©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
A Role for 3AB Protein in Poliovirus Genome Replication (*)

Juan Lama (§) , Miguel A. Sanz , Pedro L. Rodrguez

From the (1)Centro de Biologa Molecular ``Severo Ochoa,'' Universidad Autónoma de Madrid, Cantoblanco, 28049 Madrid, Spain

ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

The poliovirus polypeptide 3AB, the precursor of the genome-bound VPg protein, stimulates in vitro the synthesis of poly(U) directed by the viral polymerase 3D (Lama, J., Paul, A., Harris, K., and Wimmer, E. (1994) J. Biol. Chem. 269, 66-70), suggesting that 3AB could be modulating the activity of the viral polymerase in poliovirus-infected cells. To address the exact function of 3AB in the viral replication cycle, a biochemical and molecular genetic analysis of 3AB has been carried out. 3AB protein bound RNA probes in two different assays, and amino acid positions implicated in the RNA binding activity of 3AB were determined. Mutant proteins with reduced RNA binding activity were unable to stimulate 3D polymerase activity. Purified protein 3A showed no RNA binding or 3Dstimulatory activity, but 3A and VPg mutations conferred a synergistic effect on the 3AB functions. Polioviruses encoding for these mutant 3ABs were constructed. These mutant viruses translated their RNA genomes in vitro and processed their polyproteins as wild type virus did. Cells infected with 3AB mutant viruses showed over 90% inhibition in the accumulation of plus and minus viral RNA strands and more than 100-fold reduction of virus yield at 4 h postinfection. Our results suggest that 3AB protein functions in vivo as a co-factor of the viral polymerase and that the activity of 3AB may be regulated by proteolytic processing.


INTRODUCTION

Poliovirus, a member of the Picornaviridae family, possesses a single-stranded genome of positive polarity(1) . The mechanism of replication of poliovirus RNA is understood only in general terms. Upon infection of permissive cells, the viral RNA gains access into the cytoplasm, where the replication of the genome takes place(2, 3) . First, the viral RNA, after its translation to produce virus-specific polypeptides, serves as a template to synthesize minus strand RNA. The minus strand is used to synthesize more plus strand RNAs, which can be used as templates for new minus strand RNAs, as messenger RNAs, or as genomic virion RNAs(4) . All nascent viral RNA strands synthesized in the cell are covalently linked to the VPg protein (VPg = 3B)(5, 6) . The mechanism by which this peptide is bound to the 5`-end of the viral genome constitutes one of the puzzling unanswered questions in the poliovirus replication field. The viral polymerase (3D) is the only polypeptide isolated from infected cells with RNA-dependent RNA polymerase activity(7, 8) . Thus, it has been considered the protein responsible for the elongation of the RNA strands. In addition, most if not all of the poliovirus nonstructural proteins have been implicated in the replication of the genome(9, 10, 11, 12, 13, 14, 15) . Special interest has been focused on the 3AB protein. 3AB is the most abundant precursor of the VPg protein(16) . Protein 3AB localizes to the membranous replication complexes, where active replication takes place(17, 18) . It has been postulated that an uridylylated form of VPg or, more likely, an uridylylated form of 3AB might serve as a primer to initiate polymerase activity by 3D(19, 20) . However, the nucleotidylyl transfer reaction has never been reproduced in vitro using purified components.

The proposed function of 3AB in poliovirus RNA replication mainly arose from genetic studies. The first genetic evidence was reported by Berstein and Baltimore(21) . These authors described a cold-sensitive poliovirus mutant encoding for a 3A protein with a single amino acid insertion. This mutant showed a small plaque phenotype and severe replication defects at the restrictive temperature. Since then, multiple poliovirus 3A mutants with defects in RNA replication have been isolated(22, 23) . Many of these mutations were mapped to the hydrophobic domain of 3A, which presumably promotes the binding of the protein to the cellular membranes. Amino acid changes in the hydrophobic domain commonly result in death or mutant viruses with drastic impairments in the replication of the RNA(23) . If RNAs obtained by transcription from these mutant cDNAs are used to program HeLa cell extracts for in vitro translation, in some cases aberrant polyprotein-processing patterns are obtained. Thus, the hydrophobic region of 3A may serve to anchor the polypeptide, or part of it, to the membranous replication complex, a process that can modulate the proteolytic cleavage of the polypeptide. Recently, a key finding to understanding the function of 3AB in RNA replication has been reported(24) . 3AB protein purified to almost homogeneity possesses an intrinsic stimulatory activity of the viral polymerase 3D. Thus, in vitro poly(U) synthesis by 3D is stimulated more than 50-fold in the presence of recombinant purified 3AB. The transactivation activity occurs in the absence of any detected uridylylation and was completely dependent on the presence of a nucleic acid primer. These results suggested that 3AB could function in vivo as a co-factor of 3D in viral transcription. However, genetic evidence supporting this hypothesis is still lacking. We now report the characterization of mutant polioviruses with severe defects in viral RNA replication. These viruses encode for 3AB proteins unable to transactivate the viral polymerase.


MATERIALS AND METHODS

Cells and Viruses

Virus propagation and virus plaque assays were performed on HeLa R19 cell monolayers grown in Dulbecco's modified Eagle's medium supplemented with 10% calf serum. HeLa cell lysates for in vitro translation were made from HeLa S3 cells grown in minimum essential medium modified for suspension cells supplemented with 5% calf serum. The viruses used in this work were obtained after transfection of HeLa cell monolayers with infectious RNAs derived from the genomic cDNA cloned in the pT7XLD vector(25) . Wild type and K9E-3AB viruses were plaque-purified and amplified in HeLa cells up to titers of 0.5-1 10 pfu/ml.()Stocks of the viruses K107-3AB and K9E/K107 3AB were obtained after 3 days of transfection with the corresponding RNAs. These stocks have titers of 0.5-1 10 pfu/ml and were directly used in the experiments described. Additional plaque purification and amplification of these viruses gave place to the generation of revertant viruses with wild type plaque phenotype. The complete 3AB gene of the viral RNA of all the stock viruses was sequenced. Virus RNA purification, cDNA synthesis, and PCR from the cDNA obtained were performed to ensure that the mutations introduced had been retained in the viral genome. To create mutant cDNAs with the desired mutations the 3AB gene was PCR-amplified from the corresponding pT7lac3AB plasmid. To do so, the following primers were used: 5`-3A (5`-CCC GGG CAT ATG GGA CCA CTC CAG TAT AAA GAC-3`) and 3`-3BC (5`-GCG TAA TCG AAC CCT GGT CCT TGT ACC TTT GCT GTC CGA AT-3`). To amplify mutant DNAs K107-3AB and K9E/K107 3AB, the PCR reaction was carried out with primers 5`-3A and 3`-3BC K107E (5`-GCG TAA TCG AAC CCT GGT CCT TGT ACC TCT GCT GTC CGA AT-3`). After PCR amplification, the fragments were digested with AvaII, and the 3AB gene was exchanged with the equivalent AvaII DNA fragment of the pT7XLD plasmid. The 3AB regions of the wild type and mutant cDNAs were completely sequenced by the dideoxy method (Sequenase, U.S. Biochemical Corp.). Transfection of HeLa cells was carried out with Lipofectin reagent. Ten µg of in vitro transcribed RNA, without further purification, were mixed with 15 µg of Lipofectin and incubated at 37 °C for 2 h with HeLa cells that had been previously washed with prewarmed phosphate-buffered saline to remove calf serum. After this time, the cells were kept in 2% calf serum for 3 days at 37 °C, and the viruses were recovered by three cycles of freeze-thawing. To quantify the cytopathic effect, the percentage of cells showing clear symptoms of morphological rounding was estimated de visu.

