et al., 1991; Tao and Frankel, 1992). The Tat protein is a transcriptional activator which binds to TAR, a bulged stem-loop structure present at the 5' end of viral mRNA molecules. Tat contains a 9-amino acid region of basic amino acids that are required for the recognition of TAR. Arginine-rich motifs (ARMS) are conserved among many RNA binding proteins, including the capsid
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proteins of some RNA viruses (Frankel et al., 1991; Lazinski et al., 1989; Mattaj, 1993). The ARM sequences may be involved in recognizing specific RNA secondary structures as well, and this function has been confirmed for recognition of RNA hairpins by some bacteriophage antiterminator proteins (Lazinski et al., 1989). Although not all RNAbinding proteins contain linear ARM sequences (poliovirus does not), sequence-specific RNA binding motifs formed from clusters of arginine residues brought together in the folded protein might be envisioned as a method of RNA recognition (Calnan et al., 1991). By using the recombinant vaccinia virus systems, we analyzed two arginine residues associated with a cavity on the poliovirus capsid interior (Ansardi et al., 1994a). The dimensions of the cavity, approximately 10 A wide and 5 A deep, are sufficient to accommodate a helical segment of RNA. The cavity displays similarity to those of some of the icosahedral viruses in which nucleic acid interaction with the capsid was visible in the three-dimensional structure (canine parvovirus and +X174), including the presence of several basic amino acid residues (McKenna et al., 1992; Tsao et al., 1991). Most of the basic residues of the poliovirus cavity are well-conserved in amino acid sequence alignments of capsid proteins from various picornaviruses. 'Ibo arginine residues associated with this depression were analyzed by sitedirected mutagenesis for their functional role in capsid assembly and RNA encapsidation. One of the arginine residues (VP1-R129) is wellconserved among different picornaviruses in capsid sequence alignments, and substitution of this residue with lysine or glutamine disrupted assembly of the capsid. The second cavity-associated arginine residue targeted for mutagenesis, VP4-RO34, is not well conserved in sequence alignments of picornavirus capsid proteins. The lysine substitution for arginine at this position had no observable effects on assembly or encapsidation. In contrast, substitution of glutamine for this arginine residue rendered the mutant capsid defective for virion formation, especially at 395°C. The encapsidation defect for this mutant could not be separated in these studies from a primary defect in capsid assembly that was noted even in the absence of the defective RNA genome. Other amino acid residues within the depression provide potential targets for future mutagenesis studies. A lysine residue at VP3-041 and an arginine residue at VP1-267 both have side chains well exposed to the interior of the virus. Surprisingly, substitution of an arginine residue buried at a protomer-protomer interface (VP3-R223) affected RNA encapsidation. At 37"C,capsid proteins derived from cleavage of the VP3-R223K mutant precursor did not assemble subviral particles or virions; at 33"C, however, capsid proteins derived from the mutant precursor were
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
51
found to cosediment on sucrose density gradients with poliovirus empty capsids. RNA-containing virions derived from the mutant were not observed at either temperature. The ability of the mutant to assemble empty capsids but not virions at 33°C suggested that this substitution might have destroyed an encapsidation determinant. The side chain of this arginine residue is not well exposed on the interior surface of the capsid in the mature virion, however, and almost certainly could not make direct contact with the RNA genome. The lysine substitution for arginine at this location might have resulted in secondary structural effects that rendered the capsid incapable of forming mature virions. The VP3-R223 residue potentially forms a hydrogen bond with the side chain of threonine VP3-031. The cavity-associated residue VP1R129 potentially forms a hydrogen bond with the main-chain oxygen of this same residue. Thus, substitution of the buried residue might exert secondary defects on the cavity region. The phenotype of this mutant provides some preliminary evidence that this region of the capsid may be involved in RNA interaction. Poliovirus capsid protein-RNA binding may include nonsequencespecific interactions similar to those described for the icosahedral viruses in which nucleic acid was observed in the X-ray structure. The poliovirus capsid may also contain determinants required for recognizing a sequence-specific RNA secondary structure which acts in cis as an encapsidation signal. Just as information about poliovirus capsid protein determinants required for RNA encapsidation is limited, little is known about the cis elements of the poliovirus genome required for encapsidation. Whatever the RNA encapsidation signal of poliovirus may be, its identification is likely to be difficult. Important information has been gained from in uitro analyses of protein-RNA interactions in other systems (Calnan et al., 1991; Geigenmuller-Gnirke et al., 1993; Gott et al., 1991; Fbmaniuk et al., 1987). Unfortunately, in uitro encapsidation systems for poliovirus have not been developed. In uitro interactions between poliovirus RNA and capsid proteins are difficult to study for several reasons. Encapsidated RNA molecules are always linked to VPg (Wimmer, 1982), and linkage of VPg to the RNA genome requires that the processes of poliovirus RNA replication take place (Kuhn and Wimmer, 1987; Paul et al., 1987b; Richards and Ehrenfeld, 1990). Thus, encapsidatable RNA genomes likely cannot be generated by in uitro transcription from a cDNA copy because RNA molecules synthesized by this method are not linked to VPg. Furthermore, construction of deletions in poliovirus RNA outside of the P1 region are incompatible with RNA replication, because the replication functions of many of the P2 and P3 proteins cannot be complemented in trans (Bernstein et al., 1986; Johnson and Sarnow, 1991). For these
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DAVID C. ANSARDI et al.
same reasons, naturally occurring DI genomes of poliovirus contain deletions only in the P1 region, because propagation of these genomes requires that they retain the capacities for both replication and encapsidation. The system developed by Ansardi et al. (1993) provides a method for trans-encapsidation of a poliovirus subgenomic replicon. An important feature of this trans-encapsidation system is that it separates the supply of capsid proteins away from the subgenomic replicon. An exciting potential use for this system would be to force adaptation of the RNA genome to a mutant capsid protein. In other words, if mutant capsid proteins are identified with defects at the encapsidation stage, serial passage of the subgenomic replicon with the recombinant vaccinia virus expressing a continuous supply of the mutant capsid might result in adaptation of the replicon to be encapsidated. This strategy might provide a method for identifying otherwise elusive cis elements of the poliovirus RNA genome required for encapsidation. In fact, serial passage of PVdefSM with the recombinant vaccinia virus which expresses the VP1-G001V precursor and the VP3-R223K precursor at 33°C results in low levels of encapsidation, suggesting that the PVdefSM genome can “adapt” to this mutant capsid protein (D. C. Ansardi and C. D. Morrow, unpublished, 1994).
C . Studies on Maturation Cleavage Using Complementation System The assembly of an infectious poliovirus virion requires the proteolytic cleavage between an asparagine-serine amino acid pair in VPO after encapsidation of the viral genomic RNA. This cleavage, which results in the processing of VPO to VP4 and VP2, has been termed the maturation cleavage and is believed to occur via an intramolecular event (Arnold et al., 1987). It has been difficult to study the features of this cleavage as well as the generation of infectious poliovirions owing to the rapid cleavage of VPO and maturation into infectious virus. Studies using the recombinant vaccinia virus systems have described mutants in which a glutamine-glycine amino acid pair (VP4-QG)and a threonine-serine amino acid pair (VP4-TS)were substituted for the asparagine-serine amino acid pair in the maturation cleavage site (Ansardi and Morrow, 1995). The mutations in which a glutamine-glycine amino acid pair were substituted in the maturation cleavage site resulted in a capsid protein that could be proteolytically processed and assembled into subviral intermediates including an empty capsid-like structure in the presence of PVdefSM. However, no PVdefSM was encapsidated in virions containing a QG. In contrast, the threonine-serine substitution for the asparagine-serine at the
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
53
maturation cleavage site resulted in a capsid protein which, on proteolytic processing, could assemble into subviral intermediates and encapsidate the PVdefSM RNA. The maturation cleavage was significantly delayed compared to wild type. Interestingly, the cleavage event in these particles occurred in vitro as well, and this cleavage could be accelerated by incubation at physiological temperatures. The results of these studies support the concept that a series of conformational changes occur during the maturation cleavage of VPO. The mechanism by which this occurs throughout the entire poliovirion (i.e., the 60 copies of VPO to be cleaved) is not clear. It is possible that a cooperative effort exists between the subunits during the maturation cleavage, as has been suggested for nodaviruses (Zlotnick et al., 1994). Whether this is the case for the maturation cleavage of poliovirus is currently under investigation utilizing a different set of mutants in combination with the complementation system. IX. PERSPECTIVES ON POLIOVIRUS ASSEMBLY
A proposed ordered pathway for poliovirus morphogenesis was discussed in earlier sections. The phenotypes of mutants generated from our studies have afforded some valuable tools with which to assess the validity of this pathway (Table I). The first step in the proposed pathway of poliovirus morphogenesis is cleavage of the precursor protein P1. The cleavage site mutants described were not completely blocked in processing at the altered (QV) sites, making it difficult to determine whether P1 precursors cleaved at only one site would assemble. However, no evidence was found for assembly of specific stable structures from P1 cleavage intermediates. Using sucrose density gradients, no evidence was found that uncleaved P1 precursor assembled stable subviral particles, as most uncleaved P1 was localized in fractions of the gradients above the 14s pentamer fractions. It is possible that uncleaved precursors are associated together as labile oligomers, and the rapid assembly of P1 cleavage products suggests that some type of P1-3CD assembly complex may exist. A significant proportion of the mutants analyzed failed to assemble subviral particles (Pl-myr-, VP3-G001V, VP3-R223K at 37"C, VP1R129K, VP1-R129Q). A common feature of all of these mutants was that they were stable in the precursor form but on proteolytic cleavage were degraded. A similar observation was made with FMDV capsid proteins expressed by a recombinant vaccinia virus (Belsham et al., 1991) and with an assembly-defective attenuated mutant of poliovirus
54
DAVID C. ANSARDI et al. TABLE I ASSEMBLY PHENOTYPES OF POLIOVIRUS CAPSIDMUTANTSEXPRESSED BY RECOMBINANT VACCINIA VIRUSES
Precursor P1 (wild type) PlmyrVP3-GOOlV VP1-G001V VP1-A1-4 VP4-RO34K VP4-RO34Q VP3-R223K VP1-R129K VP1-R129Q VP4-ND69T VP4-NO69Q VP2-SOlG
Cleavage by 3CD.
14s VVP3b
EC VVP3c
EC PVdefSMd
RNA Encapsidatione
++++f
++++
++++
++++ ++
++++
++++ ++++ ++++
+++ ++++
++++ ++ ++ +++ ++++ ++++
++++ ++++ ++++ ++++ ++++
-
++++ ND
++++ +++ -
37°C
NDg ND ND
++++ +++ - 37°C
++ 33°C
++ 33°C
ND
ND ND
++++ +
++++ -
-
-
+
- 37°C
+ + + 33°C
+*
+++ 37°C + 39.5”C - 37°C - 33OC*
-
++++ ++
Cleavage of precursor in cells coinfected with VVP3. Assembly of 1 4 s pentamers in cells coinfected with VVP3. Assembly of 75s empty capsids in cells coinfected with VVP3. d Assembly of 755 empty capsids in cells coinfected with PVdefSM. Formation of mature RNA-containing virions in cells coinfected PVdefSM; reduced yields may result from a defect at a n earlier step. Mutants with defects a t the encapsidation step are marked (*). f + + + +, wild type; + + +, 50-70% wild type; + +, 25-50% wild type; + ,lo-25% wild type; -, none detected. 8 ND, Not determined. Delayed cleavage of VPO resulting in accumulation of poliovirions. a
type 3 (Macadam et al., 1991). In the P1 precursor form, the recombinant vaccinia virus-expressed FMDV capsid proteins were stable, but on cleavage of the precursor, the capsid proteins, which failed to assemble subviral particles, were rapidly turned over. Together, these results suggest that picornaviral capsid precursors are recognized by the cell as correctly folded proteins, but on cleavage of the precursor, the capsid proteins must rapidly assemble into their oligomeric forms or are targeted for degradation. The stability of the P1 precursors may also provide further evidence that P1 precursors or P1‘and 3CD proteins assemble protein complexes prior to cleavage. P1 or P1-3CD oligomers may be recognized by the cell as “correctly folded.” However, on cleavage, mutant capsid proteins that fail to assemble stable 14s pentamers may dissociate and be recognized as “misfolded.” In preliminary experiments the P1 capsid precursor has been found to interact
POLIOVIRUS ASSEMBLY AND RNA ENCAPSIDATION
55
with the hsp 72/73 members of the 70-kDa family (Hsp 70) of heatshock proteins (Beckman et al., 1990; Pelham, 1988; D. C. Ansardi and C. D. Morrow, unpublished, 1993).A published report has described the interaction of poliovirus capsid proteins with proteins of the hsp 70 family of chaperones (Macejak and Sarnow, 1992). It is not clear if the interaction with hsp 72/73 plays a role in targeting assembly-defective mutant capsid protomers for degradation (Beckman et al., 1990; Pelham, 1988). In most cases in which cleaved poliovirus capsid proteins failed to assemble in cells coinfected with the mutant P1-expressing recombinant vaccinia and VVP3, they also failed to assemble in cells coinfected with the P1-expressing recombinant and PVdefSM. These results might be taken as evidence that the ability to assemble subviral particles is a prerequisite for interaction with the RNA genome. A potential deviance from this idea was noted by the different assembly phenotypes of the capsid proteins derived from the nonmyristylated precursor and the precursor containing a QG substitution at the VP4/VP2 junction. In cells coexpressing 3CD as the only other poliovirus component, the cleavage products derived from these precursors failed to assemble. However, in the presence of the defective poliovirus genomic RNA, cleavage products of these precursors assembled low levels of subviral particles. We have speculated that these particles may represent uncondensed capsids assembled around a nucleating RNA genome. The phenotype of these mutants were unique among those studied and provide preliminary evidence that the RNA genome might play a nucleation role in assembly and facilitate interactions among capsid protomer subunits. Clearly, extensive further studies are needed t o address these issues. Both 14s pentamers and empty capsids have been proposed to be the direct precursor of the poliovirus virion. In our studies, three capsid mutants, nonmyristylated P1 in VVPlmyr-/PVdefSM-coinfected cells, the VP4-QG mutant in VV-VP4QG/PVdefSM-coinfected cells, and VP3-R223K at 33"C, assembled structures consistent with empty capsids but did not assemble RNA-containing virions at detectable levels. The phenotypes of the mutants suggest that empty capsid formation is not sufficient to ensure assembly of RNA-containing virions. In the course of these studies, one mutant, VP1-GOOlV, was identified which assembled empty capsids in excess over virions, whereas a second mutant, VP4-R034Q, assembled virions over empty capsids at a higher ratio than normal. The assembly phenotype of the VP4-RO34Q mutant can be traced to a diminished pool of assembly-competent protomers in comparison to wild type, suggesting that empty capsids and virions assemble separately from common pools of capsid subunits.
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However, it is possible that assembly ofempty capsids with this mutant was a rate-limiting step in virion formation and that, once this step occurred, RNA encapsidation proceeded rapidly. The identity of the direct precursor to the poliovirus virion thus remains an open question. The RNA encapsidation step of poliovirus assembly remains the most elusive in viral morphogenesis. It is clear that the amino terminus of VP1 plays a role in this process, giving merit to the future analysis of this region of the capsid as a possible determinant for encapsidation. The arrangement of basic amino acid side chains within an interior depression in the capsid makes this region of the capsid an attractive candidate for further analyses. Finally, mutations in the maturation cleavage site have profound effects on the capacity of the processed viral proteins to assemble and encapsidate genomic RNA. The existence of the provirion as an assembly intermediate was supported from the analysis of a capsid mutation containing a threonineserine mutation cleavage site. In summary, the use of recombinant vaccinia virus vectors for analyzing the processes of poliovirus capsid assembly and RNA encapsidation overcomes limitations of previous intracellular assembly analyses which required isolation of mutant polioviruses subject to the potential for reversion. The systems described offer the dual capability of analyzing P1 capsid precursor cleavage and subviral particle formation separately from the encapsidation step. Information from the three-dimensional structure of poliovirus and its picornavirus relatives provides a rational basis for targeting regions of the capsid for mutagenesis studies. The use of recombinant vaccinia viruses with defined mutations in P1 in combination with PVdefSM might allow for scaleup and recovery of sufficient virus particles for structural analysis. Generation of additional poliovirus capsid mutants with defined mutations to be analyzed by these new methods will, it is hoped, further an understanding of the molecular mechanisms of poliovirus morphogenesis.
ACKNOWLEDGMENTS We thank Dee Martin for the preparation of the manuscript. M.J.A. was supported by a National Institutes of Health training grant (T32-A1 07150). Research was supported by a grant from the National Institutes of Health, National Institute for Allergy and Infectious Disease (A1 25005). to C.D.M.
