Open Collections

UBC Theses and Dissertations

UBC Theses Logo

UBC Theses and Dissertations

Molecular characterization of rubella virus nonstructural proteins and viral RNA synthesis Liang, Yuying 2000

Your browser doesn't seem to have a PDF viewer, please download the PDF to view this item.

Item Metadata

Download

Media
831-ubc_2000-565769.pdf [ 10.74MB ]
Metadata
JSON: 831-1.0099576.json
JSON-LD: 831-1.0099576-ld.json
RDF/XML (Pretty): 831-1.0099576-rdf.xml
RDF/JSON: 831-1.0099576-rdf.json
Turtle: 831-1.0099576-turtle.txt
N-Triples: 831-1.0099576-rdf-ntriples.txt
Original Record: 831-1.0099576-source.json
Full Text
831-1.0099576-fulltext.txt
Citation
831-1.0099576.ris

Full Text

M O L E C U L A R CHARACTERIZATION OF RUBELLA VIRUS NONSTRUCTURAL PROTEINS AND VIRAL RNA SYNTHESIS  By Yuying Liang B. Sc., Sichuan University, P. R. China, 1992 M . Sc., Shanghai Institute of Biochemistry, Academia Sinica, 1995  A thesis submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy in the Faculty of Graduate Studies, Genetics Program  We accept this thesis as conforming to the required standard  The University of British Columbia © Yuying Liang,. .. J 0 September 2000.  In presenting degree  this  at the  thesis  in  partial fulfilment  of  University of  British Columbia,  I agree  freely available for reference copying  of  department  this or  publication of  and study.  thesis for scholarly by  this  his  or  her  Department of The University of British Columbia Vancouver, Canada  requirements that the  I further agree  purposes  representatives.  may be It  thesis for financial gain shall not  permission.  DE-6 (2/88)  the  is  that  an  advanced  Library shall make it  permission for extensive  granted  by the  understood be  for  allowed  that without  head  of  my  copying  or  my written  ABSTRACT  Rubella virus (RV) is a single-strand positive RNA virus. Its 9762-nucleotide (nt) genomic RNA contains two large open reading frames (ORFs) at the 5'- and 3'-termini, encoding s  nonstructural and structural proteins, respectively. The nonstructural proteins (NSPs) are first translated as a 200-kDa polyprotein (p200), which undergoes a single proteolytic cleavage into pl50 and p90 by an intrinsic protease, the RV nonstructural protease (NS-pro).  The purpose of this thesis work is to characterize RV NSPs and viral RNA synthesis at the molecular level, including the characterization of the RV NS-pro domains, the roles of NSPs in viral RNA synthesis, and the regulatory mechanism for viral RNA synthesis.  RV NS-pro is an M-group papain-like cysteine protease (PCP) with both trans- and ciscleavage activities. Employing an in vitro translation system, I mapped the regions of NS-pro required for trans- and cis- activities, and demonstrated the importance of an X domain in rrarcs-cleavage activity. RV NS-pro catalytic region was also predicted to have a papain-like folding from primary and secondary structure analyses.  The time course of RV RNA synthesis was precisely determined. Synthesis of negative-strand RNA stops after 10 h postinfection (hpi), and subsequently switches to highly efficient synthesis of positive-strand RNAs (genomic and subgenomic RNA). The roles of respective NSPs and the underlying regulatory mechanism for RV RNA synthesis were investigated by mutational analysis and complementation experiments. Processing of p200 was found to be crucial for virus replication. Uncleaved p200 was shown to be functional in negative-strand RNA synthesis but not for positive-strand RNA. In contrast, the complex formed by the ii  cleavage products pl507p90 is not an active replicase for negative-strand RNA, but is required for the efficient generation of positive-strand RNAs. Syntheses of both negative- and positivestrand R V RNAs were found to be cw-preferential from complementation experiments.  A mechanism for R V NSP translation, processing and R V R N A replication was proposed. Newly translated p200 functions in cis to synthesize a full-length negative-strand RNA. The subsequent processing of p200 into pl50/p90 converts this replicase into one with positivestrand R N A specificity, which functions in cis to produce genomic and subgenomic R N A efficiently.  iii  T A B L E OF CONTENTS  ABSTRACT LIST OF TABLES LIST OF FIGURES LIST OF ABBREVIATIONS ACKNOWLEDGEMENTS 1. INTRODUCTION  ii vi vii viii x 1  1.1. Rubella virus (RV) biology 1 1.1.1. Classification 1 1.1.2. Clinical aspects 1 1.1.3. Virion structure and virion proteins 2 1.1.4. Genome organization and sequence information 4 1.1.5. Replication features and infection cycles 6 1.1.6. Attachment, entry and uncoating 7 1.1.7. R V nonstructural proteins (NSPs) 8 1.1.8. Replication complexes and viral R N A synthesis 9 1.1.9. Expression and processing of R V structural proteins (SPs) 10 1.1.10. Virus assembly and budding 13 1.2. Viral encoded papain-like cysteine proteases (PCPs) 16 1.2.1. Viral proteases 16 1.2.2. The papain-like cysteine protease (PCP) family 17 1.2.3. The M - , and L - group PCPs 18 1.2.4. Alphavirus NS protease 19 1.2.5. R V NS protease (NS-pro) 20 1.3. Replication of positive-strand RNA viruses 20 1.3.1. R N A replication studies on some model positive-strand R N A viruses: poliovirus, B M V , and alphavirus 21 1.3.2. Common replication strategies of positive-strand R N A viruses 27 1.3.3. Cw-preferential replication 30  2. PROJECT RATIONALE AND THESIS OBJECTIVES 3. MATERIALS AND METHODS 3.1. Materials and supplies 3.2. Methods 3.2.1. Growth of cells and viruses 3.2.2. Propagation of bacterial strains 3.2.3. Preparation of competent cells and transformation 3.2.4. Mini preparation of plasmid D N A 3.2.5. Restriction endonuclease digestions and D N A modifications 3.2.6. Polymerase Chain Reaction 3.2.7. Plasmid Construction 3.2.8. In vitro transcription 3.2.9. In vitro translation 3.2.10. Vero cells transfected by R N A transcripts using Lipofectin 3.2.11. BHK-21 cells transfected by R N A transcripts using electroporation  33 35 35 35 35 36 36 36 37 38 39 46 47 47 48 iv  3.2.12. 3.2.13. 3.2.14. 3.2.15. 3.2.16. 3.2.17. 3.2.18. 3.2.19.  Plaque assay and virus growth analysis Metabolic labeling Immunoprecipitation Total RNA preparation RNase protection assay (RPA) Electrophoresis Image analysis and cleavage efficiency comparison Sequence analysis  4. RESULTS AND DISCUSSIONS  48 49 49 50 50 53 54 55  56  4.1. Characterization of domains involved in cis- and trans-cleavage activities of RV NS-pro 56 4.1.1. Processing of RV NSP by in vitro translation 56 4.1.2. Construction of truncated NS-pro cDNA clones 59 4.1.3. Defining the NS-pro domain required for trans cleavage 61 4.1.4. Domains required for cis cleavage 64 4.1.5. Effect of N-terminal regions on cleavage efficiency 66 4.1.6. Secondary structure prediction for RV NS-pro 69 4.1.7. Discussion 1 72 4.2. Effects of NSP cleavage on virus replication and RNA synthesis 76 4.2.1. Construction of mutants 76 4.2.2. Effects of mutations on NSP processing 77 4.2.3. Effects of mutations on virus growth 79 4.2.4. Effects of mutations on viral RNA synthesis 83 4.2.5. Discussion II : 91 4.3. Molecular characterization of RV RNA synthesis 96 4.3.1. RV RNA replication is as-preferential 96 4.3.2. Time course of RV RNA synthesis 103 4.3.4. Discussion III 105  5. SUMMARY AND PERSPECTIVE REFERENCES  110 112  V  LIST O F T A B L E S  Table Table Table Table  1. 2. 3. 4.  Sequence information on oligonucleotide primers used in this work Mutations created at the catalytic site and around the cleavage site Effects of R V NSP cleavage mutants on virus replication Comparison of the relative amounts of RNAs produced in the WT and mutants  40 77 80 89  vi  LIST OF FIGURES  Figure 1. Genome organization of RV and SIN 5 Figure 2. Replication strategy of RV 10 Figure 3. Predicted membrane topology of the RV SPs 12 Figure 4. Polio virus genome organization 22 Figure 5. Genome organization of brome mosaic virus (BMV) 24 Figure 6. Locations of PCR primers for serial deletions on RV genome 41 Figure 7. Locations of PCR primers for site-directed mutagenesis on RV genome 45 Figure 8. Probes used in RNase protection assay (RPA) and the relative positions and lengths of the protected regions 51 Figure 9. RV NSP processing in in vitro translation systems with or without addition of Zn . 57 Figure 10. RV p200(G1301S) and pl50 cleave substrate protein in trans 59 Figure 11. Plasmids and protease-encoding constructs 60 Figure 12. In vitro translation of protease constructs ( A 3 4 8 / G 1 3 0 1 , M 8 2 7 / G 1 3 0 U V 9 2 0 / G 1 3 0 1 , A974/G1301, A1020/G1301, and G1102/G1301) and examination of their frans-cleavage activities. 62 Figure 13. In vitro translation of protease constructs (V920/H1290, ^92^x295, V920/P1296, V920/L1297, and V920/R1299) and examination of theirtams-cleavageactivities 63 Figure 14. Autolytic processing of protease constructs V920/I1773, A974/I1773, A1020/I1773, and 2+  G1102/I1773  65  Figure 15. Cleavage efficiencies of protease constructs 68 Figure 16. Comparison of primary and secondary structures between RV NS-pro and papain. 71 Figure 17. Functional domains of RV NS-pro and comparison with complete papain sequence. 73 Figure 18. Effects of mutations on NSP processing in an in vitro translation system 78 Figure 19. Comparison of processing ratios between the WT and mutant NSPs 79 Figure 20. Immunoprecipitation of RV-specific SPs from cells transfected by WT and mutant RNAs 81 Figure 21. Growth curves of the WT, G1300A, and R1299A mutant viruses 83 Figure 22. Sensitivity of RPA for detection of both positive- and negative-strand RV RNAs.. 85 Figure 23. RNA analysis of the WT and mutant constructs 88 Figure 24. Comparison of WT and mutant G1301S on the levels of negative-strand RNA synthesis 90 Figure 25. Trans complementation of RV replication 98 Figure 26. Schematic diagrams of constructed replication-defective mutants 99 Figure 27. Synthesis of positive-strand RNA is ds-preferential 102 Figure 28. Accumulated amounts of viral RNAs during virus infection 104 Figure 29. Proposed mechanism for RV NSP translation, processing, and viral RNA synthesis. 109  vii  LIST O F A B B R E V I A T I O N S A ATCC ATP BHK BMV bp BSA BVDV CIP CRS cpm CTP DI DMEM DMSO DNA DNase dNTPs DTT EAV EDTA EM EMBL ER FCS FMDV GTP h hpi HAV HI ICTV IRES kb kDa kV Lpro M m.o.i MEM mg MHV min ml mM  absorbance American Type Culture Collection adenosine triphosphate baby hamster kidney cell line brome mosaic virus base pair bovine serum albumin Bovine viral diarrhea virus calf intestinal alkaline phosphatase congenital rubella syndrome count per minute cytidine triphophate defective-interfering Dulbecco's modified Eagle's medium dimethyl sulfoxide deoxyribonucleic acid deoxyribionuclease deoxynucleoside triphosphates dithiothreitol equine arteritis virus ethylene diaminetetraacetic acid electron microscopy European Molecular Biology Laboratory endoplasmic reticulum fetal calf serum the foot-and-mouse virus guanosine triphosphate hour hour post infection hypovirulence-associated virus hemagglutination inhibition International Committee on Taxonomy of Viruses Internal ribosomal entry site kilobase kilodalton kilovolt leader protease molar multiplicity of infection minimal essential medium milligram Mouse hepatitis virus minute milliliter millimolar  MOPS ng NIH nm NSP NS-pro nt  3-[N-morpholino]-propanesulfonic acid nanogram National Institutes of Health nanometer nonstructural protein nonstructural protease nucleotide  OD260 ORF PAGE PBS PCP PCR PEG PFU pg PIPES PMSF RdRp RNA RNase RPA rpm RRV RT RV s S SDS SFV SIN SL SP ss TM Tris ts TYMV ug UTP UTR VLP WT YFV  absorbance at 260 nm wavelength open reading frame polyacrylamide gel electrophoresis phosphate-buffered saline papain-like cysteine protease polymerase chain reaction polyethylene glycol plaque forming unit picogram piperazine-N, N'-bis[2-ethane-sulfonic acid] phenylmethylsulfonyl fluoride RNA-dependent R N A polymerase ribonucleic acid ribonuclease RNase protection assay rotation per minute Ross River virus room temperature rubella virus second Svedberg unit sodium dodecyl sulfate Semliki Forest Virus Sindbis Virus stem-loop structural protein single-stranded transmembrane domain trishydroxymethylaminomethane temperature sensitive turnip yellow mosaic virus microgram uridine triphosphate untranslated region Virus-like particle wild type yellow fever virus  ix  ACKNOWLEDGEMENTS  First, I would like to thank my supervisor, Dr. Shirley Gillam, for giving me an opportunity to pursue the graduate study, for her advice, support, and encouragement throughout. I would also like to thank members in my supervisory committee, Drs. Caroline Astell, Peter Candido, and Ann Rose for their advice, suggestion, and discussions. Thank you to Jiansheng, Dawei, Zhiyong (Joe), Karen, and Hanwei for all the helps, and to many people in the B.C. Research Institute for a wonderful working environment. I was supported by the studentship award from B C Research Institute for Children's and Woman's Health. This thesis is dedicated to my parents.  1. I N T R O D U C T I O N  1.1.  1.1.1.  Rubella virus (RV) biology  Classification  Rubella virus (RV) is the single member of the Rubivirus genus of the Togaviridae family, which consists of two genera - Alphavirus and Rubivirus (Francki et al., 1991). Members of this family are small enveloped viruses with a single-strand positive R N A genome. The Alphavirus genus contains 27 distinct arthropod-transmitted viruses, which can replicate in both arthropod and vertebrate hosts. In contrast, the only known natural reservoir for R V is in humans and R V has no known invertebrate host. Extensive studies have been carried out on the structure and replication of Sindbis virus (SIN), the prototype of alphaviruses (for a review, see Strauss and Strauss, 1994).  1.1.2.  Clinical aspects  RV is the causative agent of a benign disease, German measles (also known as rubella, 3-day measles) characterized by symptoms including maculapapular rash, lymphadenopathy, lowgrade fever, conjunctivitis, sore throat, and arthralgia (for a review see Wolinsky, 1996). The major health concern with R V is its teratogenicity. The incidence of birth defects in babies born to women infected during the first trimester approaches 80%. The birth defects, collectively known as congenital rubella syndrome (CRS), involve heart defects, cataracts, deafness, and mental retardation (Frey, 1994). Immunization of infants  and susceptible  women of  childbearing age against R V infection has dramatically reduced the incidence of both rubella and CRS. However, R V remains a significant human pathogen as shown by the resurgence of 1  rubella despite a vaccination program, the significant incidence of joint involvement (transient arthralgia and arthritis, chronic arthritis) following virus infection or vaccination, and its association with chronic diseases including autoimmune diseases (Frey, 1994).  1.1.3.  Virion structure and virion proteins  RV is a small spherical particle of 60-70 nm diameter (Bardeletti et al, 1975). Thin-section electron microscopy of the virion reveals a 30-35 nm electron-dense core surrounded by an electron-lucent zone between the core and the virion envelope (Murphy et al., 1968; Murphy, 1980). The central core, suggested to have a T = 3 icosahedral symmetry (Frey, 1994), contains a single copy of an R N A genome protected by multiple copies of capsid protein. On the lipid bilayer envelope, the glycoproteins form rather poorly resolvable 6- to 8-nm surface spikes (Trudel et al., 1980). Following degradation of virions, spherical subunits that form hexagonal and pentagonal arrays were observed, possibly representing an end-on view of the virion spike (Payment et al., 1975; Frey, 1994). It is unknown whether the glycoproteins on the R V envelope form a similar T = 4 icosahedral lattice to those of alphavirions (Strauss and Strauss, 1994).  RV is composed of three structural proteins (SP), the capsid protein C and two envelope glycoproteins, E l and E2 (Oker-Blom et al, 1983; Clarke et al, 1987). The non-glycosylated C protein has 299 amino acids and is rich in proline and arginine residues (Clarke et al,  1987;  Frey and Marr, 1988). The cluster of positively-charged residues may be involved in binding to the genomic R N A in nucleocapsids. With a molecular mass of 35 kDa, C migrates as a doublet on polyacrylamide gel electrophoresis, the lower band being of greater intensity than the upper one (Oker-Blom et al, 1983; Marr et al, 1991). In addition, the capsid protein in virions has 2  been reported to be a disulfide-linked dimer (Waxham and Wolinsky, 1983; Baron and Forsell, 1991). Nevertheless, dimerization of capsid protein does not seem to be required for virus particle formation (Lee et al, 1996).  From the deduced sequence, the large surface glycoprotein E l is a 412-amino-acid protein and migrates as a discrete band of 58 kDa (Clarke et al., 1987). Monoclonal antibodies against E l exhibit hemagglutination inhibition (HI) and/or neutralization activity, suggesting that E l contains hemagglutination activity and neutralization domains (Green and Dorsett, 1986; Gerna et al, 1987; Ho-Terry and Cohen, 1985; Trudel et al, 1985; Umino et al, 1985; Waxham and Wolinsky, 1983, 1985b). Although most monoclonal antibodies against E l recognize nonlinear epitopes, the linear epitopes recognized by some anti-El monoclonal antibodies have been mapped. Epitopes that react with monoclonal antibodies specific for hemagglutination and neutralization have been defined between amino acids 245 and 285 (Terry et al, 1988), and between amino acids 202 and 283 (Wolinsky et al, 1991). Chaye et al. (1992) further mapped the hemagglutination epitope to amino acids between 214 and 240, and two neutralization epitopes to amino acids between 214 and 233, and between 219 and 233, respectively.  E2, the heavily glycosylated envelope protein, migrates as a smear between 42 and 54 kDa (Oker-Blom et al, 1983). These forms of E2 derive from a single protein (deduced 281 amino acids) with different extents of glycosylation (Clarke et al,  1987; Waxham and Wolinsky,  1985b). The biological role of E2 remains obscure, although it appears to contain one neutralization epitope (Green and Dorsett, 1986) and a strain-specific epitope (Dorsett et al, 1985). E2 is required for cell surface expression of E l (Hobman et al, 1990). Very few E2specific monoclonal antibodies are isolated compared to the large number of El-specific when mice are immunized with whole R V (Green and Dorsett, 1986; Waxham and Wolinsky, 1983), 3  possibly due to poor immunogenicity of E2 in native virions. In fact, E2 epitopes seem to be buried under E l in the glycoprotein spike complexes (Ho-Terry and Cohen, 1984; Katow and Sugiura, 1988a; Waxham and Wollinsky, 1985a; Green and Dorsett, 1986).  In  the alphavirions, E l and E2 form a heterodimer by noncovalent bonds and three  heterodimers form each surface spike (Vogel et al,  1986; Anthony and Brown, 1991).  Although the structure of the R V surface spikes has not been defined, there is evidence that E l and E2 form a complex in the virion. E1-E2 heterodimers were observed when virions were disrupted with nonionic detergent (Dorsett et al., 1985; Baron and Forsell, 1991) as well as when E l and E2 were coexpressed from cDNA constructs (Baron and Forsell, 1991). A small fraction of the glycoproteins in the virion is disulfide-linked, including E l - E l homodimer and E1-E2 heterodimer (Waxham and Wolinsky, 1983; Dorsett et al, 1985). Moreover, the E l - E l homodimers seem to be present in a complex with E2 since they can be immunoprecipitated by both anti-El and anti-E2 antiserum (Baron and Forsell, 1991).  1.1.4.  Genome organization and sequence information  The R V genome (Fig. 1) is a single-strand positive RNA of 9762 nucleotides (nt) (Pugachev et al,  1997a) encoding two large open reading frames (ORFs). The 5'-proximal nonstructural  protein-ORF (NSP ORF) from nt 41 to nt 6388 encodes nonstructural proteins involved in viral RNA synthesis (Pugachev et al, 1997a; Yao et al, 1998); the 3'-proximal structural proteinORF (SP ORF) from nt 6512 to nt 9701 encodes structural proteins required for virus assembly (Clarke et al, 1987; Dominguez et al, 1990; Pugachev et al, 1997).  4  R  |  i |xHll  p150  V  5'cap^B  • 24SsgRNA  p90 C  E2  E1  •  26S s g R N A  p200  SIN 5'  nsP1 V  cap-  •UH  n s P 2  "V  I llllll HH  n s P 3  X  V  n s P 4  E2  C E3  |  | Me thy Itransf erase  ill  [ x ] X-domain  RNA-dependent RNA polymerase  E1 6K  "poly(A) 3'  protease helicase  Figure 1. Genome organization of R V and SIN. Both genomes contain two large ORFs, NSP and SP ORFs, at the 5' and 3' ends, respectively. NSPs of the two viruses are first translated as polyprotein precursors, which are processed into final cleavage products by virus-encoded proteases. R V NSP polyprotein p200 is cleaved once into pl50 and p90, whereas SIN NSP is cleaved into four final products, nsPl to nsP4. R V and SIN NSPs share a conserved domain with unknown function called the X domain. Four conserved enzyme motifs identified in a number of positive-strand R N A viruses, the methyltransferase, protease, helicase and RNA-dependent RNA polymerase (RdRp), were also identified in R V and SIN. However, the order of these domains in R V differs from that of SIN. SPs of both viruses are translated as polyproteins from subgenomic RNAs, which are produced during virus infection, and posttranslationally cleaved into C, E2 and E l , with an additional two small peptides E3 and 6K in SIN. (Modified from Frey, 1994).  The  5' and 3' ends of the R V genome can bind specific host proteins (Pogue et al., 1993;  Nakhasi et al., 1994; Chen and Frey, 1999) and are presumed to be critical cis elements for viral RNA  synthesis. The 5' end contains a 14-nt single-stranded leader (ss-leader) followed by  a stem-and-loop structure [5'(+)SL] (nt 15 to nt 65), the complement of which at the 3' end of the negative-strand RNA  [3'(-)SL] has been proposed to function as a promoter for synthesis of  positive-strand genomic RNA.  Pugachev and Frey (1998) examined the effects of the 5' end  structure on virus replication by making mutations and deletions in an infectious cDNA clone. It was revealed that the A A dinucleotide at nt 2 and 3 is of critical importance whereas mutations in the other positions caused variable but not lethal effects. In general, mutations 5  within the stem had a more profound effect on viral phenotype than did mutations in either the ss-leader or upper loop. The 5'(+)SL was found to be important for efficient translation of the NSP ORF. In contrast, positive-strand R N A synthesis was unaffected by the 5'(+)SL mutations as well as the ss-leader mutations, which argues against the proposed function of the 3'(-)SL as a promoter for initiation of the genomic positive-strand RNA.  The 3' end of the R V genome contains four SL structures. SL1 and SL2 are both located in the E l coding region, while SL3 and SL4 are within the 59-nt 3'-untranslated region (UTR) preceding the poly(A) tract. SL2 is a structure shown to interact with human calreticulin (Singh et al., 1994), an autoantigen potentially involved in R V R N A replication and pathogenesis. Site-directed mutagenesis of the 3' U T R with an R V infectious cDNA clone (Chen and Frey, 1999) revealed that most of the 3' UTR is required for viral viability except for the 3'-terminal 5 nt and the poly(A) tract, although poly(A) was rapidly regenerated during subsequent replication. Maintenance of the overall SL3 structure, the 11-nt single-stranded sequence between SL3 and SL4 and the sequences forming SL4 were all important for viral viability. In contrast to a previous proposal, maintenance of the SL2 stem rather than the U - U bulge was critical in calreticulin binding. However, mutations in the SL2 stem or the U - U bulge gave rise to viable viruses when introduced into an infectious cDNA clone, indicating that binding of calreticulin to SL2 is independent of viral viability.  1.1.5.  Replication features and infection cycles  Compared to the rapid and efficient replication of alphavirus, R V infection has a long latency period, slow replication cycle, and low virus production (Hemphill et al., 1988). Differences in the replication features between R V and alphavirus are summarized as follows. (1) Alphavirus  6  can infect a wide range of host cells, including an acute, usually short-duration infection in vertebrates and a persistent lifelong infection in arthropods. This situation is mirrored in cell culture (Strauss and Strauss, 1994). R V replicates in a number of primary cell cultures and continuous cell lines of vertebrates, but fails to infect mosquitoes (Tesh and Rosen, 1975). (2) Alphavirus infection reaches a peak by 4 to 8 h after a 2 to 3-h latent period, whereas R V infection has a latent period of 24 h and reaches peak level by 48 to 72 h. (3) Alphavirus can grow to high titers (10 PFU/ml) in cultured vertebrate cells. In contrast, R V normally grows to 9  a very.low titer in most cell lines except for Vero and BHK-21 cells, the two most used permissive cell lines to grow R V with a titer of 10 -10 PFU/ml (Frey, 1994). (4) Alphavirus 7  8  infection shuts down host R N A and protein synthesis, whereas inhibition of host protein synthesis by R V infection is indistinguishable until 40 hpi and reaches 40% at 44 hpi (Hemphill etal., 1988).  Some other features of R V replication include the profound inhibitory effects of interferon a and Y on the synthesis of viral RNAs and proteins, and the time-dependent inhibition by actinomycin D and a-amanitin (Nakhasi et al., 1988).  1.1.6.  Attachment, entry and uncoating  RV attaches to susceptible cells rapidly, mostly within 1 h. The R V receptor on the host cell surface has not been identified. It is believed that R V enters cells through receptor-mediated endocytosis. The low pH environment of endosomes triggers the fusogenic activity of the viral glycoproteins, causing fusion of the virus membrane with the endosomal membrane and resulting in the release of the viral nucleocapsid into the cytoplasm. Consistent with this hypothesized entry mechanism, exposure of the R V glycoproteins to low pH causes a fusogenic 7  activity (Katow and Sugiura, 1988b). Katow and Sugiura (1988a) suggested that R V E l plays a vital role in membrane fusion. Sequence analysis revealed an internal hydrophobic domain within E l , which was shown to be involved in membrane fusion by site-directed mutagenesis (Yang et al, 1998). The mechanism for uncoating of the genome from nucleocapsid remains obscure. C protein was found to undergo a conformational change at low pH that renders it hydrophobic (Mauracher et al., 1991), indicating that a low pH environment may also trigger the uncoating process.  1.1.7.  R V nonstructural proteins (NSPs)  The uncoated R V genome first serves as the mRNA for the translation of NSPs. A 200-kDa polyprotein, p200, is first produced and processed by intrinsic protease activity into the 150kDa N-terminal product (pi50) and the 90-kDa C-terminal product (p90) (Bowden and Westaway, 1984; Chen et al, 1996; Forng and Frey, 1995; Marr et al, 1994; Yao et al, 1998). Sequence analysis has predicted four conserved enzyme motifs in the NSP sequence, ordered from the N terminus to the C terminus as methyltransferase, protease, helicase and RNAdependent R N A polymerase (RdRp) (Fig. 1) (Frey, 1994; Koonin and Dolja, 1993). Therefore, the cleavage product pi50 carries methyltransferase (Rozanov et al, (Gorbalenya et al,  1992) and protease  1991) motifs on the N and C termini respectively, whereas p90 contains  helicase (Gorbalenya et al, 1990) and RdRp motifs (Kamer and Argos, 1984) on the N and C termini, respectively. These four enzyme motifs have also been identified in a number of animal and plant positive R N A viruses, possibly representing universal enzymatic requirements for their viral R N A synthesis (Koonin and Dolja, 1993; Strauss and Strauss, 1994). The protease domain is discussed in more details in section 1.2. The exact role of each R V NSP in viral R N A synthesis remains to be determined.  8  1.1.8.  Replication complexes and viral R N A synthesis  Replication complexes are formed as membrane-bound cytoplasmic vacuoles to synthesize viral R N A during R V infection (Lee et al., 1992, 1994). From immunogold labeling electron microscopy (EM) studies, Magliano et al. (1998) identified these membrane-bound structures as virus-modified lysosomes. The replication complexes presumably consist of R V NSPs and host factors. However, no successful isolation of functional replication complexes has been reported. Several host factors specifically binding to the 5' and 3' ends of R V R N A have been identified (Pogue et al., 1993; Nakhasi et al., 1991, 1994) and might be components of replication complexes. Two host proteins, 59 and 52 kDa in size, specifically interact with the RV 5'(+)SL R N A (Pogue et al., 1993), and were identified as Ro/SSA-associated antigens (Nakhasi et al., 1994). The 3'(-)SL structure (complementary negative-strand equivalent of the 5'-(+)SL) is thought to serve as a promoter for the initiation of positive-strand R N A synthesis, although this hypothesis was not confirmed from mutational analysis (Pugachev and Frey, 1998). Nevertheless, three cytosolic proteins with relative molecular masses of 97, 79, and 56 kDa were found to bind specifically to the 3'(-)SL structure (Nakhasi et al., 1991). Host protein binding specifically to the 3' end was found to be a homologue of human calreticulin (Nakhasi et ah, 1994), and the binding site was mapped to SL2 (Singh et al., 1994). In addition, Chen and Frey (1999) identified three 3'-end RNA-protein complexes by gel mobility shift assay, and detected six host protein species, with molecular masses of 120, 80, 66, 55, 48, and 36 kDa, interacting with the 3'-UTR.  RV employs a similar R N A replication mechanism to alphavirus (Fig. 2). Upon infection, NSPs are translated from input genomic R N A and form functional replication complexes with host 9  factors to replicate viral RNA. A full-length negative-strand R N A is first produced on the template of input genomic RNA and serves as the template for the replication of 40S positivestrand genomic RNA and for the transcription of 24S subgenomic RNA, which is initiated at an internal site  (U6436)  in the negative-strand RNA (Hemphill et al, 1988; Pugachev et al., 1997a).  ribozyme  O  O'  translation of NSPs  NSPs  (-) gRNA <«  host factors  40S gRNA,  5' 24S sgRNA  production of full-length negative-strand RNA  replication of genomic RNA and transcription of subgenomic RNA  Figure 2. Replication strategy of R V . Upon infection, NSP is translated from the input genomic RNA and associates with host factors to form replication complexes, which function to replicate viral RNAs. First, a full-length negative-strand RNA is produced and serves as the intermediate for the replication of positivestrand genomic RNA and transcription of subgenomic RNA (Modified from Wolinsky, 1996).  1.1.9.  Expression and processing of RV structural proteins (SPs)  R V SPs are translated from the 24S subgenomic RNA as a 110-kDa polyprotein in the order of N H - C - E 2 - E l - C O O H (Fig. 1) (Oker-Blom, 1984; Clarke et al, 1987). The p i 10 is processed 2  into C, E2, and E l , the latter two being further posttranslationally modified during intracellular transport. C is a cytoplasmic protein, whereas E l and E2 are type I membrane glycoproteins, consisting of an N-terminal large ectodomain, a single transmembrane domain near the C -  10  terminus, and a short C-terminal cytoplasmic tail (Clarke et al,  1987; Hobman and Gillam,  1989; Hobman et al, 1988; Singer et al, 1987).  