RNA Binding Assays

To determine the RNA binding capabilities of different 3AB proteins, we routinely used a method recently described(26) . This procedure is based on the binding of biotinylated RNA probes (riboprobes) to renatured proteins bound to nitrocellulose sheets (Northwestern). Briefly, the proteins (purified or crude extracts) were separated in 20% SDS-polyacrylamide gels and transferred to nitrocellulose. After washing the nitrocellulose sheets in a renaturation buffer, the nitrocellulose was first incubated with a biotinylated riboprobe (40 ng/ml). Then unbound RNA was washed with phosphate-buffered saline-Tween 20, and the membranes were incubated with streptavidin-conjugated peroxidase. After washing off the peroxidase, the nitrocellulose was soaked in luminol-luciferin solution and exposed to x-ray films. The biotinylated riboprobes were obtained by in vitro transcription of the appropriate DNAs in the presence of 0.5 mM 21-UTP. The riboprobe ``5-leader'' consists of the genomic RNA sequence (plus strand) between nucleotides 1 and 745. The riboprobe 3`-end contains nucleotides 6516-7440 of the 3`-end of the genomic RNA, whereas the riboprobe ``2.5 BamHI'', of negative polarity, encompasses the complementary region to nucleotides 2099-4600 of the genome.

To confirm the data on RNA binding, a gel retardation assay was used in some instances to detect RNA-protein complexes. The purified proteins were incubated with 10 ng of a 51-nucleotide-long riboprobe of positive polarity, which contains a region of the 2B poliovirus gene (nucleotides 3925-3952) in addition to some plasmid-derived nucleotides. The analysis of the free and bound probe forms was carried out as described previously(26) . To quantify the amount of 3AB protein present in crude extracts, we used an anti-3A antiserum that recognizes an epitope located between amino acids 30-44 of the 3A protein.

Expression and Purification of Recombinant Proteins

Expression in E. coli of 3A and 3AB proteins was carried out from the plasmids pT7lac3A and pT7lac3AB, respectively, as described previously(27) . The mutant 3AB proteins used in this work were obtained by PCR-based random mutagenesis. To purify wild type and K9E-3AB proteins, we followed a previously described procedure(24) , but in this case, during the elution of proteins from the S-Sepharose resin, the 100 mM salt wash was omitted. The protein K107E-3AB behaved as the wild type through the DE-52 column; however, at the S-Sepharose binding step the pH of the column had to be changed from MOPS (pH 7.2) to MES (pH 6.5). The posterior elution was carried out in 200 mM NaCl, 25 mM MES (pH 6.5). As control, wild type 3AB protein was also purified in an identical way, that is at pH 6.5 during the binding and elution of the S-Sepharose column. K9E/K107E-3AB was purified as described above, but after the DE-52 column the protein was concentrated with ultrafiltration membranes (Amicon). Since this protein did not bind to the S-Sepharose column under any of the conditions tested, this fraction was no longer purified. Purification of 3A was stopped after concentration of DE-52 fractions. However, this protein was not recovered from the unbound proteins originally passed through the DE-52 resin, since under equilibration conditions the protein bound to the column and was later recovered with a 50 mM NaCl wash.

Enzymatic Assay for Poliovirus RNA Polymerase

Incorporation of [H]UMP was measured using a poly(A)-dependent, oligo(dT)-primed poly(U) polymerase assay in the presence of purified 3D. Recombinant 3D, in a poly(U)-free form, completely dependent on the presence of a nucleic acid primer for its activity, was kindly provided by Dr. S. Plotch. Reaction conditions for the polymerase assay have been described before(24) . Standard transactivation reactions were carried out in the presence of 20 ng/µl of 3A or 3AB proteins. Poliovirus polymerase was used at 0.2 ng/µl.

Quantification of Plus and Minus Viral RNA

To quantify the levels of viral specific RNA, HeLa cells were infected at a multiplicity of infection of 0.1 pfu/ml. After 4 h of infection at 37 °C the cells were lysed, and the total cytoplasmic RNA was isolated by the Nonidet P-40 lysis method (28) and used to synthesize a cDNA strand with avian mieloblastosis virus retrotranscriptase (Promega). Retrotranscriptase reactions were carried out in a 40-µl mixture containing 10 mM Tris-HCl (pH8.3), 50 mM KCl, 1.5 mM MgCl, 0.01% gelatin, 10 mM dithiothreitol, 0.1 mM dNTPs, 3 units of avian mieloblastosis virus retrotranscriptase, 15 units of RNAsin inhibitor, 0.25 µM of the first primer, and 250 ng of total cytoplasmic RNA. After incubation at 42 °C for 30 min, the retrotransciptase was destroyed by boiling, and 0.25 µM of the second primer and 1 unit of Taq DNA polymerase were added (Perkin-Elmer). PCR amplification took place during either 30 cycles (minus RNA) or 25 cycles (plus RNA) under the following conditions: 1 min at 93 °C, 1 min at 37 °C, and 5 min at 72 °C (last cycle, 10 min at 72 °C). The amount of amplified fragment was quantified in 1.0% agarose gels stained with ethidium bromide. To detect plus strand viral RNA, the retrotranscriptase reaction was carried out with the primer 3`-2B (GGC CCG GAT CCT TAT TAT TGC TTG ATG ACA TAA GGTA). The primer 5`-2A (GGC CGG CCC GGG ATT CGG ACA CCA AAAC) was later added in order to proceed with the PCR reaction. To detect minus strand polio RNA, the order of addition of the primers was reversed. As an internal standard for the reaction the -actin mRNA was amplified by RT-PCR in the same test tubes. To this end, the primer 3`--actin (GGA AGC TTC TAG AAG CAT TTG CGG TGG ACG ATG GAG GGG CC) was included into the retrotranscriptase reaction, and the primer 5`--actin (GGG AAT TCA TGG ATG ATG ATA TCG CCG CG) was later added during the PCR reaction. -actin primers were used at 1 µM. Amplification of polio and -actin cDNAs results in fragments of 727 and 1170 nucleotides in length, respectively.