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Rossman, M. G., and Johnson, J. E. (1989).Icosahedral RNA virus structure. Annu. Rev. Biochem. 58, 533-573. Rossman, M.G., Arnold, E., Erickson, J. W., Frankenberger, E. A., Griffith, J. P., Hecht, H. J., Johnson, J. E., Kamer, G., Luo, M., Mosser, A. G., Rueckert, R. R., Sherry, B., and Vriend, G. (1985).Structure of a human common cold virus and functional relationship to other picornaviruses. Nature (London) 317, 145-153. Rothberg, P. G., Harris, J. J. R., Nomoto, A,, and Wimmer, E. (1980).The genome-linked protein of picornaviruses. V. 0-4-(5’-UridylyI)-tyrosineis the bond between the genome-linked protein and the RNA of poliovirus. Proc. Natl. Acad. Sci. U.S.A. 75, 4868-4872. Rueckert, R. R. (1990).Picronaviridae and their replication. In “Virology” (B. N. Fields, D. Knipe, et al., eds.), pp. 507-547. Raven, New York. Rueckert, R., and Wimmer, E. (1984).Systematic nomenclature for picornavirus proteins. J. Virol. 50, 957-959. Sabin, A. B., and Boulger, L.R. (1973).History of Sabin attenuated poliovirus oral live vaccine strains. J. Biol. Stand. 1, 115-118. Salk, J. E. (1960).Persistence of immunity after administration of formalin-treated poliovirus vaccine. Lancet 2,715-723. Schultz, A. M., and Rein, A. (1989).Unmyristylated Moloney murine leukemia virus Pr65 gag is excluded from virus assembly and maturation events. J. Virol. 63,23702373. Schultz, A. M., Henderson, L. E., and Oroszlan, S. (1988).Fatty acylation of proteins. Annu. Rev. Cell Biol. 4,611-647. Semler, B. L.,Anderson, C. W., Hanecak, R., Dorner, F., and Wimmer, E. (1982).A membrane-associated precursor to poliovirus VPg identified by immunoprecipitation with antibodies directed against a synthetic heptapeptide. Cell (Cambridge, Mass.) 28, 405-412. Semler, B. L., Dorner, A. J., and Wimmer, E. (1984).Production of infectious poliovirus from cloned cDNA is dramatically increased by SV40 transcription and replication signals. Nucleic Acids Res. 12, 5123-5141. Smith, T. J., Kremer, M. J., Luo, M., Vriend, G., Arnold, E., Kamer, G., Rossmann, M. G., McKinlay, M. A,, Diana, G. D., and Otto, M. J. (1986).The site of attachment in human rhinovirus 14 for antiviral agents that inhibit uncoating. Science 233, 12861293. Sonenberg, N. (1987).Regulation of translation by poliovirus. Adv. Virus Res. 33, 175204. Sonenberg, N. (1990). Poliovirus translation. Curr. Top. Microbiol. Immunol. 161,23-47. Spector, D. H., and Baltimore, D. (1975).Polyadenylic acid on poliovirus RNA. 11.Poly(A) on intracellular RNAs. J. Virol. 15,1418-143. Tao, J., and Frankel, A. D. (1992).Specific binding of arginine to TAR RNA. Proc. Natl. Acad. Sci. U.S.A.89,2723-2726. Towler, D. A., Adams, S. P., Eubanks, S. R., Towery, D. S., Jackson-Machelky, E., Glaser, L., and Gordon, J. I. (1987).Purification and characterization of yeast myristoy1 CoA:protein N-myristoyltransferase. Proc. Natl. Acad. Sci. U.S.A. 84, 27082712. Towler, D. A., Gordon, J. I., Adams, S. P., and Glaser, L. (1988).The biology and enzymology of eukaryotic protein acylation. Annu. Rev. Biochem. 57,69-99. Toyoda, H., Nicklin, J. W.,Murray, M. G., Anderson, C. W., Dunn, J. J., Studies, F. W., and Wimmer, E. (1986).A second virus-encoded proteinase involved in proteolytic processing of poliovirus polyprotein. Cell (Cambridge, Mass.) 45,761-770. Trono, D., Pelletier, J., Sonenberg, N., and Baltimore, D. (1988).Translation in mam-
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DAVID C. ANSARDI et ul.
malian cells of a gene linked to the poliovirus 5’ noncoding region. Science 241,445448. Troxler, M., Egger, D., Pfistner, T., and Bienz, K. (1992). Intracellular localization of poliovirus RNA by in situ hybridization a t the ultrastructural level using singlestranded riboprobes. Virology 191, 687-697. Tsao, J., Chapman, S., Agbandje, M., Keller, W., Smith, K., Wu, H., Luo, M., Smith, J., Rassmann, M. G., Compans, R. W., and Pai-rish, C. R. (1991). The three-dimensional structure of canine parvovirus and its functional implications. Science 251, 14561464. Tucker, S. P., Thornton, C. L., Wimmer, E., and Compans, R. W. (1993). Vectorial release of poliovirus from polarized human intestinal epithelial cells. J. Virol. 67,4274-4282. Turner, P. C., Young, D. C., Flanegan, J. B., and Moyer, R. W. (1989).’Interference with vaccinia virus growth caused by insertion of the coding sequence for poliovirus protease 2A. Virology 173, 509-521. Van der Werf, S., Bradley, J., Wimmer, E., Studier, F. W., and Dunn, J. J. (1986). Synthesis of infectious poliovirus RNA by purified T7 RNA polymerase. Proc. Natl. Acud. Sci. U S A . 83,2330-2334. Watanabe, Y., Watanabe, K., and Hinuma, Y. (1962). Synthesis of poliovirus-specific proteins in HeLa cells. Biochim. Biophys. Actu 61, 976-977. Wilcox, C., Hu, J. S., and Olson, E. N. (1987). Acylation of proteins with myristic acid occurs contranslationally. Science 238, 1275-1278. Wimmer, E. (1982). Genome-linked proteins of viruses. Cell (Cambridge, Muss.) 28,199201. Wycoff, E. E., Hershey, J. W. B., and Ehrenfeld, E. (1990). Eukaryotic initiation factor 3 is required for poliovirus 2A protease-induced cleavage of the p220 component of eukaryotic initiation factor 4F. Proc. Natl. Acad. Sci. U.S.A.87, 9529-9533. Yafal, A. G., and Palma, E. L. (1979). Morphogenesis of foot-and-mouth disease virus. I. Role of procapsids as virion precursors. J. Virol. 30, 643-649. Yin, F. H. (1977). Involvement of viral procapsid in the RNA synthesis and maturation of poliovirus. Virology 83, 299-307. Yogo, Y., and Wimmer, E. (1975). Sequence studies of poliovirus RNA. 111. Polyuridylic acid and polyadenylic acid as components of purified poliovirus replicative intermediate. J . Mol. Biol. 92, 467-477. Ypma-Wong, M. F., Dewalt, P. G., Johnson, V. H., Lamb, J. G., and Semler, B. L. (1988a). Protein 3CD is the major poliovirus proteinase responsible for cleavage of the P1 capsid precursor. Virology 166, 265-270. Ypma-Wong, M. F., Filman, D. J., Hogle, J. M., and Semler, B. L. (1988b3. Structural domains of the poliovirus polyprotein are major determinants for proteolytic cleavage at Gln-Gly pairs. J. Biol. Chem. 263, 17846-17856. Yu, S. F., and Lloyd, R. E. (1991). Identification of essential amino acid residues in the functional activity of poliovirus 2A protease. Virology 182, 615-625. Zlotnick, A., Reddy, V. S., Dasgupta, R., Schneemann, A., Ray, W., Jr., Rueckert, R. R., and Johnson, J. E. (1994). Capsid assembly in a family of animal viruses primes an autoproteolytic maturation that depends on a single aspartic acid residue. J. Biol. Chem. 269,13680-13684. Zoller, M. J., and Smith, M. (1983). Oligonucleotide-directed mutagenesis of DNA fragments cloned into M13 vectors. In “Methods in Enzymology” (R. Wu, L. Grossman, and K. Moldave, eds.), Vol. 100, pp. 468-500. Academic Press, New York.
ADVANCES IN VIRUS RESEARCH, VOL. 46
GENOME REARRANGEMENTS OF ROTAVIRUSES
Ulrich Desselberger Clinical Microbiology and Public Health Laboratory Addenbrooke’s Hospital Cambridge CB2 2QW, England
I. 11. 111. IV.
V.
VI. VII. VIII. IX. X.
Discovery of Genome Rearrangements Extent of Genome Rearrangements in Rotaviruses Sequence Data of Rearranged Genes Genome Rearrangements Generated in Vitro in Cultured Cells Mechanisms of Genome Rearrangements Biophysical Data Function of Rearranged Genes and Their Products Genome Rearrangements and Evolution of Rotaviruses Genome Rearrangements in Other Genera of Reoviridae Outlook References
I. DISCOVERY OF GENOME REARRANGEMENTS Rotaviruses are the main cause of viral gastroenteritis in infants and young children and in the young of a large variety of animal species (Kapikian and Chanock, 1990).There are at least five different groups, named A-E (Pedley et al., 1986). Group A rotaviruses are responsible for the vast majority of human infections. Rotaviruses have a genome consisting of 11 segments of double-stranded RNA (dsRNA) of approximately 18,500 nucleotide pairs in total size (Estes, 1990). The RNA segments can be easily extracted from virus particles, separated by polyacrylamide gel electrophoresis (PAGE),and visualized by silver staining, ethidium bromide staining, or radiolabeling. Typical RNA profiles show four size classes (I,segments 1-4; 11, segments 5 and 6; 111, segments 7-9; and IV, segments 10 and 11)(Estes, 1990). However, these profiles are not always seen. Pedley et al. (1984) investigated rotaviruses isolated from chronically infected children with severe combined immunodeficiency (SCID). Rotavirus infections in the immunocompetent host are normally overcome within 1 week, but in SCID children rotaviruses and many other viruses establish chronic infections that result in virus shedding over many weeks, months, and even several years (Saulsbury et al., 1980; Booth et al., 1982; Chrystie et al., 1982). Rotaviruses obtained from serial fecal specimens of such children produced abnormal RNA profiles: normal 69
Copyright 0 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
70
ULRICH DESSELBERGER
RNA segments were decreased in their relative concentration or even completely lost from the profiles, but additional bands of RNA were seen migrating between RNA segments 1and 7 (Fig. 1). The intensity of the additional bands varied. It was quickly established that the additional bands consisted of dsRNA and also that they had not arisen by noncovalent linkage (Pedley et al., 1984). Northern blots of the RNAs were probed with segment-specific radiolabeled cDNA clones of bovine rotaviruses under conditions to give segment-specific reactions in controls. Blots of atypical profiles often showed multiple hybridizations: that of the homologous RNA segment of standard size and several bands of dsRNA which always migrated higher up in the gel. This hybridization pattern was maintained when blots of RNA separated under denaturing conditions (Bailey and Davidson, 1976) were probed. An example for segment 9-specific hybridization is given in Fig. 2. This indicated that the additional bands of dsRNA contained segmentspecific sequences in the form of covalently bonded concatemers.
FJG. 1. RNA profiles of serial rotavirus specimens obtained from chronically infected patient A.K. (dates of specimens are indicated a t top). All specimens were 3’-endlabeled with [32PlpCp as described by Clarke and McCrae (1981),separated on a nondenaturing polyacrylamide gel, and autoradiographed. Bovine rotavirus and human rotavirus obtained from acute infections served as controls. Order numbers of segments are indicated on both sides, and additional bands are marked by arrowheads. From Pedley et al. (1984), with permission of the authors and publisher.
GENOME REARRANGEMENTS OF ROTAVIRUSES
71
FIG.2. Hybridization of rotavirus RNA samples of patient A.K. (dates of specimens are indicated at top) and of human and bovine control RNAs on DPT paper blots to RNA segment 9-specific radiolabeled cDNA probe. 3'-End-labeled bovine rotavirus RNA (L bovine) and unlabeled bovine and human rotavirus RNAs served as controls. Autoradiogram. From Pedley et al. (1984), with permission of the authors and publisher.
When several specimens sequentially obtained from the same person were subjected to such investigation, extra bands were found over a wide range of the profile. These bands varied in intensity and appeared and disappeared on passing through the chronological series (Pedley et al., 1984;Fig. 3).Where segment derivation could be established, the molecular weights of the additional bands were not simple multiple integers of the segments from which they were derived. The variable intensity of the additional bands and of some of the normal RNA segments lead to the hypothesis that either parts of the RNA genome occurred in abnormal configuration in single virus particles or that subpopulations of viruses possessing normal and abnormal genomes coexisted and cocirculated (Pedley et al., 1984). The question also arose whether viruses possessing such genomes were defective interfering (DI) particles (Holland et al., 1980).By contrast to DI RNAs that are characterized by internal deletions (Davis et al., 19801,the larger size and migrational pattern of the additional rotavirus bands, which were maintained under denaturing conditions, excluded such a possibility (Pedley et al., 1984).Mosaic structures as
72
ULRICH DESSELBERGER
FIG.3. RNA profiles of sequential rotavirus samples (dates are indicated at top) of patient U.H. Cenomic dsRNA was extracted, separated on a 2.8% polyacrylamide-6 M urea gel, and stained with silver. Bovine rotavirus RNA served as an internal control. Numbers of segments and positions of additional bands (arrowheads) are indicated a t right. From Pedley et al. (1984), with permission of the authors and publisher.
described by Fields and Winter (1982) remained a possibility. When RNA segments were separated by PAGE for a short period, in no case were additional bands of RNA found migrating faster than the smallest RNA segment (U. Desselberger, 1985, unpublished results). The discovery of group A rotaviruses with abnormal RNA profiles also raised the question of whether the dictum of “ atypical” RNA profiles in other rotavirus groups (B-E)could be maintained (Pedley et al., 1984).
GENOME REARRANGEMENTS OF ROTAVIRUSES
73
11. EXTENT OF GENOME REARRANGEMENTS IN ROTAVIRUSES Since the original discovery genome rearrangements have been described by several independent groups t o occur not only in human rotaviruses but also in rotaviruses of a variety of animal species (humans: Albert, 1985; Dolan et al., 1985; Eiden et al., 1985; Matsuno et al., 1985; Besselaar et al., 1986; Hundley et al., 1987; Matsui et al., 1990; Mendez et al., 1992; Gault-FrBre et al., 1995; calves: Pocock, 1987; Paul et al., 1988; Scott et al., 1989; Tian et al., 1993; rabbits: Thouless et al., 1986; Tanaka et al., 1988; piglets: Bellinzoni et al., 1987; Mattion et al., 1988; Lambs: Shen et al., 1994). Whereas the initial observation was in immunodeficient children, the observations in animals and some of those in humans were in immunocompetent hosts. In a South African hospital viruses with genome rearrangements circulated for several months, infecting apparently healthy children (Besselaar et al., 1986). DATAOF REARRANGED GENES 111. SEQUENCE Nucleotide sequences of rearranged genes of several group A rotavirus strains of different origin have been obtained, and references and nucleotide sequence accession numbers are summarized in Table I. In most cases the genome rearrangement consists of a partial duplication of sequences of the open reading frame (ORF) starting beyond the termination codon and extending then to the 3‘ end of the normal gene. This is diagrammatically shown in Fig. 4 for rearranged RNA 10 of a human rotavirus isolate (Ballard et al., 1992); similar changes were also found for rearrangements of other RNAs 10 (Matsui et al., 1990),for RNAs 11(Gonzalez et al., 1989; Gorziglia et al., 1989; Scott et al., 19891, and for one RNA 5 (Hua and Patton, 1994). In most rearranged genes, the sequence runs from a normal 5’ untranslated region (UTR) and through a normal ORF. At various nucleotide positions after the termination codon (0-23; Table I), the duplication starts reinitiating from various places within the ORF but downstream of the initiation codon and then reads through a duplicated termination codon and toward a normal 3’ UTR. As the duplication of the sequence normally starts beyond the initiation codon, it remains silent as a whole, and the resulting genes have enormously long 3’ UTRs, up to 1800-1900 bp (McIntyre et al., 1987; Hua and Patton, 19941, in contrast to the relatively short 3’ UTRs (17-185 nucleotides) of the standard length genes (Estes, 1990; Desselberger and McCrae, 1994).