R V capsid protein differs from alphavirus capsid protein in the mechanism of its cleavage from the precursor. Alphavirus capsid protein is an autoprotease and releases itself from the polyprotein precursor within the cytoplasm (Aliperti and Schlesinger, 1978; Melancon and Garoff, 1987). In contrast, R V capsid protein does not contain protease activity, and its release from the polyprotein precursor is mediated by host signal peptidase, a lumenal enzyme that cleaves signal peptide from proteins following translocation (Clarke et al, 1988; McDonald et al, 1991). The cleavage between C and E2 occurs after the signal sequence of E2, a stretch of hydrophobic amino acids at the C-terminus of capsid protein. The signal sequence of E2 is inserted into the E R membrane and mediates the translocation of E2 into the E R (Hobman and Gillam, 1989; Suomalainen et al, 1990). The C-terminal transmembrane domain of E2 stops the translocation and anchors E2 to membrane (Clarke et al,  1987; Vidgren et al, 1987).  Translocation of E l is mediated by a hydrophobic signal sequence located immediately after the E2 transmembrane domain, and continues until the C-terminal transmembrane domain of E l is anchored to membrane (Hobman et al, 1988). The cleavage between E2 and E l occurs just after the E l signal sequence (Qiu et al, 1994b). The topology of each SP is schematically shown in Fig. 3.  11  lumen  cytoplasm  n  E2 signal peptide  |  E2TM  H •  E1 signal peptide  1  E1 TM  Figure 3. Predicted membrane topology of the R V SPs. Capsid is a cytosolic protein whereas E2 and E l are membrane bound with the bulk of their N termini projecting into the lumen of microsomes. The hydrophobic signal peptides of E2 and E l (shaded and striped rectangles, respectively) are thought to remain attached to the carboxyl termini of capsid and E2, respectively. E2 and E l are anchored in the lipid bilayer via a transmembrane domain (TM) near their carboxyl termini (blank and checkered rectangles, respectively). Both E2 and E l contain three N-linked glycans (Y). The cytoplasmic domain of E2 is a short hydrophilic loop between its T M and the signal peptide of E l . In contrast, E l has a carboxyl-terminal cytoplasmic tail (C) of 13 amino acids, which may interact with capsid protein. The carboxyl (C) and amino (N) termini of the proteins are indicated (Hobman et al., 1994).  Both E l and E2 are posttranslationally modified by glycosylation and acylation during the transport from E R through Golgi complex to plasma membrane. High-mannose, hybrid-type, and complex-type N-linked glycans have been found in both E l and E2 from virions (Sanchez and Frey, 1991), whereas O-linked oligosaccharides are observed only in E2 (Lundstrom et al., 1991; Sanchez and Frey, 1991). Generally, oligosaccharides on glycoproteins play a role in initiation and maintenance of correct folding, protecting polypeptides against proteolysis, and influencing the antigenicity and immunogenicity of glycoproteins (Ng et al., 1990; Gallagher et al., 1992). E2 contains three functional N-linked glycosylation sites. Removal of any site results in slower glycan processing and lower stability, suggesting that mutated E2 proteins are not properly folded and transported, and are rapidly degraded (Qiu et al., 1992b). Moreover, 12  removal of more than one glycosylation site blocks E2 from cell surface expression (Hobman et al, 1994).  E l , containing hemagglutination and major neutralization epitopes, has three functional N glycosylation sites (Hobman et al, 1991). The single glycosylation mutants ( G l , G2, and G3), but not the double mutant (G2G3) or the triple mutant (G1G2G3), were found to be capable of inducing neutralization antibodies (Qiu et al,  1992a). However, the failure of G l mutant in  producing hemagglutinin inhibition antibody suggested an important role of oligosaccharide at the G l site in hemagglutination activity (Qiu et al, 1992a).  Formation of E1-E2 heterodimer is required for transport of E l to the Golgi complex and to the cell surface (Hobman et al,  1990). E2 can be transported to the plasma membrane in the  absence of E l (Hobman and Gillam, 1989). In contrast, unassembled subunits of E l are arrested in a post-ER, pre-Golgi compartment (Hobman et al, 1992), and require coexpression of E2 for release and targeting to the Golgi complex and to the cell surface (Baron et al, 1992; Hobman et al, 1993). An internal hydrophobic domain within E l is found to be important in the noncovalent E1-E2 interaction (Yang et al, 1998). Only a small proportion of E l and E2 is transported to the cell surface. In fact, the bulk of E l and E2 accumulate in the Golgi-like compartments, facilitating the intracellular budding of virions (Bardeletti et al, 1979; Bowden and Westaway, 1985; Hobman et al, 1990; Payment et al, 1975).  1.1.10.  Virus assembly and budding  Assembly of enveloped R N A virus includes encapsidation, the formation of nucleocapsid by specific interactions between capsid protein and genomic RNA, and budding, the acquisition of 13  viral envelope derived from viral glycoprotein-modified cell membrane. Encapsidation signal, which confers the capability of efficient binding to R N A by capsid protein, has been identified in SIN (Frolova et al, 1997; Owen and Kuhn, 1996), SFV (White et al, 1998), RRV (Frolova et al, 1997), and other viruses. Encapsidation by R V capsid protein is specific for 40S positivestrand genomic R N A (Oker-Blom et al,  1984). A region between nt 330 and nt 617 was  identified to bind capsid protein specifically. Further deletion studies narrowed this down to 29 nt between nt 347 and nt 375 (Liu et al, 1996). However, it is unknown whether this sequence is sufficient for genomic RNA packaging. On the other hand, an arginine-rich domain on capsid protein between amino acids 28 and 56 interacted specifically with R V RNA. Nevertheless, other region(s) may also be involved in efficient R N A binding, since the domain between amino acids 28 and 56 has lower R N A binding affinity than intact capsid protein (Liu et al, 1996).  R V virions have been shown to bud from a variety of sites including the Golgi complex, intracellular vacuoles,  and plasma membrane (Lee et al,  1992;  Murphy,  1980). The  intracellular budding of R V is supported by studies on virus-like particles (VLPs), in which all three SPs expressed in mammalian cells were targeted predominantly to the Golgi complex with only a small proportion of E l and E2 detectable on the plasma membrane, and facilitated the budding of VLPs from these sites (Hobman et al, 1990; Qiu et al, 1994a).  Proteins involved in the budding of enveloped viruses vary (for a review see Garoff et al, 1998). (1) For alphavirus (Simons and Garoff, 1980; Suomalainen et al,  1992) and  hepadnavirus (Brass and Ganem, 1991), budding is driven by interactions between the viral transmembrane proteins and the internal virion components (core, capsid, or nucleocapsid). (2) Some viruses accomplish budding mainly by internal components. For example, core protein 14  (the Gag protein) of retroviruses (simian immunodeficiency virus, HIV-1) is able to form enveloped particles by itself (Delchambre et al,  1989; Gheysen et al,  1989). Budding of  negative-strand R N A viruses (rhabdoviruses, orthomyxoviruses, and paramyxoviruses) is directed predominantly by the matrix protein, although spike proteins (not required absolutely) are also engaged in this process (Li et al, 1993; Liu et al, 1995; Liu and Air, 1993; Jin et al., 1994; Justice et al, 1995; Mebatsion et al, 1996; Mitnaul et al, 1996; Pattnaik et al, Schnell et al,  1986;  1994). (3) Budding of coronavirus particles depends solely on the envelope  glycoproteins. Coexpression of the M H V envelope protein genes (S, M , and E) resulted in the release of membrane particles morphologically indistinguishable from authentic virions, suggesting that the membrane proteins can assemble into virtually bona fide envelopes without the nucleocapsid (Vennema et al, 1996).  Like alphavirus, budding of R V also depends on the interactions between glycoproteins and capsid protein. VLPs morphologically indistinguishable from virus particles could be released by coexpression of the three SPs, capsid C, and glycoproteins E l and E2, but not by expression or coexpression of any one or two of them (Qiu et al, 1994a). Using mutational analysis, Yao and Gillam (1999) demonstrated that the E l transmembrane domain and the cytoplasmic tail of 13 amino acids play an important role in late stages of virus assembly, possibly during virus budding, consistent with earlier studies (Hobman et al,  1994)  indicating that the E l  cytoplasmic domain may interact with nucleocapsids and that this interaction drives virus budding.  After budding into the lumen of the Golgi complexes, R V virions are released through microsome-mediated exocytosis. The E l cytoplasmic domain was found to modulate virus  15  release in a sequence-dependent manner, possibly through interactions with other proteins (Yao and Gillam, 2000).  1.2.  1.2.1.  Viral encoded papain-like cysteine proteases (PCPs)  Viral proteases  In order to use their small coding capacity economically, viruses often encode proteins in the form of polyproteins that are processed into final products by host and viral encoded proteases. Viral proteases also function to cleave host cell proteins for the benefit of virus life cycles (Roehl et al.,  1997). Therefore, virus encoded proteases generally regulate viral gene  expression and pathogenesis, and are often good targets for drug design.  There are four classes of proteases, named after the chemical nature of their catalytic sites, cysteine, serine, aspartate, and metallo-proteases. Each class can be further divided into protein families that have common origins and structural folding (Rawlings and Barrett, 1993).  RNA viral proteases have diverged to the point that sequence similarity between homologues can barely be detected except for the close vicinity of catalytic sites, possibly because of the high mutation rate of R N A viruses. Nevertheless, distant relationship between R N A viral proteases and cellular ones, mostly chymotrypsin-like or papain-like proteases, can still be detected from their common catalytic sites and similar three-dimensional structures. Viral encoded chymotrypsin-like proteases include proteases with different catalytic triads, such as Ser-His-Asp,  Cys-His-Asp,  and Cys-His-Glu,  while  adopting  homologous  folding  to  16  chymotrypsin (Bazan and Fletterick, 1988; Gorbalenya et al., 1989a; Choi et al., 1991b; Allaire et ah, 1994; Matthews et al., 1994). Viral encoded papain-like proteases are discussed below.  1.2.2.  The papain-like cysteine protease (PCP) family  The PCP family include a group of cellular and viral proteases which employ the catalytic Cys and His dyad. Viral PCPs include members with highly variable protein size and sequence except for the close vicinity of catalytic residues (Gorbalenya et ah, 1991). Until recently, the relationship between viral and cellular papain-like proteases had been based on primary sequence analysis (Gorbalenya et al., 1989b, 1991; Hardy and Strauss, 1989; Oh and Carrington, 1989) and predicted secondary structure (Skern et ah, 1998). The first crystal determination of a viral PCP, the foot-and-mouth disease virus (FMDV) leader protease (Lpro), confirmed the previous prediction that it adopts a modified compact version of the cellular papain folding (Guarne et al., 1998).  Viral PCPs contain required residues of Cys and His, a single conservative change of which abolishes all protease activity, suggesting their roles in catalytic reaction (Hardy and Strauss, 1989; Chen et al., 1996; Marr et al., 1994). The involvement of Asn as the third catalytic sites, however, is not very clear. No Asn was absolutely required for alphavirus nsP2 protease activity (Strauss et al, 1992).  Using the Schechter and Berger (1967) nomenclature for the description of cleavage sites, the amino acid residues of the N-terminal side of the cleaved bond are numbered P3, P2, PI and those residues of the C-terminal side are numbered PI', P2', P3' (PI or PI' near the cleaved bond). Alignment of the cleavage sites recognized by viral PCPs revealed some common 17  features. Most uniform is the presence of a residue with short side chain (Gly or Ala) in the PI position. The PI' position is also usually a short-side-chain residue (Chen et al., 1996).  1.2.3.  The M - . and L - group PCPs  Two groups of viral PCPs have been distinguished on the basis of their positions in polyproteins and their processing behaviors (Gorbalenya et al., 1991). Leader or L-group PCPs include potyvirus proteinase HC-Pro (Oh and Carrington, 1989), the F M D V Lpro (Piccone et al., 1995), the p29 and p48 proteases of hypo virulence-associated virus (HAV) of chestnut blight fungus (Choi et al., 1991a; Shapira and Nuss, 1991), the equine arteritis virus (EAV) nsPl protease (Snijder et al., 1992) and others. They are located at the N-terminus of the respective polyprotein and mediate a single cleavage at their own C termini (function in cis). These protease domains are located outside the domains directly involved in genome replication and expression.  Main or M-group PCPs include the NS protease of alphavirus (Hardy and Strauss, 1989), the murine coronavirus mouse hepatitis virus (MHV) PLP-1 (Baker et al., 1993; Bonilla et al., 1997), and R V NS protease (NS-pro) (Chen et al., 1996; Yao et al, 1998). They seem to be the main, and possibly the only enzymes responsible for the processing of the respective polyproteins, and function in both cis and trans. These protease domains are located in the central region of the polyprotein and therefore constitute parts of the molecules involved in genome replication and expression. It is interesting to note that an outside X domain (a novel conserved domain of unknown function) is associated with M-group PCPs. It is speculated that the X domain might function in regulating polyprotein processing. Elimination of the X domain from PLP-1 of M H V reduced cleavage by 22 to 63% (Bonilla et al., 1995).  18  1.2.4.  Alphavirus NS protease  Alphavirus NS protease was mapped to the C-terminal domain of nsP2, specifically from residue 460 to 807 in SIN, using an in vitro translation system (Hardy and Strauss, 1989). This NS protease functions both in cis and trans (Hardy and Strauss, 1989). Both sequence analysis (Hardy and Strauss, 1989) and mutational analysis (Strauss et al., 1992) demonstrated alphavirus NS protease to be a viral M-group PCP, in which  C481  and  H558  form the catalytic  dyad. A remarkable mutation, N614D, was found to enhance cleavage (Strauss et al., 1992). Its effects on virus replication and viral R N A synthesis suggested a specific function of polyprotein PI23 in negative-strand R N A synthesis (Lemm et al., 1994). Regulation of viral RNA synthesis by NSP processing is discussed in section 1.3.1.3.  Both nsP2 and polyproteins containing nsP2 have been found to be active proteases, but the cleavage site preferences of different polyproteins are different (de Groot et al., 1990). In general, (1) polyproteins containing nsPl (i.e., P12, P123, or P1234) could not cleave the nsP2/nsP3 site, but cleave the nsP3/nsP4 site fairly efficiently and the nsPl/nsP2 site inefficiently; (2) only polyproteins containing nsP3 (i.e., P123, P1234, P23 and P234) could cleave the nsP3/nsP4 site; (3) polyproteins lacking nsPl (i.e., P23 and P234), as well as nsP2 under some conditions, could cleave the nsP2/nsP3 site very efficiently; (4) the presence of nsP4 in the polyprotein does not affect cleavage specificity. These cleavage site preferences result in different processing pathways for NSPs early and late in infection (de Groot et al., 1990). Early in infection, SIN genomic RNA is translated into P123 and P1234. P123 could not autoproteolyze and P1234 cleaves at the nsP3/nsP4 site in cis, resulting in the presence of P123 and nsP4. With the increase of P123, cleavage in trans of P123 at the nsPl/nsP2 site produces  19  proteases that are active in cleaving the nsP2/nsP3 site. By 3 to 4 hpi, cleavage of the nsP2/nsP3 site occurs so rapidly that the nascent polyprotein is cut to produce PI2 and P34. P12 can be cleaved both in cis and trans into nsPl and nsP2, whereas P34 is stable. Therefore, the major NSP proteins after 4 hpi are nsPl, nsP2, nsP3 (produced by termination at the opal termination codon), and P34. Things are a little different in SFV, which lacks an opal codon between nsP3 and nsP4, in which nsP3 is produced throughout. However, SFV was also shown (Takkinen et al., 1991) to produce PI23 only early in infection and to produce PI2 and P34 late in infection, similarly to the case of SIN.  1.2.5.  R V NS protease (NS-pro)  By comparative sequence analysis, Gorbalenya et al. (1991) proposed that R V NS-pro is an M group PCP, and identified an X domain (from residue 834 to 940) adjacent to the R V NS-pro domain. Our previous studies demonstrated that R V NS-pro could function both in cis and in trans (Yao et al., 1998), a feature consistent with M-group PCPs. The catalytic dyad residues of NS-pro,  Cn52  and  H1273,  have been proposed by sequence alignment (Gorbalenya et al., 1991)  and supported by site-directed mutagenesis (Chen et al., 1996; Marr et al., 1994). However, the lack of precisely mapped domain for NS-pro and its highly variable sequence from other PCPs make the structure and functions of NS-pro poorly characterized.  1.3.  Replication  of positive-strand  RNA viruses  From the Sixth Report of the International Committee on Taxonomy of Viruses (ICTV) (Murphy et al., 1995), viruses can be classified by genome structure into double-strand D N A  20  virus, single-strand D N A virus, double-strand R N A virus, single-strand positive R N A virus, single-strand negative R N A virus, D N A and R N A Reverse Transcribing Viruses, and the Subviral Agents. Among known viruses the single-strand positive R N A virus group is most abundant. Many of these cause human, animal, and plant diseases including encephalitis, hemorrhagic fevers, and hepatitis. Their genomic RNAs act directly as viral messenger R N A and are thus infectious. Replication of positive-strand R N A viruses is of fundamental interest to understanding their pathogenesis.  A variety of approaches has been employed in studying viral R N A replication. Biochemical analyses of viral-encoded NSPs provide details of their enzymatic features. Mutational analyses were used to study their replication, including temperature-sensitive (ts) mutants, isolated variants, and mutants constructed from infectious cDNA clones.  In vivo and in vitro  reconstituted systems were constructed to reflect authentic viral R N A replication by using RNA templates and expressed viral NSPs. These varied methods were employed to characterize viral NSPs, cis R N A elements, and host factors in the replication process. Below are examples of replication studies on some model viruses like poliovirus, brome mosaic virus (BMV), and alphavirus.  1.3.1.  R N A replication studies on some model positive-strand R N A viruses: poliovirus.  B M V . and alphavirus.  1.3.1.1.  Poliovirus  Poliovirus is a member of the Picornaviridae family of positive-strand R N A viruses. The poliovirus genome (Fig. 4) is a 7500-nt positive-strand R N A with 5'-terminal covalently linked  21  protein VPg and 3'-terminal poly(A) sequence. It contains a single ORF, translated into a 220kDa polyprotein and processed by intrinsic proteases into final cleavage products. Interestingly, initiation of translation of its genomic R N A is cap-independent and controlled by a long segment within the 5' non-translated region, termed the internal ribosomal entry site (IRES) (for a review, see Schmid and Wimrner, 1994). The proteins 1A, IB, 1C, and ID, located at the N terminus of the polyprotein, are capsid proteins. The proteins 2A, 3C, and 3CD are proteases. 3B is VPg, covalently attached to the 5' end of all newly synthesized viral RNAs. Viral R N A replication requires 3D (the RdRp), and other viral proteins 2B, 2C, 3A, 3B, 3AB, and 3CD as well (for a review, see Wirnmer et al., 1993).  Poliovirus  3' NC  1  3B  5' NC 1A  1B  1C  1D  2A 2B  2C  3A  3C  3D  |—AAAAA  VPg  Figure 4. Poliovirus genome organization.  Molla et al. (1991) developed a cell-free, de novo system for the production of infectious poliovirus, in which newly synthesized proteins from poliovirion R N A using HeLa cell extract can replicate and encapsidate viral R N A to form infectious poliovirus. This is a complete system for studying poliovirus replication, i.e., translation, polyprotein processing, R N A replication, and virus assembly. This system efficiently yields virus titers of 4 x 10 PFU/ml 7  and also reflects the authentic replication of poliovirus RNA, producing the VPg-linked RNAs asymmetrically with more positive than negative strands (Barton and Flanegan, 1993; Barton et al., 1995; Barton et al., 1996). The ability to form authentic, functional replication complexes 22  and infectious virus in vitro provides direct access to the replicative process. Using this system, initiation of negative-strand R N A synthesis was found to require the activity of a guanidineinhibited viral protein 2C (Barton et al, 1995; Barton and Flanegan, 1997). The switch from translation to initiation of negative-strand R N A synthesis was also characterized. The initiation of negative-strand synthesis appears to be coordinately regulated with the natural clearance of translating ribosomes to avoid the dilemma of ribosome-polymerase collisions (Barton et al, 1999).  1.3.1.2.  Brome mosaic virus (BMV)  B M V , the type of the Bromovirus genus, is a member of the alphavirus-like superfamily (Ahlquist, 1992). The B M V genome (Fig. 5) consists of three separately encapsidated RNAs, 1, 2, and 3. B M V RNAs do not contain the 3' poly(A) of cellular RNA, but have instead a tRNAlike structure, required for efficient negative-strand R N A synthesis (Dreher & Hall, 1988a, 1988b; Miller et al, 1986; Sun et al, 1996). RNAsl and 2 encode NSP la (109 kDa) and 2a (94-kDa), respectively, which are essential R N A replication factors, la contains domains implicated in R N A helicase and R N A capping functions (Ahola and Ahlquist, 1999; Kong et al,  1999) and 2a contains an RdRp domain (Ahlquist, 1992). RNA3 encodes cell-to-cell  movement and coat proteins. Coat protein is translated from a subgenomic RNA, RNA4, which initiates internally from the negative-strand RNA3 (Janda and Ahlquist, 1993).  Janda and Ahlquist (1993) found that yeast expressing the NSPs la and 2a supports B M V RNA replication and mRNA synthesis. This yeast system reproduces all known features of B M V RNA replication in natural plant hosts, including localization to the ER, dependence on la, 2a, and the same cis R N A signals, and similar ratios of positive- to negative-strand R N A 23  (Restrepo-Hartwig and Ahlquist, 1996, 1999; Janda and Ahlquist, 1993; Sullivan and Ahlquist, 1999; Krol et al., 1999). This is a powerful system in characterizing viral NSPs (Chen and Ahlquist, 2000; Restrepo-Hartwig and Ahlquist, 1999), cis R N A elements (Sullivan and Ahlquist, 1999), and especially cellular factors essential for viral R N A replication (Diez et al., 2000; Ishikawa et al., 1997) in combination with yeast genetics.  BMV  RNA1 m7G-  RNA2  1a  2a  m7G-  RE  RNA3 m7G"  Movement  Coat  Hh  Hh  subgenomic RNA  Figure 5. Genome organization of brome mosaic virus (BMV). B M V genome contains three R N A molecules, each with 5' cap and 3' tRNA structure. The subgenomic RNA, translated into coat protein, is produced during virus infection (Diez et al, 2000).  B M V replication was also studied from a biochemical approach. The partially purified RdRp from BMV-infected barley leaves allowed synthesis of negative-strand RNAs from input positive-strand templates in a sequence-specific manner (Miller et al., 1986; Quadt and Jaspars, 1990; Kao and Sun, 1996), and also synthesized subgenomic RNA4 by initiating internally within the negative-strand RNA3 (Miller et al., 1986). This in vitro system was used to analyze the initiation of negative-strand R N A synthesis, which starts at the penultimate cytidylate (Kao and Sun, 1996; Miller et al., 1986), generates abortive initiation products (Sun et al., 1996), and  24  requires the tRNA-like structure present at the 3' end (Chapman and Kao, 1999). It was also used to study the precise initiation of positive-strand R N A synthesis, in which a 27-nt R N A from B M V was found to direct correct initiation of genomic positive-strand R N A synthesis by the B M V replicase (Sivakumaran and Kao, 1999), and initiation of subgenomic R N A (Adkins et al, 1997; Siegel et al, 1997).  1.3.1.3.  Alphaviruses  The Alphavirus genus, containing 27 currently recognized members, is the only other genus of the Togaviridae family in addition to Rubivirus (Calisher and Karabatsos, 1988; Strauss and Strauss, 1994). The alphavirus genome (Fig. 1) encodes two large ORFs. The 5'-terminal twothirds of the genome encodes NSPs and the 3'-terminal one-third encodes SPs (for a review, see Strauss and Strauss, 1994). The NSPs of alphaviruses are translated from the input genomic R N A upon infection as one or two polyproteins, P123 or P1234, depending on the virus. Cleavage of these polyproteins into nsPl, nsP2, nsP3, nsP4, as well as a number of cleavage intermediates plays an important role in regulating viral R N A synthesis. The SPs are translated as a polyprotein from a 26S subgenomic RNA, and processed into the capsid protein C, the envelope proteins E l and E2, and two small polypeptides E3 and 6K. Both the 49S genomic and the 26S subgenomic RNAs are capped and polyadenylated. During R N A replication, a fulllength negative-strand RNA is produced that serves as a template for replication of the genomic RNA and for transcription of the subgenomic RNA. Synthesis of negative-strand RNA stops at 3 to 6 hpi whereas that of positive-strand RNAs (the 49S genomic and the 26S subgenomic RNAs) continues (Sawicki and Sawicki, 1980; Sawicki et al,  1981). Several studies using  different approaches characterized the underlying regulatory mechanism (Lemm and Rice, 1993a, 1993b; Lemm et al, 1994, 1998; Shirako and Strauss, 1994). 25  Analyses of cleavage mutants suggest that virus replication is regulated by the extent of polyprotein cleavage. Mutant N614D has such a high protease activity to process nsPl/nsP2 and nsP2/nsP3 cleavage sites that polyprotein PI23 cannot exist. This mutant was lethal at 37 °C, suggesting that P123 or an intermediate polyprotein was required for R N A synthesis (Strauss et al., 1992). A further mutant, in which both nsPl/nsP2 and nsP2/nsP3 cleavage sites were blocked, allowing only PI23 and nsP4 to be produced, carried out the early negativestrand R N A synthesis normally, while it had greatly impaired positive-strand R N A synthesis (Shirako and Strauss, 1994). Cleavage at the nsP3/nsP4 site to release nsP4 was found to be absolutely required as a mutation blocking this site was lethal (Shirako and Strauss, 1994). These results demonstrated that polyprotein PI23 and nsP4 are required for negative-strand R N A synthesis, whereas cleavage products from PI23 and nsP4 are responsible for efficient positive-strand R N A synthesis.  Lemm and Rice (Lemm and Rice, 1993a, 1993b; Lemm et al., 1994) used a reconstitution system to identify the NSP requirements for RNA replication, in which various combinations of NSP cleavage products and intermediates expressed from vaccinia recombinants were tested for the ability to replicate a template RNA. They found that polyprotein PI23 and nsP4 could support negative-strand R N A synthesis, whereas the four final cleavage products or any combination of polyproteins not including P123 could not. The requirement for P123 is further supported by the fact that a mutant polyprotein P123(N614D), which contains accelerated protease activity as described above, was inactive in R N A synthesis unless the nsPl/nsP2 and nsP2/nsP3 cleavage sites were blocked (Lemm and Rice, 1993b). The authentic nsP4 was also found to be necessary for R N A synthesis, and it can be supplied from cleavage of either P34 or  26  truncated forms of P34 or as a ubiquitin-nsP4 fusion protein rapidly cleaved in the cell (Lemm and Rice, 1993a, 1993b; Lemm et al, 1994).  All these studies come to common conclusions. A model for the composition of replication complexes and the temporal regulation of negative- and positive-strand RNAs has thus been proposed (Lemm et al, 1998). Alphavirus contains three forms of replication complexes, one formed by uncleaved P123 and nsP4 generating only negative-strand RNA, one composed of nsPl, P23, and nsP4 active in both negative-strand R N A and 49S positive-strand genomic RNA syntheses, and one consisting of the final cleavage products nsPl, nsP2, nsP3, and nsP4 producing only 49S positive-strand genomic R N A and subgenomic RNA. Therefore, cleavage at the 1/2 and 2/3 sites switches the template preference of the replication complex from negative-  to positive-strand R N A and also inactivates  negative-strand R N A synthesis,  explaining the shutoff of negative^strand R N A synthesis after 4 to 6 hpi.  1.3.2.  Common replication strategies of positive-strand R N A viruses  Studies on disparate groups of plant and animal positive strand R N A viruses have revealed remarkably similar replication strategies. (1) Negative-strand R N A is the intermediate for the replication of genomic R N A and the transcription of subgenomic RNA. Upon infection, viral genomic R N A is first translated to generate replication factors that, along with host factors, form replication complexes to replicate viral RNA.  Viral R N A replicates through a negative-strand R N A intermediate,  produced first from the template of input genomic R N A and serves as the template for the replication of genomic RNA and/or the transcription of subgenomic RNA.  27  (2) Four conserved functional domains, methyltransferase, helicase, protease, and RdRp, have been identified within virus-encoded NSPs (Koonin and Dolja, 1993), suggesting that although the precise character of these and interacting host components varies for each virus they employ similar mechanisms for RNA replication. Methyltransferase is one of the enzyme activities needed for capping RNA transcripts (Bisaillon and Lemay, 1997; Rozanov et al., 1992). Helicase is capable of enzymatically unwinding duplex RNA structures by disrupting the hydrogen bonds that hold the two strands together. This activity is coupled with the hydrolysis of an NTP by NTPase activity (for a review, see Kadare and Haenni, 1997). Protease processes nonstructural polyprotein to final products. RdRp is the enzyme responsible for the de novo synthesis of RNA from the end of an RNA template (for a review, see Buck, 1996; O'Reilly and Kao, 1998). All viral RdRps lack the proof-reading function of DNA polymerase, which leads to a high rate of random mutations during replication of viral RNAs.  (3) Os-acting RNA sequences within 5' and 3'-ends, and internal regions of template RNA were identified as essential for RNA replication and transcription. Effects of these cis RNA elements on virus replication and viral RNA synthesis have been studied using mutational analysis. Both the secondary structure and the nucleotide sequence can be critical for recognition. Mutations of the stem-and-loop structure at the 5' end of RV [5'(+)SL] caused profound effects on viral phenotype (Pugachev and Frey, 1998). The 3' end of RV containing four stem-loop (SL) structures SL1 to 4 is important for virus viability (Chen and Frey, 1999). Four cis sequence elements have been identified in alphavirus RNAs; the first 44 nt and a 51-nt section in the coding region of nsPl forming stem-loop structures, a 24-nt unit found within the junction between NSP and SP ORFs, and a 19-nt span at the 3' end (Strauss and Strauss, 1986). In a 27-nt promoter for positive-strand RNA synthesis of BMV, the nucleotide sequence rather  28  than the secondary structure was shown to be important for function (Sivakumaran et al, 1999).  (4) Host factors are required participants in RNA replication. Direct and indirect evidence has confirmed that host factors (most likely proteins) are involved in positive-strand RNA virus replication. A number of SIN ts mutants were identified as host range dependent (Kowal and Stollar, 1981). Mutations in SIN RNA promoter elements affected RNA synthesis differently in different host cells (Kuhn et al., 1990; Niesters and Strauss, 1990). Partially purified replication complexes from virus-infected cells contained cellular proteins (Andino et al., 1990, 1993; Barton et al., 1991). Many cellular proteins were found to bind specifically to the presumed promoter region of viral RNA (Blyn et al., 1995; Ito and Lai, 1997; Furuya and Lai, 1993; Yu and Leibowitz, 1995; Nakhasi et al, 1990, 1994; Pogue et al, 1993; Tsuchihara et al,  1997). Identified binding cellular proteins include autoantigen La and its homolog  (Pardigon and Strauss, 1996; Pogue et al, 1996; Svitkin et al, 1994; Spangberg et al, 1999), autoantigen calreticulin (Singh et al, 1994), polypyrimidine tract-binding protein (Ito and Lai, 1997; Tsuchihara et al, 1997), heterogeneous nuclear ribonucleoprotein A l (Li et al, 1997), and others.  (5) The process of viral RNA replication is well regulated, as shown by the switch of synthesis from negative- to positive-strand RNA and the asymmetric production of excess positive-strand RNAs over negative-strand RNA. Two regulatory mechanisms have been proposed. One involves the recruitment of additional host factors (Pogue et al, 1994). The other possibility is that NSP cleavage results in distinct NSP components in replication complexes that possess different capabilities in the synthesis of different viral RNA species. A prominent example of the second mechanism is alphavirus as described above. 29  1.3.3.  Cfa-preferential replication  The genomes of positive-strand RNA virus contain essential cis sequence elements that act as recognition signals for viral RNA recognition and encodes NSPs participating in the replication process, normally in trans. Therefore, in complementation studies a replication-defective R N A template lacking a functional protein component can be rescued by a helper R N A expressing the active component in trans. However, mutations in some coding sequences of the viral genome are noncomplementable in trans, revealing their cis-acting functions. This has been reported for poliovirus (Novak and Kirkegaard, 1994), mouse hepatitis virus (MHV) (de Groot et al., 1992), clover yellow mosaic virus (White et al., 1992), cowpea mosaic virus (van Bokhven et al., 1993), turnip yellow mosaic virus (TYMV) (Weiland and Dreher, 1993), barley stripe mosaic virus (Zhou and Jackson, 1996), tomato bushy stunt virus (Scholthof and Jackson, 1997), tobacco etch virus (Mahajan et al., 1996; Schaad et al., 1996), and alfalfa mosaic virus (Neeleman and Bol, 1999). These cw-acting coding sequences are presumed to reflect a coupling between translation and replication of viral R N A or a ds-preferential function of the encoded protein in virus replication.  One of the three ORFs of T Y M V encodes an NSP precursor, which is proteolytically processed into pi50 and p70. Several genomic RNAs with different internal deletions and frameshift mutations failed to replicate detectably  in the presence  of wild-type helper genome,  demonstrating that replication of T Y M V RNA is strongly ds-preferential (Weiland and Dreher, 1993). The authors proposed a model in which the cw-preferential replication is due to the interaction of newly synthesized pi50 and p70 preferentially with the R N A genome from which they have been made, resulting in the channeled formation of a replication initiation  30  complex in cis. The cis replication depends on the pl50/p70 complex as an entity rather than on either protein alone.  Novak and Kirkegaard (1994) constructed amber mutations in the poliovirus genome and examined their rescue by wild-type proteins provided by a helper genome. Amber-suppressing cell lines were used to ensure that the defects in the amber mutants arose from failure to be translated, not from defects in RNA sequence or structure. An internal region of the poliovirus genome, located between coding region 2A-am66 and 3D-am28, was thus identified whose translation is required in cis. Such a requirement for translation in cis could, as proposed by the authors, result from either the preferential cis action of a protein, or a requirement for the act of ribosomal passage itself in cis. Several potential mechanisms were proposed by the authors to explain the cis action of a protein. (1) The newly synthesized protein is required for template establishment. A ds-acting protein might interact with the positive-strand RNA from which it is being translated to enable that RNA to be a template for negative-strand RNA synthesis. (2) Diffusion or integrity of protein could be restricted. Several poliovirus proteins and viral RNA synthesis were found to be associated with cytoplasmic membrane surfaces (Bienz et al., 1990) that restrict the diffusion of a viral protein. In addition, a short-lived intermediate in processing of the polyprotein could be present at high concentration only near the RNA from which it was translated. (3) A viral RNA molecule was only transiently competent for both translation and template establishment. The RNA molecule from which a protein was translated would be one of the few RNA molecules in the vicinity available at that time as a template for RNA synthesis. On the other hand, two potential mechanisms for effects of ribosomal passage in cis on RNA replication were also proposed. (1) Ribosomal passage could alter the RNA structure in the cw-required region to facilitate negative-strand synthesis. (2) A ribosome-associated  31  protein or subcellular structure, required for RNA synthesis, can be obtained or used by the viral genome only while ribosomes pass through the as-required region.  The benefits of as-preferential replication of positive-strand RNA viruses were suggested (Weiland and Dreher, 1993; Novak and Kirkegaard, 1994) to facilitate the assembly of replication complexes when few viral RNAs and proteins are present, and to help prevent the replication of host RNAs or defective-interfering (DI) RNAs.  32  2. PROJECT RATIONALE AND THESIS OBJECTIVES  RV NSP cleavage is critical in viral RNA synthesis and the responsible enzyme, NS-pro, is an interesting viral PCP member for study. Because of the high diversity of primary sequence among PCP members, more information is needed for a clear understanding of this protease family. Like other M-group PCPs, R V NS-pro is located in the central region of polyprotein and the functional proteolytic domains of NS-pro, in either cis- or rrans-cleavage activity, have not been defined. R V NS-pro also constitutes part of the domains mediating viral R N A synthesis. Therefore, characterization of NS-pro functional domains contributes not only to the knowledge of the viral PCP family, but also to the understanding of the biological roles of RV NSPs.  Viral R N A synthesis is a critical step for virus propagation. It is believed that R V NSPs, associated with host factors, form replication complexes to synthesize three RV-specific R N A species, full-length negative-strand  RNA, 40S  positive-strand genomic  R N A and 24S  subgenomic R N A (Frey, 1994; Wolinsky, 1996). The components of active replication complexes required for different viral R N A species have not been characterized, nor have the roles of p200, pl50 and p90 in this process been studied. The synthetic process of different viral R N A species and the underlying regulatory mechanism are yet to be studied.  The purpose of this thesis is to characterize R V NSPs and viral R N A synthesis at the molecular level with the following specific objectives: (1) characterization of the domains involved in cisand frans-cleavage activities of R V NS-pro and comparative analysis of NS-pro with papain in both primary and secondary structures; (2) investigation of the effects of NSP cleavage on virus  33  replication and viral RNA synthesis; (3) molecular characterization of viral RNA synthesis and its regulatory mechanism.  To characterize the related domains for NS-pro, a panel of in-frame deletion mutants from either the N - or C - terminus of the pi50 coding region were constructed and their respective enzyme activities examined in vitro. Comparative analysis of R V NS-pro to papain were made using the ALIGN program for primary sequence analysis and the E M B L protein structure prediction service for secondary structure.  To study the effect of NSP cleavage on virus replication and viral R N A synthesis, site-directed mutations were introduced into an infectious cDNA clone at the protease catalytic site and around the cleavage site. The constructed NSP cleavage mutants were examined for NSP processing efficiency and the levels of virus replication and viral RNA synthesis.  To characterize R V RNA synthesis and regulatory mechanisms, the time courses of three viral RNAs were determined. Trans-complementation experiments were conducted to examine the respective roles of p200, pi50, and p90 in synthesis of distinct viral R N A species.  These experiments lead to a better understanding of the structure and functions of R V NSPs and the process of R V R N A replication at the molecular level. Knowledge on this aspect not only contributes to the clarification of R V propagation and pathogenesis, but also provides a comparative model for studies on related positive-strand RNA viruses.  34  3. MATERIALS AND METHODS  3.1.  Materials and supplies  D N A modifying enzymes and restriction endonucleases were purchased from Bethesda Research Laboratories (BRL), Promega, New England Biolabs, Boehringer Mannheim, Sigma, Pharmacia and United States Biochemical Corporation. All enzymes were used as specified by the manufacturer unless indicated otherwise. L-[ S]methionine (10 mCi/ml) and oc-[ S]-CTP 35  35  (12.5 mCi/ml) were from New England Nuclear (NEN). Tissue culture reagents were from Gibco B R L . G E N E C L E A N (BiolOl) was from Promega. Peptide antibodies raised against peptides located within R V NSPs were generated in this lab. Human polyclonal anti-rubella serum was provided by Dr A. Tingle (B.C. Children's Hospital, Vancouver, B . C ) . BHK-21, Vero, and R V (M33 strain) were obtained from the American Type Culture Collection (ATCC).  3.2. Methods  3.2.1.  Growth of cells and viruses  Vero cells were cultured in Eagle's rninimum essential medium ( M E M , Gibco B R L ) supplemented with 5% fetal calf serum (FCS). BHK-21 cells were grown in M E M containing 10% FCS and 10% tryptose phosphate broth. R V (M33 strain) was propagated in Vero cells.  35  3.2.2.  Propagation of bacterial strains  E. coli strain WM1100 was used for the propagation of recombinant clones. WM1100 containing recombinant plasmids were grown in 2 x Y T medium (16 g/1 tryptone; 10 g/1 Yeast extract; 5 g/1 NaCl) containing 100 U-g/ml ampicillin for selection of antibiotic resistance.  3.2.3.  Preparation of competent cells and transformation  E. coli cells (1 ml) grown overnight were transferred into 100 ml of Psi broth (5 g/1 Yeast extract; 20 g/1 tryptone; 5 g/1 magnesium sulfate, pH7.6) and incubated at 37 °C until O D  5 5 0  reached 0.48. Cells were incubated on ice for 15 min and pelleted by centrifuge at 5000 rpm at 4 °C for 5 min. The bacterial pellet was suspended in 40 ml of Tfbl (30 mM potassium acetate; 100 mM rubidium chloride; 10 m M calcium chloride; 50 m M manganese chloride; 15% (v/v) glycerol, pH5.8) and incubated on ice for 15 min. Cells were pelleted, resuspended in 4 ml of Tfbll (10 mM MOPS; 75 mM calcium chloride; 10 mM rubidium chloride; 15% (v/v) glycerol, pH6.5). Aliquots of prepared competent cells were stored at -70 °C.  For transformation, 50 u.1 of competent cells were incubated on ice with plasmid D N A or 5 |il of ligation reaction for 30 min. After a 90-sec heat shock at 42 °C, the transformation mixture was added 1 ml of 2 x Y T medium and recovered at 37 °C by gentle shaking before plating onto selective plates.  3.2.4.  Mini preparation of plasmid D N A  Plasmid D N A mini-prep was performed by QIAprep Spin Plasmid Kit. Pelleted bacterial cells grown overnight were suspended in 250 u,l of Buffer PI. Cells were lysed by adding 250 u.1 of Buffer P2 with gentle inversion. Chromosomal D N A and proteins were precipitated by 350 ul 36  of Buffer P3 added and thoroughly mixed. The solution was centrifuged at 12000 rpm for 5 min, and the supernatant was applied to QIAprep spin columns. The columns were centrifuged briefly and the flow-throughs were discarded. The columns were washed by 0.5 ml of Buffer PB, followed by 0.75 ml of Buffer PE. The D N A was eluted by 100 ul of H 0 and collected by 2  centrifuging for 1 min. Buffers PI, P2, P3, PB, and PE were reagents provided by the kit.  A quick mini preparation of plasmids for screening by restriction enzyme digestion was done as follows. Colonies were inoculated in 3 ml of 2 x Y T medium and grown overnight at 37 °C. Take 0.2 ml of ceU culture, add 0.2 ml of solution II (1% SDS; 0.2 N NaOH) and mix gently. Add 0.2 ml of solution III (3M potassium acetate, pH5.5) and mix well. The reaction mixture was centrifuged at 14000 rpm for 2 min and the supernatant was transferred into 0.5 ml of isopropanol. The precipitated plasmids were collected by centrifuging at 14000 rpm for 1 min. After discarding supernatants completely, the pellet was resuspended by mixing (vortex) in 50 ul of T E buffer (10 mM Tris-HCI, pH 8.0; 1 mM EDTA) containing 10 ug/ml of RNase A. Use 5 \i\ for restriction enzyme digestion.  3.2.5.  Restriction endonuclease digestions and D N A modifications  All restriction digestion reactions were performed according to assay conditions specified by the suppliers.  D N A fragments were ligated using T4 D N A ligase in 50 mM Tris-HCI (pH7.6), 10 mM MgCl , 2  1 mM ATP, 1 mM D T T , and 5% (v/v) P E G at 16 °C overnight.  D N A fragments with 5' overhangs were blunt ended with E. coli D N A polymerase I Klenow fragment in the same restriction enzyme digestion buffer and incubated for 30 min at RT. 37  D N A fragments with 3' overhangs were blunt ended with T4 D N A polymerase in the presence of 2 mM dNTPs in the same restriction enzyme digestion buffer and incubated for 20 min at 16 °C.  Removal of terminal 5' phosphates from D N A fragments was done using calf intestinal alkaline phosphatase (CIP) in 50 mM Tris-HCI (pH 9.0), 1 mM M g C l , 0.1 mM ZnCl , and 1 2  2  mM spermidine for 30 min at 37 °C (for fragments with 5' overhangs), or for 30 min at 37 °C followed by 45 min at 55 °C with additional CIP (for fragments with blunt ends or 3' overhangs).  CIP reactions were terminated by phenol/chloroform extraction and D N A  fragments were ethanol precipitated or recovered by G E N E C L E A N (BIO 101).  Purification of D N A fragments from agarose gels was done using G E N E C L E A N (BIO 101). Desired fragments were excised from ethidium bromide stained agarose gels and the gel matrix was solubilized in 3 volumes of saturated sodium iodide at 55 °C for 5 min. Glassmilk was incubated with agarose solution at 55 °C for another 5 min. The formed DNA-glass-bead complex  was  pelleted  by  brief  centrifuging  and  washed  three  times  with  cold  NaCl/ethanol/water solution. The D N A was eluted from the glass beads with H 0 by 2  incubating at 55 °C for 3 min.  3.2.6.  Polymerase Chain Reaction  PCR reactions were carried out in 25 cycles of 98°C for 30 s, 50°C for 2 min, and 70 °C for 2 min using either 2.5 U of ExTaq temperature-stable D N A polymerase (TaKaRa L A PCR kit) or Native Pfu D N A polymerase (Strategene) in buffers provided by the manufacturers (10 mM 38  KC1, 10 mM (NH4) S0 ; 20 mM Tris-Cl, pH 8.75; 2 mM M g S 0 ; 0.1% TritonX-100; 100 2  4  4  jiig/ml BSA) and supplemented with 8% dimethyl sulfoxide (DMSO). The resulting PCR fragments were purified with a QIAquick Spin PCR purification kit (QIAgen).  3.2.7.  Plasmid Construction.  An R V infectious cDNA clone (pBRM33) (Yao and Gillam, 1999) was used in the plasmid construction. Site-directed mutations and deletions were accomplished by PCR using primers that contain designed substitutions and restriction enzyme sites. All primer sequences and relative positions on the R V genome are given in Table 1.  39  Table 1. Sequence information on oligonucleotide primers used in this work.  Primers  Pola rity a  Positions'*  Amino acid changes  PCR 5' and 3' primers 2770-2790 + JSY-13  JSY-12 JSY-16 JSY-7  -  -  -  JSY-5 YL-9 YL-10 YL-15 YL-16 YL-13 YL-14 YL-17 YL-18  -  + -  + + -  + +  5' ATCCATGGCGTACTATAGCGAGCGCGT 3' 5* ATATCCATGGACCCACCGCCT 3' 5'ATTCCCATGGTCGCGCTAGCCGCC 3' 5' ATTCCATGGCCACGCTGACGCACGCC 3' 5' ATATCCATGGCGACCCCCCTCGGGGAT-3' 5' ATATCCATGGGCATGTGCGGGAGTGAC-3' 5' ATTAGGCCTTAGTGGGGGCGGTCCGAGAC 3' 5'ATTAGGCCTTAGACCGCGAGCCAAAGGTG 3' 5' ATTAGGCCTTAGGGGACCGCGAGCCA 3' 5' ATTAGGCCTTACAGGGGGACCGCGAG 3' 5' ATTAGGCCTTACCGAGACAGGGGGACCGC 3'  3893-3910 3908-3925 3914-3928 3917-3931 3920-3937  Mutagenic primers + 3482-3508 JSY-2 3482-3508 JSY-3 3929-3955 + JSY-4  3929-3955 3929-3961 3929-3961 3920-3952 3920-3952 3920-3952 3920-3952 3941-3955 3948-3958  0  5' GCTGCTCGAGCGCGCCTACCG 3' 5' GTAGGTGGCGGCGTTCTTGAT 3' 5' GGTGGGCGGGGTGGCGGTAGA 3' 5' GCTTCGCTCAGGGCGCG 3'  4220-4240 1737-1757 3584-3600  N-terminal deletion primers + 1082-1101 YL-11 + 2518-2533 YL-30 2798-2812 + JSY-25 + 2960-2977 YL-25 + 3098-3115 YL-21 + 3344-3361 YL-22 C-terminal deletion primers  YL-23 YL-26 YL-28 YL-27 YL-24  nucleotide sequences  C1152S C1152S G1301S G1301S G1302stop G1302stop R1299A R1299A G1300A G1301A  5' GACCCAAACACCAGCTGGCTCCGCGCC 3' 5' GGCGCGGAGCCAGCTGGTGTTTGGGTC 3' 5' CTGTCTCGGGGCAGCGGCACTTGTGCC 3' 5' GGCACAAGTGCCGCTGCCCCGAGACAG 3' 5'CTGTCTCGGGGCGGCTAAACTTGTGCCGCCACC3' 5'GGTGGCGGCACAAGTTTAGCCGCCCCGAGACA3' 5'GCGGTCCCCCTGTCTGCAGGCGGCGGCACTTGT3' 5'ACAAGTGCCGCCGCCTGCAGACAGGGGGACCGC3' 5' GCGGTCCCCCTGTCTAGAGGCGGCGGCACTTGT 3' 5' ACAAGTGCCGCCGCCTCTAGACAGGGGGACCGC 3' 5' CTGTCTAGAGCAGGCGGCACTTGTGCC 3' 5' CTGTCTAGAGGCGCAGGCACTTGTGCCGCC 3'  polarity of primers on M33 genome. +, forward; -, backward. Positions of primers on M33 genome. Sequences of primers. The mutated nucleotides are in boldface. The restriction enzyme Xba I sites are underlined. a  b  c  40  Truncated plasmids containing in-frame deletions from either the N- or C- terminus of pl50 A series of in-frame deletions from either the N - or C-terminus of the pi50 coding region were generated by amplifying the corresponding D N A fragment from pBR-150 (containing a stop codon between pi50 and p90) using available restriction sites in R V cDNA and the Nco I site containing the initiation codon of p200. The relative positions of PCR primers on R V NSP ORF are schematically shown in Fig. 6.  RV N S P 1  348  827  920  1301  1020  974  1102  2116  129rTl295 ^296^1297 1299  YL-11 YL-30 JSY-25 YL-25 YL-21 YL-22 YL-23 YL-26 YL-28 -4 YL-23  Figure 6. Locations of PCR primers for serial deletions on RV genome. R V NSP ORF encodes a 200-kDa polyprotein (p200) to be processed into pi50 (residues 1 to 1301) and p90 (residues 1302 to 2116). PCR primers used for making deletions from either end of pi50 are shown by arrows, suggesting their polarities and relative amino acid positions on NSP ORF.  41  To make deletions from the N-terminus of pi50, amplification reactions were carried out on pBR-150 D N A template  using individual N-terminal deletion primers paired with an  appropriate downstream 3'-end primer (Table 1 and Fig. 6). The N-terminal deletion primers carry an initiation codon A T G (Nco I site) followed by a string of nucleotide sequences downstream from nt 1082, 2518, 2798, 2960, 3098, or 3344, respectively (the start nt position for A  3 4 8  ,  M 7 , V920, A974, A1020 8 2  or G1102, respectively). In brief, to clone the construct encoding  residues from A348 to G1301, primer YL-11  was paired with JSY-16 in PCR and the resulting  675-bp product was used to replace the Nco l-Not I fragment (nt 39 to 1686) of pBR-150 to make  PBR-A348/G1301. To  clone a construct encoding residues from Mg27 to  G1301,  primer YL-30  was paired with JSY-7 in PCR and the resulting 1.1-kb fragment was used to replace the Nco ISph I fragment (nt 39 to 3391) of pBR-150 to generate pBR-lVWGnoi. To make the construct encoding sequences from  V 9 o , A974, A1020 2  or G1102 to  G1301,  a primer JSY-25, YL-25, YL-21 or  YL-22 was respectively paired with JSY-12 in individual PCR and the resulting product was used to substitute the Nco I-Nco I fragment (nt 39 to 4023) of pBR-150 in the correct orientation. The constructed plasmids were named A1020/G1301  PBR-V920/G1301, PBR-A974/G1301,  pBR-  and PBR-G1102/G1301, respectively.  To make deletions from.the C-terminus of pi50, 5'- PCR primer JSY-25 was paired with each C-terminal deletion primer (YL-23, YL-24, YL-26, YL-27 or YL-28) in individual PCR (Table 1 and Fig. 6). The C-terminal deletion primers were complementary to nucleotide sequences consisted of sequences encoding the desired C-terminal amino acid (H1290, R1299)  V1295, P1296, L1297, or  plus its upstream residues, followed by stop codon T A A and the Stu I restriction site.  Each of the five PCR fragments was subsequently used to replace the Nco l-Stu I fragment (nt 39 to 6965) of pBR-150 to generate plasmid: pBR-V o/Hi29o, PBR-V920/V1295, pBR-V o/Pi296, 92  92  PBR-V920/L1297 or pBR-V o/Ri29992  42  To make protease constructs encoding sequences from nested N-termini (V920, A974, A1020 and G1102) to I1773, amplifications using N-terminal deletion primers (JSY-25, YL-25, YL-21, and YL-22) and the subsequent cloning were described as above for making N-terminal deletions of pi50, except that pBR-NSP (encoding R V wild-type NSP, without a stop codon between pi50 and p90) rather than pBR-150 was used for both PCR template and cloning vector. The resultant plasmids were further modified to remove the C-terminal half of p90 sequence by Bgl II (nt 5355) / Stu I (nt 9336) digestion, end-gap-filling and religation. The derived ORFs, being terminated by a stop codon at nt 9466, encode protein products starting from residue V o, A 92  9 7 4  ,  A1020 or G1102 to I1773 of NSP sequence, followed by a 43-amino-acid sequence resulted from the shift in reading frame after deletion from nt 5355 to 9336. The plasmids were named pBRV o/Ii773, PBR-A974/I1773, pBR-Ai 2(/Ii773 and pBR-Gno2/Ii773- All protease constructs are 92  0  schematically shown in Fig. 11.  Site-directed mutations C1152S, G1301S, R 1299A, G1300A, and G1301A A panel of site-directed mutations was introduced into pBRM33 by PCR-mediated mutagenesis with primers containing the desired nucleotide changes (all of the primer sequences are given in Table 1).  To substitute S for catalytic C n , to mutate G i i to S, or to change R1299 to A, fusion PCR 5 2  3 0  (Yao and Gillam, 1999) was employed with pBRM33 D N A as the template and two pairs of primers (schematically shown in Fig. 7). In brief, to make C1152S mutation, two pairs of primers, JSY-13 plus JSY-3 and JSY-2 plus JSY-12, were used in two PCRs to generate two products of 738-bp and 758-bp, respectively; the two partially overlapping PCR products were  43  annealed to serve as the template for amplification of the 1.47-kb fragment using JSY-13 and JSY-12. To make G1301S mutation, two pairs of primers, JSY-13 plus JSY-5 and JSY-4 plus JSY-12, were used in two PCRs to generate products of 1.19-kb and 311-bp, respectively; the two partially overlapping PCR products were annealed to serve as the template for amplification of the 1.47-kb fragment containing the G1301S mutation using JSY-13 and JSY12. To construct the R1299A mutation, two pairs of primers, JSY-13 plus YL-16 and YL-15 plus JSY-12, were used in two PCRs to produce 1.19-kb and 311-bp, which were annealed for amplification of the 1.47-kb containing the R1299A mutation using JSY-13 and JSY-12. In the end, the resulting 1.47-kb PCR products containing desired mutations (C1152S, G1301S, and R1299A) were cut with Nhe I and EcoR V and inserted into pBRM33 (minus the Nhe HEcoR V fragment) (Fig. 7) to generate pBRM33(C1152S), pBRM33(G1301S) and pBRM33(R1299A), respectively.  To facilitate mutagenesis, a silent mutation was introduced into pBRM33 to create a new Xba I site by changing C G G to A G A at nt 3935 to 3937. Fusion PCR was employed using pBRM33 D N A as the template and two paired primers, JSY-13 plus YL-14 and YL-13 plus JSY-12. The PCR product was used to replace the Nhe 1-EcoR V fragment (nt 2803 to 4213) of pBRM33, and the resultant construct was named pBRM33-X. To construct the G1300A and G1301A mutations, PCR amplifications were performed using pBRM33-X as the template and mutagenic primers containing the desired mutations: YL-17 for mutation G1300A and YL-18 for mutation G1301A. The PCR products were used to replace the corresponding Xba 1-EcoR V (nt 3933 to 4213) fragment of pBRM33-X. The constructs were named pBRM33(G1300A) and pBRM33(G1301A), respectively.  44  A/he I  J.  5'. 2803 JSY-13  •  3495  (Xba I)  EcoRV  3935 3945  4213  JSY-2  JSY-4  JSY-3  JSY-5  pBRM33  (pBRM33-X)  G1301S  C1152S YL-9  G1302stop YL-10 YL-15  R1299A YL-16 YL-13 YL-14 YL-17 YL-18  Xbal silent mutation — • G1300A — • G1301A JSY-12  Figure 7. Locations of PCR primers for site-directed mutagenesis on RV genome. Fusion PCR employed pBRM33 D N A as the template and two pairs of primers. For the C1152S mutation, paired primers, JSY-13 plus JSY-3 and JSY-2 plus JSY-12, were employed. For the G1301S mutation, paired primers, JSY-13 plus JSY-5 and JSY-4 plus JSY-12, were used. For the G1302stop mutation, paired primers, JSY-13 plus YL-10 and YL-9 plus JSY-12, were used. For the R1299A mutation, paired primers, JSY-13 plus YL-16 and YL-15 plus JSY12, were used. To create a new Xba I site at nt 3935, paired primers, JSY-13 plus YL-14 and YL-13 plus JSY-12, were used. The amplified 1.47-kb fragments containing desired mutations were replaced back into the pBR-NSP using Nhe I and EcoR V sites indicated above the genome. To make the G1300A and G1301A mutations, pBRM33-X (containing a new Xba I site at nt 3935) was used as the template in PCRs, with mutagenic primers YL-17 and YL-18 paired respectively with JSY-12. All primers are shown as arrows indicating their polarities and nucleotide positions on R V genome RNA (numbered from the 5' end of M33 genome). Primer sequences are given in Table 1.  45  Plasmid pBR-150 To change the G1302 codon to stop codon T A A , fusion PCR was employed using pBRM33 D N A as the template and two paired primers, JSY-13 plus YL-10 and YL-9 plus JSY-12 with the products of 1.19-kb and 311-bp, respectively. The two partially overlapping PCR products were annealed to serve as the template for amplification of the 1.47-kb fragment containing the G1302stop mutation using JSY-13 and JSY-12. The resulting 1.47-kb PCR product containing the G1302stop mutation was cut with Nhe I and EcoR V and inserted into pBRM33 (minus the Nhe HEcoR V fragment) (Fig. 7) to produce pBR-150.  pBRM33ASS, pBRM33AMM, and pBRM33(C1152S)ASS Plasmid pBRM33ASS was generated from pBRM33 by removing most of the structural protein ORF with the deletion from nt 6966 to 9336 after StuI digestion and reliagation. Its derived RNA, M33ASS, was a structural-protein-deleted R V R N A replicon. Construct pBRM33AMM, encoding a NSP-deleted R V RNA, was generated by the deletion of fragment between nt 1081 and  5106  from  pBRM33  using  MM  sites.  pBRM33(C1152S)ASS, encoding  the  M33(C1152S)ASS RNA, was constructed by deleting fragment between nt 6966 and 9336 from pBRM33(Cl 152S) using StuI sites.  3.2.8.  In vitro transcription.  The cDNA clones were linearized at the unique Hind III site and transcribed with SP6 R N A polymerase (Promega) in 40mM Tris-HCI (pH 7.9), 6mM MgC12, lOmM D T T , lOmM NaCl, 2mM spermidine, 0.05% Tween-20, 0.5mM each of NTPs, 1 m M cap analog 7mG5' 5'G, and PPP  1 U RNasin Ribonuclease Inhibitor. The reaction mixture was incubated at 37 °C for 2 h. R N A  46  transcripts were extracted once with phenol/chloroform, precipitated with ethanol and resuspended in H2O.  3.2.9.  In vitro translation  In vitro translation was performed according to the manufacture's (Promega) protocol in 50-u.l reaction mixtures containing 35 ui of nuclease-treated rabbit reticulocyte lysates, 20 mM amino acid mixture minus methionine, 0.4 TJ RNasin (ribonuclease inhibitor) and in vitro R N A transcripts, in the presence of either 400 uCi/ml of [ S]methionine (NEN) or 20 p-g/ml of cold 35  methionine. Translation reactions were carried out at 30 °C for desired times and terminated by the addition of SDS-PAGE loading buffer (62.5 mM Tris-HCI, pH7.4; 2% SDS; 5% pmercaptoethanol; 0.2% bromophenol Blue Dye; 500 mM sucrose). Radiolabeled proteins were visualized by fluorescence  autoradiography after SDS-polyacrylamide gel  electrophoresis  (PAGE) analysis.  In vitro translation using T N T Quick coupled transcription-translation system (Promega) was performed in 50 ul of reaction mixtures containing 40-ul T N T Quick Master Mix, 400 uCi/ml of [ S]methionine (NEN), 0.4 U RNasin (ribonuclease inhibitor), and 1 u,g of plasmid DNA. 35  Reactions were carried out at 30 °C for desired times.  3.2.10.  Vero cells transfected by RNA transcripts using Lipofectin.  About 20 (xg of in vitro transcribed R N A (in 20-u.l transcription reaction) and 10 p.g of Lipofectin (Gibco BRL) were suspended in 0.5 ml of FCS-free M E M and incubated for 20 min at RT. The formed Lipofectin-RNA mixtures were applied to a 35-mm-diameter dish of Vero 47  cell monolayer, which had been washed with FCS-free M E M twice. After incubation for 2 to 3 h at 37°C, the mixtures were removed and replaced with the culture medium. At day 6 posttransfection, culture fluids were harvested and the virus released into the culture medium was quantitated by plaque assay on Vero cells.  3.2.11.  BHK-21 cells transfected by RNA transcripts using electroporation  BHK-21 cells were harvested by trypsin treatment and washed twice with cold PBS (145 mM NaCl; 7 mM Na HP04; 3 mM NaH P04, pH7.0) and resuspended at a concentration of 10  7  2  2  cells/ml. 0.5 ml of cell suspension was mixed with about 20 u.g of in vitro transcribed RNA (in 20-ul transcription reaction), and transferred to a 2-mm-diameter cuvette. Electroporation utilized two consecutive 1.5-kV, 250 uF pulses with a Gene-Pulser (Bio-Rad). The cells were diluted with the culture medium, and distributed into 4 x 35-mm-diameter dishes. Culture fluids were collected at 48 h postelectroporation and the released virus particles were quantitated by plaque assay on Vero cells.  3.2.12.  Plaque assay and virus growth analysis.  