In Vitro Translation of Transcripts Derived from Mutant cDNAs

Translations were carried out in HeLa S3 cell extracts treated with micrococcal nuclease and programmed with infectious RNAs synthesized by in vitro transcription of wild type or mutant cDNAs. The preparation of extracts was essentially as described by Molla et al.(29) . Reactions were incubated at 30 °C for 15 h with 100 ng of transcript RNA in the presence of 0.7 µCi/µl of [S]methionine. The synthesized proteins were analyzed by autoradiography of SDS-polyacrylamide gels.


RESULTS

3AB Binds RNA in a Nonspecific Manner

Previous experiments have shown a stimulating activity of poly(U) synthesis directed by the viral polymerase 3D in the presence of purified 3AB protein (24). These results suggested that 3AB could be a co-factor of 3D in viral transcription. This stimulating activity was completely dependent on the presence of the 3AB polypeptide but was not due to a nucleotidylylation reaction on the 3AB protein, since the polymerase activity of 3D was dependent on the presence of a nucleic acid primer in the reaction mixture. An alternative hypothesis to explain the stimulatory effect of 3AB might rely on the ability of the protein to increase the affinity or the binding of 3D to either the template or the primer RNA. Paul et al.(30) have shown that under conditions where 3D and poly(A) template were reduced to a minimum, the degree of transactivation was maximal, promoting more than 100-fold stimulation. These results suggest that 3AB may enhance the interaction between the template RNA and the viral polymerase. A prediction of this model is the interaction of 3AB with RNA and 3AB with 3D. As a first step in the elucidation of the mechanism of action of 3AB, the RNA binding activity of purified 3AB was assayed (Fig. 1). Wild type purified 3AB was used to analyze the binding activity to different biotinylated RNA probes. As shown in Fig. 1A, purified 3AB bound three different heteroligomeric RNA probes. The signal given by the binding reactions increased linearly with the amount of 3AB protein used. The 5`-end of the genome has been involved in the translation and replication of the genome, and different independent domains have been identified for each of these functions(31, 32) . A pseudo-knot structure has been identified in the 3`-end of the genome(33) . To test the binding of 3AB to these structures, riboprobes spanning both ends of the genome were synthesized. The 3AB protein bound both probes, suggesting that either 3AB binds specifically to both ends of the genome or 3AB binds nonspecifically to polyribonucleic acid molecules. 3AB also bound to a riboprobe of negative polarity encompassing the complementary region between nucleotides 2099 and 4600 of the viral genome (Fig. 1A). Indeed, this binding reaction took place to a greater degree than in the other cases, although these differences may be due to differences in the level of incorporation of biotinylated UTP in the larger, minus strand RNA probe. Thus, purified 3AB binds to different RNA molecules and does not seem to show any sequence specificity for binding to the poliovirus genome. Further corroborating these results, homopolymeric RNAs, either poly(A) or poly(U), efficiently compete with the binding of the 2.5-BamHI biotinylated riboprobe to 3AB (Fig. 1B). In fact, we have tested other riboprobes, finding similar RNA binding capabilities in all cases (data not shown). In order to confirm the RNA binding activity found with the Northwestern assay, we carried out a gel retardation assay to check the effect of 3AB on a P-labeled riboprobe. As shown in Fig. 1C, this procedure allow us to detect a retardation effect on the mobility of a 51-nucleotide-long riboprobe. This binding was also avoided in the presence of competitor cold poly(A) RNA.


Figure 1: RNA binding activity of purified 3AB protein. A, poliovirus 3AB protein was purified from a recombinant E. coli clone, and different amounts of protein were electrophoresed in SDS-polyacrylamide gels. After transferring to nitrocellulose sheets, the proteins were incubated with biotinylated RNA probes, and the bound RNA was detected by the streptavidin-peroxidase-luciferin method after exposure of the gels to x-ray films. RNA binding activity was assayed with a minus strand RNA probe (2.5-BamHI) or with two different plus strand probes encompassing both extremes of the genome (5`-leader and 3`-end). B, 5 µg of purified 3AB were used to estimate the RNA binding capability in the presence of competitive amounts of nonlabeled poly(A) or poly(U) RNA, following the procedure described above. C, different amounts of purified 3AB were incubated with a 51-nucleotide-long P-labeled riboprobe in the absence or in the presence of competitive cold poly(A) RNA. The positions of the free riboprobe and the 3AB-RNA complex were determined after autoradiography of a 6% acrylamide gel.



Identification of Amino Acid Residues in 3AB Involved in the Binding of RNA

The Northwestern procedure makes use of proteins separated on polyacrylamide gels and transferred to nitrocellulose sheets. Given sufficient quantities of the recombinant protein, this assay permits the analysis of mutated proteins in total lysates without further purification. Fig. 2shows the RNA binding activity of a set of 14 different 3AB variants, including the 3A protein, which lacks the VPg sequence. A new binding protein is observed when wild type 3AB protein is expressed in Escherichia coli cells (Fig. 2, lane3). This band does not appear in the control extract without the recombinant protein ( Fig. 2, lane2). Amino acid substitutions at positions 9, 13, 54, 62, 63, 67, 75, 77, 79, 81, and 97 did not affect the binding of the biotinylated riboprobe. Neither the introduction of positively charged residues nor -helix-breaking proline residues into the hydrophobic domain seems to affect the interaction of 3AB with RNA. However, amino acid substitutions in this region produce mutant viruses with defects in RNA replication(22) . Amino acid substitution at position 107 (lysine 107 to glutamic acid) seems to abolish the RNA binding capability of 3AB. Even though similar amounts of K107E-3AB and, for instance, K97R-3AB, were contained in the E. coli extracts, no new band was found in the position expected for the recombinant K107E-3AB protein (Fig. 2, lane6), whereas a clear band was revealed in the position of the K97E-3AB protein. No RNA binding activity was detected in the K9E/K107E-3AB protein, which contains a double amino acid change in the 3A and VPg regions of the polypeptide. No binding was detected with the 3A protein.


Figure 2: RNA binding activity of mutated 3AB proteins in crude E. coli extracts. In the upperpanel (Northwestern) E. coli BL21(DE3) cells were induced to synthesize either wild type or 3AB mutated proteins. Fifty micrograms of total protein from crude extracts were electrophoresed in SDS-polyacrylamide gels and transferred to nitrocellulose to test the RNA binding capability, as shown in Fig. 1A. As a negative control, total proteins from bacteria without the expression plasmid were run in lane2. Lane1 contains 2.5 µg of purified 3AB. In the lowerpanel, after processing to measure the binding of the biotinylated riboprobe, the same nitrocellulose sheets were washed and incubated with an anti-3A antibody to determine the position (labeled with an arrow in the upperpanel) and the amount of the recombinant proteins.