TABLE I SEQUENCED GENOMEREARRANGEMENTS OF ROTAVIRUSES
RNA segment=
5 5 6 7 10
10 11 11 11
Strain brv E brv A Lp 14 H 57 A64
VMFU C71183 C60 X1 Alabama
Origin
Start of reiteration in relation to termination codon
Bovine Bovine Ovine Human Human Human Bovine Pig Lapine
-596 -52 23 0 2 0 0 6 4
Number of point mutations compared to standard geneb ND 16 6 ND 11
23 NA 33 NA
GenbankIEMBL accession number Standard gene
Rearranged gene
224735 L12248 L11596 NAc DO1146 NA NA NA NA
212108 L11575 L11595 NA DO1145 NA NA NA NA
Refs. Tian et al. (1993) Hua and Patton (1994) Shen et al. (1994) Mendez et a1. (1992) Ballard et al. (1992) Matsui et al. (1990) Scott et al. (1989) Gonzalez et al. (1989) Gorziglia et al. (1989)
a Genome rearrangements have also been observed in segment 6 of a human strain (Pedley et al., 1984) and in segments 7,8, and 9 of human strains (coding for NSP2 and NSP3) (Pedley et al., 1984; Hundley et al., 1987; Gault-Frere et al., 1995), but they have not been sequenced so far. b ND, Not determined NA, not applicable. c Partial sequence (junction region).
75
GENOME REARRANGEMENTS OF ROTAVIRUSES Normal gene 10 I4181
569
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Rearranged gene 10 FIG.4. Structures of normal and rearranged genes 10 of a human rotavirus (isolates A28 and A64, respectively).The solid bar represents the complete ORF, and the open bar symbolizes the duplicated part of the ORF of the normal gene (untranslated).Solid lines indicate 5' and 3' untranslated regions as well as sequences between the normal and duplicated ORFs. From Ballard et al. (19921, with permission of the authors and publisher.
However, RNA segment 5 of a bovine rotavirus (brv) variant was rearranged in several different ways with variants being called brv A and brv E (Tian et al., 1993). In the case of segment 5 of the brv E variant, the duplication had started before the termination codon, and an extended ORF ensued encoding segment-5-specific amino acids as the reiteration had started in frame (Fig. 5). The extended ORF codes for a protein VP5E of 728 amino acids, which was verified by PAGE of [36Slmethionine-labeled proteins of brv E-infected cells (Hundley et al., 1985; Tian et al., 1993). In contrast, rearranged segment 5 of the brv A variant possesses a different structure. The reiteration starts 52 nucleotides before the stop codon (in position 1454), but one of several additional point mutations changes the picture further: a mutation in position 808 results in a new termination codon (TAG) allowing an ORF of only 258 amino acids k e . , of 31 kDa size, slightly more than half the size of the normal product of 491 amino acids, i.e., 58 kDa). Thus a gene of 2693 bp results in only 774 (positions 33-806, i.e., 28.7%) coding for a protein! The abnormal product (Fig. 6) was detected by Hua and Patton (1994) after it had escaped screening by Hundley et al. (1985) and Tian et al. (1993), apparently because it comigrates with cellular gene products. A point mutation in the ORF of a rearranged gene 6 also had profound consequences for protein stability (see below).
76
ULRICH DESSELBERGER
FIG. 5. Structures of normal and rearranged forms of RNA segment 5 of bovine rotavirus (brv UKtc and brv E, respectively). The junction sequence is spelled out at the bottom, showing 6 amino acids on either side. From Tian et al. (19931,with permission of the authors and publisher.
Some time ago it was found that RNA segment 10 of a “short” electropherotype human group A rotavirus codes for a protein which corresponds to the product of RNA 11 of “long” electropherotype rotaviruses (Dyall-Smith and Holmes, 1981). The observations by Matsui et al. (1990) on gene 11 equivalents of rotavirus genomes yielding
FIG. 6. Diagram of standard gene 5 of bovine rotavirus and of gene A of the brv A variant. Gene duplication in gene A starts 2 positions after the termination codon. The point mutation in position 808 giving rise to an additional termination codon in gene A is indicated. The ORFs of the gene products are also shown. From Hua and Patton (1994), with permission of the authors and publisher.
GENOME REARRANGEMENTS OF ROTAVIRUSES
77
“short” and “supershort” PAGE profiles were most intriguing: whereas “supershort” strain VMRI clearly contained a partial duplication at its 3’ end, the RNA segments 10 of “short” strain DS-1 and of “supershort” strain M69 have sequences at their 3’ ends that were similar to one another but not related to any other available rotavirus gene sequence. Finally, it is remarkable that direct repeats of nucleotide sequences were observed closely upstream of the start of the duplications in a number of cases (Gorziglia et al., 1989; Ballard et al., 1992; Shen et al., 1994), but not in others (Scott et al., 1989; Matsui et al., 1990). The numbers of point mutations in the rearranged compared to the normal genes varied widely: between 6 and 33 have been counted (Table I). No genome rearrangement has been described so far which had resulted in a mosaic of sequences donated from several different RNA segments, in contrast to the DI mosaic structures of influenza viruses described by Fields and Winter (1982). IV. GENOME REARRANGEMENTS GENERATED in Vitro IN CULTURED CELLS Before nucleotide sequences of rearranged genes and biological properties of the viruses carrying them were known (see below), the phenomenon of genome rearrangements appeared to be related to that of the formation of DI RNAs. As serial passage in uitro of virus at high multiplicity of infection (MOI) has been found to be the most efficient way to generate viruses with DI genomes, this method was used to propagate rotaviruses (Hundley et al., 1985).Surprisingly, viruses with genome rearrangements (i.e., partial duplications) but not genome deletions emerged (Fig. 7). Bovine rotavirus with a standard genome transformed into brv variants with rearranged RNA segments 5 , among others variants brv A and brv E (Fig. 7, lanes 2 and 3; see also Section 111). As in virus cultures with DI RNAs, yields in virus increased and decreased in a periodic manner, and the absolute yields in viral infectivity were inversely correlated with the ratios of numbers of virus particles (nop) over infectivity [nvp/pfu (plaque-forming units)]. A t passages 7-8, viruses with genome rearrangements appeared and overgrew the virus with standard genome. This was a reproducible phenomenon and was obtainable after repeated plaquepurifications of standard virus (Fig. 8). The outcome of repeat experiments, however, was not identical in that RNA 5 equivalents with apparently different forms of rearrangements were found. The in uitro generation of viruses with rearranged genomes was reproduced with Chinese lamb rotaviruses by Shen and Bai (1990).
78
ULRICH DESSELBERGER
FIG. 7. RVA profiles of plaque-purified bovine rotaviruses obtained after serial passage at high MOI. RNA segment numbers are indicated on the right-hand side. Open arrowheads denote missing RNA segments, closed arrowheads additional RNA bands. Lanes 1and 6 show standard bovine rotavirus; lane 2, brv A; lane 3, brv E; lane 4, brv F; lane 5; brv G/H (likely to be a mixture). Analysis in 2.8% polyacrylamide-6 M urea gel, stained with silver. From Hundley et al. (1989, with permission of the authors and publisher.
Viruses possessing genome rearrangements could be plaque-purified very easily, and six times plaque-to-plaque purified virus grew perfectly well without showing the appearance of virus with standard ge-
GENOME REARRANGEMENTS OF ROTAVIRUSES RNAsegmentS+ RNAbandA-
+
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106 J n.v.p./p.f.u.
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9
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6
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Passage number FIG.8. Plot of yield of infectious virus (pfuhl) and of nvp/pfu in harvested tissue culture fluids against passage number (at high MOI). The presence or absence of RNA segment 5 and RNA band A (Fig. 7, lane 2) is indicated at the top. From Hundley et al. (1985),with permission of the authors and publisher.
nomes and remained genetically stable. These and other experiments (see below) proved that bovine rotaviruses with rearranged genes are not DI viruses. The nvp/pfu ratios were equally low for brv standard and brv A viruses (Table 11). It was also shown that genome rearrangements were a continuous phenomenon. When six times plaque-purified brv A was again serially propagated a t high MOI, second generation
80
ULRICH DESSELBERGER TABLE I1 INFECTIVITY, CONCENTRATION OF VIRUSPARTICLES, AND nvplpfu STOCKS OF STANDARD BOVINEROTAVIRUS.AND RATIOOF CLONED brv A VARIANT WITH REARRANGED GENOME~,~ Stock and preparation Standard brva 1 2 3
Infectivity (pfulml, x 107)
4 15 10 10 +. 5 brv A with rearranged genomeb 1 3 10 15 6 2 3 6 3 6 k 4
Concentration (nvpiml, x 106)
nvpipfu
1.2 9 11 7 t 4
3 6 11 7 2 3
8 5 8 10 6 2 4 4 6?3
27 5 5 16 30 7 7 13 14 9
*
aN=3. bN=8. c From Hundley et al. (1985), with permission of the authors and publisher. The brv A had a genome missing RNA segment 5 and possessing RNA band A. The arithmetic means +- standard deviation are indicated. The corresponding arithmetic means of standard brv and of brv A with rearranged genome did not differ significantly one from another ( t test, p < 0.05).
rearrangements resulted (viruses brv K and brv L; Fig. 9). When cells were infected with brv standard and brv A at different MOIs, the outcome depended on whether passage was at low or high MOI: in the first case, standard brv overgrew; in the latter, the brvA variant (Hundley et al., 1985). The effect of genome rearrangements on growth in cell culture will be discussed below.
V. MECHANISMS OF GENOMEREARRANGEMENTS The sequence data available (see Section 111) allow a formal description of genome rearrangements as partial duplications (concatemer formation) with varying consequences relative to their expression. Start of the duplication after the termination codon (excluding the
GENOME REARRANGEMENTS OF ROTAVIRUSES
81
FIG.9. RNA profiles of viruses with second generation genome rearrangements, brv K and brv L, obtained after repassage of plaque-purifiedbrv A at high MOI.(A) Analysis in a 10%polyacrylamide gel, stained with ethidium bromide. (B,C) Autoradiograms of Northern blots probed with 32P-labeled cDNA produced from RNA segment 5 (B)or RNA band L (C) according to Hundley eb al. (1987).
initiation codon) leads to long 3' UTRs, whereas start of the duplication before the termination codon leads to longer than normal ORFs and normal 3' UTRs. It is not clear, however, at which step of the replication cycle the duplication event occurs. It has been suggested that the RNAdependent RNA polymerase of rotaviruses (associated with the particle core and coded for by RNA 1; Estes, 1990) can fall back on its template at various steps of transcription (plus strand synthesis) and reinitiate and retranscribe from that template a t different places (Fig. 10A). Messenger RNAs of the rearranged size are transcribed in uitro from particles containing rearranged genes (Hundley et al., 1985), and rotaviruses with genome rearrangements are genetically stable (see Section IV). Alternatively, the primary duplication event could occur at the level of replication (negative strand synthesis) (Fig. 10B).Whereas occurrence of duplication at the transcription stage would mean that the abnormal mRNA is packaged and a strand of negative sense replicated from it in the new precore particles (Gallegos and Patton, 1989), occurrence of rearrangements at the replication stage would imply that a rearranged negative strand forms a heterohybrid with the normal positive strand, and that at the next round of infection this rearranged negative strand is transcribed at full length. As packaging of the RNA genome is very tightly controlled, a virus particle will contain only one form of one segment each.
ULRICH DESSELBERGER
82
A
B
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FIG.10. Possible mechanisms for emergence of genome rearrangements (A) during plus strand synthesis and (B) during minus strand synthesis (in precore particles). Bold lines represent the minus strand; fine lines, plus strand; dashed lines, newly synthesized strands; open circle, RNA-dependent RNA polymerase; arrows, direction of synthesis.
GENOME REARRANGEMENTS OF ROTAVIRUSES
83
Most genome rearrangements that have been sequenced can be described as intramolecular recombination events, and direct repeats close to the recombination site are often but not always found. This is similar to what has been observed in the phi (4) 6 system (Mindich et al., 1992; Onodera et al., 1993).In the case of poliovirus recombination, it was shown that recombination favored the step of secondary transcription (from the negative strand of the replicative intermediate, or RI) (Kirkegaard and Baltimore, 1986).In the phi 6 system where intermolecular recombination between the three different segments of dsRNA of the viral genome can be observed, it was shown very elegantly that recombination also occurs at the step of negative strand synthesis (Onodera et al., 1993). However, under the special conditions of those experiments only phi 6 recombinants would survive and would therefore be positively selected for in the surviving viruses. It is of interest to note that direct repeats favor genome rearrangements of rotaviruses although they do not seem to be an absolute requirement. They were found in sequenced genes by Gorziglia et al. (1989), Ballard et al. (19921, and Shen et al. (19941, but not by Scott et al. (1989) or Matsui et al. (1990). Onodera et al. (1993) in their system show very nicely that sequence identity of the landing pad for the donor strand-polymerase complex compared to the lift off point is not a prerequisite but is preferred. Although the mechanism of recombination in genome rearrangements of rotaviruses has not been elucidated, the data are consistent with the copy choice model (Kirkegaard and Baltimore, 1986; Lai, 1992) in which specific sequence homologies or secondary structures are involved in directing the switch of the polymerase (Romanova et al., 1986). Mechanisms of genome rearrangements should be explored further in in uitro transcription (Cohen et al., 1979) and replication (Chen et al., 1994) systems. Some electron microscopy data on rotaviruses are of interest in this context. Using the Kleinschmidt technique the lengths of rotavirus RNA segments have been determined, and the measurements were very precise when compared to the length obtained by sequence data (Rixon et al., 1984). Whereas viruses with standard genomes show less than 2%RNA molecules which are larger than RNA 1,viruses with rearranged genomes show about 15%RNA concatemers longer than RNA 1 and of varying length (U. Desselberger and F.Rixon, 1985, unpublished data). This suggests that rearrangements which are amplified to amounts of normal segments (see below) are only part of numerous other recombination events which did not survive. Some of the results obtained by Matsui et al. (1990) are difficult to explain; these workers obtained long 3’ UTRs of RNA segments of
84
ULRICH DESSELBERGER
larger than normal size without the evidence of an intramolecular duplication. These sequences could have “mutated away” from original duplications (being under no functional constraint) or could have been picked up from as yet unidentified cellular sequences (Qian et al., 1991),or identifiable cellular sequences as found for influenza viruses (Khatchikian et al., 1989). VI. BIOPHYSICAL DATA Once it became possible to grow human rotaviruses with genome rearrangements (Hundley et al., 1987;see below), various variants with different combinations of genome rearrangements were found. The viruses had between 450 and 1790 bp of additional RNA packaged, amounting to 1.4 to 9.6% of the standard genome size. By electron microscopy such particles were indistinguishable in size or shape from viruses possessing a standard genome (Hundley et al., 1987;McIntyre et al., 1987).Examples of the RNA profiles of such viruses are given in Fig. 11A;rotaviruses of such RNA profiles had 450,1070,1570,and 1790 bp additional RNA packaged. The viruses differed in density as determined by analytical ultracentrifugation (Fig. llB),and the differences in density were directly proportional to the number of additionally packaged base pairs (Fig. 11C;McIntyre et al., 1987).Thus, packaging of rotavirus genomes is flexible in terms of the size of packaged segments, and additionally packaged base pairs amounting to up to 10% of the total genome size were tolerated in the variant with rearrangements. Particles of viruses with up to 10% additional base pairs packaged were morphologically indistinguishable from standard rotavirus (Hundley et al., 1987).In recombinants of the phi 6 system up to 16.7% of the genome size were additionally packaged without apparent effect on the procapsid (L. Mindich, 1994,personal communication). VII. FUNCTION OF REARRANGED GENESAND THEIRPRODUCTS For some time after the first observation of viruses with genome rearrangements it was not possible to grow them in tissue culture. Therefore, it was not clear whether those viruses were functionally defective, possibly due to rearrangements of genes. Secondary rhesus monkey kidney (RMK) cell cultures infected with a human rotavirus isolate (U.H.) showing genome rearrangements by PAGE (Pedley et al., 1984)showed no cytopathic effect, although common primary group A rotavirus isolates grow very well on RMK cells (Ward et al., 1984). However, on superinfection with the tissue culture-adapted bovine ro-
FIG.11. (A) RNA profiles of bovine rotavirus and of human rotavirus variants with rearranged genomes of genotypes 2,3,7,and 9 (Hundley et al., 1987). Segment numbers (1-11) are indicated on the left-hand side, and the position and origin of rearranged bands identified on the right-hand side (bands a and f were derived from RNA 8, band d from RNA 10, and bands c and e from RNA 11; Hundley et al., 1987). The number of additionally packaged base pairs is indicated at the bottom. Analysis in 2.8% polyacrylamide-6 M urea gel, stained with silver. (B) Scans after analytical equilibrium centrifugation in CsCl of mixtures of single-shelled particles containing RNA of standard bovine rotavirus and of human rotaviruses with rearranged genomes of genotypes 2,3, and 9. Numbers of additionally packaged base pairs are indicated below scans. (C) Plot of difference in density as determined from data shown in B against number of additionally packaged base pairs. From McIntyre et al. (1987), with permission of the authors and publisher.