For viral plaque assay, Vero cells infected by a serially diluted virus stock were overlaid with 0.5% agarose in M E M containing 5% FCS, incubated at 35°C for 6 or 8 days, and stained with 5% neutral red diluted in M E M supplemented with 5% FCS. For virus growth rate analysis, Vero cell monolayers (35-mm-diameter dish) were transfected with the W T or mutant RNAs mediated by Lipofectin as described above. After removing the RNA-Lipofectin mixtures, the cells were washed with PBS, overlaid with fresh medium and incubated at 37 °C. The culture  48  medium was harvested and replaced with fresh medium every 24 h. The released virus was quantitated by plaque assay.  3.2.13.  Metabolic labeling  Labeling of Vero cells transfected with R N A was performed according to Hobman and Gillam (1989). 6 days posttransfection, RNA-transfected Vero cells (in 35-mm-diameter dishes) were washed twice with PBS, incubated with methionine-deficient D M E M for 30 min, and labeled for 2 h with 0.5 ml of methionine-deficient D M E M containing 100 uCi [ S]methionine and 35  5% FCS dialyzed against PBS. Cells were washed with cold Tris-saline and lysed with 500 ul of lysate buffer (1% Triton X-100; 1 mM E D T A ; 50 mM Tris-HCI, pH7.5; 0.15 M NaCl). After incubating on ice for 5 min, lysates were scraped off the plates and centrifuged at 4 °C for 5 min at 13000 rpm to remove the nuclei and debris. The supernatants were subjected to immunoprecipitation.  3.2.14.  Immunoprecipitation  Cell lysates diluted with lysate buffer (1% Triton X-100; 1 mM E D T A ; 50 mM Tris-HCI, pH7.5; 0.15 M NaCl) were mixed with Human polyclonal anti-rubella serum for at least 2 h at RT with constant rotation. Protein A-Sepharose beads (Pharmacia) were washed three times with lysate buffer, and mixed with the serum-lysates overnight at 4 °C with constant rotation. The beads were washed three times with lysate buffer. Antigen-antibody complexes were dissociated from the Protein A-Sepharose by boiling in 1 x SDS dissociation buffer (62.5 mM Tris-HCI, pH6.8; 10% glycerol; 2% SDS; 2% p-mercaptoethanol)  for 5 min, and the  supernatants were used in SDS-PAGE analysis.  49  3.2.15.  Total R N A preparation  Total cytoplasmic R N A was prepared from cell cultures using TRIzol reagent (GIBCO BRL). Cells cultured in 35-mm-diameter dish were lysed by 0.8 ml of TRIzol reagent and the lysates were passed through a pipette several times and transferred to a 1.5-ml microtube. The samples were incubated at R T for 5 min before 160 u,l of chloroform were added and mixed well by vigorous shaking for 15 s. The samples were incubated at R T for another 3 min and centrifuged at 14000 rpm for 15 min at 4 °C. The aqueous phase was transferred to 0.4 ml of isopropanol. RNA was precipitated by incubating at R T for 10 min and collected by centrifuging at 14000 rpm for 10 min at 4 °C. The pelleted R N A was washed with 1 ml of 75% ethanol and briefly dried before dissolving with formamide. R N A was quantitated by measuring A 6o (one A 6o 2  2  unit equals 40 U,g/ml RNA).  3.2.16.  RNase protection assay (RPA).  RPA was employed to analyze the synthesis of viral-specific RNAs during virus replication. For synthesis of plus- or minus-polarity RNA probe in vitro, a D N A fragment (nt 6323 to 6623) of pBRM33, representing the region covering the subgenomic R N A initiation site (nt 6436), was separately cloned into vector pSPT18 or pSPT19 (Pharmacia Biotech) at the EcoR I and Xba I sites to make construct pSPT-pbl8 or pSPT-pbl9. A 328-bp minus-polarity R N A probe (pbl8), synthesized with SP6 R N A polymerase from EcoR I-linearized pSPT-pbl8, can protect 301-nt positive-strand genomic R N A and the 188-nt subgenomic RNA. A 328-nt plus-polarity RNA probe (pbl9), synthesized with SP6 R N A polymerase from Hind Ill-linearized pSPTpbl9, can protect 301-nt R V negative-strand genomic RNA. For synthesis of another pair of plus- and minus-polarity R N A probes in vitro, a D N A fragment (nt 9175 to 9336) of pBRM33, in the E l coding region, was cloned into vectors pSPT19 and 50  pSPT18 respectively at the Hind III and Sma I sites to make constructs pSPT-pb20 and pSPTpb21. A 187-nt minus-polarity R N A probe (pb20), synthesized with SP6 R N A polymerase from Hind Ill-linearized pSPT-pb20, can protect 162-nt positive-strand R N A (including both genomic and subgenomic RNA). A 187-nt plus-polarity R N A probe (pb21), synthesized with SP6 R N A polymerase from EcoR I-linearized pSPT-pb21, can protect 162-nt negative-strand RNA.  6323  3'^  6623 • • 301 nt  9174 9336  • 5' (-) RV RNA  163 nt  163 nt 163 nt  • 3 ' (+) gRNA • 3 ' (+) sgRNA 6436  Figure 8. Probes used in RNase protection assay (RPA) and the relative positions and lengths of the protected regions. The plus-polarity probe pbl9 protects a 301-nt region from nt 6323 to nt 6623 of negativestrand R V RNA. The minus-polarity probe pbl8, complementary to pbl9, protects a 301-nt region of positive-strand genomic R N A and a 188-nt region from nt 6436 to nt 6623 of subgenomic RNA. The plus-polarity probe pb21 protects a 162-nt region from nt 9175 to nt 9336 of negative-strand R V RNA, and its counterpart minus-polarity probe pb20 protects a 162-nt region of both genomic and subgenomic RNAs.  The  35  S-labeled R N A probe was synthesized with SP6 polymerase in a 20-/xl in vitro  transcription reaction mixture containing buffer (40mM Tris-HCI, pH 7.9; 6mM MgCl ; lOmM 2  51  D T T ; lOmM NaCl; 2mM spermidine; 0.05% Tween-20), RNasin, 0.5 mM each of ATP, GTP and UTP, 12.5 /xM of CTP and 2.5 mCi/ml of [a- S]CTP (NEN). 35  After incubation for 90 min  at 37 °C, 2-/xl DNase I (7500 U/ml, Pharmacia Biotech) was added to the reaction mixture and incubated  for  a  further  15  min.  The  probe  was  precipitated  with  ethanol  after  phenol/chloroform extraction and resuspended in H 2 O .  For RNA analysis, total cytoplasmic RNAs were extracted with TRIzol reagent (GIBCO BRL) at indicated times posttransfection. Negative-strand R N A was analyzed by a two-cycle RPA essentially as described by Novak and Kirkegaard (1991). Approximately 20 fig of cytoplasmic R N A was incubated with 20 ng of unlabeled probe pbl9 (or pb21) (approximately 10  11  molecules) in 30 jul of hybridization buffer (40 mM PIPES; 400 mM NaCl; 1 mM E D T A ; 80% deionized formamide, pH6.4) overnight at 55 °C. RNase digestion was for 60 min at 30 °C in RNase mixture (300 mM sodium acetate; 10 mM Tris-HCI, pH 7.5; 5 mM E D T A ; 10 jUg/ml of RNase A; 70 U/ml of RNase T l ) . The reaction mixture was treated with SDS-proteinase K for 15 min at 37 °C, extracted with phenol-chloroform, and ethanol precipitated with 5 jit g/ml of tRNA. Samples were resuspended in 30 \x\ of hybridization buffer, and lx 10 cpm of  S-  labeled R N A probe (pbl9 or pb21) was added. The samples were denatured at 95 °C for 5 min, hybridized overnight at 55 °C, and subjected to RNase treatment as described above. The digestion products were resuspended in loading buffer (80% (v/v) formamide; 0.1% (v/v) xylene cyanol; 0.1% (v/v) bromophenol blue; and 2 mM E D T A ) and analyzed on a 5% polyacrylamide-urea gel, which was fixed in 10% acetic acid, infiltrated with Enhancer (DuPont), dried and exposed to X-ray film.  52  Positive-strand genomic and subgenomic RNAs were analyzed by a single round of RPA. 2 ag of total cytoplasmic RNAs were hybridized with 1 x 10 cpm of S - labeled R N A probe (pbl8 6  35  or pb20) overnight at 55°C. The samples were treated as described above.  3.2.17.  3.2.15.1.  Electrophoresis  Separation of D N A fragments by agarose gel electrophoresis  The buffer used in agarose gel electrophoresis was 1 x T A E (40 mM Tri-acetate, pH8.0; 1 mM EDTA). The gel with concentration varied from 0.8 to 1% agarose was prepared using 1 x T A E buffer with 1 (xg/ml ethidium bromide for visualization. D N A samples were mixed with loading buffer (8% sucrose, 20 mM E D T A , pH8.0; 0.05% bromophenol blue; 0.05% xylene cyanol) and separated by electrophoresis on 10 cm horizontal agarose gels at 100 volt.  3.2.15.2.  Separation of short RNA fragments by urea-PAGE  Short R N A fragments (100 to 500 nt) in RPA were separated by 7 M urea-5% polyacrylamide gel  electrophoresis  (PAGE).  The gel  contained  5% polyacrylamide (acrylamide:N,N'-  methylenebisacrylamide = 19:1) and 7 M urea in the presence of T B E buffer (89 mM Tris; 89 mM boric acid; 2 mM E D T A ; pH8.0). Samples were dissolved in the loading buffer (80% formamide; 0.1% xylene cyanol; 0.1% bromophenol blue; 2 mM E D T A ) and boiled for 5 min before loading. Gels were run in 1 x T B E buffer at constant voltage of 200 volts until the markers moved to the desired positions. Gels were fixed in 10% acetic acid for 15 min, immersed in the fluorographic reagent Enhancer (DuPont) for 45 min, vacuum dried, and exposed to X-ray film at -70 °C.  53  3.2.15.3.  Separation of proteins by SDS-PAGE  Proteins were separated using a discontinuous gel system described by Laemmli (1970). Stacking  gels contained  4%  polyacrylamide (acrylamide:N,N'-methylenebisacrylamide =  30:0.8) in the stacking buffer (0.125 M Tris-HCI, pH6.8; 0.1% SDS). Separating gels contained either 8 or 10% polyacrylamide (19:1) in the separating buffer (0.375 M Tris-HCI, pH8.8; 0.1% SDS). Protein samples were dissolved in 1 x SDS-PAGE loading buffer (62.5 m M Tris-HCI, pH6.8; 10% glycerol; 2% SDS; 2% P-mercaptoethanol) and boiled for 3 min before loading. Gels were running in the buffer (0.025 M Tris-HCI, pH8.3; 0.192 M glycine; 0.1% SDS) at constant voltage of 100 volts until the markers have run to the desired positions. The separating gel was fixed in 10% acetic acid, immersed in the fluorographic reagent Enhancer (Dupont) or Amplify (Amersham) for 30 min, vacuum dried, and exposed to X-ray film at -70 °C.  3.2.18.  Image analysis and cleavage efficiency comparison.  Image analysis was performed on a PC computer using the Scion Image program for Windows (Beta 3b) (http://www.scioncorp.com/frames/fr_download_now.htm),  the PC version of the  public domain NIH Image program (developed at the U.S. National Institutes of Health and available on the Internet at http://rsb.info.nih.gov/nih-image/). The cleavage ratio at certain incubation times for each protease construct was calculated as the percentage of the quantity of the cleaved products over the total of remaining substrate and cleaved products. The cleavage ratio was plotted against the incubation time for each construct.  54  3.2.19.  Sequence analysis.  Initial alignment of primary sequences between papain (SWISS-PROT access number P00784) and catalytic region of R V NS-pro (M33 strain GenBank access number S38480 with corrections by Pugachev et al., 1997), Therien strain GenBank access number P13889 with corrections by Pugachev et ah, 1997) was done by ALIGN program (Myers and Miller, 1988) and modified manually. Secondary structure predictions were performed using the E M B L protein structure prediction service  (http://www.emblheidelberg.de/predictprotein/predictprotein.html).  The service was described by Rost et al. (1993a, 1993b, 1994a, 1994b).  55  4. R E S U L T S A N D D I S C U S S I O N S  4.1.  Characterization of domains involved in cis- and trans-cleavage activities of RV  pro  4.1.1.  Processing of RV NSP by in vitro translation.  The R V NS-pro is a papain-like cysteine protease (PCP) encoded in the NSP ORF that cleaves the NSP ORF translation product (p200) at a single site to produce pi50 and p90. Many viral PCPs were found to be active following in vitro translation using rabbit reticulocyte lysates (Bonilla et al., 1997; Choi et al., 1991a; Den Boon et al, 1995; Hardy and Strauss, 1989). I therefore monitored the activity of RV-pro after translating R V NSP in vitro using the genomic-length R N A transcripts synthesized from an R V infectious cDNA clone derived from RV strain M33 (pBRM33) (Yao and Gillam, 1999). pBRM33 was linearized with Hind III and full-length R N A transcripts were synthesized using SP6 R N A polymerase in the presence of cap analog. In vitro translation and processing of NSP were programmed using rabbit reticulocyte lysates with synthesized R N A transcripts. A time course experiment was performed to monitor the kinetics of p200 processing. Translation of p200 was completed after 40 min (Fig. 9A, lane 2); cleavage was observed at 60 min (Fig. 9A, lane 3) and continued efficiently (Fig. 9A, lanes 4 to 6). Liu et al. (1998) suggested that activity of R V NS-pro in vitro depended on the addition of Z n . However, in my in vitro translation system, addition of 2+  Zn  2+  was not found to be required for NS-pro activity, nor did it increase the processing  efficiency of R V NSP (Fig. 9A, comparing lanes 1 to 6 to lanes 7 to 12). The presence of Z n  2+  even seemed to have a certain inhibition on the NSP processing efficiency, which, however, may come from experimental variation without statistic significance. Furthermore, using the T N T Quick coupled transcription-translation system (Promega), I also observed efficient  56  NS-  processing of R V NSP without the addition of Z n  2+  findings of Liu et al. (1998) that addition of Z n  was essential for R V NS-pro activity in the  2+  (Fig. 9B, lanes 1 to 3), in contrast to the  same translation system. In addition, I observed the efficient processing of NSP from strain Therien in vitro without the addition of Z n , using infectious cDNA clone Robo302 (Pugachev 2+  et al., 1997) or its derived R N A in either T N T transcription-translation system (Fig. 9B, lanes 4 to 6) or rabbit reticulocyte lysate (Fig. 9B, lanes 7 to 9). Liu et al. (1998) used a different Therien strain cDNA construct, Robol02, and its derived subclones in their studies. However, Robol02 has a substantially lower infectivity than Robo302 (Pugachev et al., 1997b), which might account for the observed discrepancy. Nevertheless, the exact reasons behind these contradictions need further investigation.  A.  time (min)  kDa  rabbit reticulocyte lysate M33 ~  Z  n  +  Z  B.  n  20 40 60 90 120 180 20 40 60 90 120 180  I  l  l  *  *  TNT Quick coupled transcription-translation M33  Therien  90 1 20 1 80 90 120 1 80  ret  ^ulocyte lysate  Therien 60 120 180 time (min)  t.zz  i l l i ^ p i s o  : pi 50  ip ip ™  lanes 1 2 3 4 5 6 7 8 9 10 11 12  lanes 1 2 3 4 5 6  7 8 9  Figure 9. R V NSP processing in in vitro translation systems with or without addition of Zn . 2 +  (A) In vitro translation reactions using rabbit reticulocyte lysates were programmed with genome-length R V R N A of M33 strain (transcribed in vitro from pBRM33) in the absence (lanes 1 to 6) or presence (lanes 7 to 12) of 200 fiM Z n . Aliquots were removed at indicated times during incubation and subjected to SDS-8% polyacrylamide gel electrophoresis (PAGE) analysis. (B) In vitro translation reactions using T N T Quick coupled transcription-translation system were programmed with R V full-length infectious cDNA clones based on M33 strain (pBRM33, lanes 1 to 3) or Therien strain (Robo302, lanes 4 to 6). Genome-length R V RNA of Therien strain (transcribed in vitro from Robo302) was also translated in vitro using rabbit reticulocyte lysates (lanes 7 to 9). All Reactions were carried out without the addition of Z n . Aliquots were removed at indicated times and subjected to SDS-8% P A G E analysis. Protein products were visualized by fluorescence autoradiography. Positions of molecular mass markers and cleavage products are indicated. Images were scanned using a U M A X Astra 1220U scanner with Adobe Photoshop 5.0 software. 2+  2+  57  It has been shown previously (Yao et ah, 1998) that NS-pro can function in trans in vivo by coexpression within BHK-21 cells of a construct p200(G1301S) that contains a cleavage site mutation (to serve as a protease) together with a construct p200(C1152S) that contains a protease mutation (to serve as a substrate). To demonstrate the NS-pro rrans-cleavage activity in vitro, three full-length mutants with alterations in the NSP ORF were constructed. pBR200(G1301S) is a cleavage site mutant carrying a G-to-S mutation at residue 1301; pBR200(C1152S), a protease-inactive mutant carrying a C-to-S mutation at its catalytic C1152 residue; and pBR-150 is a mutant carrying a stop codon corresponding to residue 1302.  To assay for frarcs-cleavage activity of NS-pro, two separate translation reactions were carried out. Radiolabeled p200(C1152S) was synthesized in vitro in the presence of [ S]methionine to 35  serve as a source of substrate for protease, and p200(G1301S) or pl50 was synthesized in the absence of [ S]methionine to serve as a source of protease. After a 1-h incubation at 30°C, 35  both in vitro translation reactions were terminated by the addition of RNase A and cycloheximide.  When  the  radiolabeled p200(C1152S)  was  added  to  the unlabeled  p200(G1301S) or pl50 translation reaction mixture, the cleavage products of pl50 and p90 were detected after a 1-h incubation (Fig. 10A and B), indicating that trans cleavage of p200(C1152S) catalyzed by p200(G1301S) or pl50 protease had occurred. No cleavage product was observed when p200(C1152S) was incubated alone for 5 h (Fig. 10C). For a control, p200(G1301S) or pl50 was synthesized in the presence of [ S]methionine at 30°C and 35  incubated alone for 5 h (Fig. 10A and B, lane 1).  58  B  A time(h) 5 0 1 2 3 4 kDa  5  c  5 0 1 2  3 4 5  0 1 2 3 4  5  »«»4tifi«-p200 «-pl50  200- ^ t R £ _ £ £ £  <-p90 1  lanes  1 2 3 4 5  2  3 4 5 6 7  1 2 3 4 5 6  6 7  Figure 10. R V p200(G1301S) and pl50 cleave substrate protein in trans. Protein products were labeled by synthesis in the presence of [ S]methionine or were unlabeled. After incubation at 30 °C for 1 h, translation was terminated by the addition of RNase A and cycloheximide to final concentrations of 1 and 0.6 mg/ml, respectively. Protein product p200(G1301S) or pl50 (synthesized in the absence of [ S]methionine) were mixed with radiolabeled substrate p200(C1152S) and incubated at 30 °C for up to 5 h. Samples removed at various times were subjected to SDS-PAGE analysis. (A) R V p200(C1152S) was labeled in the presence of [ S]methionine and incubated alone for 5 h (lane 1). After translation reactions were terminated, unlabeled p200(G1301S) was mixed with S-labeled substrate p200(C1152S) and incubated at 30 °C. Aliquots were removed from 0 to 5 h and subjected to SDS-PAGE analysis (lanes 2 to 7). (B) R V pl50 was labeled in the presence of [ S]methionine and incubated alone for 5 h (lane 1). After translation reactions were terminated, unlabeled pi50 was mixed with S labeled substrate p200(C1152S) and further incubated at 30 °C. Aliquots were removed from 0 to 5 h (lanes 2 to 7) and subjected to SDSPAGE analysis. (C) In vitro transcription and translation of p200(C1152S) were carried out as described. Samples were taken at the indicated times and subjected to SDS-PAGE analysis. Positions of molecular mass markers and cleavage products are indicated. Images were scanned using a U M A X Astra 1220U scanner with Adobe Photoshop 5.0 software. 35  35  35  35  35  35  4.1.2.  Construction of truncated NS-pro cDNA clones.  In order to identify the minimal domains required for functional protease activities, a panel of protease constructs was generated and expressed in vitro for protease activity analysis. The cDNA fragments encoding NS protease constructs were generated by PCR as described in section 3.2.7. The constructed plasmids were inserted downstream of the SP6 RNA polymerase promoter and R V 5' untranslated region (5-UTR). The poly(A) addition site of the R V fulllength cDNA clone (pBRM33) was preserved. The plasmids, protease products and their relative positions on the RV NSP ORF are shown in Fig. 11. 59  Plasmids  Protease constructs  pBR-NSP  p200  Schematics of protease ORFs c  1 |  H  cleavage ratio (%)  ^mmmm  1301  2  1  1  70 6  pBR-p200(G1301S)  pBRp200(G1301S)  pBR-p1 50  p150  pBR-A348/Gl301  A348/G1301  pBR-M827/Gi3oi  M827/G1 301  PBR-V920/G1301  V92o/Gl301  PBR-A974/G1301  A974/G1301  PBR-A1020/G1301  Al020/Gl301  PBR-G1102/G1301  Gl102/Gl301  PBR-V920/H1290  V920/H1290  con  I M M ^^ ^ ^ 1 2 9 0  0  PBR-V920/V1295  V920/V1295  920  tS^m  1295  0  PBR-V920/P1296  V920/Pl296  920  flHMNH  1296  8  PBR-V920/L1297  V920/L1297  Sa^SM^  1297  10  PBR-V920/Rl299  V920/Rl299  920 wasawwwS S S M M 1 2 9 9  PBR-V920/|1773  V920/I1773  9?n I--  PBR-A974/I1773  A974/ll773  PBR-Al020/Il773  Al020/ll773  pBR-Gl102/ll773  Gl102/ll773  1 1  1 •iiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiiniiiiii  iiiiinih iiiiinih - IIIIII  K  1 2116  83  1301  73  « ^ ^ i a n i B97  ISSMiSM^  70  1301  77  « M ^ 1 3 0 1  15  974 BSSSS^« M ^ 1 3 0 1  0  1020  0  H  1102  974  0  fSSPPM S M 3 1 3 0 1  i  1020 I 1102 |  5  1773  52  • • • 1773  53  1773  23  ^ M i 1773  0  Figure 11. Plasmids and protease-encoding constructs. All protease constructs were given a two-letter name; their lengths and relative positions on the RV NSP ORF are schematically shown, labeled with starting and ending protein residues (numbered according to NSP residues). The respective cleavage ratio of each construct, calculated by quantitating the cleavage products over total proteins, is given in percentage. Positions of the catalytic sites C1152 and H1273 on p200 are indicated by vertical solid lines.  60  All of the protease constructs are designated by a two-letter name indicating starting and ending amino acid positions.  A348/G1301, M 8 2 7 / G 1 3 0 1 , V  are protein fragments starting from extending to V920/R1299  G1301,  9 2 0  / G i 3 o i , A974/G1301, A i o / G i 3 o i 0 2  A348, M 8 7 , V 9 o , 2  2  A  9 7 4  the end of pl50. Fragments V920/H1290,  extend from  V920  to positions  ,  A1020,  and  and  G1102/G1301  respectively, and  G1102,  V 9 o / V i 5 , V920/P1296, V920/L1297, 2  H1290, V1295, P1296, L1297,  2 9  and  and Ri299, respectively. These  fragments did not contain a cleavage site and were examined for their rrarcs-cleavage capacity. Fragments and  V920/I1773, A974/I1773, A1020/I1773,  G1102,  respectively, to  I1773  and Gn /Ii773 extend from 02  A  3 4  8,  V  9 2 0  , A  9 7 4  ,  A1020,  of the NSP sequence. They contain cleavage sites and C-  terminal tails to be used for czs-cleavage analysis.  4.1.3.  Defining the NS-pro domain required for trans cleavage.  To determine the rrans-cleavage activity of the generated protease constructs, six protease constructs with nested N-terminal deletions A1020/G1301,  and  G1102/G1301)  were translated  (A348/G1301,  M827/G1301,  V920/G1301,  A974/G1301,  in vitro separately, producing protein products  with apparent molecular masses of 102, 50, 41, 35, 30, and 21 kDa, respectively (Fig. 12A, B, C, D, E and F, lane 1). Each was examined for trans protease activity analysis against substrate by cotranslation with p200(C1152S). In the cases of  A348/G1301, M827/G1301,  and  V920/G1301,  cleavage products pi50 and p90 were detected after a 1-h incubation and increased with incubation time, suggesting that these constructs possess trans cleavage activity (Fig. 12A, B and C, lanes 2 to 6). No detectable cleavage products (pl50 or p90) could be observed in the reactions of  A^JGnox,  Aio2o/Gi oi, 3  and  G1102/G1301  (Fig. 12D, E and F, lanes 2 to 6), suggesting  that they could not form active protease to cleave in trans. My data suggested that the domain containing active  trans protease starts at V920 or after, but at least upstream of  A 74. 9  61  A  time(h) 1 0  B  12 3 4 5  "°'  C  lime (h) 1 0 1 2 3 4 5  •>  -pl50  • 1 1 1 1 wKI  W Hr  68"  WWWW  1 2 3 4 5 6 7  3  1 2 3 4 5  ... , M827/G, 30,  lanes  3 43 5 01 2 1 2 4 5 P  z u u  2  lanes  A W G  -  §-  P  200  29 ~ ~  lanes  0  43-  4—  1 2 3 4 5 67  0  68  43 -  t l l H  1 2 3 4 5 67  97 •  6 8  f  *•• * V920/GI3OI  time (h) 1 0 1 2 3 4 5 kDa  fFPRPI  43—  ft mmm  F  time (h) 1 kDa  ~  f  4 3  i r  9 7  9 7  -p90  68-  1 2 3 4 5 6 7  E 10  —_  _ »  •fttt^plSO  97-  68-  lanes  D time (h) kDa  ^  • ••-p90  ' A348/G ! SOI -p90 4  lanes  ™  •••IP-P200  P200 ~ * g «  time (h) 1 0 1 2 3 4 5  1 2 3 4  "  <  A  ,  0  2  0  /  G  ,  3  0  5 6 7  - GIIM/GIXH  ,  lanes 1 2 3 4 5 6 7  Figure 12. / n v#ro translation of protease constructs (AmfGimi MszifGtwu Vsao&hraoij A974/G1301, A1020/G1301, and G1102/G1301) and examination of their frans-cleavage activities. A 48/Gi3oi 3  (A),  M 2 7 / G i 3 o i (B),  V920/G1301 (C),  8  A 7 4 / G i 3 o i (D), 9  A1020/G1301 (E),  and  G1102/G1301  (F) were translated individually to give 102-, 50-, 41-, 35-, 30-, and 21-kDa products (lanes 1). Cotranslation of each construct with substrate p200(C1152S) was carried out at 30 °C for 0 to 5 h. Samples were removed at each time point and subjected to SDS-PAGE analysis, (lanes 2 to 7). Positions of molecular mass markers and cleavage products are indicated. Images were scanned using a U M A X Astra 1220U scanner with Adobe Photoshop 5.0 software.  Five protease  constructs (V o/Hi29o, 92  V920/V1295,  V /Pi296, V o / L i 9 7 , 920  9 2  2  and  V920/R1299)  extending from V o to different C termini, were of the same molecular mass, 41 kDa, when 92  expressed in vitro (Fig. 13, lanes 1). As described above, each of them was examined for potential rrans-cleavage activity by cotranslation with p200(C1152S) for up to 5 h (Fig. 13). The appearance of cleavage products (pl50 and p90) in the reactions of V 2o/Pi296, 9  V920/L1297,  and V 2o/Ri299 demonstrated that they preserve protease activity (Fig. 13C, D and E , lanes 2 to 9  62  6). In contrast, neither V920/H1290 nor V920/V1295 was able to cleave substrate p200 (Fig. 13A and B, lanes 2 to 6). I therefore mapped the C terminus of the active R V NS-pro domain exactly to Pi 96- The weak frans-cleavage activity observed in V920/P1296, V 9 2 o / L i 9 7 , and 2  2  V920/R1299 could be due to the absence of the X domain in these constructs (see section 4.1.5. below).  A  B  time(h) 1 0 1 2 3 4 5 kDa _ JH 200 • • • § • -P20Q  C  time(h) 1 0 1 2 3 4 5 kDa  9 7  6843  "m„.  f»»«»-p200  200^  97-  time(h) 1 0 1 2 3 4 5 kDa  ~  |  .  97  68-  1 #» •M*M« -V - 9V 0/H 29 o2/H l2l2 9900  l m  6  43-ii  mm*mmm~v™rvno<  8  9  f  | f .- 150 m m - p90 P  "  4 3  A  * • A A A * VWP.**  29 -  lanes 1 2 3 4 5 6 7  lanes  D  lanes  1 2 3 4 5 6 7  E  time (h) kDa 200-  10 1 2 3 4 5  time (h) 1 0 1 2 3 4 5 kDa  n l l i : S  -  97 -  nr.  97 68  1 2 3 4 5 6 7  -  - 90  68  »V«20/Ll297  4 3  29lanes  y /  p  43  1 2 3 4 5 67  f  p200  t t t f l pl50 — p90 . *V920/R!299  lanes 1 2 3 4 5 6 7  Figure 13. In vitro translation of protease constructs (V920/H1290, V920/V1295, V920/P1296, V920/L1297, and V920/R1299) and examination of their frans-cleavage activities. V 2o/Hi29o (A), V 9 2 0 / V 1 2 9 5 (B), V o/Pi296 (C), V 9 2 0 / L 1 2 9 7 (D), and V 9 2 0 / R 1 2 9 9 (E) were translated m v/fro separately to yield a 40-kDa product (lanes 1). Cotranslation with substrate p200(C1152S) was carried out for 0 to 5 h (lanes 2 to 7), and samples were subjected to SDSP A G E analysis. Positions of molecular mass markers and cleavage products are indicated. Images were scanned using a U M A X Astra 1220U scanner with Adobe Photoshop 5.0 software. 9  92  63  4.1.4.  Domains required for cis cleavage.  R V NS-pro is known to possess both  trans- and ds-cleavage activities (Yao et al, 1998). The  cotranslation experiments described above identified the NS-pro domain required for trans cleavage within the fragment from V o to Pn96- This domain may or may not be the exact 92  domain required for cis cleavage. Since the C terminus of the cis protease construct must extend beyond the cleavage site, I therefore examined the N-terminal domain requirement for  cis cleavage by analysis of in vitro translation of the protease constructs Aio2o/Ii773>  V920/I1773, A974/I1773,  and Gi 102/11773- Processing of these protein constructs would accumulate a cleavage  product with an apparent molecular mass of 58 kDa (p58, the C-terminal fragment extending from residue 1302 to 1773) throughout the incubation time. I found that V920/I1773 and A974/I1773 underwent efficient processing. V920/I1773 was translated as a 98-kDa polyprotein, which was then cleaved into 58-and 41-kDa products within a 1-h incubation (Fig. 14A). A974/I1773 was translated as a 92-kDa product and cleaved into 58-kDa and 35-kDa products efficiently (Fig. 14B). Self-cleavage of A1020/I1773 (87 kDa) into 58-kDa and 29-kDa products was less efficient (Fig. 14C). The band above p29 seems to be a translation by-product rather than a cleavage product since its amount did not increase with time as the p29 band did. No cleavage could be observed in the case of G1102/I1773 (78 kDa) (Fig. 14D). Of these constructs, only V920/I1773 contains a protease domain (V920 to Gnoi) that can cleave in trans at low efficiency (Fig. 12C). Therefore, the highly efficient processing of V o/Ii773 represents cw-cleavage activity rather 92  than  trans-cleavage activity. The other two constructs,  the necessary domain (V o to R973) for 92  A974/I1773  and A1020/I1773, do not contain  trans cleavage and can function only in cis. To confirm  that constructs V920/I1773, A974/I1773, and A1020/I1773 function in  cis, a dilution experiment was  performed (Fig. 14E). Translation reactions with V920/I1773 (Fig. 14E, lanes 1 to 5),  A Jli 9Ji  in  (Fig. 14E, lanes 6 to 10), and A1020/I1773 (Fig. 14E, lanes 11 to 15) were carried out using serial  64  dilutions (0, 1:20, 1:100, 1:200, and 1:500) of each RNA transcript. For each of them, the total translation products decreased correspondingly when more-diluted R N A was added. However, cleavage products were clearly demonstrated, and the cleavage ratio (determined as in section 3.2.15.) remained roughly unchanged from that for nondiluted samples, suggesting that these cleavages occur in cis.  B time(h)0 1 2 3 4 5  time(h)0 1 2 3 4 5  kDa  kDa  97-  mmmjJk ^AvMm  •V920/L773 97_  68-  ^ p58  43-  lanes 1 2 3 4 5 6  D  •••••  68- ~ZSml^p5S 97-* p35  29-  29-  0 1 2 3 4 5  time(h) kDa  • A1020/11773 -p58  43 -  p29  29-  lanes 1 2 3 4 5 6  lanes  1 2 3 4 5 6  E  time(h)0 1 2 3 4 5 kDa 200-  200  V920/Il773 000 oSS^S  A974/Il773 _ooo o S ° S S  A1020/11773 oo © oSSSK  97  eg" — » - «  I G I W  r  m| f -* -P-> -P^I** S  _p41  43lanes  12 3 4 5 6  lanes 1 2 3 4 5  P  §#§ 6 7 8 9  Figure 14. Autolytic processing of protease constructs  10  5  •••• 8  -  g '-5 ~  -P58  1112131415  V920/I1773, A974/I1773, A1020/I1773,  and  Gll02/Il773-  V920/I1773 (A), A 74/Ii77 (B), A1020/I1773 (C), and G1102/I1773 (D) were transcribed and translated as described above for 5 h in the presence of [ S]methionine. Samples were removed at the indicated times and subjected to SDS-PAGE analysis. (A) V92o/Ii77 was translated as a 98-kDa protein and subsequently processed into 58- and 41-kDa fragments. Positions of V920/I1773, p58, and p41 are indicated by arrows. (B) A /Im3 gave a 92-kDa product, which was autocleaved into p58 and p35, as indicated by arrows. (C) A1020/I1773 was translated into an 87-kDa product, whose processing generated p58 and p29 as indicated. (D) G1102/I1773 gave a 78-kDa protein, whose autolytic processing was undetectable. (E) Translation reactions of V920/I1773, A^^nn, and A1020/I1773 were each programmed with input R N A at 0, 1:20, 1:100, 1:200, and 1:500 dilutions, and mixtures were incubated for 4 h. Positions of molecular mass markers and cleavage products are indicated. Images were scanned using a U M A X Astra 1220U scanner with Adobe Photoshop 5.0 software. 