The level of expression of different 3AB proteins in E. coli was quite variable, making it difficult to compare the binding activities of different mutated proteins. Thus, we decided to purify the wild type 3AB and 3A proteins and the mutants K9E, K107E, and K9E/K107E-3AB proteins (Fig. 3). Wild type and K9E-3AB proteins were purified following a previously published protocol(24) , although the mutated 3AB proteins changed their chromatographic behavior to different degrees and small modifications in the procedure were necessary. In the case of the proteins K107E and K9E/K107E-3AB, only 70 and 40% respectively, of the recombinant protein remains in the unbound fraction after the DEAE-cellulose purification step (data not shown). This result suggests that both residues at positions 9 and 107 are in the outer surface of the molecule, since both proteins seem to interact more strongly with the positively charged DEAE resin, as compared with either the wild type or the single mutant K9E. As expected, the 3A protein, which does not contain the highly basic VPg moiety, bound to the DEAE resin under conditions where no wild type 3AB protein remained bound to the column (data not shown). Proteins K107E and wild type 3AB were further purified through S-Sepharose resin. However, to allow efficient binding of the former, the pH of the column was reduced to 6.5. To assure that this change did not alter the RNA binding and transactivating activities of the protein, the wild type protein was purified at pH 6.5 also. Fig. 3A shows the degree of purity of the recombinant proteins. The proteins purified through DE-52 and S-Sepharose resins were more than 95% pure, whereas the proteins 3A and K9E-K107E-3AB were obtained at 90 and 70% of purity, respectively, as estimated by densitometric analysis of SDS-polyacrylamide gels. In Fig. 3B equal amounts of the purified proteins were used to estimate their binding to the biotinylated RNA probe. In the upperpanel the proteins purified through S-Sepharose resin were analyzed. Wild type protein and K9E-3AB mutant bound RNA to a similar degree. However, the amino acid substitution at position 107 (K107E) greatly reduced the binding activity, as compared with the wild type protein purified in an identical way (pH 6.5). In the lowerpanel, the effect of the double substitution (K9E/K107E) was assayed. To compare with accuracy the RNA binding activity of all the variants, the proteins recovered after the DE-52 elution step were assayed in Northwestern assays. As can be seen, the binding of the biotinylated probe produced an intense signal with wild type 3AB, whereas the K107E-3AB protein showed a drastic decrease in the RNA binding capability. No binding was detected either with the double mutant K9E/K107E-3AB, or with the wild type 3A protein. These results demonstrate that amino acid position 107 in 3AB is important for RNA binding activity. The results shown in Fig. 3B also suggest that, even though the amino acid substitution at position 9 (K9E) does not produce any effect by itself, this change seems to cause a synergistic effect on amino acid substitutions more than 100 residues apart in the 3AB molecule. Introduction of a positively charged residue at position 9 may alter the structure of the K107E-3AB protein, making it less active. This structural alteration may not take place with the natural residue (lysine) at position 107. Otherwise, a lysine at position 9 may form part of the RNA binding domain but play a minor role, only detectable if other important positions (e.g. amino acid 107) are altered. To assure that the differences observed with the biotinylated probes were reliable, the RNA binding activity of wild type and K107-3AB proteins were tested by a gel retardation assay with a P-labeled riboprobe (Fig. 3C). Incubation of the RNA probe with identical amounts of either wild type or K107E-3AB showed a different pattern for the RNA-protein complex. Thus, incubation with 1 µg of K107E-3AB gave place to an RNA-protein complex migrating slightly farther than the one observed with 1 µg of wild type protein. Higher concentrations of wild type protein produced a strong band corresponding to the retarded probe, whereas incubation with the mutant protein only elicited a weak retardation. These results corroborate the data presented in Fig. 3B demonstrating that the RNA binding activity is not completely abolished in the K107E-3AB mutant. No retardation was found either with 3A or with the K9E/K107E-3AB (data not shown).


Figure 3: Purification and RNA binding activities of 3A and 3AB mutant proteins. A, five micrograms of the purified 3AB proteins or 3.5 µg of 3A protein were submitted to SDS-polyacrylamide gel electrophoresis and stained with Coomassie Blue. Recombinant 3AB proteins with single amino acid substitutions at positions 9 (K9E) and 107 (K107E) were purified from E. coli-expressing cells through DEAE- and S-Sepharose chromatographic steps. The recombinant 3AB protein with a double amino acid substitution at positions 9 and 107 (K9E/K107E) and the wild type 3A protein were purified only through DEAE-cellulose (DE-52). The pH of the buffers used to elute the S-Sepharose columns is indicated in the top of the panel. B, five micrograms of purified 3AB and 3A proteins were assayed for RNA binding activity as shown in Fig. 1A. The upperpanel shows the activity of the fraction eluted from the S-Sepharose column at either pH 7.2 or 6.5 (S-200 fractions). The lowerpanel shows the activity of the proteins eluted from the DE-52 column. This gel has been overexposed to detect minimal amounts of bound riboprobe. C, different amounts of wild type and K107E-3AB proteins purified through DEAE- and S-Sepharose resins (the latter at pH 6.5) were used to carry out a gel retardation analysis with a P-labeled probe, as indicated in Fig. 1C. The positions of free and protein-bound probes are shown. Competitor poly(A) RNA was included in some reactions.



Binding of RNA by 3AB Is Required for the Stimulation of 3D

The following step was to ascertain how the RNA binding defects of 3AB affect the stimulatory activity on the viral polymerase. Fig. 4shows the poly(U) synthesis mediated by 3D in the absence or presence of different 3A and 3AB proteins. As described previously, the wild type 3AB protein induces a potent stimulation in the synthesis of poly(U)(24) . Thus, addition of 10 ng/µl of wild type 3AB induced a 30-fold stimulation on 3D polymerase activity. A small reduction of the stimulating activity was found with the K9E-3AB protein. This effect became clear at low concentrations of K9E-3AB. Nevertheless, when high concentrations were assayed (16 ng/µl) the mutated protein produced more than a 25-fold activation of 3D (Fig. 4). On the other hand, amino acid substitution at position 107 induced an important decrease in the 3D stimulatory activity. Thus, while 8 ng/µl of the wild type protein gave place to a 30-fold stimulation of 3D, the poly(U) synthesis in the presence of the same amount of K107E-3AB was only 3-fold stimulated, whereas no transactivation occurred either with the double K9E/K107E-3AB mutant or with the 3A protein. The results show a clear correlation between the RNA binding and 3Dstimulatory activities of 3A and 3AB proteins and, therefore, suggest that the binding of RNA by 3AB is an essential step to exert its stimulatory effect on the viral polymerase.