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ULRICH DESSELBERGER
taviruses (brv, UK Compton strain), some of the rearranged RNA segments of the U.H. virus were disproportionally amplified (Fig. 12, lane M; Allen and Desselberger, 1985). Plaques obtained from the original yield of such cultures were grown, and the RNA was extracted and analyzed by PAGE. Extensive reassortment had taken place (Fig. 12; Allen and Desselberger, 1985) occurring between standard segments of the brv and U.H. virus (no segments 5 and 6; segment 6 not shown) but also the rearranged RNAs. Standard length RNA segment 11 of brv was replaced by RNA bands F (not shown) or G and standard RNA
FIG. 12. RNA profiles of rotaviruses grown from 12 individual plaques derived from a mixed infection of bovine rotavirus and human rotavirus U.H. with rearranged genome. The RNA profile of the direct yield of the mixed infection (M)is shown in the right-hand lane. Segments 1-11 of brv and segment 5 (h5) and several rearranged bands (D,G) of the human rotavirus are denoted at right. Analysis in 2.8% polyacrylamide-6 M urea gel, stained with silver. From Allen and Desselberger (1989, with permission of the authors and publisher.
87
GENOME REARRANGEMENTS OF ROTAVIRUSES
segment 9 by bands B (not shown) or D. The segmental origin of the rearranged bands was confirmed by Northern blotting followed by hybridization of segment-specific radiolabeled probes (Allen and Desselberger, 1985). The PAGE profiles of proteins from infected cells demonstrated that the rearranged RNA bands produced normal-sized length virus-coded proteins (Allen and Desselberger, 1985), indicating that the rearranged RNAs replaced the normal RNA segments structurally and functionally. The reassortants grew well on their own in uitro, could be plaque-to-plaque purified multiple times, and remained genetically stable. Rotaviruses with genome rearrangements that had arisen after serial passage a t high MOI in uitro were equally able to reassort with human rotaviruses carrying a standard genome (Biryahwaho et al., 1987). In contrast, in cases when the normal ORF was extended (brv variant E; Hundley et ul,., 1985; Tian et al., 1993), abrogated (brv variant A; Hua and Patton, 1994), or mutated (Chinese lamb rotavirus; Shen et al., 19941, functional changes were observed. Bovine rotavirus variants E and A showed 9- to 60-fold lower yields, respectively, in single-step growth experiments and produced smaller plaques, with brv E giving plaques 40%and brv A 2% the size of plaques of standard brv (Table 111; Tian et al., 1993). The analysis of rearrangements of RNA segment 5 were of particular interest as both extension (brv E) and abrogation (brv A) of the normal ORF were found. Bovine rotavirus variant A had a truncated VP5 of 258 amino acids (due to a termination codon at positions 806808) instead of the authentic size of 491 amino acids, but was viable, nondefective, and genetically stable (Hundley et ul., 1985; Hua and
TABLE I11 IN VITROGROWTH PROPERTIES OF STANDARD BOVINE F~OTAVIRUS AND VARIANTS brv E AND brv Aa Log pfu/ml at time postinfectionb Virus
30 hr
46 hr
Plaque diameter (mm) at 7 days (mean ? SD)
Mean plaque size (mm2)
Standard brv brv E brv A
8.9 7.8 7.2
8.5 7.5 7.1
7.6 2 0.8 ( n = 9) 4.8 & 0.7 ( n = 16) 1.0 2 0.7 ( n = 50)
45.4
~~
~
18.1
0.4
~
From Tian et al. (1993), with permission of the authors and publisher. Single-step growth experiments were carried out in MA-104 cells infected at an MOI of 10 pfu per cell. Q
88
ULRICH DESSELBERGER
Patton, 1994). It was also found to be associated with the cytoskeleton of the infected cell like its normal size counterpart, demonstrating that the carboxyl-terminal half of VP5 (NSP1, NS53) is not required for rotavirus replication in vitro (Hua and Patton, 1994). Tian et al. (1993) described the even more drastically deleted VP5 gene product of rotavirus P9A5 (originally isolated from a foal) in which a deletion occurred between nucleotides 460 and 768 of the normal gene sequence. This deletion then caused a frameshift such that a stop codon was introduced 8 amino acids downstream of the deletion point, giving a predicted size of the gene product of 150 amino acids instead of the authentic size of 491 amino acids. Taniguchi et al. (1994,1995) recently described deleted VP5 genes of bovine rotavirus isolates from Thailand which had additional termination codons predicting ORFs of only 40-50 amino acids in length. The predicted protein products have so far not been found. The overall requirement of VP5 for rotavirus replication is under discussion. In a Chinese lamb rotavirus, rearrangement of RNA segment 6, the gene coding for the inner capsid protein VP6, was observed in a similar way (Shen et al., 1994) as shown by Ballard et al. (1992)for segment 10. However, the rearranged RNA6 was found to be accompanied by a point mutation in nucleotide position 949 (within the normal ORF), giving rise to a change in amino acid position 309 (from a proline to a glutamine) as the only amino acid difference compared to the VP6 of the standard genome virus, which was also available from the same lamb isolate. Proline in position 309 of VP6 is highly conserved in all group A rotavirus strains. The amino acid difference in position 309 occurred in a region of VP6 previously implicated as being important for trimerization and the formation of single-shelled particles (Clapp and Patton, 1991).The VP6 protein carrying the 309 mutation was found to be less stable than the corresponding standard VP6. Under mild denaturing conditions it did not separate on gels as a trimer (Sabara et al., 1987)but as a monomer, and it was less stable toward acid pH by almost a whole pH unit compared to the standard VP6 (Shen et al., 1994). The nvp/pfu ratio of virus possessing normal VP6 was significantly lower than that of virus carrying the mutated VP6 (Shen et al., 1994). Analysis of over 500 plaque isolates of a reassortant mixture of human viruses with genome rearrangements and standard bovine rotaviruses showed that reassortment was nonrandom, that there was linkage of occurrence of certain genes (i.e., RNA segments 5,9, and 11) in reassortants, and that the host cells on which plaque isolates were obtained (MA104 or BSC-1 cells) influenced the frequencies with which certain reassortants were recovered (Graham et al., 1987).These findings were not different from those established for other viruses
GENOME REARRANGEMENTS OF ROTAVIRUSES
89
with segmented genomes (reoviruses; Wenske et al., 1985; influenza viruses, Lubeck et al., 19791, and RNA segments with rearrangements participated in this process like standard RNA segments (Graham et al., 1987). VIII. GENOMEREARRANGEMENTS AND EVOLUTION OF ROTAVIRUSES Initially, genome rearrangements of rotaviruses were seen only in rare cases of immunodeficient human hosts (Pedley et al., 1984; Albert, 1985; Eiden et al., 1985; Dolan et al., 1985) and were thought to be more a curiosity than of particular significance. However, when rotaviruses with genome rearrangements were found to circulate for months in immunocompetent children as a nosocomial infection (Besselaar et al., 19861, and also freely circulating in a variety of otherwise healthy animal hosts (rabbits: Thouless et al., 1986; Tanaka et al., 1988; calves: Pocock, 1987; pigs: Bellinzoni et al., 19871, it became clear that the phenomenon was more frequent than originally anticipated. The various forms of rearrangements occurred mainly in genes coding for nonstructural proteins [RNA segments 5 , 8, and 9 (depending on strain), 10, and 111,but were also found for gene 6 (Pedley et al., 1984; Shen et al., 1994). These rearrangements produced RNA profiles of great diversity that were highly atypical for group A rotaviruses (Fig. 13; Desselberger, 1989). The data presented in Section VII demonstrated that genome rearrangements alone (or combined with point mutations) were able to change the structure and function of encoded proteins. It had been shown that within a single individual various forms of genome rearrangements (e.g., affecting RNA segments 8, 10, and 11) and various combinations thereof in plaque-purified viruses coexisted. At least 12 subpopulations were identified in one isolate (Fig. 14; Hundley et al., 1987) and changed in relative prevalence when observed over time in chronically infected hosts (Pedley et al., 1984; Hundley et al., 1987).Thus, multiple rearrangement variants coexisted in a constantly varying (dynamic) equilibrium, fulfilling the criteria for the presence of a quasispecies as has been described for the coexistence of various point mutants for a number of RNA viruses (Holland et al., 1982; Holland, 1984; Doming0 et al., 1985).In summary, it is therefore proposed that genome rearrangements, besides genetic point mutations (Sabara et al., 1982; Desselberger et al., 1986) and a reassortment continuum (Palese, 1984), are a third principle of the evolution of rotaviruses and can contribute to the diversity of rotaviruses in the field (Hundley et al., 1987; Desselberger, 1989; Tian et al., 1993).
90
ULRICH DESSELBERGER Group A Rotaviruses
1-
23 4-
5-
~
B
-
C
D
E
-
-
6 -
-4
1 1 0 11
-+
Rotaviruses of Groups
'Atypical'
'TvDical' human
-
- -
4
'long' ' s h o d
4 a
- -
,I b
c
d
d
1
! e
-
e
-
GI f
f
FIG.13. Diagram of RNA profiles of various group A rotaviruses with genome rearrangements and of typical RNA profiles of group A and group B-E rotaviruses. Open arrowheads denote missing normal RNA segments: closed arrowheads show various rearranged equivalents in viruses a-f. From Desselberger (1989),with permission of the publisher.
IX. GENOME REARRANGEMENTS IN OTHERGENERA OF Reouiridue Genome rearrangements have also been found involving different RNA segments of several genotypes of bluetongue virus, members of the orbivirus family (Ramig et al., 1985; Eaton and Gould, 19871, and Joklik, 1992, personal comare likely to occur in orthoreoviruses (W. munication). Thus, this mechanism of genome change seems to be possible for most animal dsRNA viruses although much less is known for viruses other than rotaviruses.
X. OUTLOOK Since the original observation of genome rearrangements in rotaviruses, much has been learned about the detailed structure of rearranged genes and their products, their functions, and their significance for the overall diversity of rotaviruses. There are still gaps in our knowledge about the exact mechanismb) by which these genome forms emerge, and it remains to be seen to what extent they occur in other double-stranded RNA viruses.
91
GENOME REARRANGEMENTS OF ROTAVIRUSES
- I - =- - -- --- --- - -- - - - - -
l o - - - - - - 11
-
-
FIG. 14. Diagram of 12 subpopulations (lanes 1-12)of human rotaviruses with various forms of genome rearrangements isolated from a single individual with chronic infection. The bovine rotavirus standard genome is shown for comparison. RNA segments (1-11) are denoted on the left-hand side, as are rearranged bands (bands c and e derived from RNA 11,band d from RNA 10,and bands a, b, f, and g from RNA 8).From Hundley et al. (1987),with permission of the authors and publisher.
ACKNOWLEDGMENTS The author thanks M. K. Estes, H. Greenberg, M. A. McCrae, L. Mindich, and J. Patton for stimulating discussions and critical reading of the manuscript.
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Shen, S., Burke, B., and Desselberger, U. (1994). Rearrangements of the VP6 gene of a group A rotavirus in combination with a point mutation affecting trimer stability. J . Virol. 68, 1682-1688. Tanaka, T. N., Conner, M. E., Graham, D. Y.,and Estes, M. K. (1988). Molecular characterization of three rabbit rotavirus strains. Arch. Virol. 98, 253-265. Taniguchi, K. (1995). “Sequence analysis of VP5 genes of porcine rotaviruses from Thailand.” International Symposium on Viral Gastroenteritis, Sapporo, Japan. [Abstract] Taniguchi, K., Kojima, K., Kobayashi, N., Urasawa, T., and Urasawa, S. (1994). F’roperties of a bovine rotavirus variant with gene 5 having a deletion of 500 base pairs. ‘henty-eighth Joint Working Conference on Viral Diseases, Japan-US Cooperative Medical Science Program, Tokyo, Japan. [Abstract] Thouless, M. E., DiGiacomo, R. F., and Neuman, D. S. (1986). Isolation of two lapine rotaviruses: Characterization of their subgroup, serotype and RNA electropherotypes. Arch. Virol. 89, 161-170. Tian, Y.,Tarlow, O., Ballard, A., Desselberger, U., and McCrae, M. A. (1993). Genomic concatemerization/deletion in rotaviruses: A new mechanism for generating rapid genetic change of potential epidemiological importance. J. Virol. 67, 6625-6632. Ward, R. L., Knowlton, D. R., and Pierce, M. J. (1984). Efficiency of human rotavirus propagation in cell culture. J. Clin. Microbiol. 19, 748-753. Wenske, E. A., Chanock, S. J., Krata, L., and Fields, B. N. (1985). Genetic reassortment of mammalian reoviruses in mice. J. Virol. 56, 613-616.
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ADVANCES IN VIRUS RESEARCH, VOL. 46
HUMAN IMMUNODEFICIENCY VIRUS TYPE 1 REVERSE TRANSCRIPTASE AND EARLY EVENTS IN REVERSE TRANSCRIPTION Eric J. Arts and Mark A. Wainberg McGill University AIDS Centre The Sir Mortirner B. Davis-Jewish General Hospital Montreal, Quebec H3T 1E2, Canada
I. Introduction 11. Overview of Human Immunodeficiency Virus Qpe 1 Replication A. Initial Events in HIV-1 Replication B. Virus Assembly and Maturation of HIV-1 111. Human Immunodeficiency Virus Q p e 1 Reverse Transcriptase A. Structure of HIV-1 Reverse Transcriptase B. Interaction of HIV-1 Reverse Transcriptase with Primer and Template C. Polymerase Active Site and Deoxynucleoside 5'-Triphosphate Binding Site of HIV-1 Reverse Transcriptase IV. Human Immunodeficiency Virus Q p e 1 Reverse Transcription A. Overview of Reverse Transcription Scheme of Retroviruses B. Origin of HIV-1 Reverse Transcription C. Host tRNALys3 Primer in HIV-1 Reverse Transcription D. RNA- and DNA-Dependent DNA Polymerization E. Fidelity of Polymerization by HIV-1 Reverse Transcriptase F. Ribonucleases of HIV-1 Reverse Transcriptase G. First Template Switch References
I. INTRODUCTION The first cases of acquired immunodeficiency syndrome (AIDS) were reported in 1981 (Gottlieb et al., 1981; Masur et al., 1981).By this date, the disease had already rapidly spread in the homosexual community and among intravenous drug users. The causative agent thought to be responsible for this syndrome showed a pattern of blood-borne transmission, but it was not until 1983 that the pathogen was isolated and partially characterized (Barre-Sinoussi et al., 1983; Popovic et al., 1984). A unique human retrovirus, lymphadenopathy-associated virus or human T-cell lymphotropic virus type I11 [later termed the human 97
Copyright 0 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
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immunodeficiency virus type 1 (HIV-111, was the proposed etiological agent of AIDS. There are now five known human retroviral species: Human T-cell lymphotropic virus (HTLV) type I, HTLV type 11, HIV type 1,HIV type 2, and the human foamy virus. The human foamy virus belongs to the subfamily Spumavirinae and is not a human pathogen (Hotta and Loh, 1987), whereas HTLV-I and -11 are classified as Oncouirinae and are shown to cause chronic and sometimes fatal leukemia (Wong-Stahland Gallo, 1985). Infections with HIV-1 or HIV-2 showed the same latent progression to disease as occurs with animal lentiviruses such as Visna and equine infectious anemia viruses (EIAV) (reviewed by Cheevers and McGuire, 1985; Davis et al., 1987).The latter viruses are classified as Lentiuirinae and differ from classic retroviruses in genomic organization. Lentiviruses and HTLV/bovine leukemia viruses contain small open reading frames (ORF) found 3’ of the gag and pol genes and surrounding the enu gene. Many of these ORFs encode for viral accessory proteins, some of which are essential for virus replication. In the case of HIV-1, there are at least six ORFs in addition to the gag, pol, and enu genes (reviewed by Cullen, 1991). These accessory genes vary in position and code for proteins that differ in structure and function in different retroviruses. However, the pol genes of all retroviruses encode three enzymes, reverse transcriptase, integrase, and an aspartic proteinase, all of which are highly specific in function. The pol gene of EIAV and possibly other lentiviruses such as Visna virus and caprine arthritis-encephalitis virus (CAEV)contains a ORF encoding a deoxyuridine triphosphatase, similar to that found in herpesvirus (Threadgill et al., 1993). The reverse transcriptase (RT) enzyme was discovered independently by Howard Temin and David Baltimore (Temin and Mizutani, 1970; Baltimore, 1970). The discovery of an RNA-dependent DNA polymerase challenged a central scientific dogma which stated that the key to reproduction of any entity was limited to progression from DNA to RNA to protein. Since the discovery of RT in RNA tumor viruses, RNA-dependent DNA polymerization activities have been characterized in the telomeric DNA of nearly all eukaryotes (Greider and Blackburn, 1985), in retrotransposons (Boeke et al., 1985; Temin, 1985), in cauliflower mosaic virus (Guilley et al., 1983; Pfeiffer and Hohn, 1983), in bacteria such as Myxococcus xanthus and Escherichia coli (Dhundale et al., 1987; Inouye et al., 1989), and in hepadnaviruses (Summers and Mason, 1982).This review focuses on the properties and inhibition of early events in HIV-1 reverse transcription.