9  3  35  3  914  65  My results suggest that a construct starting from A1020 to a residue after the cleavage site such as to include the N terminus of p90 is sufficient for cis processing. Comparison between domains required for cis- and rrans-cleavage activities of NS-pro indicated that the domains involved in cis- and frans-cleavage activities are different. Obviously, the cis protease domain must contain a cleavage site and C-terminal tail for cis cleavage to occur. This is not the case for the trans protease domain. However, it is interesting that the N-terminal domains are different between cis and trans cleavage for R V NS-pro. The domain from V o to A1020 is 92  required for trans cleavage but is dispensable for cis cleavage. It will be of interest to examine the functions of the domain from V o to At. 020 in trans cleavage. 92  4.1.5.  Effect of N-terminal regions on cleavage efficiency.  The protease constructs examined in this study were found to have variable cleavage efficiencies, depending on the region and length deleted. Cleavage efficiency was compared among different protease constructs by using the percentage of cleaved products with respect to total proteins, expressed as a cleavage ratio. The cleavage ratio for each protease construct at certain incubation time was calculated as described in section 3.2.15, and plotted against time.  Fig. 15A Compares the rrans-cleavage efficiency among positive controls [p200(G1301S) and pl50],  A348/G1301,  M827/G1301,  V 2o/Gi3oi, 9  V 2o/Pi296>  V 2o/Li297,  9  9  and V 2o/Ri299- For 9  p200(G1301S) and pl50, the function time began when the protease and substrate were mixed. However, for each of A348/G1301, M827/G1301, V920/G1301, V o/ P1296, V920/L1297 and V920/R1299, 92  the protease functioned only after it had been translated, which took about 40 to 60 min. Therefore, the effective function time was taken as the real incubation time minus 60 min. The eight protease constructs could be separated into two groups according to their proteolytic  66  activity: one group with high cleavage ratio (70 to 90%), including p200(G1301S), pl50, A348/G1301,  and  V920/G1301,  V  /Pi296,  9 2 0  and the other group with low cleavage ratio (5 to 17%), including  M827/G1301,  V9 /Li 97, 20  2  and  V 2o/Ri 999  2  A348/G1301  and M  827  / G i o i had cleavage 3  efficiencies comparable to those of p200(G1301S) and pl50, the positive controls for transcleavage activity, whereas  V920/G1301,  differing from  substantially lower cleavage ratio (17%) than  M 7/Gi3oi 82  M 7/Gi3oi 8 2  in lacking an X domain, had a  (82%). These results suggest an  important role of the X domain in trans cleavage.  As discussed above, the processing of of  A974/I1773  or  efficiencies of  A1020/I1773  is largely by cis cleavage, and the processing  is the consequence of cis cleavage only. Therefore, the processing  V o/Ii773, A974/I1773, 92  V920/I1773  and  A1020/I1773  (Fig. 15B) reflected their respective cis-  cleavage abilities. The constructs compared in Fig. 15B can be classified into two groups: one with high processing ratios (60-70%) including WT NSP, (A1020/I1773)  V920/I1773,  with a processing ratio as low as 35%. The fact that  and  V920/I1773  A974/I1773,  the other  and A974/I1773 had as  efficient cis cleavage as WT NSP suggested that the lack of an X domain in  V920/I1773  and  A974/Ii773 had no significant influence on their self-processing. However, the domain from residue 974 to 1020, though not required absolutely, had a substantial effect on cis cleavage.  67  A.  trans cleavage  120  180  incubation time (min)  B.  c/s processing  Br—  0  60  120  180  - B  240  300  incubation time (min)  Figure 15. Cleavage efficiencies of protease constructs. Protein bands of substrate and cleavage products on SDS-PAGE gel were quantitated using the Scion image program. The cleavage ratio for each protease construct was calculated, and plotted against incubation time. (A) trans-cleavage efficiency comparisons. 0, p200(G1301S); • , pl50; O , A 4 /Gi3oi; X , M / G i 3 n i ; • , V o/Gi3 i; A , V9 /P 9 ; A , V o/Li 97; •, V o/Ri 99. (B) cz's-leavage efficiency comparisons. • and O , W T NSP in the absence and presence, respectively, of Z n ; • , V /Ii773; * , A974/I1773; • , A /Ii773 • 3  92  8  92  827  0  20  12  6  92  2  2  2+  920  1020  68  4.1.6.  Secondary structure prediction for R V NS-pro.  Gorbalenya et al. (1991) reported sequence similarity between papain, a cellular cysteine protease, and R V NS-pro in the vicinity of catalytic C and H residues through local alignment. The catalytic C and H residues are separated by 133 residues in papain and 120 residues in R V NS-pro (Gorbalenya et al., 1991). There are 24 residues upstream of the catalytic C residue in papain. The active R V NS-pro domain identified in this work was larger than papain, with about 230 residues upstream of the catalytic C1152 that are required for trans cleavage or about 130 residues upstream of Cn5 that are required for cis cleavage. It has been reported that 2  through sequence alignment and secondary structure comparison to known protein structures, topologic prediction of uncharacterized proteins is possible (Skern et al., 1998). Skern et al. (1998) proposed a papain-like fold for the F M D V Lpro, a viral PCP, from the analysis of predicted secondary structure. This prediction was confirmed by a recent crystallographic analysis of F M D V Lpro showing a globular papain-like catalytic domain with adaptation for the specific requirements of the virus (Guarne et al., 1998). In the hope of obtaining initial structural information on R V NS-pro, I compared the primary and secondary structures of RV NS-pro to those of papain. The analyses were performed with strains M33 and Therien, both of which are W T isolates of R V and differ from each other by two residues (A1140V and R1201W, M33 versus Therien) within the examined NS-pro catalytic region (from residue 1128 to 1301), with identical results. Only the result for R V strain M33 is presented here (Fig. 16).  To determine the global similarity between papain and R V NS-pro with respect to their catalytic sites, sequence alignment between papain and the R V NS-pro catalytic region (residues 1128 to 1296) was made using the ALIGN program (Myers and Miller, 1988) with  69  manual modification (Fig. 16). The alignment gave an identity of 18.1%, and, as expected, the two sequences exhibited most similarity around C and H , the catalytic sites. The derived alignment was further supported by the analysis of the predicted secondary structure of RV NSpro (illustrated in Fig. 16). R V NS-pro was predicted to have the a-p structural organization found in cellular PCPs. This prediction has three cc-helices (rxLl, c«L2/3, and ccRl) and six Psheets (A-F) in R V NS-pro. Most of these were present in the papain structure at corresponding positions including PA, a L l , a R l , P C PD, PE, and PF (Kamphuis et al, 1984, 1985). The match was highest in the catalytic C and H regions, with differences occurring in the linker regions and the C end. In NS-pro, an oc-helix, ocL2/3, took the place where two helices, ocL2 and ccL3, occurred in papain. PB1 and PB2 for NS-pro did not match the position of PB for papain well. Furthermore, the ccR2 in the linker region and PG at the C end were missing in the NS-pro prediction. R V NS-pro had a shorter linker region (120 residues) between catalytic C and H residues than papain (133 residues) and a shorter tail after the catalytic H residue (23 residues) than papain (53 residues), which explained the discrepancies. It was proposed (Skern et al., 1998) that loops between the secondary elements that define the papain topology can be modified, by insertions or deletions, without interfering with the overall folding of the molecule. Therefore, the global similarity of their secondary structures suggested that R V NSpro might maintain a papain-like topology for its catalytic region, whereas those differences may come from the adaptive changes for the viral specificity. Crystallographic data are necessary for precise structure determination for RV NS-pro.  70  PA i=> 10  20  papain IPEYVDWRQKGAVTPVKNQGSC  • :•  NS-pro  ocLl AAAAAAAAAAAAAA 30 40  aL2 AAAA 50  GSlWAFSAVVTIEGIIKI  :...:.  . .[!J:  . :....  RTGNLNEYSEQEL  .  HAALCRTGVPPRVSTRGGELDPNTCW-LRAAANVAQAARACGAYTSAGCPKCAYGRA  1128  1150  1=^>  1170  AAAAAAAAAAAAAAAA  PA  AA  aLl  aL3 PB AAAAAAAAA •=: 60 70 80 90 100 110 LDCDRRSYGCNG-GYPWSALQLVAQYGIHYRNTYP-YEGVQRYCRSREKGPYAAKTDGVR  AA papain  NS-pro LSEARTHEDFAALSQWWSASHADASPDGTGDPLDPLMETVGCAC-SR 1190  VWVGSEH  1210  1230  AAAAAAAAAAAAAAAAAA  •=>-<  aL2/3  PB1  PB2  aRl PC aR2 PD pE =0—AAAAAAAAA—i=> AAAA 1 120 130 140 150 160 170 p a p a i n QVQPYNEGALLYSIANQPVSWLEAAGKDFQLYRGGIFVGPCGNKVDHAVAAVGYGPNYI NS-pro EAPPDHLLVSLHRAPNGPWGWLEVRARP 1250 1  AAAA aRl  papain  EGG  NPTG 1270  >  HFVCAVGGGPRRV < = > PD  PC  PF  PG  c=>  <==>  1=0 PE  180 190 200 210 LIKNSWGTGWGENGYIRIKRGTGNSYGVCGLYTSSFYPVKN  NS-pro SDRPHL WLA 1290 1  VPLSRG-G >  PF  Figure 16. Comparison of primary and secondary structures between RV NS-pro and papain.  71  Figure 16. Comparison of primary and secondary structures between R V NS-pro and papain. The protein sequence of papain (SWISS-PROT accession number P00784) was aligned to that of NS-pro (strain M33; GenBank accession number S38480 with corrections by Pugachev et al., 1997) using ALIGN software, with manual adjustment. The papain sequence is numbered as for mature protease, and RV-NS-pro sequence is numbered as for the NSP ORF. Identical residues are marked conserved amino acids with Cys and His at the catalytic sites are in boldface and shaded. Features of the secondary structure of papain (Kamphuis et al., 1984) are illustrated above the sequence. Secondary structure for R V NS-pro was calculated by the E M B L protein prediction server (Rost et al., 1993, 1994) and is illustrated below the NS-pro sequence, a-helices are shown as curves, and P-sheets are shown as arrows, cc-helices are named according to the nomenclature of Kamphuis et al. (1984); P-sheets are named as described by Skern et al. (1998).  4.1.7.  Discussion I  I have used an in vitro translation system to identify domains important for cis- and transcleavage activities of R V NS-pro. The results are summarized in Fig. 17. Through analysis of protease activity using R N A transcripts from cloned material with serial deletions from either end of pi50,1 have demonstrated that R V NS-pro requires a region from residue 920 to 1296 to perform functional trans cleavage (Figs. 12 and 13). The N-terminal region of NS-pro was roughly determined to reside between residues 920 and 974 (Fig. 12C and D). The C end was precisely determined to be  P1296  (Fig. 13B and C). However, the minimal NS-pro domain  (residues 920 to 1296) for trans cleavage processed only 5 to 17% of substrate after 4 h of incubation, compared to the 70 to 96% for the positive controls, p200(G1301S) and pl50 (Fig. 15A). The NS-pro domain that maintains as high rrans-cleavage ability as the positive controls is found in the construct  M827/G1301  (80% cleavage at 4 h), starting from around residue Mg27  (Fig. 15A).  72  residue  834  920 940  1020  1128  1296 RV NS-pro  •o  X-domain N-terminal boundary for minimal trans minimal cis cleavage cleavage  Cys  N-terminal boundary for  1 19  134 pro domain  n  Cys  345'  ^jmmmd papain His  signal peptide  Figure 17. Functional domains of RV NS-pro and comparison with complete papain sequence. Residues of R V NS-pro are numbered as for the R V NSP ORF. The X domain (residues 834 to 940) is shown by the dotted line, positioned outside the essential protease domains of NS-pro shown as a solid heavy line. The N-terminal boundary for the minimal domain of either transox ds-cleavage activity is indicated by the arrow starting from residue 920 or 1020, respectively. The region from residue 1128 to 1301 is compared to the mature papain protease in this work, and the boundary is indicated by vertical dotted lines. The predicted catalytic residues Cys and His of R V NS-pro and catalytic residues Cys and His of papain are represented by open squares. For a complete papain sequence, residues 1 to 18 encode the signal peptide shown as a heavy line, residues 19 to 133 are the pro region, and residues 134 to 345 contain the mature papain sequence.  R V NS-pro possesses both cis and trans activities (Yao et al., 1998). Both use the same cysteine protease cleavage mechanism and thus should employ the same core catalytic structure (Kamphuis et al., 1985). They may vary in other external domain requirements. In addition to the protease domain, the cis protease must include intact cleavage site and substrate regions. The most significant difference between cis and trans cleavage for R V NS-pro lies at the N terminal domains. When a panel of protease constructs with nested N-terminal deletions (V920/I1773, A974/I1773, Aio2o/Ii773>  examined for autoprocessing,  and G1102/I1773), with weak or no rrans-cleavage activity, was  V920/I1773, A974/I1773,  and  A1020/I1773  were active in cis, while  73  G1102/I1773  was not (Fig. 14). Thus, protease active in trans starts after residue 920 but before  974, while that in cis begins after residue 1020. These results suggested that the core protease domain (required for both cis and trans activity) ranges from around residue 1020 to residue 1296, and that the fragment from residue 920 to 1020 is required only for trans cleavage while being dispensable for cis processing (Fig. 17). My data are the first to show that R V NS-pro uses different domains for cis and trans cleavage. Since trans cleavage is a bimolecular interaction, it is likely that the domain of residues 920 to 1020 is involved in protein-protein interaction, required to position substrate protein into the trans protease catalytic site. For cis cleavage, this protein-protein interaction is not necessary in order to hold substrate and enzyme together. Identification of the frans-specific domain facilitates the future studies on the biological significance of rrans-cleavage activity.  Sequence analysis on M-group PCPs of several virus families (such as alphavirus and coronavirus, etc.) had identified a conserved X domain near the protease domain (Gorbalenya et al., 1991; Strauss and Strauss, 1994). In RV, this X domain lies N-terminal to the protease domain, ranging from residue 834 to 940 (Fig. 17) (Gorbalenya et al., 1991). Functions of the X  domain remain to be characterized. Association of X domain with M-group PCPs  (possessing both cis and trans activity) rather than L-group PCPs (containing only cis activity) encouraged the speculation that the X domain might be involved in the regulation of polyprotein processing (Gorbalenya et al., 1991). Elimination of the X domain from PLP-1 of M H V reduced cleavage by 22 to 63% (Bonilla et al,  1995; Teng et al,  1999). In my  experiments, R V NS-pro remained enzymatically active after all or most of the X domain had been removed. V920/G1301 cleaved substrate in trans, and A974/I1.773 processed itself efficiently (Figs. 12C and 14B). However, cleavage efficiencies differed considerably. In trans cleavage, the absence of the X domain in V920/G1301 caused a substantially decreased cleavage ratio 74  (17%) compared to that for  M827/G1301  (82%), which contains the X domain (Fig. 15A). In  contrast, cis cleavage was not affected significantly by the presence of the X domain, since V920/I1773  and  A974/I1773  (both missing the X domain) processed themselves almost as efficiently  as the positive control, WT-NSP (Fig. 15B). My results demonstrate the importance of the X domain in trans cleavage. Although it is unclear at present what function it could play in R V NS-pro trans cleavage, I speculate that this proline-rich region might provide a protein-protein interaction domain that enhances the opportunity for protease to meet its trans cleavage substrate and thus decreases the Km of protease. Further studies of the biologic significance and functional mechanism of the X domain in NSP processing and virus replication are indicated.  The PCP family include a group of cellular and viral proteases which employ the catalytic C and H dyad. The distant relationship between viral and cellular PCPs was suggested from many primary sequence comparisons (Berti and Storer, 1995; Gorbalenya et al., 1991) and from the crystal structure of F M D V Lpro, the only structure determined on a viral PCP (Guarne et al., 1998). Sequence alignment showed that the catalytic region of R V NS-pro (from residue 1128 to 1296) has global sequence similarity with papain (Fig. 16). Secondary structure comparison also supported their topologic relationship (Fig. 16). It is possible that the catalytic region of RV NS-pro exhibits a papain-like folding with adapted modifications. To obtain the full tertiary structural information of RV NS-pro will require crystallographic analysis.  The additional N-terminal region of the R V NS-pro core domain (from  A1020  to  A1127)  has no  corresponding sequence in papain and was excluded from alignment with it. It is likely that this N domain may not contain sequences directly required for protease activity. Rather, it may serve other subsidiary functions, such as folding assistance, conformational stability, and/or protein-protein interactions. Papain, as well as other proteases, is translated as a pro-protease 75  with an additional N-terminal region (115 residues of pro region for papain) (SWISS-PROT accession number P00784) (Fig. 17). In many proteases, the pro region plays an active role in protein folding. Subtilisin (Bryan et al.,  1995), oc-lytic protease (Baker et al., 1992),  carboxypeptidase A l (Phillips and Rutter, 1996), and carboxypeptidase Y (Ramos et al., 1994) do not fold into active conformations in the absence of their pro regions. The pro region is essential for folding of at least one PCP (cathepsin L) (Tao et al., 1994). The pro regions of PCPs can also perform other biological roles, such as stabilization (Mach et al., 1994; Tao et al., 1994) and subcellular targeting (Mao et al., 1998; Mclntyre and Erickson, 1993). It would be interesting to determine whether the region from A1.020 to A1127 of R V NS-pro serves subsidiary roles similar to those of the pro regions for many other proteases.  4.2.  4.2.1.  Effects of NSP cleavage on virus replication and RNA synthesis  Construction of mutants.  In order to study the effects of R V NSP processing on virus replication, the R V genome was altered by mutations that affect only NSP processing while avoiding any other possible structural or functional alterations in the NSP or in the viral genome. A panel of site-directed mutations was generated by PCR mutagenesis. These included catalytic-site mutation C1152S and cleavage-site mutations G1301S, R1299A, G1300A, and G1301A. Most of the introduced mutations are conservative alterations, such as C to S and G to A , and thus are unlikely to affect the overall structure of the protein or functions of NSP other than proteolytic processing. To facilitate the process of mutagenesis for construction of the G1300A and G1301A mutations, a silent mutation was introduced into the R V infectious cDNA clone pBRM33 (Yao and Gillam, 1999) to create a new Xba I site at nt 3935. The resultant cDNA clone was named pBRM33-X.  76  In  terms  of  virus  growth,  plaque  size,  and  specific  infectivity,  pBRM33-X  was  indistinguishable from pBRM33 (Table3, Fig.20, and Fig. 21). Amplified PCR fragments containing the desired mutations were reintroduced into pBRM33 or pBRM33-X, and the respective  cDNA  clones  were  named  after  their  mutations:  pBRM33(C1152S),  pBRM33(G1301S), pBRM33(R1299A), pBRM33(G1300A), and pBRM33(G1301A). The plasmid constructs encoding these mutations are listed in Table 2.  Table 2. Mutations created at the catalytic site and around the cleavage site. Mutation  Catalytic site  Around cleavage site  None(WT)  C 2  Ri299-Gi3oo-G oi*Gi302  C1152S  S  8  n 5  13  R1299A  A  G1300A  A  G1301A  A  G1301S  S  a  Cleavage occurs after G1301, as indicated by the vertical arrow.  4.2.2.  To  Effects of mutations on NSP processing.  determine whether the p200 polyprotein itself can function in R N A replication, I  constructed a panel of cleavage-defective  mutations. The effect of mutations on NSP  processing was determined using time course analysis of in vitro translation reactions programmed with full-length R N A transcripts from cDNA clones as described in section 4.1. The extent of NSP processing differed greatly among the W T and mutant RNAs (Fig. 18). To compare their cleavage efficiencies more precisely, the processing ratio, calculated as the percentage of the cleavage products in the total proteins, was assessed for each mutant and plotted against the incubation time (Fig. 19). From both gel analysis and the calculated processing ratio, WT NSP processing (Fig. 18A) was almost complete at 3 h of incubation (Fig. 77  19). The NSP processing of the G1300A mutant was slightly delayed and decreased (Fig. 18B and 19); its processing ratio at 3 h was 75% of the WT level (Fig. 19). Mutation R1299A substantially impaired NSP processing, since the cleavage products were detected only after 3 h of incubation (Fig. 18C) and the cleavage ratio at 3 h was approximately 20% of the WT level (Fig. 19). Mutation G1301A resulted in minimally detectable cleavage of p200, with a minute amount of p90 detected after a 3-h incubation time (Fig. 18D). Mutations C1152S and G1301S abolished NSP processing completely (Fig. 18E and F). Thus, my generated mutations either abolished (C1152S and G1301S) or blocked (R1299A, G1300A and G1301A) NSP processing to various degrees.  A. WT  B. G1300A  time (min) kDa  2  time (h)  °' °' 4  6  I  200 ""'  97lanes  1  °' °' °' 9  1 2  0  1 8 0  1  PP P t H «»«—  Wf  2  3  4  5  0  1  3  01  6  1  2  2  3  4  0  1  3  4  5  ___ ___ _ I  AM  1  4  5  6  v  2  3  4  5  6  4  5  F. G1301S time (h)  2  3  4  5 i  A3  0  1  2  3  W i i p i ^ l W - p l O O .  -  o200  3  E. C1152S 5  2  mm IP mm mm mm m~ piso  time (h)  • 97lanes 1  5  wm ^ —— —  a  200-  4  • H HI  time (h) D  time (h)  2  4_H_. ___ ___ ___  D. G1301A k  C. R1299A  P200  -»pl50 - 90 p  2  3  4  5  6  12  3  4 5 6  12  3  4 5 6  Figure 18. Effects of mutations on NSP processing in an in vitro translation system. Full-length R V RNAs containing site-directed mutations were generated from corresponding Hind III-linearized cDNA templates with SP6 R N A polymerase. In vitro translation reaction mixtures containing rabbit reticulocyte lysates were prepared at 30°C. Aliquots were removed at the indicated times, and the protein products were resolved by sodium dodecyl sulfatepolyacrylamide gel electrophoresis (SDS-PAGE). Panels: A, WT NSP; B, G1300A mutant NSP; C, R1299A mutant NSP; D, G1301A mutant NSP; E , C1152S mutant NSP; F, G130IS mutant NSP. Positions of molecular mass markers and protein products are indicated. Images were scanned using a U M A X Astra 1220U scanner with Adobe Photoshop 5.0 software.  78  o 0.9 -  'I 0.8 o) 0.7 •I 0.6 3 0.5 -  o 0.4 -  «• 0.3 -  W 0.2 z  0.1 0-  0  3  1 2  4  5  incubation time (h)  Figure 19. Comparison of processing ratios between the WT and mutant NSPs. Protein bands of p200 and cleavage products, pi50 and p90, were each quantitated at indicated times using SCImage software as described in Materials and Methods. The cleavage ratio was calculated as the percentage of the cleaved products over the total proteins, and plotted against incubation time. • , WT NSP; • . NSP(G1300A); A , NSPCR1299A); O , NSP(G1301A); • , NSP(C1152S) and NSP(G1301S).  4.2.3.  Effects of mutations on virus growth.  In order to examine the effects of mutations on R V replication, W T or mutant RNA, transcribed from respective full-length cDNA clones, was used to transfect either Vero or BHK-21 cells. The transfected Vero cells were incubated for 5 days and assayed for infectious virus released into the culture medium (Table 3). Infectious virus particles could be harvested from Vero cells transfected by the WT R N A and those with the G1300A and R1299A mutations, yielding virus titers of 3.4 x 10 , 1.2 x 10 , and 1.0 x 10 PFU/ml, respectively. In 6  6  3  contrast, no plaques were detected in the medium containing RNAs with the mutations G1301A, G1301S and C1152S, indicating that they are noninfectious (Table 3). Transfected Vero cells were also analyzed for the production of RV-specific SPs by immunoprecipitation with human anti-RV serum. R V SPs were readily detectable for the W T and the G1300A mutant but in a substantially lower quantity for the R1299A mutant. No R V SPs could be 79  detected from cells transfected by the G1301A, G1301S and C1152S RNAs (Fig. 20), indicating that they are defective in replication. To confirm that the different amounts of virus produced by the W T and mutant RNAs are not dependent on the host cells used, I also transfected BHK-21 cells with RNA transcripts by electroporation. At 48 h postelectroporation, the culture medium was collected and the released virus particles were quantitated. Consistent with the results obtained with Vero cells, infectious virus particles were only detected from BHK-21 cells transfected with the WT, G1300A and R1299A RNAs. The virus titers were 1.5 x 10 , 6 x 10 , and 5 x 10 PFU/ml, respectively. Again, no infectious virus could be harvested 7  6  3  from BHK-21 cells transfected with transcripts of G1301A, C1152S, or G1301S mutant R N A (Table 3). R N A transfection of BHK-21 cells by electroporation resulted in higher virus titers than that of Vero cells using Lipofectin, most likely due to the higher transfection efficiency of electroporation.  Table 3. Effects of RV NSP cleavage mutants on virus replication. Virus titer (PFU/ml) at: 3  Day 5 posttransfection of Vero cells  48 h posttransfection of BHK-21 cells  WT  3.4 x 10  6  1.5 x 10  WT M33-X  2.6 x 10  6  N/A  G1300A mutant  1.2 x 10  6  6.0 x 10  R1299A mutant  3  1.0 x 10  5.0 x 10  G1301A mutant  0  b  G1301S mutant  0  b  C1152S mutant  0  b  RV  7  Plaque phenotype on Vero cells 0  Big, clear, 6 dpi Big, clear, 6 dpi  6  Big, clear, 6 dpi  3  Tiny, unclear, 8 dpi  0  b  None  0  b  None  0  b  None  ^he virus titers, determined by plaque assay on Vero cells, are the means of at least two independent experiments. N/A, data not available. ''The culture medium was tested without dilution and at 1:10 and 1:100 dilutions for the plaque assay. 0, no plaque formation was observed. Vero cells infected by the WT or mutant virus initiated by R N A transfection were overlaid with agarose medium and incubated for 6 days (for the WT and G1300A mutant) or 8 days (for the R1299A) before being stained with neutral red. dpi, days postinfection.  c  80  kDa 97 — 68  43  -  El  _ •  SK 3E SK  E2 C  29 — lanes  1 2  3  4  5  6  7  8  Figure 20. Immunoprecipitation of RV-specific SPs from cells transfected by W T and mutant RNAs. Vero cells in 35-rnm-diameter dishes were transfected with WT or mutant R V RNAs mediated by Lipofectin for 2 h at 37 °C and incubated with fresh medium for 6 days. Transfected cells were labeled with [ S]-methionine for 2 h and lysed with lysate buffer. R V specific proteins were immunoprecipitated using human anti-RV serum and separated by 10% SDS-PAGE. Lane 1, mock transfection. Lane 2, WT M33 RNA. Lane 3, WT M33-X RNA. Lane 4, M33(G1300A) RNA. Lane 5, M33(R1299A) RNA. Lane 6, M33(G1301A) RNA. Lane 7, M33(G1301S) RNA. Lane 8, M33(C1152S) RNA. The positions of molecular mass marker and RV structural proteins were indicated. Images were scanned using a U M A X Astra 1220U scanner with Adobe Photoshop 5.0 software. 35  The results obtained with the two cell types are comparable. The amount of virus produced from each infectious R N A (WT, G1300A, or R1299A) varies with its NSP processing efficiency. In both cases, W T RNA, having the most efficient NSP processing, produced the highest virus titer. The G1300A mutant RNA, with 75% of the W T level of NSP processing, produced viruses at 30 to 40% of the WT level. The R1299A mutant RNA, with NSP  81  processing at 20% of the WT, released viruses at a level 2 x 10 to 3 x 10 -fold lower than that 3  3  of the WT. The G1301A mutant RNA, with minimally detectable NSP processing in vitro (processing ratio of less than 10% at 5 h of incubation), and the C1152S and G1301S mutant RNAs, abolishing NSP processing completely, released no infectious virus particles in either Vero or BFfK-21 cells and thus are effectively lethal. In addition to the differences in virus titer, the W T and mutants also have different plaque phenotypes. The W T and the G1300A mutant produced large, clear plaques at day 6 postinfection while the R1299A mutant resulted in tiny, unclear plaques only after day 8 postinfection.  To further analyze the influences of NSP cleavage on virus replication, growth rates were determined for the WT and infectious mutant (R1299A and G1300A) RNAs. Vero cells were transfected with the respective full-length RNAs mediated by Lipofectin. Culture medium was harvested every 24 h and replaced with fresh medium. Virus titers in the culture medium were quantitated by plaque assay on Vero cells and are shown in Fig. 21. For the WT, the amount of virus produced was about 3 x 10 PFU/ml at day 1 and reached a peak of 5 x 10 PFU/ml at day 3  6  4. The G1300A mutant had growth kinetics similar to those of the W T but yielded a 10-fold lower amount of released virus (2 x 10 PFU/ml) at day 1 and a 3-fold lower amount (1.6 x 10 2  6  PFU/ml) at day 4. The R1299A mutant virus was not detectable until after day 3, and its titer at day 5 was 2 x 10 -fold lower than that of the WT. The failure to detect R1299A virus plaques 3  before day 3 posttransfection may be due to their small size at early stages of infection. My results demonstrate that NSP cleavage plays an important role in virus replication.  82  time posttransfection (h)  Figure 21. Growth curves of the W T , G1300A, and R1299A mutant viruses. Vero cells in 35-mm-diameter dishes were transfected with W T or mutant RNAs mediated by Lipofectin for 2 h at 37 °C. The cells were overlaid with culture medium after removal of the Lipofectin-RNA mixtures. The culture medium was changed every 24 h, and the virus particles released into the medium were quantitated by plaque assay. The results shown are the means of at least two independent experiments. Symbols: O , W T M33 RNA; • , WT M33-X RNA; • , G1300A mutant; • , R1299A mutant.  4.2.4.  Effects of mutations on viral RNA synthesis.  The reduction in virus yield due to defects in NSP cleavage presumably occurred at the level of viral R N A synthesis. To determine at which step(s) R N A synthesis is impaired in the mutant viruses, I examined the synthesis of three viral R N A species in the W T and NSP cleavage mutant viruses at the early stage of virus replication using an RNase protection assay (RPA).  To evaluate the sensitivity of the RPA for detection of positive-strand R V R N A , various amounts of positive-strand R V RNA, transcribed in vitro from pBRM33, were subjected to an  83  RPA using 10 cpm of S-labeled probe pbl8, which contains 301 nt of the R V sequence and 6  35  27 nt of the vector sequence. The negative-polarity R V RNA, transcribed from a cDNA clone encoding an R V genome of reverse polarity, was also used as a negative control. As shown in Fig. 22A, a protected band of 301 nt was present in reaction mixtures containing 100 pg (lane 3), 1 ng (lane 4), 10 ng (lane 5), and 20 ng (lane 6) of positive-strand RNA, but was absent in reaction mixtures containing 10 pg of positive-strand R N A (lane 2) and 20 ng of negativestrand genomic R N A (lane 7). These data indicate that this assay is strand specific and sensitive enough to detect at least 100 pg of positive-strand R V R N A (approximately 10 molecules). 7  Furthermore, quantitative analysis of the protected probe suggested that the signal was proportional to the amount of positive-strand R N A used.  