Figure 4: Effect of RNA binding-defective 3AB proteins on the polymerase activity of 3D. Poly(A)-dependent, oligo(dT)-primed poly(U) synthesis catalyzed by purified poliovirus RNA polymerase was assayed in the presence of different amounts of wild type 3AB, K9E, K107, K9E/K107-3AB, or wild type 3A proteins. The mixture was incubated for 80 min as indicated under ``Materials and Methods.'' The data are plotted as the -fold stimulation compared with the synthesis of poly(U) in the absence of 3AB or 3A proteins.



Characterization of 3AB Poliovirus Mutants with Severe Defects in RNA Replication

In order to understand the role of 3AB in viral replication, the 3AB mutations were introduced into the viral cDNA genome. In vitro transcription to produce infectious RNAs was carried out with T7 RNA polymerase. The transcript RNAs were mixed with Lipofectin and used to transfect HeLa cell monolayers. Then the appearance of the cytopathic effect was followed. Wild type RNA produced around 50% CPE by 2 days after transfection (). The mutant K9E-3AB showed a small delay in the appearance of the CPE. However, the mutant RNA encoding for the 3AB protein with the lysine to glutamic acid change at position 107 showed almost 2 days of delay in the appearance of the cytopathic effect, and this was still greater in the case of the double K9E/K107E-3AB mutant. The titer of the viruses recovered after 72 h of transfection was estimated. Transfection with either wild type or K9E-3AB mutant gave place to virus titers over 1 10 pfu/ml. However, the viruses recovered after transfection with K107E-3AB, and K9E/K107E-3AB showed titer reductions of 3 log units, suggesting that the defect of the encoded 3AB proteins severely impaired the replication of these viruses. As expected for replication defects, the viruses recovered showed a drastic plaque size phenotype reduction (Fig. 5). Thus, viruses recovered after 3 days of transfection with K107-3AB RNA produced plaques with an average diameter of 2.4 mm (small-plaque phenotype), in contrast to the wild type and the K9E-3AB counterparts, with an average plaque diameter of 4.9 mm after 72 h of incubation at 37 °C (data not shown). Interestingly, the double mutant showed a minute plaque size phenotype (0.8 mm in diameter). These results suggest that, as in the RNA binding and 3Dstimulatory functions, mutations at positions 9 and 107 of 3AB have a synergistic effect on the replication of the virus. The RNA sequence of each of these viruses was verified by RT-PCR sequencing of the virion RNA. The same plaque phenotype was observed when the amplified 3AB regions of each mutant were reintroduced in the wild type cDNA clone and used to produce new infectious RNAs (data not shown). Attempts to amplify and grow the K107E-3AB mutants repeatedly failed. When these viruses were further grown in HeLa cell monolayers, the obtained viruses rapidly reverted to the normal plaque size phenotype. As an example, the plaque sizes of the viruses recovered 3 or 6 days post-transfection can be compared in Fig. 5. This was not surprising, since although the RNA sequence of the pooled virion RNA was shown to be correct for any of the viruses recovered after 3 days of transfection, the analysis of the plaque phenotype showed the co-existence of large plaque revertant viruses among the small plaque mutant ones. The proportion of revertants was as much as 1% of the total population in the case of the double mutant after 3 days of transfection (data not shown). None of the mutant viruses was thermosensitive, and identical plaque size phenotypes were observed in different cell lines (data not shown). The failure to grow these viruses above 1 10 pfu/ml was a great handicap in subsequent work. The growth pattern of the isolated mutants was studied in one-step growth curves. HeLa cells were infected at the maximal multiplicity of infection (0.1 pfu/cell) permitted by the low titer stocks. The intracellular virus yield was monitored at different times postinfection (Fig. 6). The mutants K107E and K9E/K107-3AB showed different kinetics as compared with the wild type virus. This effect was only patent at early times of infection. Thus, at 4 h postinfection, the production of viruses was 100-1000-fold greater in wild type virus than in any of the K107E mutants. This effect disappeared later in the infection, and the rate of virus production between 4 and 8 h was identical for all the tested viruses. Despite this partial recovery, the total viral production at 8 h postinfection was 10 times lower in both K107E-3AB mutant polioviruses. These results suggest that the viruses that encode 3AB proteins defective in RNA binding and 3D transactivating activities suffer an impairment in virus growth during an early step of the infection. No deviation from the wild type behavior was found with the K9E-3AB mutant, suggesting that the stimulatory function of 3D provided by the mutated 3AB was enough to permit the replication of the virus at wild type levels.


Figure 5: Plaque phenotype of poliovirus mutants encoding for 3AB proteins with defects in the in vitro stimulation of poly(U) synthesis catalyzed by 3D. Infectious RNAs from wild type, K9E, K107E, and K9E/K107E-3AB pT7XLD constructs were obtained by invitro transcription of full-length cDNA clones. HeLa cells were transfected with infectious RNAs by the Lipofectin procedure. At 3 or 6 days post-transfection the cells were scraped and lysed to recover viruses. The plaque phenotype of these viruses was determined after infection of HeLa cells overlaid with 0.7% agar. Two different dilutions are shown for each virus.




Figure 6: One-step growth curves of wild type and 3AB poliovirus mutants. HeLa cell monolayers were infected with a multiplicity of infection of 0.1 pfu/cell and incubated at 37 °C. At different times postinfection the cells were scraped, and the intracellular virus production was assayed. Virus yield is given as the log pfu/ml.