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II. OVERVIEW OF HUMAN IMMUNODEFICIENCY VIRUSTYPE 1REPLICATION
A . Initial Events in HIV-1 Replication The outer envelope glycoprotein (gp120) of HIV-1 specifically interacts with the extrinsic CD4 protein on the host cell plasma membrane (Maddon et al., 1986; McDougal et al., 1986). The CD4 receptor is expressed on macrophages, monocytes, and a subset of T lymphocytes (Thelper cells), but the V3 domain of gp120 may control the tropism of viral entry into the latter cells (Hwang et al., 1991). On CD4-gp120 binding, the viral and cellular membranes fuse in a pH-independent manner (Stein et al., 1987). This fusion is thought to be facilitated by the viral gp41 transmembrane protein, which has a hydrophobic amino-terminal domain that bears a high degree of amino acid sequence similarity to the amino termini of the fusion and hemagglutinin proteins of paramyxoviruses and orthomyxoviruses, respectively (Bosh1 et al., 1989). Virus-host cell membrane fusion or endocytosis permits HIV-1 core entry into the cytoplasm. The fate of the viral core remains uncertain, but only partial dissolution is required for deoxynucleotide 5’-triphosphate (dNTP) entry and initiation of reverse transcription (Zhang et al., 1993). In HIV-1 reverse transcription, proviral double-stranded DNA (dsDNA) is synthesized from the ( +) RNA genome by the viral RT enzyme (Di Marzo Veronese et al., 1986; reviewed by Skalka and Goff, 1993). As will be described, reverse transcription may be initiated in virions prior to host cell entry (Arts et aZ., 1994a; Lori et al., 1992; Trono, 1992; H. Zhang et al., 1993, 1994),but more recent results suggest that this virion DNA may not be required for infection. In addition, completion of HIV-1 reverse transcription in a quiescent CD4+ lymphocyte may require cell activation (Zack et al., 1990). This stall in HIV-1 reverse transcription during infection of a quiescent host cell may be an initial step leading to latent viral infection. The 5’ ends of both strands in double-stranded proviral DNA are subjected to endonucleolytic dinucleotide cleavage by HIV-1 integrase in the nucleus or during transport of the viral DNA to the nucleus (Bushman et al., 1990; Engelman et al., 1991).Integrase, possibly associated with a nucleoprotein complex (HIV-1 nucleocapsid and matrix proteins) (Bukrinsky et al., 1993131, remains bound to the ends of proviral DNA, thus permitting a nucleophilic attack at a single, nonspecific site on host genomic DNA (Engelman et aZ., 1991).Integration of proviral DNA of nearly all retroviruses is only possible during mitosis of a cycling cell (Lewis et al., 1992; Roe et al., 1993; Peters et al., 1977).
/
I I I I
I
I I I
C
I I I
I I I I
Mutation
Frameshift
I I
I
G-C A 4-C-G 4-G-C C-G GGGAAACGGAGUG GAACAAACGGAGAA 3'
~~-LWUGUC
SL
+
H
+
As
+
AB
-
I 1592
FIG.5. Proposed frameshift site in BWYV RNA. The slippery heptanucleotide is underlined, and the pseudoknot is indicated by the joining of the loop structure with the downstream sequence. The diagram shows the deletion mutations AB and A S made by BglII and Sac11 digestion respectively. Point mutations (Hor SL) ae indicated by arrows. The effects of the mutations on frameshift during translation of transcript RNA in wheat germ extracts is shown on the right-hand side.
GGGAAAC sequence (Garcia et al., 1993), it also occurs elsewhere between the downstream Sac11 and BglII sites (Fig. 5). This region contains two GGGAAAG sequences and one CCCAAAG. In summary, there is good evidence for in uiuo ribosomal frameshifting, but the mechanism by which it happens is far from being understood. In uitro experiments with mutagenized sequences on and around the slippery site have yielded somewhat discrepant results. Probably it will only be from studies on full-length transcripts that a full analysis of the sequences involved in this mechanism will be possible. Indeed, it has been shown that, in cells infected with full-length PAV cDNA, there is proportionately more of the protein resulting from frameshifting than is found in in vitro translation experiments or when truncated cDNA is expressed in uitro or in uiuo (C. P. Paul and W. A. Miller, 1994, personal communication).
B . Internal Initiation of Translation Most eukaryotic mRNAs are monocistronic, and translation initiation occurs according to the scanning model (Kozak, 1989). The 40s
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ribosomal subunit binds to the cap structure at the 5’ end of the mRNA and then scans in the 3’ direction until the first AUG is reached. The 60s ribosomal subunit then binds to it, and protein synthesis begins. However, initiation efficiency can be modulated by the nucleotide context around the AUG, the most favorable being an A at position -3 and a G at +4 (Cavener and Ray, 1991; Kozak, 1989; Lutcke et al., 1987). If the first AUG is in an unfavorable context, some of the scanning ribosomes do not initiate until the second AUG if its context is more favorable. This leaky scanning mechanism (Kozak, 1989) can be utilized by bicistronic mRNAs like those of several animal viruses (see Samuel, 1989, for a review). In uitro translation of PLRV RNA or BWYV RNA in wheat germ extracts resulted in the synthesis of two major products with M, of 28,000 and 70,000 (PLRV; Mayo et al., 1989) or 25,000 and 66,000 (BWYV; Veidt et al., 1992). The apparent M,values correspond to the predicted sizes of the proteins encoded by the respective ORF 0 and ORF 1, indicating that initiation for ORF 1 occurs at an internal AUG. In reticulocyte lysate, the shorter protein encoded by ORF 0 was poorly expressed (Mayo et al., 1982, 1989; Veidt et al., 1992). A third minor polypeptide of apparent M,. 125,000 (PLRV) or 100,000 (BWYV) was also detected, which could represent the fusion protein resulting from frameshift from ORF 1 to ORF 2 (see Section IV, A). The same approach of translating transcript RNA from cloned cDNA has been used to investigate the expression of ORF 3 and ORF 4 of BWYV. The translation products included coat protein (identified by immunoprecipitation) and a protein of apparent M , 22,000 (which was not immunoprecipitated) that could be the translation product of ORF 4 (Veidt et al., 1988). In similar experiments with a transcript of PAV cDNA, Dinesh-Kumar et al. (1992) showed that the coat protein and the M , 17,000 polypeptide encoded by ORF 4 are produced in uitro from a single mRNA, by initiation at the first two AUG codons. The ratio between the amounts of coat protein and P4 produced varied from 1:l to 1:7 depending on the salt concentrations used in the translation. To determine the relative expression of ORF 3 and ORF 4 of PLRV in uiuo, the GUS gene was translationally fused to a cDNA fragment containing one or the other initiation codon. The resulting constructs expressed a GUS protein with an N-terminal part of either 29 PLRV amino acids (translation initiation at the coat protein AUG) or 20 PLRV amino acids (translation initiation at the P4 AUG) (Tacke et al., 1990). Tobacco or potato protoplasts were then electroporated with the construct and GUS activity in the protoplasts measured. Initiation at the internal AUG of ORF 4 was very efficient in this system. The amount of P4 synthesized exceeded by sevenfold that of the coat protein. When a similar approach was used for PAV ORF 3 and ORF 4, the
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M. A. MAY0 A N D V. ZIEGLERGRAFF
ratio of initiation of P4 synthesis to that of coat protein was about 2 (Dinesh-Kumar and Miller, 1993).Thus, although the ratios differed with both PLRV and PAV, initiation at the second AUG was more common than at the first AUG. This may be because the AUG of ORF 4 is flanked by a more favorable context for initiation than is the coat protein initiation codon. Except for SDV and BMYV, all luteoviruses have a better initiation context around the ORF 4 AUG than around the ORF 3 AUG. The same is true for the initiation codons of ORF 0 and ORF 1. Dinesh-Kumar et al. (1992)reported that a secondary structure could be formed by the leader sequence of PAV subgenomic RNA, which would locate the coat protein AUG in a stem-loop structure, possibly making it less accessible to ribosomes. By site-directed mutagenesis, Dinesh-Kumar and Miller (1993)showed that improving the context of the coat protein AUG resulted in increased expression of ORF 3,compared to that of wild-type ORF 3 both in uitro and in uiuo. Destabilization of the secondary structure increased simultaneous expression of both ORFs, irrespective of the sequence context. Unexpectedly, for a given coat protein AUG context, changes that decreased initiation at the downstream AUG also reduced initiation at the first codon. Therefore, they proposed a new model in which pausing of the ribosomes at the second AUG enhances initiation at the upstream AUG codon. The 80s ribosomes formed at the P4 AUG are proposed to melt some base pairing upstream of this AUG, and therefore cause stacking of the upstream scanning 40s subunit, leaving it time to interact with the coat protein initiation codon and commence translation (Dinesh-Kumar and Miller, 1993).
C . Subgenomic mRNA Synthesis As with many RNA viruses, the ORFs in luteovirus RNA located in the 3’half of the genome are translated from subgenomic messenger RNAs. A major subgenomic species of about 2.6 to 2.9 kb was detected in cells infected with PAV (Dinesh-Kumar et al., 1992),PLRV (Smith and Harris, 1990;Tacke et al., 1990;Miller and Mayo, 1991),BWYV (Falk et al., 1989;Veidt et al., 1992)or CABYV (Guilley et al., 1994). These subgenomic RNAs allow the expression of the gene cluster of ORFs 3,4,and 5 (Tacke et al., 1990;Dinesh-Kumar et al., 1992). A second species of 0.8 kb has been detected in PAV-infected plants (Dinesh-Kumar et al., 1992)which could account for the expression of ORF 6. An extra small RNA of 0.7 kb was found in BWYV-infected plants (Falk et al., 1989), and one of 0.3 kb was detected in plants infected with PAV (Kelly et al., 1993).None of these RNAs was encap-
MOLECULAR BIOLOGY OF LUTEOVIRUSES
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sidated (Smith and Harris, 1990; Miller and Mayo, 1991; DineshKumar et al., 1992). The positions in the genome RNA of the 5’ ends of the larger subgenomic RNAs of several luteoviruses have been mapped precisely (PLRV, Miller and Mayo, 1991;CABYV, Guilley et al., 1994;BWYV, V. Ziegler-Graff, 1994,unpublished observations). The leader sequences are unusually long (212nucleotides for PLRV and CABYV, 224 nucleotides for BWYV) and the 5‘ ends are located 12,8,or 19 nucleotides upstream of the termination codon of ORF 2. The 5’4erminal sequences of the subgenomic RNA are identical to those of the respective 5’4erminal sequences of the genomic RNA (Fig. 6).Moreover, the first eight nucleotides of the subgenomic and genomic RNA of BWYV, PLRV, and CABYV are identical, which may reflect a conserved replicase recognition signal in the minus-strand RNA. Comparisons among the intergenic regions of different luteovirus RNAs indicate the presence of two conserved regions between nucleotides -102 to -91 and -46 to -24 (Mayo et al., 1989;Vincent et al., 1991).They are particularly A-U-rich and could represent the “core region” of the subgenomic promoter as defined for brome mosaic virus RNA by Marsh et al. (1987).This region may therefore be involved in the synthesis and CABYV
BWYV
PLRV
C A G G A G A A A U U G - N,,
--
FIG.6. The 5’-terminal sequence similarities between the genomic and subgenomic RNAs of CABYV, BWYV, and PLRV. For each virus, the genomic sequence [(l) indicates the 5’-nucleotidel is aligned against the place in the genomic RNA where the subgenomic RNA commences. Vertical lines show the sequence matches between the termini of genomic and subgenomic RNA, the box indicates sequences common to the three viruses. Open boxes indicate the 3’ extremity of ORF 2 (P2) or the 5’ extremities of ORFs 0 or 3 (PO, P3); N, signifies intervening sequence.
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M. A. MAY0 AND V. ZIEGLERGRAFF
regulation of the larger subgenomic RNA of these luteoviruses. The 5‘ end of the subgenomic RNA of a German isolate of PLRV has been mapped to 40 nucleotides upstream the coat protein AUG codon (Prufer et al., 1992). In contrast to these subgroup I1 viruses, the 5’ end of the subgenomic RNA of PAV is 89 nucleotides upstream of the coat protein initiation codon (Dinesh-Kumar et al., 1992). In this case, the initiation site for the synthesis of the subgenomic RNA is different from the 5’ proximal sequence of the genomic RNA. However, with PAV from Australia, L. Kelly (1994, personal communication) found the subgenomic leader sequence to be 188 nucleotides long, which would mean that, as for subgroup 11 viruses, the 5’ end of the subgenomic RNA is in ORF 2.
D . Readthrough or Leaky Translational Termination Readthrough occurs when a stop codon is suppressed by binding a suppressor tRNA. The action of a suppressor tRNA has been demonstrated in the readthrough of termination codons in RNAs of tobacco mosaic virus (Beier et al., 1984) and tobacco rattle virus (Zerfass and Beier, 1992). The result is a relative abundance of the smaller product and a small proportion of a larger fusion protein that includes the translation product of the next in-frame ORF. In luteoviruses, ORF 5 is expressed as an ORF 3 + ORF 5 fusion protein by translational readthrough of the coat protein UAG termination codon. The sequence context surrounding the leaky stop codon is identical for all luteoviruses: AAAUAGGUAGAC (termination codon in bold type). In the RNA of tobacco mosaic virus (TMV),the two codons downstream of the suppressible UAG codon at the end of the ORF encoding the M, 126,000 protein have been shown to form part of the signal promoting readthrough (Skuzeski et al., 1991). Although the sequence context of the luteovirus leaky stop codon is different from that in TMV RNA (CCAUAGCAAUUA),it is likely that the conserved sequence context in which the UAG is embedded in luteovirus RNAs is similarly important in conferring leakiness. The readthrough protein has been detected in potato plants and tobacco protoplasts infected with PLRV (Bahner et al., 1990) and in C. quinoa protoplasts infected with BWYV (Reutenauer et al., 1993) by using antibodies raised against P5 (see Section II1,F for more details). Readthrough protein has also been detected in preparations of purified particles of PLRV (Bahner et al., 1990),PAV (Waterhouse et al., 19891, RPV (Vincent et al., 1991), and BWYV (see Section V,O. In in uitro translation experiments, the efficiency with which the ORF 3 UAG in PAV RNA was suppressed by readthrough was reported
MOLECULAR BIOLOGY OF LUTEOVIRUSES
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to vary from 7 to 15% depending on the salt concentration (DineshKumar et al., 1992).The frequency of in uiuo suppression of the UAG stop codon separating the PLRV ORFs 3 and 5 was determined in a transient expression system using a clone in which the 18 nucleotides upstream and 21 nucleotides downstream of the stop codon were translationally fused to the GUS gene. When GUS activity was measured in tobacco and potato protoplasts electroporated with this construct, the efficiency of suppression was about 1% (Tacke et al., 1990). Readthrough was about 5% when the stop codon was in the context of the leaky stop codon in TMV RNA (Skuzeski et al., 1991). Miller et al. (1995)reported that they were unable to observe suppression of the PAV RNA stop codon using the same approach, but they could detect readthrough in uiuo when the construct was a full-length clone of PAV in which the GUS gene was inserted downstream of the coat protein stop codon. Miller et aZ. (1995)suggested therefore that readthrough in PAV RNA requires a viral or virus-induced transacting factorh) or distant cis-acting signals not present in the initial construct. A sequence rich in C residues consisting of 16 uninterrupted CCxxxx repeats located downstream of the coat protein stop codon has been shown to be involved in efficient suppression in protoplasts infected with PAV transcript RNA; 10 such repeats were sufficient for suppression, but elimination of them all resulted in no suppression (S. P. Dinesh-Kumar and W.A. Miller, 1994,personal communication). A similar pattern of from 7 to 16 such repeats is present in the sequences of RNA of all luteoviruses (Miller et al., 1995). The amino acid sequence encoded by this sequence is not involved in the mechanism because the introduction of a frameshifting mutation between the stop codon and the repeats in PAV RNA had no effect on the extent of readthrough (Miller et al., 1995).