In virus-infected cells, R V negative-strand genomic R N A exists mostly as a double-stranded intermediate form with positive-strand R N A present in large molar excess. To prevent interference between the probe and negative-strand R N A by the large molar excess of positivestrand RNA, a two-cycle RPA was employed to detect negative-strand genomic R N A (Novak and Kirkegaard, 1991). To determine the sensitivity of the two-cycle RPA for negative-strand RNA, various amounts of negative-strand genomic RNA, transcribed in vitro from a cDNA clone encoding an R V genome of reverse polarity, were hybridized with 20 ng of unlabeled RNA probe pbl9 (the complementary sequence of pbl8) in the first-cycle RPA. The products were subsequently hybridized with 10 cpm of S-labeled pbl9 and subjected to a second cycle 6  35  (Fig. 22B). The probe pbl9 is negative-strand specific, since no signal band was observed in the reaction mixture containing 10 ng of positive-strand genomic R N A (lane 7). The 301-nt signal band was apparent in a reaction mixture containing 1 ng (lane 4), 10 ng (lane 5), or 20 ng (lane 6) of negative-strand genomic RNA. A longer exposure (3 days) also detected the existence of this band in a reaction mixture containing 100 pg of negative R N A (lane 3). 84  Therefore, this two-cycle R P A is sensitive enough to detect more than 100 pg of negativestrand genomic R N A (approximately 10  7  R N A molecules). The signal intensity was  proportional to the amount of negative-strand RNA.  B  (+) R N A 00 -O  00  CL,  0  ft 00  OO 60  0  o o ^* ©  G  ©  (-)RNA  W> G  o  OH  <N <N  O H d  o o o  328 nt 301 nt lane 1 2 3 4 5 6 7  00  ft  lane 1 2  i—i  „. '—  M  o  B 1  i—i  M  00  o  CS  II  3 4 5 6  328 nt 301 nt 7  Figure 22. Sensitivity of RPA for detection of both positive- and negative-strand RV RNAs. RPA reactions were carried out as described in section 3.2.14. The full-length R V RNAs of either positive or negative polarity used in this experiment were transcribed in vitro from the respective R V cDNA clone, pBRM33 or pBRNM33, encoding a full-length R V genome downstream of the SP6 R N A polymerase promoter in either the forward or the backward orientation. (A) Standard RPA reactions were performed on 10 pg, 100 pg, 1 ng, 10 ng, and 20 ng (lanes 2 to 6) of positive-strand R V RNAs in the presence of 1 x 10 cpm of S-labeled probe pbl8. A reaction mixture containing 20 ng of negative-strand R V R N A (lane 7) was also included as a negative control (lane 7). The products of RPA reactions along with 2 x 10 cpm of pbl8 (lane 1) were resolved on a 5% polyacrylamide-7 M urea gel, which was treated with Enhancer (DuPont), dried, and exposed to X-ray film. (B) Two-cycle R P A reactions were performed with various amounts of negative-strand R V R N A . Negative-strand R V R N A was hybridized with 10 ng of transcribed unlabeled pbl9 and subjected to the first-cycle R P A reaction. The products of the first-cycle RPA reaction were hybridized with 1 x 10 cpm of S labeled pbl9 and subjected to the second-round R P A reaction. To examine the strand specificity of probe pbl9, 10 ng of positive-strand R V R N A (lane 7) was analyzed in parallel. The control reaction mixture contained no R V R N A (lane 2). Lanes 3 to 6 represent reaction mixtures containing 100 pg, 1 ng, 10 ng and 20 ng of negative-strand R V RNA, respectively. Lane 1, pbl9. The autoradiographs were exposed for 1 day. The positions of the 328-nt and the 301-nt bands are indicated. Images were scanned using U M A X Astra 1220U scanner with Adobe Photoshop 5.0 software. 6  35  3  6  35  85  I then examined viral R N A synthesis in BHK-21 cells transfected by electroporation with WT or mutant R N A transcripts. I consider virus-infected cells a less ideal system for the analysis of viral R N A synthesis because (1) revertants or second-site mutations may exist in virus stocks, (2) the system might be affected by the early steps of virus entry prior to viral RNA synthesis (e.g., virus entry and nucleocapsid uncoating), (3) the percentage of cells initially infected by RV is quite low (10 to 20%) (Frey, 1994; Hemphill et al,  1988), and (4) studies of  noninfectious mutants are impossible. In contrast, electroporation of viral R N A transcripts into BHK-21 cells provides an advantageous system for the analysis of R N A synthesis at an early stage of virus replication. This system bypasses the steps of virus entry and nucleocapsid disassembly; has a high efficiency of transfection, allowing detection of low levels of negativestrand RNA; and allows studies of noninfectious mutants.  To determine R N A synthesis of WT and mutant viruses, the respective viral RNAs transcribed in vitro were used to transfect BHK-21 cells by electroporation. At the indicated times, total cellular R N A was extracted and subjected to an RPA. A 2-|ig sample of R N A was used for positive-strand R N A detection, and 20 |xg was used for negative-strand R N A detection. At 0 h postelectroporation, the 301-nt protected fragment representing the positive-strand genomic R N A was apparent for all constructs (Fig. 23A, lanes 3, 6, 9, 12, 15, 18), representing the input genomic R N A transfected into cells. By 8 h postelectroporation, the intensity of the 301-nt band decreased to a low level (Fig. 23A, lanes 4, 7, 10, 13, 16, 19), suggesting that the input genomic R N A had mostly been degraded at that time. At 24 h postelectroporation, accumulation of both the protected 301-nt fragment and the 188-nt fragments (representing subgenomic RNA) was apparent for the WT and mutant G1300A (Fig. 23A, lanes 5 and 8). Much less of these bands was found with the R1299A mutant R N A (Fig. 23A, lane 11). The  86  G1301S and C1152S mutant RNAs exhibited decreased levels of the 301-nt band (compared with the amounts detected at 8 h; Fig. 23A, lanes 17 and 20). Subgenomic R N A represented by the 188-nt fragment was scarcely detectable for the two mutants. Thus, they show no evidence of production of positive-strand RNA. The G1301A mutant showed the presence of low levels of the 301-nt band with little increase at 24 h over the level found at 8 h. By 24 h, a trace of the 188-nt fragment was detected (Fig. 23A, lane 14), indicating some slight synthesis of positivestrand RNA. These results confirm that the more-infectious constructs produce more positivestrand RNAs and the noninfectious ones produce little (G1301A) or no (G1301S and C1152S) positive strands. Detailed quantitation of the amount of positive-strand R N A is presented below.  Fig. 23B shows the levels of negative-strand R N A produced by the constructs. At 0 h postelectroporation, no protected negative-strand RNA fragment (301 nt) was observed for any construct (Fig. 23B, lanes 3, 7, 11, 15, 19, and 23). By 4 h postelectroporation, all constructs had produced detectable negative-strand RNA (Fig. 23B, lanes 4, 8, 12, 16, 20, 24), with the WT showing the lowest level. Negative-strand R N A continued to accumulate at 8 h in all constructs (Fig. 23B, lanes 5, 9, 13, 17, 21, 25). At 24 h, the amount of negative-strand RNA had further increased in the WT and the G1300A and R1299A mutants (Fig. 23B, lanes 6, 10, and 14), but had not increased (or even decreased) in the G1301A, G1301S and C1152S mutants (Fig. 23B, lanes 18, 22, 26). The last three mutants are those that showed little or no positive-strand R N A synthesis (see above), suggesting that the absence of positive-strand RNA accumulation prevents continued synthesis of negative-strand R N A or allows its degradation. The increased amount of negative-strand RNA in the WT, G1300A and R1299A at 24 h may mean that the presence of newly synthesized positive-strand R N A allows negative-strand RNA to accumulate further. Whether this occurs in new cells from reinfection or in the same cells is 87  unclear. In general, all of the constructs, including those without synthesis of positive-strand RNA, produced negative-strand RNA at early stages of infection.  £ P  b l 8  1  0  WT 8  24  G1300A 8 24  0  R1299A 8 24  0  G1301A 0 8 24  G1301S 8 24  0  C1152S 0 8 24  time (h) 328nt •* 301nt  n m  -  m,  n  i»  f - - |  - • m ••  m  I -1 -1 -1 I  lane 1 2 3 4  5  6  7  8  9  I  -«188nt  10 11 12 13 14 15 16 17 18 19 20  B WT pbl9 §  0  lane 1 2 3  4 8  4  G1300A 24 0 4 8  5 6  7 8  R1299A  24 0 4  G1301A  8 24 0 4 8  G1301S  24 0 4  C1152S  8 24 0  4 8  24  time(h)  9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26  Figure 23. R N A analysis of the W T and mutant constructs. (A) Positive-strand R N A analysis. BHK-21 cells were electroporated with RNAs transcribed in vitro from cDNA clone containing the WT (lanes 3 to 5), mutant G1300A (lanes 6 to 8), R1299A (lanes 9 to 11), G1301A (lanes 12 to 14), G1301S (lanes 15 to 17), or C1152S (lanes 18 to 20). At indicated time of postelectroporation (0, 8, and 24 h), total cytoplasmic RNAs were extracted using TRIzol reagent and subjected to RPA using S-labeled probe pbl8, which was loaded in lane 1. BHK-21 cells electroporated with no R V R N A served as control (lane 2). The autograph was exposed for 1 day. (B) Negative-strand genomic R N A analysis. Total cytoplasmic RNAs extracted at 0, 4, 8 and 24 h postelectroporation of BHK-21 cells transfected respectively with the WT (lanes 3 to 6), G13000A (lanes 7 to 10), R1299A (lanes 11 to 14), G1301A (lanes 15 to 18), G1301S (lanes 19 to 22), or C1152S (lanes 23 to 26), were subjected to the two-cycle RPA, as described in Materials and Methods. The S-labeled probe pbl9 was loaded in lane 1. BHK-21 cells electroporated with no R V R N A served as control (lane 2). The autoradiograph was exposed for two days. The positions of the 328, 301, and 188-nt were indicated by arrowheads. Images were scanned using U M A X Astra 1220U scanner with Adobe Photoshop 5.0 software. 35  35  88  To compare the efficiencies of RNA production between the WT and mutants more precisely, I assessed the amount of negative-strand RNA, positive-strand genomic RNA, and subgenomic R N A for all the constructs and normalized the results for mutants against that for the WT. The molar ratio of subgenomic R N A to positive-strand genomic R N A was also calculated for the WT, G1300A, and R1299A. Results are summarized in Table 4.  Table 4. Comparison of the relative amounts of RNAs produced in the WT and mutants. Construct  Negative-strand RNA  b  Positive-strand RNA Genomic Subgenomic c  SG/G molar ratio  4h  8h  WT  1  1  1  1  4.70 ± 1.00  G1300A  2.54 ± 0 . 3 7  1.13 ± 0 . 4 0  0.88 ± 0 . 1 3  0.82 ± 0 . 0 8  4.45 ± 1.20  R1299A  2.05 ± 0.50  1.41 ± 0 . 3 5  0.19 ± 0 . 1 3  0.24 ± 0 . 1 0  5.15 ± 1 . 0 6  G1301A  2.21 ± 1.11  1.39 ± 0 . 3 7  N/A  N/A  N/A  G1301S  2.36 ± 0 . 5 2  0.97 ± 0 . 1 1  N/A  N/A  N/A  e  d  N/A N/A N/A 1.43 ± 0 . 3 3 1.90 ± 0 . 5 7 The values shown are the results of at least two independent experiments. The amount of negative-strand R N A at the respective time point (4 or 8 h postelectroporation) was assessed as described in Materials and Methods and normalized against that of the WT (value of 1.0). Positive-strand RNA, either genomic or subgenomic, produced at 24 h postelectroporation was assessed and normalized against that of the WT (value of 1.0). The SG/G molar ratio is the calculated molar ratio of subgenomic R N A to genomic RNA. N/A, data not available. C1152S  a  b  c  d  e  Negative-strand R N A was compared at 4 and 8 h, because these were the points when synthesis of negative-strand R N A was not complicated by new synthesis of positive-strand R N A or by reinfection. In general, the mutants produced significantly more negative-strand RNA than did the WT at 4 h, and almost equally with the WT at 8 h. I also compared the amount of negativestrand R N A between the W T and mutant G1301S every 2 h after electroporation (Fig. 24). Mutant G130IS produced substantially more negative-strand R N A at early stage from 2 to 6 h 89  than WT (Fig. 24, comparing lanes 3 to 5 to lanes 8 to 10), whereas both reached nearly the same level at 8 h (Fig. 24, lanes 6 and 11). In what way the WT differs from the mutants at early stages is unclear. One explanation might be that more efficient processing of the WT-NSP decreases the available negative-strand R N A replication complex. However, the widely varied NSP cleavage among the mutants (from 0% to 75% cleavage ratio) did not result in correlated production of negative-strand R N A at 4 h. A likely reason is that a limiting host factor(s) might define the number of replication complexes formed, so further increased amounts of p200 had no effect on levels of replication complexes available. I propose that, at 4 h, the WT p200 has not saturated the limited host factor(s) in the formation of negative-strand R N A replication complex. Increased amounts of p200 in G1300A, due to delayed and less efficient NSP cleavage, result in higher production of negative-strands. Yet further increased amounts of p200 in other mutants would not further increase negative-strand R N A synthesis because such host factor(s) are limited. At 8 h, all the constructs produced similar amount of negativestrands,  suggesting  that  they contained comparable negative-strand  R N A replication  complexes, possibly limited by host factor(s).  WT pbl9 0 2  G1301S  4 6  8 0 2 4  6 8  (h) |^328„t  lllllkllll  lanes 1 2 3 4  5 6  7 8 9  10 11  Figure 24. Comparison of WT and mutant G1301S on the levels of negative-strand RNA synthesis. BHK-21 cells were electroporated with either WT R N A (lanes 2 to 6) or mutant G130IS R N A (lanes 7 to 11). Total cellular RNAs were prepared every 2 h postelectroporation and subjected to the detection of negative-strand RNA using two-cycle RPA with pbl9 (lane 1).  90  It was assumed that mutants G1301S and C1152S synthesized no positive-strand RNA. The average density of the 301-nt band for the two at 24 h was taken to represent the level of input genomic RNA present in other mutants and was subtracted from their measured values. Although G1301A produced a low level of subgenomic RNA and possibly some genomic RNA as well (Fig. 5A), these were not subjected to quantitation or molar ratio calculation because of the likely presence of background genomic RNA. It is evident that all the mutants produced less positive-strand RNA than did the WT (Table 4). Positive-strand RNA levels in mutants are in accord with their respective NSP processing efficiencies and virus yields. The more the NSP cleavage is impaired, the less positive-strand RNA is produced. The slightly impaired mutant G1300A produced 88% of the level of genomic RNA and 82% of the subgenomic RNA produced by the WT. The substantially impaired mutant R1299A transcribed levels of 19% of genomic RNA and 24% of subgenomic RNA produced by the WT. Production of the genomic and subgenomic positive-strand RNAs were almost equally lowered. The molar ratio of subgenomic/genomic RNA, ranging from 4.5-5.2 for the mutants did not vary dramatically from the WT. These results indicate that NSP cleavage may not be the cause for the differential production of subgenomic and genomic RNA in virus-infected cells.  4.2.5.  Discussion II  The cleavage of p200 is essential for virus replication. In RNA viruses, processing of nonstructural polyprotein is temporally regulated such that the ratio of polyprotein and cleavage products changes over the course of infection. Nonstructural polyprotein processing has been demonstrated to be essential for virus replication in alphavirus (Shirako and Strauss, 1994), Flavivirus yellow fever virus (YFV) (Chambers et al., 1993, 1995; Nestorowicz et al., 1994) and Bovine viral diarrhea virus (BVDV) (Xu et al., 1997) by examining the effects of  91  mutations that inactivate the protease function or block the cleavage site with the use of infectious cDNA clones. In this study, I generated a panel of cleavage mutants and demonstrated that the cleavage of nonstructural polyprotein p200 is essential for R V replication. The effects on R V replication were found to correlate with the efficiency of p200 polyprotein processing. Mutations that completely abolished (G1301S and C1152S) or nearly abolished (G1301A) p200 cleavage shut down virus replication (Table 3). A mutation with a minor influence on NSP processing (mutant G1300A) produced infectious virus with a growth rate decreased by 3 - 10 fold. A mutation with a profound effect on NSP cleavage (mutant R1299A) was viable but with a growth rate lowered by 2000 - 3000 fold. Examination of RNA synthesis suggested that defective production of positive-strand R N A in the mutants, including both positive-strand genomic R N A and subgenomic RNA, may be the cause for reduced virus production (Table 4). More infectious constructs produced more positive-strand RNAs than less infectious ones, with noninfectious constructs producing little or none. However, given its 20% of p200 processing level and 20% of positive-strand genomic R N A , R1299A produced an unexpectedly lower level of virus yield (10 -fold lower than the WT). Although alternative 3  explanations may exist, a likely reason is reinfection. The positive-strand R N A level was compared at 24 h postelectroporation, whereas virus yields were compared at 48 h (BHK-21 cells) or 5 days (Vero cells) posttransfection. It also seems that productive release of infectious virus particles may require a threshold level of positive-strand R N A synthesis. For example, mutant G1301A is noninfectious by plaque-assay, although a rmnimal level of positive-strand RNA could be detected from RPA. I believe that these mutations act by impairing p200 cleavage rather than by directly affecting the activity of the R N A replicase. First, the introduced mutations, except for R1299A, are conserved substitutions. Second, the mutations at cleavage sites are unlikely to be important for the replicase activity. Third, for all the mutants,  92  there is a good correlation between p200 cleavage efficiency, virus production, and viral RNA synthesis.  Uncleaved polyprotein p200 can produce negative-strand RNA, whereas cleavage products from p200 are required for efficient positive-strand RNA synthesis. I have shown that all the mutants, including the noninfectious cleavage-defective mutants G1301S and C1152S, accumulated negative-strand RNA as efficiently as the WT at 8 h (Fig. 23B and Table 4), suggesting that uncleaved p200 is sufficient to produce negative-strand RNA from the input genomic RNA. Interestingly, mutants even produced more negative-strand RNA at 4 h than the WT, providing further evidence for the role of p200 in negative-strand RNA synthesis. However, the amount of negative-strand RNA did not increase proportionally to the amount of p200 among mutants, suggesting that limiting host factor(s) may also play a role in regulating the number of replication complexes for negative-strand RNA synthesis.  The capacity to synthesize positive-strand RNA differed greatly between the WT and mutants. All mutants produced lower levels of positive-strand RNA, both genomic and subgenomic RNA, than the WT. Mutants more defective in cleaving p200 produced less positive-strand RNA (Fig 21A and Table 4). The cleavage-defective mutants G1301S and C1152S showed accumulation of positive-strand RNA barely detected by RPA (Fig. 23A, lane 8). This suggests that cleavage products from p200 (i.e., pi50 and p90) are responsible for efficient synthesis of both positive-strand genomic RNA and subgenomic RNA. In view of the limited sensitivity of RPA used in the study, the possibility of an inefficient synthesis of positive-strand RNA by uncleaved p200 cannot be ruled out. For the two infectious mutants (G1300A and R1299A), the molar ratios of subgenomic RNA to positive-strand genomic RNA were not significantly  93  different from the WT, indicating that p200 cleavage does not contribute to the differential synthesis of positive-strand genomic and subgenomic RNA.  My studies suggest a strong similarity between RV and alphavirus (for a review, see Strauss and Strauss, 1994), a well-characterized positive-strand RNA virus genus, in NSP processing and viral RNA synthesis. Alphavirus NSP contains three cleavage sites generating four cleavage products (nsPl to nsP4) and a number of intermediates. A model for the composition of replication complexes and the temporal regulation of negative- and positive-strand RNAs has been proposed from several lines of studies (Lemm and Rice, 1993a, 1993b; Lemm et al., 1994, 1998; Shirako and Strauss, 1994). Three forms of replication complexes are involved in alphavirus replication: the uncleaved P123 and nsP4 generates only negative-strand RNA; the complex composed of nsPl, P23, and nsP4 is active in both negative-strand RNA and 49S positive-strand genomic RNA syntheses; the complex consisting of the final cleavage products nsPl, nsP2, nsP3, and nsP4 produces only 49S positive-strand genomic RNA and subgenomic RNA. Cleavage at the 1/2 and 2/3 sites respectively switches the product preference of the replication complex from negative- to positive-strand RNA, and also inactivates its capacity for negative-strand RNA synthesis, which explains the shutoff of negative-strand RNA synthesis after 4 to 6 hours post infection. In RV, p200 is cleaved into pi50 and p90, giving a much simpler NSP composition. This work demonstrates that the replication complex composed of polyprotein p200 is active in negative-strand RNA synthesis but incapable of efficient positivestrand RNA synthesis, while cleavage of p200 is required for efficient positive-strand RNA synthesis. For both alphavirus and RV, cleavage of polyprotein or intermediates causes the switching of negative-strand to more efficient positive-strand RNA synthesis. However, it has yet to be determined that synthesis of negative-strand RNA ceases after an early stage of RV replication. My RNase protection assay results are complicated by reinfection as well as the 94  fast growth rate of BHK-21 cells. Studies using virus-infected Vero cells are under way. It also remains to be examined whether or not the cleaved products of RV NSP, pi50 and p90, transcribe negative-strand RNA.  My work provides the first experimental data demonstrating the relationships between RV NSP cleavage and virus replication, and particularly, between NSP cleavage and viral RNA synthesis. From my results and the studies of alphavirus replication, I hypothesize that uncleaved p200 forms the replication complex for negative-strand RNA synthesis and that cleavage of p200 into pi50 and p90 converts the complex into one with the capacity for efficient positive-strand RNA synthesis. Whether pi50 and p90 also produce negative-strand RNA remains to be investigated. It is of interest that p200 is capable of negative-strand RNA synthesis but incapable of positive-strand RNA synthesis. One possibility is that recognition of positive-strand RNA promoters or initiation of positive-strand RNA synthesis needs a specific component or conformation not present in p200 but generated by its cleavage. Positive-strand RNA viruses replicate through negative-strand intermediates, the regulatory mechanism of which is worthy of study. Previous studies on alphavirus and the present one on RV may indicate a possible mechanism of RNA replication for a group of viruses, namely that the change from synthesis of negative-strand RNA to positive-strand RNA is mediated by NSP cleavage.  95  4.3.  4.3A.  Molecular characterization ofRV RNA synthesis  R V RNA replication is ris-preferential.  In order to study the roles of NSPs in synthesizing distinct viral RNAs, I employed complementation experiments, in which engineered R N A transcripts were coelectroporated into BHK-21 cells. Resulting production of viral RNAs was monitored by RPA. M33ASS is a modified R V R N A transcript with most of the SP region deleted, from nt 6966 to 9334 (Fig. 25A). M33AMM is a replication-defective R N A transcript with a frame-shift deletion in the NSP ORF from nt 1081 electroporation,  separately  to 5106 or  (Fig. 25A). R N A molecules were introduced by  together,  into  BHK-21  cells.  At  different  times  postelectroporation, total RNAs were isolated and subjected to RPA using either probes specific to both molecules (pbl8 and pbl9), or probes specific to M33AMM only (pb20 and pb21) (Fig. 25A). pbl8 is positive-strand specific and differentiates genomic from subgenomic RNA by the size of protected bands, 301 nt for genomic and 188 nt for subgenomic RNA. pb20 is also positive-strand specific but fails to differentiate genomic from subgenomic RNA, as both generate 162-nt protected bands. pbl9 and pb21, complementary to pbl8 and pb20 respectively, are negative-strand specific giving protected bands of 301 nt and 162 nt, respectively. When the M33ASS R N A was electroporated into B H K cells alone, the 301-nt band, representing negative-strand RNA, was detected by pbl9 at 4, 7, and 24 h (Fig. 25B, lanes 13 to 15). The 301-nt product, representing genomic R N A , and the 188-nt band, representing subgenomic RNA, were detected by pbl8 at 24 h (Fig. 25B, lane 4). The results demonstrated that the M33ASS R N A replicates itself in essentially the same way as does the WT M33 R N A (Fig. 23). When the M33AMM R N A was introduced alone, no signal band representing negative-strand R N A was detected by either pbl9 (Fig. 25B, lanes 17 to 19) or by  96  pb21 (Fig. 25C, lane 12), nor was the band specific to positive-strand RNAs detected by pbl8 (Fig. 25B, lane 7) or by pb20 (Fig. 25C, lane 5) at 24 h. This demonstrated that the M33AMM RNA was defective in both negative- and positive-strand R N A syntheses. When both RNAs were coelectroporated, no signal band specific to negative-strand R N A was detected at 7 and 24 h using pb21 (Fig. 25C, lanes 15 and 16), nor were bands specific to positive-strand RNAs detected by pb20 at 24 h (Fig. 25C, lane 8), indicating that replication of the M33AMM RNA was not rescued by the M33ASS RNA. The positive-strand RNAs detected by pbl8 (Fig. 25B, lane 10) and the negative-strand R N A detected by pbl9 (Fig. 25B, lanes 21 to 23) were therefore generated solely by the self-replication of the M33ASS RNA.  97  ft*- s g R N A 6966 M 3 3 A S S  5  9336  5106  1081  y  5*.  M 3 3 A M M  pbl8/pbl9  B  •y  "  pbl8  pb20/pb21  pbl9  M33ASS  M33ASS-  r  +  M33ASS M33AMM M33AMM pbl8  m  0  7 24 0  M33ASS  pbl9  0 4 7  7 24 0 7 24 328 nt 30 Int  If -  +  M33AMM 24 0  4 7  M33AMM  24 0  m  M  4 7  H  M  24 time(h)  I  - 328 nt -301nt  I~  188t  lanes  1  2  3  4  5  6  7  8  9  10  11  12  13  14  15  (+) gRNA & sg RNA  16  17 18 19  (-) gRNA  M33ASS-  t«  M33ASS-  +  < M33AMM M33AMM 7 24 0 7 24 time (h)  3 M33AMM M33AMM  pb21  2  0  162 nt 2  3  4  5  6  7  8  9  10  (+) gRNA & sgRNA  11  ~~ Z  7 24 0 7 24 ^  187 nt  lanes 1  22 23  pb21  pb20  pb20 S o  20 21  12  13  14 15  16  f  time (h)  187 nt 162 nt  17  (-) gRNA  Figure 25. Trans complementation of RV replication. (A) Schematic diagram of the replication-defective R N A M 3 3 A M M and the helper R N A M33ASS, with the relative positions of RPA probes (pbl8/pbl9, pb20/pb21). The start of the subgenomic R N A is indicated by the arrow. (B) RPAs using pbl8 (lane 1) and pbl9 (lane 11), specific to both M 3 3 A M M and M33ASS R N A . The two RNAs were introduced by electroporation either singly or in combination into BHK-21 cells. 4 p:g of total R N A prepared at 0, 7, and 24 h postelectroporation was used for positive-strand R N A detection by pbl8 (lanes 2 to 10), and 20 pig of those at 0, 4, 7, and 24 h was used for negative-strand R N A detection by pbl9 (lanes 12 to 23). The autograph was exposed for 1 day. (C) RPAs using pb20 (lane 1) and pb21 (lane 9), specific to the M 3 3 A M M R N A only. The M33ASS R N A produced no specific band using either pb20 (lane 2) or pb21 (lane 10). After the M 3 3 A M M R N A was electroporated alone or in combination with the M33ASS RNA, 4 u.g of total R N A prepared at 0, 7, and 24 h was used for positive-strand R N A detection by pb20 (lanes 3 to 8), and 20 (ig of those was used 98  for negative-strand R N A detection by pb21 (lanes 11 to 16). A reaction mixture containing 5 ng of full-length negative-strand R N A transcript produced in vitro was included as a positive control for RPA by pb21 (lane 17). The autograph was exposed for 1 day. Positions of specific protected bands are indicated. Images were scanned using an U M A X Astra 1220U scanner with Adobe Photoshop 5.0 software.  The failure of rrans-complementation of the M33AMM R N A does not appear to be due to the absence or disruption of critical cis-RNA elements. I created several replication-defective constructs with either frame-shift addition, deletions, or a stop codon in different NSP regions (Fig. 26). None of these could be rescued by the helper RNA. I therefore suggest that replication of R V RNA, or at least the negative-strand R N A , is cis-preferential. I then examined whether the failure of positive-strand R N A replication is caused by its ds-preferred replication or by the absence of prerequisite negative-strand R N A synthesis.  1686 frame-shift addition of 4 nt 2551  4287 frame-shift deletion 4213  5355 frame-shift deletion  ^3943^ mutation of G G C to stop codon T A A at nt 3943  Figure 26. Schematic diagrams of constructed replication-defective mutants. Mutants were constructed on pBRM33 with frameshift addition, deletions, and a stop codon within NSP ORF (nt 41 to 6388).  99  As described in section 4.2.4, both the C1152S mutant (containing mutated NS-pro catalytic site) and the G130IS mutant (containing mutated NSP cleavage site) are functional in negativestrand R N A synthesis but defective in positive-strand R N A synthesis, because they are unable to process the NS polyprotein p200. The two mutants differ in that p200(C1152S) can be processed by a functional protease, such as p200(G1301S), provided in trans, whereas p200(G1301S)  cannot be cleaved due to its mutated NSP cleavage  site. I designed  complementation experiments to examine the trans or cis function of pl50/p90 in synthesizing positive-strand RNAs. M33(C1152S) R N A was further modified with a deletion in the SP region from nt 6966 to 9334 to yield M33(C1152S)ASS RNA (Fig. 27A). Its replication can be separated from that of the M33(G1301S) R N A by RPA using different probes: pbl8/pbl9 is specific to both RNAs and pb20/pb21 is specific to M33(G1301S) only (Fig. 27A). When the M33(C1152S)ASS and M33(G1301S) RNAs were introduced in cells separately or together, the 301-nt signal band specific to negative-strand R N A was detected by pbl9 at 4 and 7 h (Fig. 27C), demonstrating the successful synthesis of negative-strand RNA. In contrast, when the M33(C1152S)ASS and M33(G1301S) RNAs were electroporated separately, no accumulation of the 301-nt product specific to genomic R N A or the 188-nt band specific to subgenomic RNA was observed at 24 h using pbl8 (Fig. 27B, lanes 4 and 7) or pb20 (Fig. 27B, lane 15), suggesting that synthesis of positive-strand RNAs was defective in both constructs. However, when the RNAs were introduced together, accumulations of both 301- and 188-nt products, representing genomic and subgenomic RNAs, were clearly detected by pbl 8 at 24 h (Fig. 27B, lane 10), indicating the efficient rescue of synthesis of positive-strand RNAs. However, a very low level of the 162-nt band was detected by pb20 at 24 h (Fig. 27B, lane 18). This probe is specific only to the M33(G1301S) RNA, suggesting that synthesis of positive-strand RNAs of the M33(G1301S) R N A was not functionally rescued. Therefore, the large amount of the  100  positive-strand RNAs generated (Fig. 27B, lane 10) derived mostly from the M33(C1152S)ASS RNA. A straightforward explanation is that p200(C1152S) was cleaved by p200(G1301S) into pi50 and p90. These functioned preferentially in cis to produce positive-strand RNAs efficiently from their own negative-strand template on the M33(C1152S)ASS RNA, but functioned inefficiently in trans using the negative-strand template of the M33(G1301S) RNA. These results strongly support the hypothesis that synthesis of the positive-strand RNAs is carried out preferentially in cis by replication complexes composed of pl50/p90. In conclusion, I have shown for the first time that both negative- and positive-strand R V RNAs are replicated preferentially in cis.  101  I  A  M33(C1152S)ASS 5'.  ^  6966  sgRNA 9336  3' 3'  M33(G1301S) 5' pbl8/pb!9  pb20/pb21  B. Positive-strand RNAs phi 8  pb20 CO  00  1  m  80  oo  O  BO  O  I/O  5  to  3  ro  so •J-J  oo  to p  V.  Q  CO  +  u  0  in  co  2,4-  —  co co  pbl8 0  7  24 0  7 24  0  I  §  i  1* lanes  1 2 3  pb20 2  7 24  0  5  7 24  8  9  time (h)  — 187 nt •  7  7 24  H  188 nt 6  0  u ro co  - 328 nt - 301 nt  *** 4  <1  on  10  11  •  *  —  1 6 2 nt  12 13 14 15 16 17 18  C . Negative-strand RNA oo oo  oo 00  <  oo  c  ro  3  CO  pbl9  CO  ro ro  0 4 7  -  y .  r n m  02 4 7S  0 4 7  time (h)  . n  lanes  2-+  y  1 2 3 4 5  - 328 nt -  6  7 8  |  |  -  301 nt  9 10  Figure 27. Synthesis of positive-strand RNA is cis-preferential. (A) Schematic diagram of M33(C1152S)ASS and M33(G1301S) RNAs with the relative positions of pbl8/pbl9 and pb20/pb21. The NS polyprotein p200(C1152S) translated from M33(C1152S)ASS R N A can be processed by p200(G1301S) translated from M33(G1301S) RNA. The start of the subgenomic R N A (sgRNA) is indicated. (B) Positive-strand RNAs detected by pbl8 (lane 1), specific to both M33(C1152S)ASS and M33(G1301S) RNAs, or by pb20 (lane 11), specific to M33(G1301S) RNA only. After the two RNAs were electroporated 102  separately or together, 4 \ig of total R N A prepared at 0, 7, and 24 h were used in RPA with pbl8 (lanes 2 to 10). RPA reactions using pb20 were also conducted on samples prepared from M33(G1301S) electroporated alone (lanes 13 to 15) and from coelectroporation (lanes 16 to 18). pb20 was not specific to the M33(C1152S)ASS transcript (lane 12). The autograph was exposed for 1 day. (C) Negative-strand R N A detected by pbl9 (lane 1). The two RNAs were electroporated separately or together into BHK-21 cells. 20 \ig of total R N A samples prepared at 0, 4, and 7 h postelectroporation were used for negative-strand R N A detection by pbl9 (lanes 2 to 10). The autograph was exposed for 1 day. Positions of specific protected bands are indicated. Images were scanned using an U M A X Astra 1220U scanner with Adobe Photoshop 5.0 software.  4.3.2.  Time course of R V R N A synthesis.  Incorporation of [ H] uridine into infected cells is not a sensitive assay for detection of R V 3  RNA synthesis (Hemphill et al., 1988), because of the very low level of R V R N A replication against the high background of cellular R N A synthesis. By analyzing [ H]uridine-labeled, 3  actinomycin D-resistant total R N A on glyoxal-agarose gel electrophoresis, Hemphill et al. (1988) were able to detect genomic and subgenomic viral RNAs and quantitated their respective amounts at each time point. RNA synthesis was found to increase dramatically from 6 through 19 hpi, with maximal RNA synthesis at 24 hpi (Hemphill et al., 1988). However, this study did not differentiate negative-strand from positive-strand R N A and was not sensitive enough to detect R N A synthesis at very early stages of virus infection. To determine whether R V negative-strand R N A synthesis stops after a certain time (as does that of alphavirus), a more sensitive assay is needed. In the above experiments, I employed RPA to quantitate three RV R N A species at early stages of replication. I then used the same technique to determine the amounts of three viral RNAs at each time point postinfection.  103  Vero cell monolayers on 35-mm-diameter dishes were infected with R V (M33 strain) at m.o.i. of 0.1 PFU/cell for 1 h. Total RNA was extracted every 2 h and subjected to RPA. Negativestrand R N A was detected using pbl9 in a two-cycle reaction and positive-strand RNAs were determined using probel8 in a one-cycle reaction. The total R N A at each time point was suitably diluted so that the probe used was not saturating. Intensities of the signal bands specific to the three viral RNAs were quantitated, standardized against the amount of total RNA, and plotted against infection time (Fig. 28). Note that the scale for negative-strand RNA (on the right) is much lower than that for positive-strand RNAs (on the left).  Figure 28. Accumulated amounts of viral RNAs during virus infection. Vero cell monolayers on 35-mm-diameter dishes were infected with R V (M33 strain) at m.o.i. of 0.1 PFU/cell for 1 h at 37 °C. At 2-h intervals, total RNAs were prepared, and subjected to detection of positive- and negative-strand RNAs using pbl8 and pbl9, respectively. Samples were diluted differently in order not to saturate the probe. The negative-strand RNA, positivestrand genomic and subgenomic RNAs were analyzed by RNase protection assay (RPA) as described below. Intensity of RPA band was quantitated by image analysis and converted to the amount of probe (cpm) loaded at the same time. The amount of viral R N A was calculated from the intensity of its corresponding signal band (cpm) to allow for dilution and standardized against the amount of total R N A (mg). Amounts of each viral R N A were plotted against infection times (hpi). • , negative-strand RNA; • , positive-strand genomic RNA; O , subgenomic RNA. Note positive- (left) and negative-strand R N A (right) use different y scales. Values presented are the results of two independent experiments.  104  Negative-strand R N A was first detected at 2 hpi and increased continuously until 10 hpi, remaining almost constant thereafter. This result suggests that synthesis of negative-strand R N A ceases after 10 hpi. In contrast, positive-strand RNAs (both genomic and subgenomic RNAs) appeared at low level after 4 to 6 hpi, and increased dramatically after 10 hpi. Syntheses of genomic and subgenomic RNAs seem to be synchronized and the ratio of subgenomic to genomic R N A (SG/G) remained fairly level between 2.3 and 3.7. It is of interest to note that the dramatic increase of positive-strand RNAs started after 10 hpi, when synthesis of negativestrand R N A had stopped.  4.3.4.  Discussion III  R V R N A replication is crs-preferential. For a number of positive-strand R N A viruses, R N A replication was found to require the translation of part or all of the NSP coding region (de Groot et al, 1992; Mahajan et al, 1996; Neeleman and Bol, 1999; Novak and Kirkegaard, 1994; Schaad et al, 1996; Scholthof and Jackson, 1997; van Bokhoven et al, 1993; Weiland and Dreher, 1993; White et al,  1992; Zhou and Jackson, 1996).  Using complementation  experiments, I demonstrated that neither negative- nor positive-strand RNAs of R V can be functionally rescued by frans-provided NSPs translated from another helper R N A molecule, suggesting that replication of R V RNAs depends on the translation of R V NSPs in cis. Although other explanations are possible, this requirement for translation in cis appears to be the result of the preferential cis action of p200 in synthesizing R V negative-strand RNA and the preferential cis action of pl50/p90 in synthesizing positive-strand RNAs. Mechanisms for the cis requirement of a protein remain obscure but several possibilities have been proposed (Novak and Kirkegaard, 1994). One explanation is the establishment of a template. The newly  105  synthesized p200 might interact with the positive-strand genomic R N A from which it is being translated to enable that RNA molecule to become the functional template for negative-strand RNA synthesis. The negative-strand RNA might interact with the replication complex by which it was produced to become a functional template for positive-strand R N A synthesis. More likely is the restriction of diffusion of NSPs, which could be limited by subcellular localization or protein stability. R V replication complexes, consisting of NSPs, host factors, and viral RNAs, are localized to membrane-bound virus-modified lysosomes (Lee et al., 1992; 1994; Magliano et al., 1998). Binding to membrane might restrict the diffusion of p200 and its cleavage products pl5G7p90. As a negative-strand RNA replicase, p200 is probably present at high concentration only near the R N A from which it is translated, because of efficient selfprocessing. Therefore, both subcellular localization and protein stability could restrict p200 from using other R N A templates. After the cleavage of p200 into pl507p90, the replication complex may still remain associated with the newly-synthesized negative-strand RNA, serving as its preferential template. The mechanisms for the ds-preferential replication of R V need further investigation.  Synthesis of RV negative- and positive-strand RNAs is regulated by NSP cleavage. In a precise determination of the time courses of three R V RNAs, I have shown that negative-strand RNA accumulates until 10 hpi and remains nearly constant thereafter. In contrast, positivestrand RNAs (both genomic and subgenomic) do not increase much before 10 hpi and accumulate rapidly thereafter. Synthesis of negative-strand R V R N A seems to stop after 10 hpi with a switch to efficient synthesis of positive-strand RNAs. This replication pattern of R V resembles that of alphavirus, in which negative-strand R N A synthesis ceases after 4 to 6 hpi and switches to the positive-strand RNA synthesis (Sawicki et al., 1981).  106  The switch from the negative-strand R V R N A synthesis to the production of positive-strand RNAs is regulated by the cleavage of NSP, converting p200 into pl50/p90. In studies described in section 4.2.4, I have shown that polyprotein p200 is functional in negative-strand R N A synthesis and the cleavage of p200 into pl50/p90 is required for efficient synthesis of positivestrand R N A (Fig. 23 and Table 4). One question to be answered is whether pl50/p90 produces negative-strand R N A as well. A straightforward approach to study the roles of pi50 and p90 in negative-strand R N A synthesis is to conduct reconstitution or complementation experiments, in which pi50 and p90 are provided separately from either expression vectors or from two R N A molecules cotransfected and examined for their capacity to replicate a template RNA. However, this approach is unsuitable for R V because its R N A replication is c/s-preferential (Fig. 25 and Fig. 27). Furthermore, no mutation has been identified in R V comparable to N614D in alphavirus, which enhances the protease activity so much that no polyprotein can be detected (Strauss et al., 1992; Strauss and Strauss, 1994). My results suggest that pl50/p90 is not an effective replicase for negative-strand R N A synthesis, because (1) synthesis of negativestrand R N A increases only in the early stages of infection and stops later when the concentration of pl50/p90 increases (Fig. 28); (2) at very early stages of infection, cleavagedefective mutants with lower NSP cleavage efficiencies and thus lower levels of pl50/p90 produce higher levels of negative-strand RNA than WT (Fig. 23 and Table 4). It is likely that RV shares a similar mechanism for regulation of R N A synthesis with alphavirus, in which different viral R N A species are produced by distinct replication complexes containing different NSP components that result from NSP processing. In alphavirus, uncleaved PI23 and nsP4 form the replicase for negative-strand RNA; nsPl, P23, and nsP4 form the replicase for both negative-strand R N A and 49S positive-strand genomic RNA; the final cleavage products, nsPl, nsP2, nsP3, and nsP4, form the replicase only for 49S positive-strand genomic R N A and subgenomic RNA. The NSP of R V is cleaved only once and thus gives much simpler 107  components. p200 is the main negative-strand R N A replicase, whereas pl50/p90 is the main positive-strand R N A replicase for both genomic and subgenomic RNAs. Therefore, the timedependent transition from negative- to positive-strand R N A synthesis is controlled by the temporal regulation of the processing of p200 into pl50/p90, which not only activates the efficient positive-strand R N A synthesis but also shuts off the negative-strand R N A synthesis. Early in infection, levels of p200 support the accumulation of negative-strand R N A while low levels of cleavage products pl507p90 produce limited positive-strand RNAs. After 10 hpi, the processing of much of the translated p200 and perhaps limited novel NSP translation significantly increase the level of pl50/p90, causing a dramatic increase in production of positive-strand RNAs. At this stage, synthesis of negative-strand R N A is largely shut down due to lack of intact p200. My work contributes to understanding the mechanism of viral R N A synthesis.  All positive-strand  R N A viruses  replicate  through a negative-strand R N A  intermediate and the regulatory mechanism of the transition from negative- to positive-strand R N A synthesis is of great interest. From previous studies on alphavirus (Lemm and Rice, 1993, 1994, Lemm et al., 1994, 1998; Shirako and Strauss, 1994) and my work on RV, the temporal regulation of polyprotein cleavage may represent a common strategy for viral R N A regulation. In addition, the binding of host factors to the replication complexes may also play a critical role in this process (Pogue et al, 1994).  Mechanism for R V NSP translation, processing, and R N A synthesis. From previous studies on other positive-strand R N A viruses, particularly alphavirus (Lemm and Strauss, 1993a, 1993b; Lemm et al, 1994, 1998; Shirako and Strauss, 1994; Novak and Kirkegaard, 1994) and my work on R V , a mechanism of R V NSP translation, processing and R N A synthesis is proposed (Fig. 29). Upon infection, the input genomic R N A serves as the template for 108  translation of polyprotein p200. This, associated with host factors, binds to the 3' end of the RNA, from which it is being translated, and functions in cis to synthesize a full-length negative-strand RNA. The subsequent processing of p200 into pl50/p90 converts the original replication complex into one with specificity for synthesis of positive-strand RNAs. This complex remains associated with the nascent negative-strand RNA template, recognizes the promoter for either genomic RNA or subgenomic RNA, and functions in cis to produce genomic or subgenomic RNA efficiently.  iif  ribosome  ^  /-»)'^^ —p ' r2 v O0 l 0 3  (-) gRNA  < -  host factors  translation of p200, which, along with host factors, generates (-) gRNA in cis.  p200 3' 5'-  p200 is processed into p150/p90, which converts the original replication complex into efficient (+) RNAs synthesis. 40S gRNA 24S sgRNA  replication of (+) gRNA and transcription of (+) sgRNA  Figure 29. Proposed mechanism for RV NSP translation, processing, and viral RNA synthesis. Upon infection, p200 is translated from the input genomic RNA (gRNA) and possibly binds to the 3' end of the template RV RNA. Along with host factors, p200 functions in cis to generate a full-length negative-strand RNA. The subsequent processing of p200 into pl50/p90 not only disables the capacity for negative-strand RNA synthesis, but also permits efficient positivestrand RNA synthesis, including both 40S genomic RNA and 24S subgenomic RNA (sgRNA).  109  5. SUMMARY AND PERSPECTIVE  In summary, my work identified the domains required by RV NS-pro for cleavages in trans and cis (Fig. 17). Both cleavages require a core catalytic domain from A1020 to  P1296,  while  containing different N and C ends. To cleave in trans protease needs an additional domain (V920  to A1020) at the N end, while cis protease contains a four-residue linker, the cleavage site  G1301 and the substrate region at the C end. I also demonstrated that the X domain is important in trans cleavage of RV NS-pro but has no significant influence on cis cleavage. Defining the regions and roles of protease-related domains of RV NS-pro clarifies our understanding of this specific viral PCP and provides a basis for comparison with other proteases. Primary sequence analysis and predicted secondary structure of RV NS-pro showed distant homology to papain. Crystallographic data are needed for more precise three-dimensional structure determination.  My work also characterized the features of RV RNA synthesis at a molecular level and its regulation by NSP cleavage. Both negative- and positive-strand RNAs are synthesized preferentially in cis. Synthesis of negative-strand RNA stops around 10 hpi and is switched to the efficient production of positive-strand RNAs. Polyprotein p200 is required for negativestrand RNA synthesis but not for positive strands, whereas its cleavage products pl50/p90 are required for efficient positive-strand RNA synthesis. A molecular mechanism is proposed for RV NSP translation, processing and RNA synthesis. Upon infection, NSP is translated from input genomic RNA into p200, which remains bound to the 3' end of the translation-template RNA and functions to generate a full-length negative-strand RNA, with the help of host factors. The cleavage of p200 into pl50/p90 leaves the replicase with little or no capacity for negativestrand RNA synthesis, but allows efficient activity for positive-strand RNA synthesis, resulting in the large amount of genomic RNA and subgenomic RNA produced thereafter. 110  Some questions still remain to be answered in future studies on R V NSPs and viral R N A replication. It will be interesting to define the protein-protein interaction domains among R V NSPs, including the proposed protein-protein interaction domains within NS-pro, and to characterize the effects of these interactions on viral R N A synthesis. The underlying reason for the differential synthesis of genomic and subgenomic R N A has not been characterized. Whether it is because of additional host factors, different promoter activities, or other reasons, is worthy of studying. Host factors identified so far as binding specifically to R V R N A are autoantigens (calreticulin, La autoantigen), whose functions for viral R N A synthesis remain elusive. Characterization of the complete components of replication complexes, including both NSPs and host factors, is needed for a clear understanding of R V R N A replication. Such knowledge may illuminate good targets for prevention of R V infection and combating this and other virus induced diseases.  Ill  REFERENCES Adkins, S., R. Siegel, J. H. Sun, and C. C. Kao. 1997. Minimal templates directing accurate initiation of subgenomic RNA synthesis in vitro by the brome mosaic virus RNA-dependent RNA polymerase. RNA 3: 634—647. Ahlquist, P. 1992. Bromovirus RNA replication and transcription. Curr. Opin. Genet. Dev. 2: 71-76. Ahola, T., and P. Ahlquist. 1999. Putative RNA capping activities encoded by brome mosaic virus: methylation and covalent binding of guanylate by replicase protein la. J. Virol. 73(12): 10061-10069. Aliperti, G., and M. J. Schlesinger. 1978. Evidence for an autoprotease activity of Sindbis virus capsid protein. Virology 90: 366-369. Allaire, M., M. M. Chernaia, B. A. Malcolm, and M. N. James. 1994. Picornaviral 3C cysteine proteinases have a fold similar to chymotrypsin-like serine proteinases. Nature 369(6475): 72-6. Andino, R., G. E. Rieckhof, and D. Baltimore. 1990. A functional ribonucleoprotein complex forms around the 5' end of poliovirus RNA. CeU 63(2): 369-380. Andino, R., G. E. Rieckhof, P. L. Achacoso, and D. Baltimore. 1993. Poliovirus RNA synthesis utilizes and RNP complex formed around the 5'-end of viral RNA. EMBO J. 12:3587-3598 Anthony, R. P., and D. T. Brown. 1991. Protein-protein interactions in an alphavirus membrane. J. Virol. 65: 1187-94. Baker, D., J. L. Sohl, and D. A. Agard. 1992. A protein-folding reaction under kinetic control. Nature 356: 263-265. Baker, S. C , K. Yokomori, S. Fong, R. Carlisle, A. E. Gorbalenya, E. V. Koonin, and M. M. C. Lai. 1993. Identification of the catalytic sites of a papain-like cysteine proteinase of murine coronavirus. J. Virol. 67: 6056-6063. Bardeletti, G., J. Tektoff, and D. Gautheron. 1979. Rubella virus maturation and production in two host cell system. Intervirology 11: 97-103. Bardeletti, G., N. Kessler, N. Aymard-Henry. 1975. Morphology, biochemical analysis and neuraminidase activity of rubella virus. Arch Virol. 49: 175-186. Baron, M. D., and K. Forsell. 1991. Oligomerization of the structural proteins of rubella virus. Virology 185: 811-819. Baron, M. D., T. Ebel, and M. Suomalainen. 1992. Intracellular transport of rubella virus structural proteins expressed form cloned cDNA. J. Gen. Virol. 73: 1073-1086. 112  Barton, D. J., and J. B. Flanegan. 1993. Coupled translation and replication of poliovirus RNA in vitro: synthesis of functional 3D polymerase and infectious virus. J. Virol. 67: 822831. Barton, D. J., and J. B. Flanegan. 1997. Synchronous replication of poliovirus RNA: initiation of negative-strand RNA synthesis requires the guanidine-inhibited activity of protein 2C. J. Virol. 71(11): 8482-8489 Barton, D. J., B. J. Morasco, and J. B. Flanegan. 1996. Assays for poliovirus polymerase, 3D(Pol), and authentic RNA replication in HeLa S10 extracts. Methods Enzymol. 275: 35-57. Barton, D. J., B. J. Morasco, and J. B. Flanegan. 1999. Translating ribosomes inhibit poliovirus negative-strand RNA synthesis. J.Virol. 73(12): 10104-10112. Barton, D. J., E. P. Black, and J. B. Flanegan. 1995. Complete replication of poliovirus in vitro: preinitiation RNA replication complexes require soluble cellular factors for the synthesis of VPg-linked RNA. J. Virol. 69: 5516-5527. Barton, D. J., S. G. Sawicki, and D. L. Sawicki. 1991. Solubilization and immunoprecipitation of alphavirus replication complexes. J. Virol. 65:1496-1506. Bazan, J. F., and R. J. Fletterick. 1988. Viral cysteine proteases are homologous to the trypsin-like family of serine proteases: structural and functional implications. Proc. Natl. Aca. Sci. USA. 85: 7872-7876. Berti, P. J and A. C. Storer. 1995. Alignment/phylogeny of the papain superfamily of cysteine proteases. J. Mol. Biol. 246: 273-283. Bienz, K., D. Egger, M. Troxler, and L. Pasamontes. 1990. Structural organization of poliovirus RNA replication is mediated by viral proteins of the P2 genomic region. J. Virol. 64: 1156-1163. Bisaillon, M., and G. Lemay. 1997. Viral and cellular enzymes involved in synthesis of mRNA cap structure. Virology 236: 1-7. Blyn, L. B., R. Chen, B. L. Semler, and E. Ehrenfeld. 1995. Host cell proteins binding to domain IV of the 5' noncoding region of poliovirus RNA. J. Virol. 69: 4381-4389. Bonilla, P. J., S. A. Hughes, and S. R. Weiss. 1997. Characterization of a second cleavage site and demonstration of activity in trans by the papain-like protease of the murine coronavirus mouse hepatitis virus A59. J. Virol. 71: 900-909. Bonilla, P. J., S. A. Hughes, J. F. Pinon, and S. R. Weiss. 1995. Characterization of the leader papain-like proteinase of MHV-A59: identification of a new in vitro cleavage site. Virology 209: 489-497. Bowden, D. S., and E. G. Westaway. 1984. Rubella virus structural and nonstructural proteins. J. Gen. Virol. 65: 933-943. 113  Bowden, D. S., and E. G. Westaway. 1985. Changes in glycosylation of rubella virus envelope proteins during maturation. J. Gen. Virol. 66: 201-206. Buck, K. W. 1996. Comparison of the replication of positive-stranded RNA viruses of plants and animals. Adv. Virus Res. 47: 159-251. Brass, V., and D. Ganem. 1991. The role of envelope proteins in hepatitis B virus assembly. Proc. Natl. Acad. Sci. USA 88: 1059-1063. Bryan, P., L. Wang, J. Hoskins, S. Ruvinov, S. Strausberg, P. Alexander, O. Almog, G. Gilliland, and T. Gallagher. 1995. Catalysis of a protein reaction: mechanistic implications of the 2.0 A structure of the subtilisin-prodomain complex. Biochemistry 34: 10310-10318. Calisher, C. H., and N. Karabatsos. 1988. Arbovirus serogroups: definition and geographic distribution, p. 19-57. In T. P. Monath (ed.), The arboviruses: epidemiology and ecology. CRC Press, Inc., Baca Raton, Fla. Chambers, T. J., A. Nestorowicz, and C. M. Rice. 1995. Mutagenesis of the yellow fever virus NS2B/3 cleavage site: determinants of cleavage site specificity and effects on polyprotein processing and viral replication. J. Virol. 69: 1600-1605. Chapman, M. R., and C. C. Kao. 1999. A minimal RNA promoter for minus-strand RNA synthesis by the brome mosaic virus polymerase complex. J Mol Biol. 286(3): 709-720. Chaye, H., P. Chong, B. Tripet, B. Brush, and S. Gillam. 1992. Localization of the virus neutralizing and hemagglutinin epitopes of E l glycoprotein of rubella virus. Virology 189: 483-492. Chen, J., and P. Ahlquist. 2000. Brome mosaic virus polymerase-like protein 2a is directed to the endoplasmic reticulum by helicase-like viral protein la. J Virol. 74(9): 4310-4318. Chen, J.-P., J. H. Strauss, E. G. Strauss, and T. K. Frey. 1996. Characterization of the rubella virus nonstructural protease domain and its cleavage site. J. Virol. 70: 4707-4713. Chen, M. H., and T. K. Frey. 1999. Mutagenic analysis of the 3' c/s-acting elements of the rubella virus genome. J. Virol. 73(4): 3386-403. Cheng, R. H., R. J. Kuhn, N. H. Olson, M. G. Rossmann, H. K. Choi, T. J. Smith, T. S. Baker. 1995. Nucleocapsid and glycoprotein organization in an enveloped virus. Cell 80(4): 621-30. Choi, G. H., D. M. Pawlyk, and D. L. Nuss. 1991. The autocatalytic protease p29 encoded by hypovirulence-associated virus of the chestnut blight fungus resembles the potyvirus-encoded protease HC-Pro. Virology 183: 747-752. Choi, H. K., L. Tong, W. Minor, P. Dumas, U. Boege, M. G. Rossmann, G. Wengler. 1991. Structure of Sindbis virus core protein reveals a chymotrypsin-like serine proteinase and the organization of the virion. Nature 354(6348): 37-43. 114  Clarke, D. M., T. W. Loo, H. McDonald, and S. Gillam. 1988. Expression of rubella virus cDNA coding for the structural proteins. Gene 65:23-30. Clarke, D. M., T. W. Loo, I. Hui, P. Chong, and S. Gillam. 1987. Nucleotide sequence and in vitro expression of rubella virus 24S subgenomic mRNA encoding the structural proteins E l , E2, and C. Nucleic Acid Res. 15: 3041-3057. de Groot, R. J., R. G. van der Most, and W. J. M. Spann. 1992. The fitness of defective interfering murine coronavirus Dl-a and its derivatives is decreased by nonsense and frameshift mutations. J. Virol. 66: 5898-5905. de Groot, R. J., W. R. Hardy, Y. Shirako, and J. H. Strauss. 1990. Cleavage-site preferences of Sindbis virus polyproteins containing the nonstructural proteinase: evidence for temporal regulation of proteinase: evidence for temporal regulation of polyprotein processing in vivo. EMBO J. 9: 2631-2638. Delchambre, M., D. Gheysen, D. Thines, C. Thiriart, E. Jacobs, E. Verdin, M. Horth, A. Burny, and F. Bex. 1989. The GAG precursor of simian immunodeficiency virus assembles into virus-like particles. EMBO J. 8: 2653-2660. Den Boon, J. A., K. S. Faaberg, J. J. M. Meulenberg, A. I. M. Wassenaar, P. G. W. Plagemann, A. E. Gornalenya, and E. J. Snijder. 1995. Processing and evolution of the Nterminal region of the arterivirus replicase ORFla protein: identification of two papain-like cysteine proteases. J. Virol. 69: 4500-4505. Diez, J., M. Ishikawa, M. Kaido, and P. Ahlquist. 2000. Identification and characterization of a host protein required for efficient template selection in viral RNA replication. Proc Natl Acad Sci U S A . 97(8): 3913-3918. Dominguez, G., C. Y. Wang and T. K. Frey. 1990. Sequence of the genome RNA of rubella virus: Evidence for genetic rearrangement during Togavirus evolution. Virology 177: 225-238. Dorsett, P. H., D. C. Miller, K. Y. Green, F. I. Byrd. 1985. Structure and function of the rubella virus proteins. Rev Infect Dis. Suppl 1: S150-6. Doxsey, S. J., F. M. Brodsky, G. S. Blank, A. Helenius. 1987. Inhibition of endocytosis by anti-clathrin antibodies. Cell 50(3): 453-63. Dreher, T. W., and T. C. Hall. 1988a. Mutational analysis of the sequence and structural requirements in brome mosaic virus RNA for minus-strand promoter activity J. Mol. Biol. 201: 31-40. Dreher, T. W., and T. C. Hall. 1988b. Mutational analysis of the tRNA mimicry of brome mosaic virus RNA J. Mol. Biol. 201: 41-55. Forng, R.-Y., and T. K. Frey. 1995. Identification of the rubella virus nonstructural proteins. Virology. 206: 843-853. 115  Francki, R. I. B., C. M. Fauquet, D. L. Knudson, and F. Brown, (eds) 1991. Classification and nomenclature of viruses. Fifth report of the international committee on Taxonomy of Viruses. Archives of Virology, Suppl 2. Springer-Verlag, Vienna. Frey, T. K. 1994. Molecular biology of rubella virus. Adv. in Virus Res. 44: 69-160. Frey, T. K., and I. D. Marr. 1988. Sequence of the region coding virion proteins C and E2 and the carboxy terminus of the nonstructural proteins of rubella virus: comparison to alphavirus. Gene 62: 85-99. Frolova, E., I. Frolov, S. Schlesinger. 1997. Packaging signals in alphaviruses. J. Virol. 71(1): 248-58. Furuya, T., and M. M. C. Lai. 1993. Three different cellular proteins bind to complementary sites on the 5'- end-positive and 3'-end-negative strands of mouse hepatitis virus RNA. J. Virol. 67: 7215-7222. Gallagher, P. J., J. M. Henneberry, J. F. Sambrock, and M. J. H. Gething. 1992. Glycosylation requirements for intracellular transport and function of the hemagglutinin of influenza virus. J. Virol. 66: 7136-7145. Garoff, H., R. Hewson, and D. J. E . Opstelten. 1998. Virus maturation by budding. Microbiol Mol Biol Rev. 62(4): 1171-1190. Review. Gerna, G., M. G. Revello, M. Dovis, E. Petruzzelli, G. Achilli, E. Percivall, M. Torsellini. 1987. Synergistic neutralization of rubella virus by monoclonal antibodies to viral haemagglutinin. J. Gen. Virol. 68: 2007-2012. Gheysen, D., E . Jacobs, F. de Foresta, C. Thiriart, M. Francotte, D. Thines, and M. De Wilde. 1989. Assembly and release of FflV-1 precursor Pr55gag virus-like particles from recombinant baculovirus-infected cells. Cell 59: 103-112 Gorbalenya, A. E., A. P. Donchenko, V. M. Blinov, and E . V. Koonin. 1989a. Cysteine proteases of positive strand RNA viruses and chymotrypsin-like serine proteases. A distinct protein superfamily with a common structural fold. FEBS Lett. 243(2): 103-114. Gorbalenya, A. E., E. V. Koonin, A. P. Donchenko, and V. M. Blinov. 1989b. Coronavirus genome: prediction of putative functional domains in the non-structural polyprotein by comparative amino acid sequence analysis. Nucleic Acids Res. 17(12): 4847-4861. Gorbalenya, A. E . , E . V. Koonin, and M. M.-C. Lai. 1991. Putative papain-related thiol proteases of positive-strand RNA viruses. FEBS Lett. 288: 201-205. Gorbalenya, A. E., E. V. Koonin, and Y. I. Wolf. 1990. A new superfamily of putative NTPbinding domains encoded by genomes of small DNA and RNA viruses. FEBS Lett. 262: 145148.  116  Gorbalenya, A. E., V. M. Blinov, A. P. Donchenko, and E. V. Koonin. 1989. An NTPbinding motif is the most conserved sequence in a highly diverged monophyletic group of proteins involved in positive strand RNA virus replication. J. Mol. Evol. 28: 256-268. Green, K. Y., and P. H. Dorsett. 1986. Rubella virus antigens: localization of epitopes involved in hemagglutination and neutralization by using monoclonal antibodies. J. Virol. 57(3): 893-898. Guarne, A., J. Tormo, R. Kirchweger, D. Pfistermueller, I. Fita, and T. Skern. 