In light of these results we decided to measure the viral RNA synthesis early in infection. An RT-PCR procedure was used to estimate the accumulation of plus and minus viral RNA strands. The cells were infected with 0.1 pfu/cell, and after 4 h the viral specific RNA levels were measured (Fig. 7). As an internal control for the retrotranscriptase and PCR reactions, the -actin mRNAs were amplified in the same test tubes. The PCR signal for both plus and minus polio RNA obtained with any of the K107E mutants was equal or less than the signal observed with a 10-fold dilution of the RNA isolated from cells infected with the wild type virus. However, the amount of -actin mRNA was not significantly altered after 4 h of infection with any of the viruses tested. These estimations mean over 90% inhibition in the accumulation of viral RNAs of K107E and K9E/K107E-3AB polioviruses. No significant difference in the amount of polio-specific RNAs was found with the K9E-3AB mutant virus. The defect observed in the K107E viruses seems to occur at a step affecting the synthesis of both plus and minus strands. In the case of the K107E mutant polioviruses, the impossibility of obtaining high titer viral stocks impeded the detection of the viral proteins synthesized in infected cells. No difference in the pattern of viral proteins was found in HeLa cells infected with either K9E-3AB or wild type poliovirus (data not shown). To test whether the amino acid substitution at position 107 affects the synthesis and processing of the polyprotein, we decided to analyze the proteins synthesized in vitro with nuclease-treated HeLa cell extracts. These lysates permit the faithful translation of poliovirus RNA. Indeed, under optimized conditions, translation of poliovirus RNA leads to de novo viral RNA synthesis and the formation of infectious viral particles(29) . Therefore, this procedure perfectly mimicked the biochemical processes that take place in vivo. Translation of mutant and wild type RNAs was performed in uninfected HeLa cell extracts programmed with viral RNAs (Fig. 8). The proteins were labeled with [S]methionine and separated in SDS-polyacrylamide gels. No difference was observed in the general pattern of synthesized proteins. Indeed, 3AB was clearly observed in all cases, suggesting that the 3AB-3C processing is not being affected by the lysine to glutamic acid change at position 107. Note that 3ABs containing a glutamic acid at position 9 migrated slightly faster than the other 3AB proteins. Taken together, these results suggest that the defects found in the K107E-3AB poliovirus mutants cannot be due to abnormal processing of the polyprotein but rather are due to defects in intrinsic activities of the 3AB protein.


Figure 7: Detection of plus and minus strand polio RNA in HeLa cells infected with 3AB poliovirus mutants. HeLa cells were infected with either wild type poliovirus or with the K9E, K107E, or K9E/K107E-3AB mutant viruses at a multiplicity of infection of 0.1 pfu/cell. After 4 h of infection the cells were lysed, and the total cytoplasmic RNA was isolated. Identical amounts of these RNAs were used as templates for an RT-PCR reaction to determine the amount of plus and minus polio RNA and -actin mRNA. The upperportion shows the 737-nucleotide-long product amplified by RT-PCR, which corresponds to the 2AB encoding region of the genome and the 1170-nucleotide fragment corresponding to the -actin mRNA. To estimate with accuracy the amount of RNA, the RT-PCR analysis was also carried out with serial dilutions of the RNA recovered after 4 h postinfection with wild type virus. RT-PCR reactions with RNA isolated from mock-infected cells were carried out in parallel. The lowerpart of the figure represents the densitometric analysis of the -actin and 2AB amplified fragments. Details are given under ``Materials and Methoeds.''




Figure 8: In vitro synthesis of poliovirus proteins in HeLa cell extracts programmed with wild type or 3AB mutant RNAs. The RNAs obtained by in vitro transcription of cDNA clones were used to program HeLa S3 cell extracts treated with micrococcal nuclease. The synthesized proteins were labeled with [S]methionine and electrophoresed in polyacrylamide gels. Control reaction was programmed with no exogenously added RNA. The positions of some viral proteins are denoted.




DISCUSSION

In order to ascertain the function of 3AB during the poliovirus replication cycle, a biochemical and molecular genetic analysis of this gene has been carried out. Recently, a novel procedure for the study of protein RNA interactions has been developed(26) . This device allowed us to demonstrate an RNA binding activity for 3AB protein. The RNA binding capability of 3AB was studied by assaying a number of mutants. Among these, five out of the nine positively charged lysine residues in the 3AB molecule were tested, leading us to the identification of amino acid residue 107 as an important part of the RNA binding domain. Interestingly, the hydrophobic domain of 3AB was not shown to be relevant to eliciting an RNA binding activity, despite the fact that this region of the protein has been involved in the replication of the genome. Probably, 3AB is endowed with different and independent domains implicated in RNA synthesis. In addition, Lama and Carrasco()have shown evidence for a membrane-permeabilizing activity of 3AB protein and proposed that this protein may be involved in the generation of the cytopathic effect observed in infected cells. Thus, 3AB could perform multiple functions during poliovirus infection, perhaps at different steps of the replication cycle.

Under our experimental conditions, the binding of 3AB seems to be nonspecific. The purified protein bound equally well to three different biotinylated riboprobes representing both extremes of the genome and an internal fragment of the complementary viral RNA (minus strand RNA). Another report has shown the binding of 3AB to P-labeled poly(A) probes that were retarded in gel shift assays(30) . The authors suggested that the binding of 3AB may be specifically directed to the 3`-end-poly(A) region of the genome, but no experimental evidence was shown to support this hypothesis. Although we cannot rule out a specific binding of 3AB to any part of the genome, the available data supports the possibility that 3AB protein by itself binds to any heteropolymeric or homopolymeric ribonucleic acid.

The RNA binding-defective 3AB proteins showed reduced stimulatory activities of the poly(U) synthesis directed by the viral polymerase. Thus, the single amino acid substitution at position 107 showed 10-20% of the wild type stimulatory activity whereas no transactivation was observed with either the double mutant K9E/K107E or the 3A protein, lacking the complete VPg sequence. A reduction was observed with the single mutant K9E-3AB, although this reduction was partially overcome at greater 3AB concentrations. This defect does not seem to be important since the K9E-3AB poliovirus mutant replicated at wild type levels. Introduction of a positively charged residue at position 9 may drastically change the electrostatic and/or structural requirements for RNA binding function. Glutamic acid at position 9, but not at position 107, drastically changed the electrophoretic behavior of 3AB in SDS-polyacrylamide gels (see for instance Fig. 3A). However, the key findings of this work were two: 1) the purified 3A protein does not show either RNA binding or transactivating activity, and 2) mutations in the 3A and VPg region cause a synergistic effect if placed together in the same 3AB molecule. This synergistic effect was also observed in the plaque size phenotype and in the time of appearance of CPE after transfection of HeLa cells with the mutant RNAs. Taken together, these results suggest that 3AB, but not 3A, plays an important role in the synthesis of virus-specific RNAs. This function must probably rely on its RNA binding and transactivating activities. Additional mutants with defects in both RNA binding and transactivation of 3D will be required to further confirm the generality of this conclusion. A more extensive analysis might identify positions in 3AB apart from the VPg domain but still involved in these functions.

The level of accumulation of both plus and minus viral RNA strands was inhibited by more than 90% at 4 h postinfection, and the virus yield was reduced more than 100-fold in HeLa cells infected with the K107-3AB poliovirus mutants. Therefore, these alterations seem to take place in the replication of both strands of the genome, probably at an early step in the infection when the amount of viral RNA is low. The replication impairment may be due to a defect in the elongation step or in a common step in the synthesis of both kinds of strands. Otherwise, we cannot rule out that the uridylylation of VPg is being affected in these mutant viruses. Nevertheless, the strong correlation between the in vitro stimulation of poly(U) synthesis produced by 3AB and the replication rate of the mutant viruses makes it likely that the RNA synthesis defect takes place in a step that requires the stimulatory effect of 3AB to be carried out. Unfortunately, we are not able to contrast both hypotheses. In vitro experiments would be necessary to demonstrate a reduced capability of the mutant 3ABs to function as substrates in the uridylylating reaction. However, the responsible enzyme has not been identified yet. On the other hand, VPg uridylylation has been studied in vitro with polio-infected HeLa cell extracts. In order to prepare these extracts, cells are infected with as much as 500 pfu/cell(19) . These experiments are incompatible with the replication-defective mutants described herein.