E . Proteolysis and Cap-Independent Translation
A protease consensus has been found in P1 proteins encoded by subgroup I1 virus RNA (Koonin and Dolja, 1993;see also Section II1,C). However, no direct evidence has been reported so far for proteolysis of luteovirus proteins. Nonetheless, proteolytic processing of luteovirus proteins, at least for PLRV and RPV, is implied because the VPg does not correspond to the translation product of an ORF. It has been suggested that part of the P1 forms the VPg (Koonin and Dolja, 1993).By analogy with other viruses which have a VPg, it is predicted that the processing to give the VPg is by a virus-coded protease. Miller et al. (1995)reported that a 500-nucleotide sequence located in the 3' end of PAV RNA was required for efficient translation of
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M. A. MAY0 A N D V. ZIEGLER-GRAFF
uncapped transcripts of PAV cDNA in wheat germ extracts. Deletion of portions of this sequence (from nucleotide 4513 to 5009)resulted in a marked reduction in translation to yield P1.Capping of these deleted transcripts restored full translational activity. These results were not observed in rabbit reticulocyte lysate. Miller et al. (1995)suggested that this cap-independent translation mechanism could be an adaptation of PAV to multiplication in cereals. V. PARTICLE STRUCTURE
A. Possible Tertiary Structure The secondary structure of luteovirus coat proteins was predicted by Dolja and Koonin (1991)on the basis of a multiple sequence alignment with coat proteins of known secondary structure. This model was used by Torrance (1992)to interpret the location of epitopes in PLRV particles (see below). There has been an improvement in the reliability of secondary structure predictions (Rost and Sander, 1993,19941,and the sequences shown in Fig. 2 were submitted to this procedure. The program HSSP/MaxHom was used to align the sequences and then predict the secondary structure of the aligned proteins. The predicted regions of p- sheet and a-helix are shown in Fig. 2.The p sheets are labeled to correspond with the letter coding used for SBMV and tomato bushy stunt virus (TBSV) (Rossmann and Johnson, 1989);the long continuous p sheet near the C terminus is assumed to correspond to p sheets H and I (Fig. 2).As a control, the coat protein of SBMV was assessed, and the program predicted all of the p sheets known to be present (AbadZapatero et al., 1980)except that the edges of the p-sheet regions were all underestimated, suggesting that lengths of p sheets in luteovirus coat proteins shown in Fig. 2 are also underestimates. Assuming that p sheets B to I are arranged in a shell domain, as they are in the coat proteins of many diverse viruses (Rossmann and Johnson, 1989),it is possible to predict the parts of the proteins which would correspond with the parts of SBMV protein known to be exposed on the exterior of the virus particles (Hermodson et al.,1982).These are also indicated on Fig. 2. The N-terminal 58 to 69 amino acids of luteovirus coat proteins to the N-terminal side of p sheet B (Fig. 2) resemble this region (R domain) of many coat proteins of viruses with isometric particles in being highly basic (Rossmann and Johnson, 1989). They contain between 19 and 21 R or K residues separated by relatively nonpolar residues such as G and N, and no acidic residues. By analogy with the
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coat proteins of SBMV or TBSV, this region is predicted to be internal, and presumably the basic residues neutralize the negative charge on the virus RNA.
B . Location of Epitopes Inspection of multiple sequence alignments, such as that in Fig. 2, suggest interpretations for the observation that epitopes are often shared between some, but not all, luteoviruses. (e.g., D’Arcy et al., 1989). For example, an epitope common t o BMYV, BWYV, and RPV particles but not on those of PLRV, PAV, MAV, or GRAV could be in the tripeptide LAG or the hexapeptide STINKF. An epitope shared by BLRV, BWYV, and RPV particles but not those of PLRV or SDV could be in INKF. Greater precision is possible with some combinations such as the possibility that an epitope shared by PAV and BMYV but not by BWYV is created by substituting a n M for an I, a T for a n A, or SA for NS or SS. The positions of these possible epitopes are indicated in the BWYV sequence in Fig. 2. The last possibility, like the other possible sites involving INKF, is in the putative E-F loop which should be on the surface of the virus particle. A less conjectural approach is to assess the reactions of the antibodies to each of a set of overlapping peptides that represent the entire protein sequence. By this method, Torrance (1992) has identified 11 epitopes in the coat protein of PLRV and has mapped them to the positions shown in Fig. 2. The major epitope was found to comprise the amino acids at the extreme N terminus. Antibodies to this epitope reacted with intact virus particles, which suggests that the epitope is external although the location of the N-terminal arms of the coat proteins of SBMV and TBSV are known to be internal (Rossmann and Johnson, 1989). Torrance (1992) suggested that the N-terminal amino acids of PLRV coat protein are exposed at the particle surface when particles swell because of changes in pH or ionic conditions.
C . Presence of Readthrough Protein (P5) As discussed in Section III,F, the readthrough protein is expressed as a fusion protein attached to the C terminus of the coat protein, and a few percent of the coat protein molecules produced have this extension. These molecules form part of the virus particles, and P5 has been detected in purified particles of PAV (Waterhouse et al., 19891, PLRV (Bahner et al., 1990), RPV (Vincent et al., 1991), and BWYV (Reutenauer, 1994) (Fig. 7). With both PLRV and RPV there is evidence for P5 being readily degraded after isolation from infected tissue. Bahner
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M. A. MAY0 AND V. ZEGLERGRAFF
FIG.7. Expression of P5 of PLRV and BWYV. Samples were protein from healthy protoplasts (A, lanes 1 and 4); PLRV-infected protoplasts (A, lanes 2 and 5); purified particles of PLRV (A, lanes 3, 6 and 7); purified particles of BWYV (B, lanes 1 and 2); leaf tissue of Nicotiana cleuelandii infected with BWYV by agroinfection (B, lane 3); or leaves of mock-inoculated N. cleuelandii (B, lane 4). Protein extracts were subjected to electrophoresis and then either staining with Coomassie blue (A, lane 7)or silver nitrate (B, lane 1)or transfer by electroblotting to nitrocellulose. Blots were reacted with mouse monoclonal antibodies to PLRV particles (A, lanes 1, 2, and 3) or rabbit polyclonal antibody to PLRV P5 (A, lanes 4 and 5), to BWYV particles (B, lanes 2, 3, and 4, lower part), or BWYV P5 (B, lanes 2,3, and 4, upper part). Antibody reaction was detected by conjugated anti-mouse or anti-rabbit antibodies. RT indicates the position of the P3 + P5 readthrough protein, RT*indicates the position of the partial degradation product of P3 + P5, and CP indicates the position of the coat protein (P3).
et al. (1990)found that whereas the fusion protein of P3
+ P5 had an
M, of about 80,000 when extracted from infected cells or particles rapidly sedimented from infected protoplasts, the C-terminal half of the P5 part of this fusion protein was rapidly degraded when infected cells were disrupted and its size in particles of purified PLRV was about M, 53,000. The fusion protein in particles of RPV had an apparent M, of 63,000 (theoretical size 66,000) but was often degraded to M, 58,000 and sometimes was undetectable (Vincent et al., 1991).It has been suggested that the readthrough domain is subject to partial proteolytic degradation in the course of virus purification (Bahner et al., 1990;Vincent et al., 1991). Detection of the truncated readthrough protein in purified virus particles of a frameshift mutant of BWYV
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indicated that the site of this cleavage was near the C end of the luteovirus conserved domain (see Fig. 4),which suggests that there is a common site for this cleavage in different luteoviruses (V. Brault, 1994,personal communication). The C termini of the coat proteins of SBMV and TBSV are at the surface of the virus particles, and the correspondence in apparent structure of luteoviruses with these viruses (Dolja and Koonin, 1991; Section V,A) makes it likely to be so in luteovirus particles. Thus, the readthrough protein is predicted to protrude on the outside of the particles. This model of luteovirus particles consisting of M, 23,000coat protein molecules and a few much larger proteins of coat plus readthrough protein might explain the occasional protuberances seen on PLRV particles in some electron micrographs (Harrison, 1984). In more recent experiments, it was possible t o demonstrate specific labeling of BWYV particles with antibodies raised against a fusion protein containing part of P5 (J. F. J. M. Van den Heuvel, 1994,personal communication).
D . Heterologous Encapsidation It has been known for some time that aphids can transmit a luteovirus which they do not normally transmit if the virus is acquired from plants also infected with a transmissible luteovirus. For example, Myzus persicae was able to transmit carrot red leaf virus (CRLV)when feeding on plants doubly infected by CRLV and PLRV (Waterhouse and Murant, 1983).Similar results were obtained with mixtures of strains of BYDV (Rochow, 1970b;H u et al., 1988).Work in which virus particles were trapped by reaction with specific antibodies and the RNA molecules contained in the particles were characterized by reaction with specific nucleotide sequence probes has demonstrated two types of heterologous encapsidations (Creamer and Falk, 1990;Wen and Lister, 1991). These are transcapsidation or genomic masking, where particles are formed from the protein of one virus and the RNA of another virus, and phenotypic mixing, where the coat protein shell contains proteins from more than one virus. Transcapsidation was demonstrated between RPV and either PAV or MAV and between MAV and the RMV strain of BYDV (RMV) (Creamer and Falk, 1990), whereas phenotypic mixing occurred with PAV and MAV and with RPV and RMV (Wen and Lister, 1991).Thus, although PAV and MAV coat proteins differ in 54 amino acids, of which 31 are in the shell domain (Section V,A), they can coassemble. Transcapsidation also suggests a lack of specificity in the interaction of virus RNA with coat protein. A more extreme example of this is
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the association between luteoviruses and several other viruses to form transmission complexes (Waterhouse et al., 1988; Murant, 1993). The dependent viruses lack coat proteins and thus depend on the luteovirus for RNA protection and a means of being transmitted by vector aphids. They have been classified in the genus Umbravirus (Murant et al., 1995). Other examples are the symbiotic association of the two RNAs comprising the genome of PEMV and the dependency of ST9-associated RNA on the ST9 strain of BWYV (see Section VII1,D).
E . Determinants for Particle Assembly Purified preparations of some viruses with isometric particles characteristically contain RNA-free shells, often called top component. Particle assembly of such viruses presumably does not require RNA. Only with one isolate of BWYV has a top component been found in preparations of luteovirus particles (Hewings and D’Arcy, 1986). Some slowly sedimenting PLRV protein structures were found in PLRVinfected tobacco protoplasts 2 days after inoculation, but no virus-like particles could be detected (Miller, 1992). Work with mutants of BWYV has shown that the readthrough protein contributes little or nothing to the assembly of virus particle (Reutenauer et al., 1993).Virus particles were made in protoplasts infected with mutants lacking almost all of the readthrough protein (see Section 111,F). More recent work in which PLRV coat protein has been expressed in heterologous systems may lead to a system for determining the factors needed for PLRV particle assembly (J. Lamb, 1994, personal communication). When Spodoptera frugiperda cells were infected with a recombinant baculovirus that contained cDNA encoding PLRV coat protein under the control of the polyhedrin promoter, they expressed large amounts of P3. The PLRV protein accumulated in the nuclei of the insect cells apparently in amorphous aggregates. When the inserted cDNA also contained sequence encoding a histidine tag at the N terminus of the coat protein, the resultant modified coat protein also accumulated in nuclei, but it formed viruslike particles that aggregated into crystallike structures. The cDNA did not encode P5, and the result therefore suggests that P5 is not needed for the assembly of particles. However, it is not known what effect the amino acid extension of the N terminus has; it may be mimicking the RNA component of PLRV particles, or it may mimic the role played by P5, even though P5 is a C-terminal extension of P3 rather than an N-terminal extension.
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VI. LOCATION OF LUTEOVIRUS REPLICATION
A . Limitation to Phloem Tissue Microscopic examination of tissues infected with luteoviruses shows that virus accumulates almost exclusively in the sieve elements and companion cells of the phloem and occasionally in phloem parenchyma. Typically, not all phloem cells contain virus particles, and movement vertically in infected plants is probably relatively rapid whereas horizontal movement is slow and inefficient (Waterhouse et al., 1988). Restriction of luteovirus multiplication to these cells is not because luteoviruses cannot replicate in other cell types, as luteoviruses have been shown to multiply in isolated mesophyll protoplasts and in a few cells outside the vascular tissue. Replication has been demonstrated in mesophyll protoplasts inoculated with PAV (Barnett et al., 1981; Dinesh-Kumar et al., 1992),PLRV (Kubo and Takanami, 19791,tobacco necrotic dwarf virus (TNDV) (Kubo, 1981), BWYV (Veidt et al., 19921, and RPV (Silver et al., 1994). TNDV was shown to multiply in inoculated epidermal cells (Imaizumi and Kubo, 1980), BYDV was sometimes found in xylem parenchyma cells (Gill and Chong, 1981), and in N . cleuelandii plants infected with PLRV a few mesophyll cells became infected (Barker, 1987). Moreover, double infection of N . cleuelandii with PLRV and potato virus Y (PVY) resulted in about a sevenfold increase in the proportion of the mesophyll cells that contained PLRV. This effect was also found in plants infected with PLRV and potexviruses, tobraviruses, or carrot mottle virus (CMoV) (type species of the genus Umbravirus) but not in plants infected with a variety of other viruses including PEMV (Barker, 1989). A similar effect was detected in plants infected with BWYV (Barker, 1989). An interesting parallel occurs with plants infected with PEMV. The luteovirus-like RNA-1 of PEMV (see Section VII1,C) multiplies in infected protoplasts but does not multiply when inoculated into plants unless these are also inoculated with RNA-2. In plants inoculated with RNA-2 alone it can spread systemically, but no virus particles are formed (Demler et al., 1994). In this way, PEMV RNA-2 resembles umbraviruses such as CMoV. It seems likely that the CMoV or PVY in the plants doubly infected with PLRV provide a factor which moderately enhances the very restricted movement of the PLRV in nonphloem cells; the RNA 2 of PEMV provides a fully functional movement protein that assists or permits PEMV to spread systemically. A different coinfection occurs with the ST9 strain of BWYV. When plants are infected with BWYV-ST9, the presence of a smaller, ST9-
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associated RNA (see Section VII1,D) enhances the amount of BWYV accumulating in infected tissue (Passmore et al., 1993). However, despite this extra accumulation, BWYV could not be detected outside the phloem tissue (Sanger et al., 1994). A distinction between BWYV-ST9 and the PLRV-CMoV system is that CMoV spreads systemically in infected plants, whereas, although the ST9-associated RNA can infect inoculated leaves, it does not spread systemically (Passmore et al., 1993).
B . Cytopathological Effects The effects that infection has on cell ultrastructure differ according to which subgroup the virus belongs (Gill and Chong, 1979). Infection of oat plants with MAV or PAV (subgroup I) resulted in the formation of single membrane-bound vesicles and dense filaments that seemed to accumulate in the nuclei. Virus particles accumulated in the cytoplasm, and late in infection the nuclei became distorted, and deteriorated. In contrast, infection of the same host species with RPV (subgroup 11)resulted in the formation of double membrane-bound vesicles and tubules. Virus particles accumulated in the nuclei, which remained intact (Gill and Chong, 1979). The effects of other subgroup I1 genome viruses were similar, although no PLRV particles were detected in nuclei (Shepardson et al., 1980). Infection of isolated protoplasts with PEMV RNA-1 induced the formation of vesicles, and virus particles were detected within the nuclei as well as in the cytoplasm (Demler et al., 1994). As discussed above, the expression in insect cells of the coat protein gene of PLRV in a recombinant baculovirus resulted in the accumulation of the coat protein in the nuclei (J.W. Lamb, G. H. Duncan, M. A. Mayo, and R. T. Hay, 1994, unpublished results). When the gene was modified to produce a tagged protein, the modified protein formed into viruslike particles. The coat protein of PLRV and most other luteoviruses contain highly basic sequences near the N termini that resemble nuclear location signals (Garcia-Bustos et al., 1991), but the sequence is probably on the inside of intact particles where it would normally interact with virus RNA (see Section V,A). These observations reinforce the suggestion made by Esau and Hoefert (1972) for BWYV that virus particles assemble in the nucleus. VII. PHYTOPATHOLOGY
A. Diagnosis of Luteovirus Infection Diagnosis of virus infection can be of two types. Either a broad specificity detection is required so that even viruses only marginally
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related to the reference standard are detected (e.g., as with polyclonal antiserum), or a highly specific detection is required in order to discriminate the virus from close relatives (e.g., as with monoclonal antibodies). The accumulation of cloned cDNA to luteovirus RNA and knowledge of the nucleotide sequences of several luteoviruses and their strains has led to the development of nucleic acid hybridizationbased probes for each of the types of diagnostic tests. Not surprisingly in the light of the data in Table 11, probes made from cDNA to different parts of the genome were either highly specific or reacted to some extent with RNA from other luteoviruses (e.g., Waterhouse et ul., 1986). Probes complementary to the coat protein gene of PLRV (Robinson and Romero, 1991) or BWYV (Herrbach et al., 1991) have proved to be the least specific. Probes to the coat protein gene of PLRV detected BWYV and RPV readily, GRAV and CRLV weakly, but did not detect PAV or MAV (Robinson and Romero, 1991); probes to the coat protein gene of BWYV detected BMYV and PLRV readily, RPV and PAV weakly, but did not detect MAV (Herrbach et al., 1991). These results do not correlate well with the amount of amino acid sequence identity between the coat proteins of the various luteoviruses (Table 11). A potentially powerful method for diagnosing luteovirus infection is the use of reverse transcription followed by polymerase chain reaction (RT/PCR) using primers designed to hybridize to all luteovirus RNAs. Robertson et al. (1991) described such a pair of primers, one of which is partially degenerate, that hybridized with RNA and cDNA of PLRV, BWYV, and PAV. The amplified product represents the 3’-terminal464 nucleotides of the coat protein gene. Detection by this method is potentially highly discriminatory, as sequence analysis of the PCR product would show how similar the detected luteovirus was to known viruses. However, the greater number of sequences available now show that, at least for some luteoviruses, the primers are unlikely to work because of mismatching. The downstream primer, which includes the termination codon, hybridizes well, but the upstream primer is a poor match to SDV RNA and also may not hybridize with BLRV RNA. A possible new upstream “universal luteovirus primer” was derived from an alignment of the coat protein genes of BLRV, BWYV, CABYV, MAV, PAV, PLRV, RPV, and SDV and should yield a PCR product corresponding to the 3’-terminal 319 nucleotides of the coat protein gene (M. A. Mayo and C. A. Jolly, 1994, unpublished results). Detection can be made more sensitive and more specific, without recourse to sequencing, by using immune-capture PCR in which virus particles are bound to a plate coated with a specific antibody and then RNA is extracted from the bound virus and subjected to RT/PCR. The method has been used for detecting potyvirus particles (Wetzel et al.,
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1992) and is also effective for PLRV; the sensitivity obtained was sufficient to detect the PLRV particles circulating in a single aphid (C. A. Jolly, 1994, personal communication).