1998. Structure of the foot-and-mouth disease virus leader protease: a papain-like fold adapted for self-processing and eIF4G recognition. EMBO J. 17: 7469-7479. Hardy, W. R., and J. H. Strauss. 1989. Processing the nonstructural polyproteins of Sindbis virus: nonstructural proteinase is in the C-terminal half of nsP2 and functions both in cis and in trans. J. Virol. 63: 4653-4664. Hemphill, M. L., R.-Y. Forng, E. S. Abernathy, and T.K. Frey. 1988. Time course of virusspecific macromolecular synthesis during rubella virus infection in Vero cells. Virology 162: 65-75. Hobman, T. C , and S. Gillam. 1989. In vitro and in vivo expression of rubella virus E2 glycoprotein: the signal peptide is located in the C-terminal region of capsid protein. Virology 173: 241-250. Hobman, T. C , L. Woodward, and M. G. Farquhar. 1992. The rubella virus E l glycoprotein is arrested in a novel post-ER, pre-Golgi complex. J. Cell Biol. 118: 781-792. Hobman, T. C , L. Woodward, and M. G. Farquhar. 1993. The rubella virus E2 and E l spike glycoproteins are targeted to the Golgi complex. J. Cell Biol. 121: 269-281. Hobman, T. C , M. L. Lundstrom, and S. Gillam. 1990. Processing and intracellular transport of rubella virus structural proteins in COS cells. Virology 178: 122-133. Hobman, T. C , N. O. Seto., and S. Gillam. 1994. Expression of soluble forms of rubella virus glycoproteins in mammalian cells. Virus Res. 31: 277-289. Hobman, T. C , R. Shukin, and S. Gillam. 1988. Translocation of rubella virus glycoprotein E l into the endoplasmic reticulum. J. Virol. 62: 4259-4264. Hobman, T. C , Z. Qiu, H. Chaye, and S. Gillam. 1991. Analysis of rubella virus E l glycosylation mutants expressed in COS cells. Virology 181: 768-772. Ho-Terry, L., and A. Cohen. 1984. The role of glycosylation on haemagglutination and immunological reactivity of rubella virus. Arch Virol. 79: 139-146. Ho-Terry, L., and A. Cohen. 1985. Rubella virus hemagglutinin: association with a single virion glycoprotein. Arch Virol. 72: 47-54.  117  Ishikawa, M., J. Diez, M. Restrepo-Hartwig, and P. Ahlquist. 1997. Yeast mutations in multiple complementation groups inhibit brome mosaic virus RNA replication and transcription and perturb regulated expression of the viral polymerase-like gene. Proc Natl Acad Sci U S A . 94(25): 13810-13815. Ito, Y., and M. M. C. Lai. 1997. Determination of the secondary structure and cellular protein binding to the 3-untranslated region of the hepatitis C virus RNA genome. J. Virol. 71: 86988706. Janda, M., and P. Ahlquist. 1993. RNA-dependent replication, transcription, and persistence of brome mosaic virus RNA replicons in S. cerevisiae. Cell 72(6): 961-970. Jin, H., G. P. Leser, and R. A. Lamb. 1994. The influenza virus hemagglutinin cytoplasmic tail is not essential for virus assembly or infectivity. EMBO J. 13: 5504-5515.  Justice, P. A., W. Sun, Y. Li, Z. Ye, P. R. Grigera, and R. R. Wagner. 1995. Membrane vesiculation function and exocytosis of wild-type and mutant matrix proteins of vesicular stomatitis virus. J. Virol. 69: 3156-3160.  Kadare, G., and A.-L. Haenni. 1997. Virus-encoded RNA helicases. J. Virol. 71: 2583-2590. Review. Kamer, G., and P. Argos. 1984. Primary structural comparison of RNA-dependent polymerase from plant, animal and bacterial viruses. Nucleic Acid Res. 12: 7269-7282. Kamphuis, I. G., J. Drenth and E. N. Baker. 1985. Thiol proteases. Comparative studies based on the high-resolution structures of papain and actinidin, and on amino acid sequence information for cathepsins B and H, and stem bromelain. J. Mol Biol. 182(2): 317-329. Kamphuis, I. G., K. H. Kalk, M. B. A. Swarte and J. Drenth. 1984. Structure of papain refined at 1.65 A resolution. J. Mol. Biol. 179: 233-256. Kao, C. C , and J. H. Sun. 1996. Initiation of minus-strand RNA synthesis by the brome mosaic virus RNA-dependent RNA polymerase: use of oligoribonucleotide primers J. Virol. 70: 6826-6830. Katow, S., and A. Sugiura. 1988a. Conformational change of rubella virus spike proteins induced by 2-mercaptoethanol. Jpn J Med Sci Biol. 41: 109-115. Katow, S., and A. Sugiura. 1988b. Low pH-induced conformational change of rubella virus envelope proteins. J. Gen. Virol. 69: 2797-2807. Kong, F., K. Sivakumaran, and C. Kao. 1999. The N-terminal half of the brome mosaic virus la protein has RNA capping-associated activities: specificity for GTP and Sadenosylmethionine. Virology 259(1): 200-210. Koonin, E. V., and V. V. Dolja. 1993. Evolution and taxonomy of positive-strand RNA viruses: implications of comparative analysis of amino acid sequences. Crit. Rev. Biochem. Mol. Biol. 28: 375-430. 118  Kowal, E. V., and V. Stollar. 1981. Temperature-sensitive host-dependent mutants of Sindbis virus. Virology 114: 140-148.  Krol, M. A., N. H. Olson, J. Tate, J. E. Johnson, T. S. Baker, and P. Ahlquist. 1999. RNAcontrolled polymorphism in the in vivo assembly of 180-subunit and 120-subunit virions from a single capsid protein. Proc Natl Acad Sci U S A . 96(24): 13650-13655.  Kuhn, R. J., X. Hong, and J. H. Strauss. 1990. Mutagenesis of the 3' nontranslated region of Sindbis virus RNA. J. Virol. 64: 1465-1476.  Laemmli, U . K. 1970. Cleavage of structural proteins during assemble of the head of bacteriophage T4. Nature 227: 680-685.  Lee, J. Y., D. Hwang, and S. Gillam. 1996. Dimerization of rubella virus capsid protein is not required for virus particle formation. Virology 216: 223-227.  Lee, J.-Y., J. A. Marshall, and D. S. Bowden. 1992. Replication complexes associated with the morphogenesis of rubella virus. Arch. Virol. 122: 95-106.  Lee, J.-Y., J. A. Marshall, and D. S. Bowden. 1994. Characterization of rubella virus replication complexes using antibodies to double-stranded RNA. Virology 200: 307-312. Lemm, J. A., and C. M. Rice. 1993a. Assembly of functional Sindbis virus R N A replication complexes: requirement for coexpression of P123 and P34. J. Virol. 67: 1905-1915.  Lemm, J. A., and C. M . Rice. 1993b. Roles of nonstructural polyproteins and cleavage products in regulating Sindbis virus R N A replication and transcription. J. Virof 67: 1916-1926.  Lemm, J. A., A. Bergqvist, C. M. Read, and C. M . Rice. 1998. Template-dependent initiation of sindbis virus RNA replication in vitro. J. Virol. 72: 6546-6553.  Lemm, J. A., T. Rumenapf, R. G. Strauss, J. H. Strauss, and C. M. Rice. 1994. Polypeptide requirements for assembly of functional Sindbis virus replication complexes: a model for the temporal regulation of minus-strand and plus-strand RNA-synthesis. E M B O J. 13: 2925-2934.  Li, H. P., X. Zhang, R. Duncan, L. Comai, and M. M. Lai. 1997. Heterogeneous nuclear ribonucleoprotein A l binds to the transcription-regulatory region of mouse hepatitis virus RNA. Proc. Natl. Acad. Sci. U S A 94: 9544-9549.  Li, Y., L. Luo, M. Schubert, R. R. Wagner, and C. Y. Kang. 1993. Viral liposomes released from insect cells infected with recombinant baculovirus expressing the matrix protein of vesicular stomatitis virus. J. Virol. 67: 4415-4420. Liu, C , and G. M . Air. 1993. Selection and characterization of a neuraminidase-minus mutant of influenza virus and its rescue by cloned neuraminidase genes. Virology 194: 403-407.  119  Liu, C , M. C. Eichelberger, R. W. Compans, and G. M. Air. 1995. Influenza type A virus neuraminidase does not play a role in viral entry, replication, assembly, or budding. J. Virol. 69: 1099-1106.  Liu, X., S. L. Ropp, R. J. Jackson, and T. K. Frey. 1998. The rubella virus nonstructural protease requires divalent cations for activity and functions in trans. J. Virol. 72: 4463-4466. Liu, Z., D. Yang, Z. Qiu, K.-T. Lim, P. Chong, and S. Gillam. 1996. Identificiation of domains in rubella virus genomic R N A and capsid protein necessary for specific interaction. J. Virol. 70: 2184-2190.  Lundstrom, M . L., C. A. Mauracher, and A. J. Tingle. 1991. Characterization of carbohydrates linked to rubella virus glycoprotein E l . J. Gen. Virol. 72: 843-850.  Mach, L., J. S. Mort, and J. Glossl. 1994. Noncovalent complexes between the lysosomal proteinase cathepsin B and its propeptide account for stable, extracellular, high molecular mass forms of the enzyme. J. Biol. Chem. 269: 13036-13040.  Magliano, D., J. A. Marshall, D. S. Bowden, N. Vardaxis, J. Meanger, and J.-Y. Lee. 1998. Rubella virus replication complexes are virus-modified lysosomes. Virology 240: 57-63.  Mahajan, S., V. Dolja, and J. C. Carrington. 1996. Roles of the sequence encoding tobacco etch virus capsid protein in genome amplification : requirements for the translation process and a ds-active element. J. Virol. 70: 4370-4379.  Maniatis, T., E. F. Fritsch, and J. Sambrook. 1992. Molecular cloning: a laboratory manual, 2  nd  ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, New York, N.Y.  Mao, P. L, Y. Jiang, B. Y. Wee and A. G. Porter. 1998. Activation of caspase-1 in the nucleus requires nuclear translocation of pro-caspase-1 mediated by its prodomain. J Biol Chem. 273(37): 23621-23624.  Marr, L. D., A. Sanchez, T. K. Frey. 1991. Efficient in vitro translation and processing of the rubella virus structural proteins in the presence of microsomes. Virology 180: 400-5.  Marr, L. D., C.-Y. Wang, and T. K. Frey. 1994. Expression of the rubella virus nonstructural protein ORF and demonstration of proteolytic processing. Virology. 198: 1-7. Matthews, D. A., W. W. Smith, R. A. Ferre, B. Condon, G. Budahazi, W. Sisson, J. E. Villafranca, C. A. Janson, H. E. McElroy, C. L. Gribskov, and S. Worland. 1994. Structure of human rhinovirus 3C protease reveals a trypsin-like polypeptide fold, RNA-binding site, and means for cleaving precursor polyprotein. Cell 77(5): 761-71.  Mauracher, C. A., S. Gillam, R. Shukin, and A. J. Tingle. 1991. pH-dependent solubility shift of rubella virus capsid protein. Virology 181: 773-777.  McDonald, H., T. C. Hobman, and S. Gillam. 1991. The influence of capsid protein cleavage on the processing of E2 and E l glycoproteins of rubella virus. Virology 183: 52-60.  120  Mclntyre, G. F. and A. H. Erickson. 1993. The lysosomal proenzyme receptor that binds procathepsin L to microsomal membranes at pH5 is a 43-kDa integral membrane protein. Proc. Natl. Acad. Sci. USA 90: 10588-10592. Mebatsion, T., M. Konig, and K.-K. Conzelmann. 1996. Budding of rabies virus particles in the absence of the spike glycoprotein. Cell 84: 941-951 Melancon, P., and H. Garoff. 1987. Processing of the Semliki Forest virus structural polyprotein: role of the capsid protease. J. Virol. 61: 1301-1309. Miller, W. A., J. J. Bujarski, T. W. Dreher, and T. C. Hall. 1986. Minus-strand initiation by brome mosaic virus replicase within the 3' tRNA-like structure of native and modified RNA templates. J. Mol. Biol. 187: 537-546. Mitnaul, L. J., M. R. Castrucci, K. G. Murti, and Y. Kawaoka. 1996. The cytoplasmic tail of influenza A virus neuraminidase (NA) affects NA incorporation into virions, virion morphology, and virulence in mice but is not essential for virus replication. J. Virol. 70: 873879 Molla, A., A. V . Paul, and E. Wimmer. 1991. Cell-free, de novo synthesis of poliovirus. Science 254: 1647-1651. Murphy, F. A . 1980. Togavirus morphology and morphogenesis. p241-316. In R. W. Schlesinger (ed.). The Togavirusese. Academic Press, New York. Murphy, F. A., C. M. Fauquet, D. H. L. Bishop, S. A . Ghabrial, A. W. Jarvis, G. P. Martelli, M. A . Mayo, and M. D. Summers. 1995. Virus Taxonomy: The Classification and Nomenclature of Viruses. The Sixth Report of the International Committee on Taxonomy of Viruses (book). Springer-Verlag. Vienna, Murphy, F. A., P. E. Halonen, A. K. Harrison. 1968. Electronmicroscopy of the development of rubella virus. J. Virol. 2: 1223-1227. Myers, E., and W. Miller. 1988. Optimal alignments in linear space. CABIOS 4:11-17. Nairn, H. Y., and H. Koblet. 1990. The cleavage of p62, the precursor of E2 and E3, is an early and continuous event in Semliki Forest virus-infected Aedes albopictus cells. Arch. Virol. 110: 221-237. Nakhasi, H. L., D. X. Zheng, I. K. Hewlett, and T. Y . Liu. 1988. Rubella virus replication: effect of interferons and actinomycin D. Virus Res. 10(1): 1-15. Nakhasi, H. L., N. K. Singh, G. P. Pogue, X. Q. Cao, and T. A. Rouault. 1994. Identification and characterization of host factor interactions with c/s-acting elements of rubella virus RNA. Arch Virol Suppl.9: 255-267. Nakhasi, H. L., T. A . Rouault, D. J. Haile, T.-Y. Liu, and R. D. Klausner. 1990. Specific high-affinity binding of host cell proteins to the 3' region of rubella virus RNA. New Biol. 2: 255-264. 121  Nakhasi, H. L., X. Q. Cao, T. A. Rouault, and T. Y. Liu. 1991. Specific binding of host cell proteins to the 3'-terminal stem-loop structure of rubella virus negative-strand RNA. J. Virol. 65(11): 5961-5967. Neeleman, L., and J. F. Bol. 1999. Cw-acting functions of alfalfa mosaic virus proteins involved in replication and encapsidation of viral RNA. Virology 254: 324-333. Nestorowicz, A., T. S. Chambers, and C. M. Rice. 1994. Mutagenesis of the yellow fever virus NS2A/2B cleavage site: effects on proteolytic processing, viral replication and evidence for alternative processing of the NS2A protein. Virology 199: 114-123. Ng, D. P., S. W. Hiebert, and R. A. Lamb. 1990. Different roles of individual N-linked oligosaccharide chains in folding, assembly, and transport of the simian virus 5 hemagglutinin neuraminidase. Mol. Cell Biol. 10: 1989-2001. Niesters, H. G. M., and J. H. Strauss. 1990. Defined mutations in the 5' nontranslated sequence of Sindbis virus RNA. J. Virol. 64: 4162-4168. Novak, J. E. and K. Kirkegaard. 1991. Improved method for detecting poliovirus negative strands used to demonstrate specificity of positive-strand encapsidation and the ratio of positive to negative strands in infected cells. J. Virol. 65: 3384-3387. Novak, J. E., and K. Kirkegaard. 1994. Coupling between genome translation and replication in an RNA virus. Genes Dev. 8: 1726-1737. Oh, C. S., and J. C. Carrington. 1989. Identification of essential residues in potyvirus proteinase HC-Pro by site-directed mutagenesis. Virology 173(2): 692-9. Oker-Blom, C. 1984. The gene order for rubella virus structural proteins is NH2-C-E2-E1COOH. J. Virol. 51: 354-358. Oker-Blom, C , I. Ulmanen, L. Kaariainen, and R. F. Pettersson. 1984. Rubella virus 40S genome RNA specifies a 24S subgenomic mRNA that codes for a precursor to structural proteins. J. Virol. 49: 403-408. Oker-Blom, C , N. Kalkkinen, L. Kaariainen, and R. F. Pettersson. 1983. Rubella virus contains one capsid protein and three envelope glycoproteins, E l , E2a, and E2b. J. Virol. 46(3): 964-973. O'Reilly, E. K., and C. C. Kao. 1998. Analysis of RNA-dependent RNA polymerase structure and function as guided by known polymerase structures and computer predictions of secondary structure. Virology 252: 287-303. Review. Owen, K. E., and R. J. Kuhn. 1996. Identification of a region in the Sindbis virus nucleocapsid protein that is involved in specificity of RNA encapsidation. J. Virol. 70(5): 27572763.  122  Pardigon, N., and J. H. Strauss.  1992. Cellular proteins bind to the 3' end of Sindbis virus minus strand RNA. J. Virol. 66: 1007-1015.  Pardigon, N., and J. H. Strauss.  1996. Mosquito homolog of the La autoantigen binds to Sindbis virus RNA. J. Virol. 70(2): 1173-81.  Pardigon, N., E. Lenches, and J. H. Strauss. 1993.  Multiple binding sites for cellualr proteins in the 3' end of Sindbis alphavirus minus sense RNA. J. Virol. 67: 5003-5011.  Paredes, A. M., M. N. Simon, D. T. Brown. 1992.  The mass of the Sindbis virus nucleocapsid suggests it has T = 4 icosahedral symmetry. Virology 187(1): 329-332.  Pattnaik, A. K., D. J. Brown, and D. P. Nayak. 1986.  Formation of influenza virus particles lacking hemagglutinin on the viral envelope. J. Virol. 60: 994-1001.  Payment, R., D. Ajdukovic, V . Pavilanis. 1975.  Rubella virus. I. Morphology and structural  proteins. Can J Microbiol. 21(5): 703-709. French.  Phillips, M. A., and W. J. Rutter. 1996.  Role of the prodomain in folding and secretion of rat pancreatic carboxypeptidase A l . Biochemistry 35: 6771-6776.  Piccone, M. E., M. Zellner, T. F. Kumosinski, P. W. Mason, and M. J. Grubman. 1995. Identification of the active-site residues of the L proteinase of foot-and-mouth disease virus. J. Virol. 69: 4950-4956.  Pogue, G. P., C. C. Huntley, and T. C. Hall.  1994. Common replication strategies emerging from the study of diverse groups of positive-strand R N A viruses. Arch Virol Suppl. 9: 181-194. Review.  Pogue, G. P., X.-Q. Cao, N. K. Singh, and H. L. Nakhasi. 1993.  5' sequences of rubella virus R N A stimulate translation of chimeric RNAs and specifically interact with two host-encoded proteins. J. Virol. 67: 7106-7117.  Pogue, G., P. Hofmann, R. Duncan, J. M. Best, J. Etherington, R. D. Sontheimer, and H. L. Nakhasi. 1996. Autoantigens interact with ds-acting elements of rubella virus RNA. J. Virol. 70: 6269-6277.  Pugachev, K. V . , and T. K. Frey. 1998. Effects of defined mutations in the 5' nontranslated region of rubella virus genomic R N A on virus viability and macromolecule synthesis. J Virol. 72(1): 641-650. Pugachev, K. V., E. S. Abernathy, and T. K. Frey. 1997a. Genomic sequence of the RA27/3 vaccine strain of rubella virus. Arch Virol. 142(6): 1165-1180  Pugachev, K. V . , E. S. Abernathy, and T. K. Frey.  1997b. Improvements of the specific infectivity of the rubella virus (RUB) infectious clone: determinants of cytopathogenicity induced by RUB map to the nonstructural proteins. J. Virol. 71: 562-568.  123  Qiu, Z., F. Tufaro, and S. Gillam.  1992a. The influence of N-linked glycosylation on the anigenicity and irnrnunogenicity of rubella virus E l glycoprotein. Virology 190: 876-881.  Qiu, Z., T. C. Hobman, H. McDonald, N . O. Seto, and S. Gillam.  1992b. Role-of N-linked oligosaccharides in processing and intracellular transport of E2 glycoprotein of rubella virus. J. Virol. 66: 3514-3521.  Qiu, Z., D. Ou, H. Wu, T. C. Hobman, and S. Gillam.  1994a. Expression and characterization of virus-like particles containing rubella virus structural proteins. J. Virol. 68: 4086-4091.  Qiu, Z., H. McDonald, J. Chen, T. C. Hobman, and S. Gillam. 1994b. Mutational analysis of the arginine residues in the E2-E1 junction region on the proteolytic processing of the polyprotein precursor of rubella virus. Virology 100: 821-825. Quadt, R., and E. M. Jaspars. 1990. Purification and characterization of brome mosaic virus RNA-dependent R N A polymerase. Virology. 178(1): 189-194. Ramos, C , J. R. Winther, M. C. Kielland-Brandt. 1994.  Requirement of the propeptide for in vivo formation of active yeast carboxypeptidase Y . J. Biol.Chem. 269: 7006-7012.  Rawlings, N . D., and A. J. Barrett.  1993. Evolutionary families of peptidases. Biochem. J.  290: 205-218.  Restrepo-Hartwig, M., and P. Ahlquist.  1996. Brome mosaic virus helicase- and polymeraselike proteins colocalize on the endoplasmic reticulum at sites of viral R N A synthesis. J. Virol. 70(12): 8908-8916.  Restrepo-Hartwig, M., and P. Ahlquist.  1999. Brome mosaic virus R N A replication proteins la and 2a colocalize and la independently localizes on the yeast endoplasmic reticulum. J. Virol. 73(12): 10303-10309.  Roehl, H. H., T. B. Parsley, T. V. Ho, and B. L. Semler.  1997. Processing of a cellular polypeptide by 3CD proteinase is required for poliovirus ribonucleoprotein complex formation. J. Virol. 71(1): 578-585.  Rost, B. and C. Sander. 1993a. Improved prediction of  protein secondary structure by use of sequence profiles and neural networks. Proc. Natl. Acad. Sci. U.S.A. 90: 7558-7562.  Rost, B. and C. Sander. 1993b. Prediction of protein  structure at better than 70% accuracy. J.  Mol. Biol. 232:584-599.  Rost, B. and C. Sander.  1994a. Combining evolutionary information and neural networks to predict protein secondary structure. Proteins 19: 55-72.  Rost, B., C. Sander and R. Schneider.  1994b. PHD - an automatic mail server for protein secondary structure prediction. CABIOS 10: 53-60.  124  Rozanov, M. N., E. V. Koonin, and A. E. Gorbalenya. 1992.  Conservation of the putative methyltransferase domain: a hallmark of the "Sindbis-like" supergroup of positive-strand R N A virus. J. Gen. Virol. 73(8): 2129-2134.  Sanchez, A., and T. K. Frey.  1991. Vaccinia-vectored expression of rubella virus structural proteins and characterization of the E l and E2 glycosidic linkages. Virology 183: 636-646.  Sawicki, D. L., and S. G. Sawicki.  1980. Short-lived minus-strand polymerase for Semliki  Forest virus. J. Virol. 34: 108-118.  Sawicki, S. G., D. L. Sawicki, L. Kaariainen, and S. Keranen. 1981.  A Sindbis virus mutant temperature-sensitive in the regulation of minus-strand RNA synthesis. Virology 115: 161-172.  Schaad, M. C, R. Haldeman-Cahill, S. Cronin, and J. C. Carrington. 1996.  Analysis of the VPg-proteinase (NIa) encoded by tobacco etch potyvirus: Effects of mutations on subcellular transport, proteolytic processing, and genome amplifiction. J. Virol. 70: 7039-7048.  Schechter, I., and A. Berger.  1967. On the size of the active site in proteases. Biochem.  Biophys. Res. Com. 27: 157-162.  Schmid, M., and E. Wimmer. 1994. IRES-controlled protein synthesis and genome replication of poliovirus. Arch Virol Suppl. 9: 279-289. Review. Schnell, M. J., T. Mebatsion, and K.-K. Conzelmann. 1994.  Infectious rabies viruses from  cloned cDNA. E M B O J. 13: 4195-4203.  Scholthof, J. B., and A. O. Jackson.  1997. The enigma of pX: A host-dependent as-acting element with various effects on tombus-virus RNA accumulation. Virology 237: 56-65.  Shapira, R., and D. L. Nuss.  1991. Gene expression by a hypo virulence-associated virus of the chestnut blight fungus involves 2 papain-like proteinase activities: essential residues and cleavage site requirements for p48 autoproteolysis. J. Biol. Chem. 266: 19419-19425.  Shirako, Y., and J. H. Strauss. 1994. Regulation of Sindbis virus R N A replication: uncleaved PI23 and nsP4 function in minus strand R N A synthesis whereas cleaved products from PI23 are required for efficient plus strand RNA synthesis. J. Virol. 68: 1874-1885. Siegel, R. W., S. Adkins, and C. C. Kao.  1997. Sequence-specific recognition of a subgenomic promoter by a viral R N A polymerase. Proc. Natl Acad. Sci. U S A 94: 1123811243.  Simons, K., and H. Garoff.  1980. The budding mechanism of enveloped animal viruses. J.  Gen. Virol. 50: 1-21.  Singer, S. J., P. A. maher, and M. P. Yaffe. 1987. On the transfer of integral membrane protein into membranes. Prot. Natl. Acad. Sci. USA. 84: 1960-1964. Singh, N. K., C. D. Atreya, and H. L. Nakhasi.  1994. Identification of calreticulin as a rubella virus R N A binding protein. Proc. Natl. Acad. Sci. U S A 91: 12770-12774 125  Sivakumaran, K. and C.C. Kao.  1999. Initiation of genomic positive strand synthesis from D N A and R N A templates by a viral RNA-dependent R N A polymerase. J. Virol. 73: 6415— 6423.  Sivakumaran, K., C. H. Kim, R. Jr. Tayon, and C. C. Kao. 1999.  R N A sequence and secondary structural determinants in a minimal viral promoter that directs replicase recognition and initiation of genomic plus-strand R N A synthesis. J Mol Biol. 294(3): 667-682.  Skern, T., I. Fita and A. Guarne. 1998.  A structural model of picornavirus leader proteinases based on papain and bleomycin hydrolase. J. Gen. Virol. 79: 301-307.  Snijder, E. J., A. L.  M . Wassenaar, and W. J. M. Spaan. 1992. The 5' end of the equine arteritis virus replicase gene encodes a papain-like cysteine protease. J. Virol. 66: 7040-7048.  Spangberg, K., L. Goobar-Larsson, M . Wahren-Herlenius, and S. Schwartz. 1999. The La protein from human liver cells interacts specifically with the U-rich region in the hepatitis C virus 3' untranslated region. J Hum Virol. 2(5): 296-307.  Strauss, E. G., and J. H. Strauss. 1986.  Structure and replication of the alphavirus genome, p. 35-90. In S. Schlesinger and M . J. Schlesinger (ed.), The Togaviridae and Flaviviridae. Plenum publishing Corp., New York.  Strauss, E. G., R. J. de Groot, R. Levinson, and J. H. Strauss.  1992. Identification of the active site residues in the nsP2 proteinase of Sindbis virus. Virology 191: 932-940.  Strauss, J. H., and E. G. Strauss.  1994. The alphaviruses: Gene expression, replication, evolution. Microbiol. Rev. 58: 491-562. Review.  Sullivan, M . L., and P. Ahlquist.  1999. A brome mosaic virus intergenic RNA3 replication signal functions with viral replication protein la to dramatically stabilize R N A in vivo. J. Virol. 73(4): 2622-2632.  Sun, J. H., S. Adkins, G. Faurote, and C. C. Kao. synthesis catalyzed by the B M V RNA-dependent oligonucleotides. Virology 226(1): 1-12.  1996. Initiation of (-)-strand R N A R N A polymerase: synthesis of  Suomalainen, M . , H. Garoff, and M. D. Baron.  1990. The E2 signal sequence of rubella virus remains part of the capsid protein and confers membrane association in vitro. J. Virol. 64: 5500-5509.  Suomalainen,  M . , P. Liljestrom, and H. Garoff. 1992. Spike interactions drive the budding of alphaviruses. J. Virol. 66: 4737-4747.  protein-nucleocapsid  Svitkin, Y. V., K. Meerovitch, H. S. Lee, J. N. Dholakia, D. J. Kenan, V. I. Agol, and N. Sonenberg. 1994. Internal translation initiation on poliovirus RNA: further characterization of La function in poliovirus translation in vitro. J. Virol. 68: 1544-1550.  126  Takkinen, K., J. Peranen, and L. Kaariainen. 1991.  Proteolytic processing of Semliki Forest virus-specific non-structural polyprotein. J. Gen Virol. 72: 1627-1633.  Tao, K., N. A. Stearns, J. Dong, Q. L. Wu, and G. G. Sahagian.  1994. The proregion of cathepsin L is required for proper folding stability, and ER exit. Arch. Biochem. Biophys. 311: 19-27.  Teng, H., J. D. Pinon and S. R. Weiss.  1999. Expression of murine coronavirus recombinant papain-like proteinase: efficient cleavage is dependent on the lengths of both the substrate and the proteinase polypeptides. J. Virol. 73: 2658-2666.  Terry, G. M., L. Ho-Terry, P. Londesborough, and K. R. Rees.  1988. Localization of the  rubella E l epitopes. Arch. Virol. 98: 189-197.  Tesh, R. B., and L. Rosen.  1975. Failure of rubella virus to replicate in mosquitos.  Intervirology. 5(3-4): 216-219.  Trudel, M., F. Nadon, C . Sequin, A. Amarouch, P. Payment, and S. Gillam. 1985. E l glycoprotein of rubella virus carries an epitope that binds a neutralizing antibody. J. Virol. Methods 12: 243-250.  Trudel, M., M. Ravaoarinoro, and P. Payment.  1980.  Reconsitution  of rubella  hemagglutinin on liposome. Can J. Microbiol. 26: 899-904.  Tsuchihara, K., T. Tanaka, M. Hijikata, S. Kuge, H. Toyoda, A. Nomoto, N. Yamamoto, and K. Shimotohno. 1997. Specific interaction of polypyrimidine tract-binding protein with the extreme 3'-terminal structure of the hepatitis C virus genome, the 3'X. J Virol. 71(9): 67206726.  Umino, Y., T. A. Sato, S. Katow, T. Matsuno, and A. Sugiura. 1985.  Monoclonal antibodies  directed to E l glycoprotein of rubella virus. Arch Virol. 83: 33-42.  Van Bokhoven, H., O. Le Gall, D. Kasteel, J. Verver, J. Wellink, and A. van Kammen. 1993. Cis- and trans-acting elements in cowpea mosaic virus R N A replication. Virology 195: 377-386.  Vennema, H., G.-J. Godeke, J. W. A. Rossen, W. F. Voorhout, M. C. Horzinek, D.-J. E. Opstelten, and P. J. M. Rottier. 1996. Nucleocapsid-independent assembly of coronaviruslike particles by co-expression of viral envelope protein genes. E M B O J. 15:2020-2028.  Vidgren, G., K. Takkinen, N. Kalkkinen, L. Kaarianen, and R. F. Pettersson. 1987. Nucleotide sequence of the genes coding for the membrane glycoproteins E l and E2 of rubella virus. J. Gen. Virol. 68: 2347-2357.  Vogel, R. H., S. W. Provencher, C . H. von Bonsdorff, M. Adrian, J. Dubochet. 1986. Envelope structure of Semliki Forest virus reconstructed from cryo-electron micrographs. Nature 320: 533-535.  127  Waxham, M. N., and J. S. Wolinsky. 1983.  Immunochemical identification of rubella virus  hemagglutinin. Virology 126: 194-203.  Waxham, M. N., and J. S. Wolinsky. 1985a. A model of the  structural organization of rubella  virions. Rev. Inf. Dis. 7 (Suppl. 1): S133-S139.  Waxham, M. N., and J. S. Wolinsky. 1985b. Detailed immunologic analysis of the structural polypeptides of rubella virus using monoclonal antibodies. Virology 143: 153-165. Weiland, J. J., and T. W. Dreher.  1993. CVs-preferential replication of the turnip yellow mosaic virus R N A genome. Proc. Natl. Acad. Sci. U S A 90: 6095-6099.  White, C. L., M. Thomson, and N. J. Dimmock.  1998. Deletion analysis of a defective interfering Semliki Forest virus R N A genome defines a region in the nsP2 sequence that is required for efficient packaging of the genome into virus particles. J. Virol. 72(5): 4320-4326.  White, K. A., J. B. Bancroft, and G. A. Mackie.  1992. Coding capacity determines in vivo accumulation of defective RNA of clover yellow mosaic virus. J. Virol. 66: 3069-3076.  Wimmer, E., C. U. T. Hellen, and X. Cao.  1993. Genetics of poliovirus. Annu. Rev. Genet.  27: 353-436.  Wolinsky, J. S.  1996. Rubella. p899-921. In B.N. Fields, D . M . Knipe, P.M. Howley, et al. (ed), Fields Virology, third edition. Lipincott-Raven Publishers, Philadelphia.  Wolinsky, J. S., M. McCarthy, O. Allen-Cannady, W. T. Moore, R. Jin, S. N. Cao, A. lovett, and D. Simmons. 1991. Monoclonal antibody-defined epitope map of expressed rubella virus protein domains. J. Virol. 65: 3986-3994.  Xu, J., E. Mendez, P. R. Caron, C. Lin, M. A. Murcko, M. S. Collett, and C. M. Rice. 1997. Bovine viral diarrhea virus NS3 serine proteinase: polyprotein cleavage sites, cofactor requirements, and molecular model of an enzyme essential for pestivirus replication. J. Virol. 71: 5312-5322.  Yang, D., D. Hwang, Z. Qiu, and S. Gillam. 1998. Effects of mutations in the rubella virus E l glycoprotein on E1-E2 interaction and membrane fusion activity. J. Virol. 72: 8747-8755. Yao, J., and S. Gillam.  1999. Mutational analysis, using a full-length rubella virus cDNA clone, of rubella virus E l transmembrane and cytoplasmic domains required for virus release. J. Virol. 73: 4622-4630.  Yao, J., and S. Gillam.  2000. A single-amino-acid substitution of a tyrosine residue in the rubella virus E l cytoplasmic domain blocks virus release. J. Virol. 74(7): 3029-3036.  Yao, J., D. Yang, P. Chong, D. Hwang, Y. Liang, and S. Gillam. 1998. Proteolytic processing of rubella virus nonstructural proteins. Virology 246(1): 74-82. Yu, W., and J. L. Leibowitz. 1995.  Specific binding of host cellular proteins to multiple sites within the 3' end of mouse hepatitis virus genomic RNA. J. Virol. 69(4): 2016-2023. 128  Zhou, H., and A. O. Jackson. 1996.  Analysis of cis-acting elements required for replication of barley stripe mosaic virus RNAs. Virology 219(1): 150-160.  129  

Cite

Citation Scheme:

        

Citations by CSL (citeproc-js)

Usage Statistics

Share

Embed

Customize your widget with the following options, then copy and paste the code below into the HTML of your page to embed this item in your website.
                        
                            <div id="ubcOpenCollectionsWidgetDisplay">
                            <script id="ubcOpenCollectionsWidget"
                            src="{[{embed.src}]}"
                            data-item="{[{embed.item}]}"
                            data-collection="{[{embed.collection}]}"
                            data-metadata="{[{embed.showMetadata}]}"
                            data-width="{[{embed.width}]}"
                            async >
                            </script>
                            </div>
                        
                    
IIIF logo Our image viewer uses the IIIF 2.0 standard. To load this item in other compatible viewers, use this url:
http://iiif.library.ubc.ca/presentation/dsp.831.1-0099576/manifest

Comment

Related Items