The function of 3AB does not seem to be essential for the virus. A null 3D transactivating 3AB protein (K9E/K107E-3AB) was able to maintain the replication of the virus, although to greatly reduced levels. This does not mean at all that the proposed function is not important for the virus. On the contrary, its importance is supported by the high phenotypic reversion rate showed by the K107E-3AB mutant viruses. Different reports have identified amino acid changes in 3AB that exert a severe impairment in the replication of the RNAs(21, 22, 23) . Despite this, the exact role of 3AB in the replication of the genome has never been understood(1, 34) . At least in one case, the altered function of 3AB was shown to be necessary only at an early step in the infection cycle(21) . Surprisingly, the synthesis of both positive and negative strands was severely depressed, as occurs with the replication mutants shown here. It would be of interest to determine whether that 3AB protein is able to stimulate poly(U) synthesis by 3D. Kuhn et al. described a poliovirus mutant (VPg26) with a double amino acid substitution at positions 107 (K107E) and 97 (K97R) of 3AB protein(35) . The authors showed a deviation from the kinetics of virus production similar to our K107E-3AB mutants, but the nature of this alteration was not studied. Other amino acid substitutions at position 107 showed virus-yield defects early in the infection, corroborating the idea that this position plays an important role in the replication of the virus.

Important considerations must be kept in mind to understand how 3AB works in vivo. We have demonstrated that 3AB seems to bind RNA probes in a nonspecific way, but then how is the specificity for polio RNA obtained? Our attempts to demonstrate such preference by, for instance, the 3`-end of the genome have failed to date. In any case, this selectivity should be extraordinarily high to ensure that the replication complexes can trap viral RNAs in a cytoplasm full of millions of molecules of cellular RNAs. Another intriguing possibility comes up if 3AB binds much faster to the messenger RNA from where it is being translated. In this way, the chance to bind to cellular RNAs would be reduced to a minimum without the requirement for any specific signal in the poliovirus genome. This ``trapping'' activity might not be so important at late steps of infection, when thousands of virus-specific RNA molecules are found in the cytoplasm. This could be an explanation for the overcoming of the growth defects observed with the K107E mutants at late steps of the infection. Otherwise, we cannot rule out the existence of factors that in conjunction with 3AB direct the viral polymerase to specific regions of the genome.

Another puzzling question arising from our results is, ``Why does 3D need a transactivator to reach full activity?'' This modular system must contribute to the poliovirus cycle with some advantages. The replication of viral RNAs can be divided into two separate reactions, initiation and elongation, which may be carried out by different enzymes(2, 36) . In some way, these reactions share some similarities with the synthesis of RNA directed by DNA-dependent RNA polymerases. In both cases initiation relies on the recognition of specific signals in the RNA or DNA template molecules. For transcription to occur, first the polymerase recognizes and stably binds to specific promoter regions forming a ``closed complex.'' Later, the closed promoter complex will isomerize to an ``open state,'' allowing the release of the polymerase to carry out the elongation of the RNA chain. The OmpR transcriptional activator protein enhances the binding of E. coli RNA polymerase to synthetic promoters. This interaction leads to the activation of transcription on weak promoters, whereas the same interaction produces a negative effect on strong promoters(37) . These findings suggest that an increase in the affinity of the polymerase to the promoter can lead to an activation of the initial step but may cause a delay in subsequent steps, probably by ``freezing'' the polymerase in the closed complex state. During RNA-dependent RNA synthesis a similar process may be taking place. Thus, after binding of the polymerase (or a precursor thereof) to the viral genome (initiation), the system must switch to the elongation step. We propose that 3AB may be involved in this switching step. Initially, 3AB may be directly implicated in the recognition of these signals, increasing the affinity of the polymerase to these regions. Then, the proteolytic processing of 3AB to VPg and 3A (without affinity to RNA) could allow the release of the viral polymerase and the commitment of the process into the elongation step. Therefore, 3AB could work as OmpR protein does, but in this case the function may be modulated by proteolytic processing. Evidence supporting this model comes from the fact that 3AB but not 3A binds RNA and that this activity seems to play an important role in the in vitro transactivation of 3D.

The above model implies that in vivo the stimulatory activity of 3AB is partially due to its interaction with either 3D or a precursor thereof and that this interaction increases the chance of the polymerase to take part in the initiation process. Interestingly, Molla et al.(38) have shown that 3AB can co-immunoprecipitate with either 3D or 3CD, suggesting a stable interaction of 3AB with these proteins. The identification of 3AB and 3D mutants unable to interact with each other will help to elucidate the interactions that modulate the poliovirus replication complex. Alternatively, a prediction of this model implies that if the cleavage of 3AB to 3A and VPg is avoided, the synthesis of RNA chains would be stopped or delayed in the initiation step, just after addition of a short oligonucleotide to the primer protein. Also, if 3AB or a precursor of this protein is priming the initiation reaction, this model would explain the impossibility of detecting elongating RNA chains with the 3AB primer protein covalently bound to the 5`-end of the genome. Nevertheless, the proofs for this model will have to await the development of in vitro systems where the different steps involved in the synthesis of poliovirus RNA can be dissociated.

  
Table: 0p4in Plaque phenotype was determined for those viruses obtained after 72 h of transfection.(119)


FOOTNOTES

*
This work was supported by Plan Nacional project BIO 92-0715, Dirección General de Investigación Cientfica y Técnica project PB90-0177, and an institutional grant of Fundación Ramón Areces. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom correspondence should be addressed. Tel.: 34-1-397-8041; Fax: 34-1-397-4799.

The abbreviations used are: pfu, plaque-forming unit(s); PCR, polymerase chain reaction; RT-PCR, reverse transcription-polymerase chain reaction; MOPS, 4-morpholinepropanesulfonic acid; MES, 4-morpholineethanesulfonic acid; CPE, cytopathic effect.

Lama, J., and Carrasco, L., FEBS Lett., in press.


ACKNOWLEDGEMENTS

The expert help of Noem Sevilla is acknowledged. We thank Luis Blanco, Margarita Salas, and Isabel Najera for critical reading of the manuscript. The invaluable help of Luis Carrasco is also acknowledged.