B . Resistance to Luteoviruses For many viruses, it has been shown that plants are more resistant to virus infection and/or multiplication when they have been transformed such that they express the coat protein gene of the target virus (Reavy and Mayo, 1992). Application of transformation methods to produce plants resistant to luteoviruses has been done with PLRV. DNA encoding PLRV coat protein (and thereby also P4) has been inserted into the genomes of potato plants (Kawchuk et al., 1990,1991; Van der Wilk et al., 1991; Barker et al., 1992) and tobacco plants (Barker et al., 1993). The transformants were often resistant to infection by the feeding activity of viruliferous aphids (Kawchuk et al., 1990) and/or showed a restriction in the amount of virus accumulating after the primary infection of potato (Kawchuk et al., 1991) or tobacco (Barker et al., 1993). Transgenic potato plants were also resistant to secondary infection (Barker et al., 1992). The resistance seemed to be independent of the production of coat protein, as little or no protein could be detected in many transgenic lines that were nonetheless resistant to PLRV. Resistance to infection was not always particularly strong, but multiplication resistance resulting in virus accumulating to about 10 to 20% of the amount in control plants is comparable to the resistance being used in current breeding programs. Antisense constructs were also effective (Kawchuk et al., 1991; Van der Wilk et al., 19911, which reinforces the view that the resistance is caused by the production of PLRV RNA sequences (or their complement) rather than coat protein. Multiplication resistance was shown t o be related to the amount of transcript synthesized, but not in a strict correlation (Barker et al., 1992, 1993). Derrick and Barker (1992) showed that in potato resistant to PLRV, either because of transformation with the PLRV coat protein gene or because of the action of a host gene(s), only the adaxial phloem was infected and fewer cells were infected than in control plants. In experiments with the same transgene, it was shown that the effects of the transgene and that of the host resistance gene were additive; transformation of relatively resistant genotypes further enhanced their resistance to levels approaching that of extremely resistant wild Solanurn species (Barker et al., 1994). The similarity in phenotypic effect of the PLRV coat protein gene and host resistance genes may be coincidence, but if not, it raises the intriguing prospect that the two types of resis-
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tance are induced by the same or similar mechanisms (H. Barker, 1994, personal communication). Plants expressing a mutated form of P4 were found to be resistant to virus spread, and it was suggested that the transgene product had interferred with the transport role of the P4 made during the course of an infection (Tacke et al., 1993b). VIII. TAXONOMY
A . Speciation The available sequences provide sufficient data to pose the question of when strains of a luteovirus should be considered different viruses. BYDV isolates have been placed in two subgroups on the basis of serological relationships, cytopathological effects, and the doublestranded RNA formed in infected tissue (Rochow 1970a; Waterhouse et al., 1988; Ueng et al.,1992). Strains MAV and PAV were placed in one subgroup, and RPV was placed in the other subgroup. The molecular evidence therefore reinforces the biological evidence in arguing for considering BYDV to consist of two species; the corollary is that one should be renamed. However, the situation is more complex when sequences of individual genes are compared. If P1 or P2 are considered, then PAV and MAV should be considered as strains because they are 98% identical; if genes in the 3’ coding block are compared then values of 73% (P3), 72% (P4), and 72% (P5) identity (Table 11) suggest dissimilarity approaching that thought of for other virus groups such as potyviruses (Shukla and Ward, 1989) to indicate that the viruses belong to different species. Although MAV and PAV could perhaps be considered as strains of one virus because of their similar host ranges and ability to cross-protect against one another (Wen et al., 1991), this seems less reasonable for CABYV and GRAV which, although they have coat proteins that are 75% identical (Table II), are transmitted by different aphid species and infect different hosts. In contrast, with the four strains of PLRV sequenced the minimum similarity was 88%between P1 of Australian PLRV and that of Canadian or Dutch PLRV. In comparisons among isolates obtained in Scotland, the sequences of P3 and P5 were 96 to 99% identical (Jolly, 1994). Thus the variation among isolates from diverse countries (Table 11) was similar to that obtained in Scotland. PLRV appears to vary very little. In a larger study, J. De Miranda and R. Hull (1994, personal communication) compared 37 isolates of BWYV-like viruses from Europe and
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Iran by sequencing RNA consisting of about 400 nucleotides of ORF 2, the intergenic region and ORF 3. The isolates could be clustered in different ways according to which regions of the sequences were considered. The ORF 3 of all isolates were 93% or more identical, the ORF 2 sequences fell into two groups with only 63% identity between the groups, but the intergenic sequences in one of these groups fell into two subgroups with only 68% identity between them. There have been a number of attempts to rationalize the classification of luteoviruses with some serological relationship to BWYV. BMYV, RPV, CRLV, and the RGV strain of BYDV have all been considered to be strains of BWYV (Casper, 1988). This lumping approach does not seem very useful for pathologists seeking to discriminate between pathogens and is not supported by the differences between the sequences of the coat proteins of BWYV and RPV (Table 11).Moreover, a hybridization probe corresponding in sequence to ORF 0 of BMYV, which is pathogenic for beet, did not cross-react with RNA of BWYV, which is not pathogenic for beet (0.Lemaire, 1994, personal communication).
B . Structure of Genus Luteovirus As discussed above, viruses in the genus Luteovirus have one of two genome arrangements, each typified by distinctive polymerase sequences (Fig. 1).The polymerases resemble those of either carmoviruses or sobemoviruses, which are considered to be very different types of RNA viruses (Dolja and Carrington, 1992; Gibbs, 1995). Thus, classification on the basis of the polymerase sequence would probably place viruses in each luteovirus subgroup in different families. But the biology of the viruses (phloem restriction, persistent aphid transmission) and the molecular features (suppression of the amber termination codon of the coat protein gene to yield a readthrough protein, encoding of P4 inside the P3 gene) are consistent with luteoviruses belonging to one genus. The pragmatic solution to this difficulty would seem to be to retain the grouping known at present as the genus Luteovirus but to consider each of the subgroups as genera within the larger grouping. This arrangement would keep biologically similar viruses together. The implication would be that building a phylogenetic tree of viruses from the sequences of polymerase domains does not necessarily lead to workable taxa. Indeed, the extent of detectable recombination within the genomes of luteoviruses should be a strong disincentive to the use of the polymerase gene to typify a virus.
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C . Problem of Pea Enation Mosaic Virus Pea enation mosaic virus resembles luteoviruses in being persistently transmitted by aphids but is distinct in being mechanically transmissible, being capable of invading mesophyll tissues in infected plants, and having a bipartite genome (Hull and Salquero, 1991). Molecular analysis of the PEMV genome has provided an explanation for these properties and has posed a taxonomic challenge (Demler et al., 1994).Figure 8 shows the arrangement of the ORFs in the two genome RNAs. The larger RNA of the genome (RNA-1)resembles that of subgroup I1 luteoviruses. It contains ORFs which are equivalent in position and in sequences of the encoded proteins to ORFs 0, 1 , 2 , 3 , and 5 of luteoviruses. Moreover, in the overlap between ORF 1 and ORF 2 there are slippery sequences, and ORF 2 is probably expressed by translational frameshift from the ORF 1 frame, just as for luteovirus subgroup I1 genomes. There is no ORF equivalent to ORF 4 of luteoviruses in PEMV RNA-1, and the ORF corresponding to ORF 5 is relatively short (Fig. 8).The ORF 2 region of RNA-1 encodes a putative polymerase that is similar to subgroup I1 luteovirus polymerases and the RNA can multiply in inoculated protoplasts independently of the smaller genome RNA (RNA-2)(Demler et al., 1994).ORF 3 encodes the coat protein, which is about 30 to 35%identical to those of luteoviruses, and virus particles accumulate in protoplasts inoculated with RNA-1 alone. However, RNA-1 is incapable of systemic movement in plants. RNA-2 also resembles the genome of other viruses, in this case carmoviruses (Fig. 8). RNA-2 encodes a polymerase and is capable of multiplying in both inoculated protoplasts and inoculated plants (Demler et al., 1993) independently of RNA-1. In plants, RNA-2 moves systemically. The combination of the two somewhat defective RNA RNA 1
FIG.8. Genome organization of pea enation mosaic virus. Boxes indicate the ORFs. Those in RNA-1 are labeled to correspond with the ORFs of PLRV in Fig. 1.
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molecules, each resembling a different type of virus, produces an agent which behaves like a virus and which is stable in nature. A major biological feature of PEMV is its persistent transmission by aphids, and a possible solution to the problem of how to classify PEMV is to regard it as a luteovirus, albeit one with anomalous properties. A further complication is that the polymerase of PEMV RNA-2 has some sequence relatedness with the polymerases of the subgroup I genomes of luteoviruses. It is conceivable that the recombinatorial origin of the polymerase part of the subgroup I genomes was a virus like PEMV RNA-2. Whatever the eventual taxonomic outcome, the elucidation of the genome structure of PEMV is a vivid example of the contribution of molecular biology to taxonomic thinking.
D . RNA Associated with Luteoviruses 1 . ST9-Associated Satellite-Like RNA Another example of a “symbiotic” association between viral RNAs is the ST9 strain of BWYV. Particles of BWYV-ST9 contain two RNA species (Falk et al., 1989). The larger resembles the genome of other isolates of BWYV, and it multiplies when inoculated into protoplasts. The smaller is a 2.8-kb RNA which encodes a polymerase (Chin et al., 1993) and is capable of replicating in inoculated plants independently of BWYV RNA (Passmore et al., 1993); however, it does not encode a coat protein (Chin et al., 1993). When plants are infected by BWYVST9, symptoms are more severe, and virus yields are about 10 times greater, than in plants infected with other isolates of BWYV (Falk et al., 1989). Aphids transmit both RNA species, each encapsidated in a different particle (Sanger et al., 1994). This association resembles that between the RNAs of PEMV but is different in that BWYV-ST9 does not invade mesophyll tissue and is not mechanically transmissible. According to the current definition of satellites (Mayo, 19911, BWYVST9-associated RNA is not a satellite as it is capable of replication independent of its associated virus (BWYV-ST9).It, and indeed PEMV RNA-2, resembles umbraviruses in being dependent on a luteovirus for encapsidation and therefore transmission (Murant et al., 1995). However, the 2.8-kb ST9-associated RNA is much smaller than the approximately 4.5-kb RNA of umbraviruses. 2 . Satellite RNA of Barley Yellow Dwarf Virus (RPV Strain) Satellite RNA has been detected in RPV cultures (Miller et al., 1991; Silver et al., 1994).The satellite is a D-type (Mayo, 1991)that occurs as circular molecules of 322 nucleotides. The RNA replicates by a rolling
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circle mechanism during which the RNA undergoes self-cleavage of both (+I and (-) sense RNA to release monomers from the polymeric products of replication. The satellite replicates in protoplasts coinfected with RPV but not in those coinfected with PAV (Silver et al., 1994).
IX. CONCLUDING REMARKS Molecular analysis has shown that the genomes of luteoviruses combine most of the strategies used to express the monopartite genomes of (+) sense ssRNA viruses (Morch and Haenni, 1987). Moreover, luteovirus genomes have clearly evolved by recombination between blocks of coding sequence derived from distinct ancestral viruses. Thus, despite the difficulties of studying these viruses, many interesting molecular biology features have been demonstrated and would repay more intensive study. The biological features of luteoviruses are also unusual in that, except in peculiar circumstances, luteoviruses are confined to the phloem of their hosts (Section VI,A) and during transmission luteovirus particles interact with surfaces in their insect vectors to cross several cell boundaries (Section 111,F).Both features can be explained in a general way, but the molecular bases for the properties are as yet poorly understood and are thus excellent candidates for more penetrating molecular study. In this review we have attempted to show progress beyond the first phase of luteovirus molecular biology in which sequences have been accumulated. It seems clear that much more progress can be anticipated on several fronts in the near future and that in many cases the knowledge gained should convey lessons applicable in the wider field of RNA virus molecular biology.
ACKNOWLEDGMENTS We thank Allen Miller for sharing a review with us prior to publication and to H. Barker, V. Brault, S. P. Dinesh-Kumar, H. Guilley, E. Herrbach, R. Hull, C. A. Jolly, L. Kelly, J. W. Lamb, 0. Lemaire, R. R. Martin, J. de Miranda, C. P. Paul, K. Scott, J. F. J. M. Van den Heuvel, and I. Veidt for allowing us to refer to unpublished results andlor personal opinions. Financial support from the Scottish Office Agriculture and Fisheries Department is also acknowledged.
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INDEX
A
Antigen e, mediating host immune responses, 174-175
B Barley yellows dwarf virus heterologous encapsidation, 447 P5 protein, 432 RNA translation, 439-440 Beet western yellows virus agroinfection experiments, 425 heterologous encapsidation, 447 infection diagnosis, 451 P5 protein, 431-434 protein roles, deduction by mutagenesis, 434-435 replication, 449-450 ribosomal frameshift, 436-437 RNA translation, 439 subgenomic mRNA synthesis, 440-442 Borna disease virus, 333, 335-336 Bragg reflections, iridoviruses, 371
C
Capsid, see also Complementation system empty, poliovirus, 25-29, 55-56 iridoviruses, 367-369 mutations affecting RNA encapsidation, poliovirus, 47-52 myristylation, poliovirus, functional significance, 37-38 precursor, poliovirus, proteolytic cleavage, 44-47 Carrot red leaf virus, 447 Central nervous system involvement, cytomegalovirus, 217218 viral persistence, 335 Chickenpox, see Varicella-zoster virus
46 1
Chloriridovirus classification, 351, 363 hybridization complexes, 357, 359 Cleavage maturation, using complementation system, poliovirus, 52-53 proteolytic, capsid precursor, poliovirus, 44-47 Complementation system poliovirus encapsidation study, 39-53 assembly phenotypes, 42 capsid mutations, 47-52 affecting RNA encapsidation, 47-52 capsid precursor, proteolytic cleavage, 44-47 capsid proteins, 43-44 maturation cleavage, 52-53 nucleating role of RNA genome, 43 P1 precursors, 39, 42-43 Cucurbit aphid-borne yellows virus, subgenomic mRNA synthesis, 440-442 Cytomegalovirus, 197-241 human infection determinants, 198216 cell culture systems, 200-201 cell differentiation, 213 gene expression early phase, 210 late, 211 permissive culture cells, 199-212 IE2 protein functions, 204-205 immediate early gene location, 202204 initial events, 214-215 monoclonal antibodies, 206-207 noninfectious enveloped particles, 211-212 nonpermissive, 212-214 polypeptides, immunoblot analyses, 207-209 replication cycle, 199, 202 permissive culture cells, 199-212 strain variabilities, 216
462
INDEX
tegument proteins, 215 virion protein role in initiating infection, 215 infection hematopoietic system and circulating cells, 228, 231-232 latent, 236-241 cell culture models, 239-241 latency site, 238-239 murine, as model, 237-238 tissue cells, 219-220, 224-230 organ tropism, 216-219 spread and pathogenesis, cell types involved, 234-236 transmission modes, 232-234 Cytopathology, luteoviruses, 450 Cytoskeleton, viral interactions affecting cell function, 337-338 Cytotoxicity assays, varicella-zoster virus, 282-283
repetitive, iridoviruses, 390-391 replication, iridoviruses, 375-376 torsion-induced bends, HIV-1 reverse transcription, 115
E Encapsidation, see also Complementation system heterologous, luteoviruses, 447-448 RNA, 30-34 capsid mutations affecting, poliovirus, 47-52 hepadnaviruses, 176-180 requirements for, poliovirus, 31 signals, 32-33 subcellular location, 33-34 Enzymatic activities, iridoviruses, 385386 Epitopes, location, luteoviruses, 445
D
F Dazaifu IV genes, 388-389 repetitive DNA, 390 Defective interfering particles, poliovirus, 31-32, 40 Deoxynucleoside 5"-triphosphate, HIV-1 reverse transcriptase binding site, 116-1 19 DNA complementary, frameshifting, luteoviruses, 437-438 covalently closed circular formation, 191-192 hepadnaviruses, 170-172 invertebrate iridoviruses, restriction endonuclease profile, 354-357 methylation iridoviruses, 376-377 transcription, 380 polymerization, RNA- and DNAdependent, 107, 131-133 priming and synthesis, hepadnaviruses minus strand, 184-189 plus strand, 189-191 proviral, double-stranded, HIV-1, 101, 104
Failure to thrive, 333 Feline leukemia virus, 333 Frameshift, translational, 435-438 G
Genes, iridoviruses, 388-390 Glutamine-glycine dipeptide sites, cleavage, poliovirus, 12-13 Growth hormone deficiency syndrome, caused by lymphocytic choriomeningitis virus persistent infection, 324-333 Guinea pig model, varicella-zoster virus, cell-mediated immunity, 303-306
H Hepadnaviral polymerase, 180-183 experimental approaches, 180-181 minus strand priming, 182-183 mutational analyses, 181-182 sequence similarities, 181
Hepadnaviruses, 167-192, see also Hepatitis B virus DNA, covalently closed circular, 170172 life cycle, 170-172 protein products, 174-176 reverse transcription, 172, 184-192 covalently closed circular DNA formation, 191-192 minus strand DNA priming and synthesis, 184-189 plus strand DNA priming and synthesis, 189-191 pregenomic RNA organization, 184 RNA packaging, 186 RNA, encapsidation cis-acting signals on pregenomic RNA, 177-180 core particle assembly, 176-177 missense mutations, 177 surface antigen forms, 175-176 transcripts, 173-174 virion, structure, 170-171 Hepatitis B virus, 167-168, see also Hepadnaviruses genome organization, 173 historical background, 168-170 minus strand priming, 182-183 mutational analyses, 181-182 open reading frames, 174-176 P protein expression, 187 transfections, 186 Herpesvirus, see Varicella-zoster virus Human foamy virus, 100 Human immunodeficiency virus type 1, 99-146 DNA double-stranded proviral, 101, 104 significance in infectiousness, 123124 LTR, 104 proteolytic processing, 106 replication, 101-106 initial events, 101, 104-105 reverse transcription first template switch, 137-146 efficiency, 142 genomic RNA dimerization, 143 interstrand and intrastand, 145 mode, 145
oligoribonucleotide size, 141 RNA-dependent DNA polymerization, 140 RNase H digestion, 141 origin, 122-125 RNA genomic, interaction with tRNAitLys3, 127-130 templates, 129 trans-activation response element, 104 virus assembly and maturation, 102103, 105-106 virus-host cell membrane fusion, 101 Human immunodeficiency virus type 1 reverse transcriptase, 107-119 association constants, DNA primerDNA template, 131 DNA polymerization, RNA- and DNAdependent, 131-133 template-DNA primer duplex, 114115 torsion-induced bends, 115 160itgag-pol precursor, 124 heterodimer, 108-109 hydroxyl-radical footprinting, 113-114 interaction with primer and template, 113-116 polymerase active site, 113 specific binding, 115-116 nuclease footprinting, 114 polymerase active site and deoxynucleoside 5-triphosphate binding site, 116-119 polymerization by, fidelity, 133-134 recombinant, 108 ribonucleases, 134-137 structure, 109-113 dimerization sites, 112 functional domains, 110-111 p51 and p66 domains, 109, 112 Human T-cell lymphotropic virus, 100 Hybridization, RNA, rotaviruses, 72-73
I Immunocompromised hosts, cytomegalovirus infection, 218, 220223
464
INDEX
Iridoviruses, 347-401 capsid, 367-369 classification, 350-366 alternative approaches, 360-361 comparative studies, 352-359 current system, 350-352 invertebrate, 362-364 new nomenclature, 359-360 new scheme, 365-366 suggested changes to current system, 362-366 vertebrate, 364-365 description, 348-349 DNA-DNA dot-blot hybridization values, 356-358 ecology, 391-399 future directions, 399-401 genes, 388-390 hybridization complexes, 357, 359 infectious particles per host, 392 invertebrate classification, 362-364 DNA restriction endonuclease profile, 354-357 iridescence phenomenon, 371-372 lipid membrane, 361, 368-370 models of host-iridovirus population dynamics, 391 molecular biology, 386-391 particle core, 370-371 persistence alternative hosts, 398-399 in host populations, 396-397 physical, 395 physicochemical properties, 366-367 repetitive DNA, 390-391 replication cell penetration and uncoating, 373 cytoskeletal manipulation, 383-385 DNA, 375-376 enzymatic activities, 385-386 host macromolecular synthesis shutdown, 373-375 methylated DNA transcription, 380 methylation of DNA, 376-377 mRNA stability and methylation, 380-381 nongenetic reactivation, 377-380 transcription, 377-381 translation, 381-382 virion packaging, 382-383
transmission, 393-395 vertebrate, 348 classification. 364-365
L Lentiviruses, 100 Lipid membrane, iridoviruses, 361, 368370 Luteoviruses, 415-457 gene expression, 435-444 internal initiation of translation, 438-440 proteolysis and cap-independent translation, 443-444 readthrough, 442-443 subgenomic mRNA synthesis, 440442 translational frameshifting, 435-438 gene function determination, 424-425 genome structure, 417-424 open reading frame arrangement, 417-420 putative recombination, 424 terminal structures and noncoding regions, 422-423 variation among coding sequences, 420-422 variation among strains, 423 infection diagnosis, 450-452 open reading frames 5 and 6, 431-434 particle structure, 444-448 epitope location, 445 heterologous encapsidation, 447-448 particle assembly determinants, 448 readthrough protein, 445-447 tertiary, 444-445 PO protein, 426-427 P1 and P2 proteins, 427-428 P3 protein, 428-429 P4 proteins, 429-431 P5 proteins, 431-434, 445-447 replication, 449-450 resistance to, 452-453 taxonomy barley yellows dwarf virus, satellite RNA, 456-457 pea enation mosaic virus problem, 455-456 speciation, 453-454
INDEX ST9-associated satellite-like RNA, 456 structure, 454 Lyrnphocystiuirus,classification, 351 Lymphocytic choriomeningitis virus, persistent infection, growth hormone deficiency syndrome, 324-333
M Memory T-lymphocyte responses, varicella-zoster virus, 285-286, 290300 natural infection, 291-296 varicella vaccine, 296-300 Methylation DNA iridoviruses, 376-377 transcription, 380 mRNA, 380-381 Microfilaments, cytoskeletal manipulation, iridoviruses, 384 Microtubules, cytoskeletal manipulation, 383-384 Moloney murine leukemia virus, 333 Mosquitoes, iridoviruses transmission, 393 Murine cytomegalovirus, as model system, 237-238 Myristylation, poliovirus capsid proteins, 18-19 functional significance, 37-38
N Neuroendocrine dysfunctions, virusinduced, in absence of cytolysis and inflammation, 333-337 Nongenetic reactivation, iridoviruses, 377-380
P Particle assembly, determinants, luteoviruses, 448 Pathogenesis cytomegalovirus, 234-236 new perspective, 314
465
Pea enation mosaic virus genome organization, 455 replication, 449 taxonomy, 455-456 Pentamer, 14S, 22-24 poliovirus, 55 Phloem tissue, luteovirus tissue, 449450 Picornuviridae,2, see also Poliovirus Poliovirus, 1-56 arginine residues, 50-51 assembly pathways, 20-21 assembly phenotypes, 53-54 assembly process, 34-38, 53-56 capsid myristylation functional significance, 37-38 P1 and 3CD expression using recombinant vaccinia virus vectors, 35-37 using recombinant vaccinia viruses, 34-35 f3-barrel, 16-17 f3 strands, 16-17 capsid cavity associated with, 50 empty, 25-29, 55-56 protein-RNA binding, 51 cascade of polyprotein processing, 3-4 coding portion, 5 genomic organization, 3-6 life cycle, 6-14 events, 8 protease 2Apr0 release, 11-12 protease 3Cpr0 release, 12-13 RNA replication, 13-14 translation, 10-11 virus entry and uncoding, 7, 9-10 morphogenesis, 19-30 empty capsid, 25-29 55 protomer, 20-22 provirion, 29-30 1 4 s protomer, 22-24 5”-NTR, 4-5 1 4 s pentamers, 55 proteolytic cleavages, 5-6 recombination, 85 RNA, encapsidation, 30-34 defective interfering particles, 3 1-32 RNA requirements, 31 signals, 32-33
studies, complementation system, 39-53 subcellular location, 33-34 RNA-protein interactions, 49-50 serological types, 2 subgenomic replicon, 52 virion, 14-19 capsid protein myristylation, 18-19 properties, 15-16 structure, 16-18 VP1, amino-terminal portion, 47-49 Polymerization DNA,RNA- andDNA-dependent,131133 by HIV-1 reverse transcriptase, fidelity, 133-134 Potato leafroll virus infection diagnosis, 451 particle assembly determinants, 448 P4 protein, 430-431 replication, 449-450 resistance to, 452 RNA translation, 439-440 subgenomic mRNA synthesis, 440-442 Procapsid, poliovirus, 25-29 Protease 2Apr0, release, poliovirus, 11-12 3CD, P1 precursor cleavage, 35-36 30'0, release, poliovirus, 12-13 Proteins, see also Luteoviruses capsid interaction with poliovirus RNA, 51-52 poliovirus myristylation, 18-19 targeted, 43 VP4, poliovirus, 9 RNA interactions, poliovirus, 49-50 TATA-binding, cytomegalovirus, 205 varicella-zoster virus, putative functions, 267 Proteolysis, cap-independent translation and, luteoviruses, 443-444 Proteolytic processing, HIV-1, 106 Protomer, 5S, poliovirus, 20-22 Provirion, poliovirus, 29-30
R Rabies virus, 333, 335 Ranavirus, classification, 351, 364-365
Readthrough, protein presence, luteoviruses, 445-447 Recombinant vaccinia virus, poliovirus assembly process studies, 34-35 vectors, P1 and 3CD expression, 35-37 Reouiridae, see also Rotaviruses genome rearrangements, 92 Replication, see also Iridoviruses HIV-2 initial events, 101, 104-105 virus assembly and maturation, 102-103, 105-106 luteoviruses, 449-450 nonlytic strategy, viral persistence, 321-322 RNA, poliovirus, 7, 13-14 Retrovirus reverse transcription scheme, 119-122 species, 100 type D, human immunodeficiency virus type 105-106 Reverse transcriptase, 100, see also Human immunodeficiency virus type 1 reverse transcriptase Reverse transcription hepadnaviruses, see Hepadnaviruses HIV-1 first template switch, 137-146 host tRNAitLys3 primer, 125-131 origin, 122-125 sites of hypermutability and pausing, 134 scheme, retroviruses, 119-122 Rhinovirus, provirion, 29 Ribonucleases, HIV-1 reverse transcriptase, 134-137 RNA encapsidation, 30-34 capsid mutations affecting, poliovirus, 47-52 hepadnaviruses, 176-180 requirements, poliovirus, 31 signals, 32-33 subcellular location, 33-34 frameshift, luteoviruses, 436-437 hybridization, rotaviruses, 72-73 luteovirus, putative recombination, 424 messenger genomic and subgenomic, hepadnaviral transcripts, 173-174
467
INDEX methylation and stability, 380-381 subgenomic synthesis, luteoviruses, 440-442 translation, poliovirus, 6 poliovirus, interaction with capsid proteins, 51-52 pregenomic cis-acting signals, hepadnaviruses, 177-180 organization, hepadnaviruses, 184 profiles, rotaviruses, 72-74 protein interactions, poliovirus, 49-50 replication, poliovirus, 7, 13-14 satellite, barley yellow dwarf virus, 456-457 ST9-associated satellite-like, luteoviruses, 456 transfer isoacceptor species, 125-127, 129 primer, retroviruses, 121 tRNAitLys3 primer, HIV-1 reverse transcription, 125-131 translation, poliovirus, 10-1 1 RNA polymerase, RNA-dependent, poliovirus, 5 Rotaviruses, 71-93 biophysical data, 86-87 genome rearrangements discovery, 71-74 duplication, 83 evolution, 91-92 extent, 75 in vitro in cultured cells, 79-82 mechanisms, 82-86 other genera of Reouiridue, 92 RNA, segments 5 and 6,89-90 sequence data, 75-79 groups, 71 in uitro growth properties, 89 rearranged genes, function, 86-91 RNA hybridization, 72-73 profiles, 72-74 plaque-purified, 79-80 second generation genome rearrangements, 82-83 segment 5, normal and rearranged forms, 77-78 standard gene 5, 77-78 3 UTR, 75,83 variants, 77
S
Sedimentation coefficient, poliovirus virion, 15-16 Severe combined immunodeficiency, rotaviruses infections, 71
T Tipula oleracae, iridoviruses transmission, 394 T-lymphocyte proliferation antigen-specific, varicella-zoster virus, 281 assay tests, varicella-zoster virus, 282 T lymphocytes, varicella-zoster virus tropism, severe combined immunodeficient hu mouse, 276-280 Trans-activation response element, HIV-1,104 Transcription, iridoviruses, 377-381 methylated DNA, 380 Translation iridoviruses, 381-382 luteoviruses cap-independent, proteolysis and, 443-444 internal initiation, 438-440 leaky termination, 442-443 poliovirus mRNA, 6 RNA, 10-11 Transmission, iridoviruses, 393-395
V Varicella vaccine maintaining cell-mediated immunity, 302-303 memory T-lymphocyte responses, 296300 primary cell-mediated immune response, 285-290 Varicella-zoster virus, 265-306 cell-mediated immune response, 280306 assessment methods, 281-283 guinea pig model, 303-306 long-term immunity, 293
468
INDEX
maintenance mechanisms, 300-303 memory T-lymphocyte, 285-286, 290-298 primary, 283-290 natural infection, 284-285 varicella vaccine, 285-290 disease correlations with cell-mediated immunity, 272-273 gene sequences, 268-269 infection peripheral blood cells in uitro, 273275 primary, viremia, 267-271 medical significance of related disease, 266 pathogenesis, 271 proteins putative functions, 267 recognized by T-lymphocytes, 291292 reactivation maintaining cell-mediated immunity, 300-302 viremia, 271-273 reexposure, maintaining cell-mediated immunity, 300 subclinical viremia, relationship between episodes, 305 tropism, T lymphocytes, severe combined immunodeficient hu mouse, 276-280 Venezuelan encephalitis virus, 334 Viral persistence, 313-339 iridoviruses alternative hosts, 398-399 in host populations, 396-397 physical, 395
requirements avoidance of recognition of infected cells by specific immune response, 316-318 block of action of nonspecific antiviral defense mechanisms on infected cells, 318-319 essential, 315 induction of suppression of host immune response, 319-321 nonlytic strategy of replication, 321322 virus-induced changes in cells, 316321 virus-induced alterations, host cellular differentiated functions in absence of cytolysis, 323-338 experimental evidence, 323-324 growth hormone deficiency syndrome, 324-333 neuroendocrine dysfunctions, 333337 viral interactions with cytoskeleton, 337-338 Viremia during primary varicella-zoster virus infection, 267-271 varicella-zoster virus reactivation, 271-273 Virions, iridoviruses, packaging, 382383
W Wasting syndrome, 333
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