REFERENCES
  1. Wimmer, E., Hellen, C. U. T., and Cao, X.(1993) Annu. Rev. Genet.27, 353-436 [CrossRef][Medline] [Order article via Infotrieve]
  2. Semler, B. L., Kuhn, R. J., and Wimmer, E.(1988) in RNA Genetics, Vol. I, (Domingo, E., Holland, J. J., and Ahlquist, P., eds) pp. 23-49, CRC Press, Boca Raton, FL
  3. Wimmer, E., Kuhn, R. J., Pincus, S., Yang, C. F., Toyoda, H., Nicklin, M. J., and Takeda, N.(1987) J. Cell Sci. (Suppl.)7, 251-276 [Medline] [Order article via Infotrieve]
  4. Kuhn, R. J., and Wimmer, E.(1987) in The Molecular Biology of Positive Strand RNA Viruses (Rowlands, D. J., Mahy, B. W. J., and Mayo, M., eds) pp. 17-51, Academic Press, Inc., New York
  5. Flanegan, J. B., Petterson, R. F., Ambros, V., Hewlett, N. J., and Baltimore, D.(1977) Proc. Natl. Acad. Sci. U. S. A.74, 961-965 [Abstract]
  6. Lee, Y. F., Nomoto, A., Detjen, B. M., and Wimmer, E.(1977) Proc. Natl. Acad. Sci. U. S. A.74, 59-63 [Abstract]
  7. Flanegan, J. B., and Baltimore, D.(1979) J. Virol.29, 352-360 [Medline] [Order article via Infotrieve]
  8. Flanegan, J. B., and Van Dyke, T. A.(1979) J. Virol.32, 155-161 [Medline] [Order article via Infotrieve]
  9. Bernstein, H. D., Sarnow, P., and Baltimore, D.(1986) J. Virol.60, 1040-1049 [Medline] [Order article via Infotrieve]
  10. Burns, C. C., Lawson, M. A., Semler, B. L., and Ehrenfeld, E.(1989) J. Virol.63, 4866-4874 [Medline] [Order article via Infotrieve]
  11. Kirkegaard, K.(1992) Curr. Opin. Genet. & Dev.2, 64-70 [Medline] [Order article via Infotrieve]
  12. Andino, R., Rieckhof, G. E., and Baltimore, D.(1990) Cell63, 369-380 [Medline] [Order article via Infotrieve]
  13. Andino, R., Rieckhof, G. E., Trono, D., and Baltimore, D.(1990) J. Virol.64, 607-612 [Medline] [Order article via Infotrieve]
  14. Johnson, K., and Sarnow, P.(1991) J. Virol.65, 4341-4349 [Medline] [Order article via Infotrieve]
  15. Molla, A., Paul, A. V., Schmid, M., Jang, S. K., and Wimmer, E.(1993) Virology196, 739-747 [CrossRef][Medline] [Order article via Infotrieve]
  16. Semler, B. L., Anderson, C. W., Hanecak, R., Dorner, L. F., and Wimmer, E.(1982) Cell28, 405-412 [Medline] [Order article via Infotrieve]
  17. Takegami, T., Semler, B. L., Anderson, C. W., and Wimmer, E.(1983) Virology128, 33-47 [Medline] [Order article via Infotrieve]
  18. Bienz, K., Egger, D., Rasser, Y., and Bossart, W.(1983) Virology131, 39-48 [Medline] [Order article via Infotrieve]
  19. Takeda, N., Kuhn, R. J., Yang, C. F., Takegami, T., and Wimmer, E. (1986) J. Virol.60, 43-53 [Medline] [Order article via Infotrieve]
  20. Takegami, T., Kuhn, R. J., Anderson, C. W., and Wimmer, E.(1983) Proc. Natl. Acad. Sci. U. S. A.80, 7447-7451 [Abstract]
  21. Berstein, H. D., and Baltimore, D.(1988) J. Virol.62, 2922-2928 [Medline] [Order article via Infotrieve]
  22. Giachetti, C., Hwang, S. -S., and Semler, B. L.(1992) J. Virol.66, 6045-6057 [Abstract]
  23. Giachetti, C., and Semler, B. L.(1991) J. Virol.65, 2647-2654 [Medline] [Order article via Infotrieve]
  24. Lama, J., Paul, A. V., Harris, K. S., and Wimmer, E.(1994) J. Biol. Chem.269, 66-70 [Abstract/Free Full Text]
  25. van der Werf, S., Bradley, J., Wimmer, E., Studier, W., and Dunn, J. (1986) Proc. Natl. Acad. Sci. U. S. A.83, 2330-2334 [Abstract]
  26. Rodrguez, P. L., and Carrasco, L.(1994) BioTechniques17, 702-705 [Medline] [Order article via Infotrieve]
  27. Lama, J., and Carrasco, L.(1992) J. Biol. Chem.267, 15932-15937 [Abstract/Free Full Text]
  28. Favaloro, J., Treisman, R., and Kamen, R.(1980) Methods Enzymol.65, 718-749 [Medline] [Order article via Infotrieve]
  29. Molla, A., Paul, A. V., and Wimmer, E.(1991) Science254, 1647-1651 [Medline] [Order article via Infotrieve]
  30. Paul, A., Cao, X., Harris, K., Lama, J., and Wimmer, E.(1994) J. Biol. Chem.269, 29173-29181 [Abstract/Free Full Text]
  31. Larsen, G. R., Anderson, C. W., Dorner, A. J., Semler, B. L., and Wimmer, E.(1982) J. Virol.41, 340-344 [Medline] [Order article via Infotrieve]
  32. Pelletier, J., and Sonenberg, N.(1988) Nature334, 320-325 [CrossRef][Medline] [Order article via Infotrieve]
  33. Jacobson, S. J., Konings, D. A. M., and Sarnow, P.(1993) J. Virol.67, 2961-2971 [Abstract]
  34. Porter, A. G.(1993) J. Virol.67, 6917-6921 [Medline] [Order article via Infotrieve]
  35. Kuhn, R. J., Tada, H., Ypma-Wong, M. F., Semler, B. L., and Wimmer, E. (1988) J. Virol.62, 4207-4215 [Medline] [Order article via Infotrieve]
  36. Richards, O. C., and Ehrenfeld, E.(1990) Curr. Top. Microbiol. Immunol.161, 89-119 [Medline] [Order article via Infotrieve]
  37. Tsung, K., Brissette, R., and Inouye, M.(1990) Proc. Natl. Acad. Sci. U. S. A.87, 5940-5944 [Abstract]
  38. Molla, A., Harris, K., Paul, A., Shin, S., Mugavero, J., and Wimmer, E. (1994) J. Biol. Chem.269, 27015-27020 [Abstract/Free Full Text]

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.