Department of Medical Microbiology and Immunology, Texas A&M University System Health Science Center, College Station, TX 77843-1114, USA1
The Institute of Advanced Studies, John Curtin School of Medical Research, PO Box 334, Canberra City, ACT 2601, Australia2
Office of Texas State Chemist, MS 2114, Texas A&M University, College Station, TX 77843-2114, USA3
Author for correspondence: Van Wilson. Fax +1 979 845 3479. e-mail v-wilson{at}tamu.ed
![]() |
Abstract |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
Although E1 interaction with both the E2TAD and the E2DBD has been established previously (Chen & Stenlund, 1998 ; Sarafi & McBride, 1995
; Yasugi et al., 1997
), the total number of interactions and the domains of each protein specifically involved are still not defined completely. While both an N-terminal E1 (E1N)E2 interaction as well as a C-terminal (E1C)E2 interaction have been reported previously (Benson & Howley, 1995
; Leng et al., 1997
; Moscufo et al., 1999
; Sarafi & McBride, 1995
; Thorner et al., 1993
), some studies have failed to detect an interaction between one or the other of these two E1 domains and E2 (Benson & Howley, 1995
; Lusky & Fontane, 1991
; Moscufo et al., 1999
). One goal of this study was to systematically identify the domains involved in the interaction between E1 and E2, both in vitro and in vivo. We verified independent E1NE2 and E1CE2 interactions and further demonstrated that E1N interacted with both the E2TAD and the E2DBD. Using single substitution mutations within E1N, we demonstrated that the amino acids involved in E2TAD and E2DBD interaction are not identical. In addition, we showed that two conserved, hydrophilic regions within the E1DBD, HR1 and HR3, are both important for E2TAD interaction. However, the E1NE2TAD interaction was not required for replication function and instead may be involved in transcription.
![]() |
Methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Protein purification.
For the expression of GSTE2 (glutathione S-transferase fused to E2), 2x YT medium containing 2% glucose and 50 µg/µl ampicillin was inoculated with an overnight culture of pGEXE2 and grown for either 3 h at room temperature or 1 h at 18 °C. Protein expression was induced with IPTG at a final concentration of 1 mM and the culture was incubated for a further 2 h at room temperature or 1218 h at 18 °C. Cells were then centrifuged at 10000 g for 15 min and the pellet was frozen at -70 °C for at least 1 h. B-per reagent (Pierce) containing 5 mM DTT and 1 mM PMSF was used for cell resuspension. Lysozyme was added to a concentration of 100 µg/ml and the suspension was incubated on ice for 1 h. The sample was then sonicated twice for 15 s using an Ultrasonics sonicator with the microtip at maximum power and then centrifuged for 30 min at 4 °C. GlutathioneSepharose beads were added to the supernatant and rotated overnight at 4 °C. Beads were washed twice with GST-C buffer (50 mM TrisHCl, pH 7·9, 250 mM NaCl, 5 mM EDTA, 10% glycerol) plus 5 mM DTT and 10 mM PMSF, once with GST-E buffer (50 mM TrisHCl, pH 8·0, 1 M NaCl, 5 mM EDTA, 10% glycerol) plus 5 mM DTT and 10 mM PMSF, and finally in GST-C buffer. Protein was eluted using 10 mM glutathione in GST-C buffer. Human thrombin (5 units) (Sigma) was added directly to 100 µg of eluted GSTE2 and incubated for 4 h at 20 °C. PMSF was added to a final concentration of 1 mM in order to inhibit the thrombin and the cleaved GST plus uncleaved GSTE2 were removed using glutathioneSepharose beads. Conversely, GSTE2 bound to glutathioneSepharose beads was cleaved using 10 units of thrombin overnight at room temperature. Beads were centrifuged at low speed and supernatant containing cleaved E2 was removed and stored at -20 °C. Protein quality was assessed by SDSPAGE and protein concentration was determined using the Bradford method. GSTE2DBD was expressed and purified as above. GSTE2TAD was purified as above except that the cell pellet was resuspended in PBS and French-pressed at 16000 PSI before centrifugation and the addition of glutathioneSepharose beads to the supernatant.
Electromobility shift assays (EMSAs).
Gel shift assays were performed as described previously (Gonzalez et al., 2000 ). Briefly, each 10 µl reaction mixture contained EMSA buffer (20 mM potassium phosphate, pH 7·0, 100 mM NaCl, 1 mM EDTA, 0·1% NP-40, 10% glycerol, 5 mM DTT, 0·07% BSA), 2·5 fmol of radiolabelled oligonucleotide, 20 ng pUC18 DNA and purified GSTE1DBD and E2. Oligonucleotide E1BS 1-4 consists of BPV-1 nucleotides 792629 (designated substrate B) and contains the authentic BPV-1 18 bp E1 binding element (E1BE) and the 12 bp E2BS12. Oligonucleotide E1BS 1-4 BS11 consists of nucleotides 789124 (designated substrate A) and includes the authentic BPV-1 E2BS11 and the 18 bp E1BE. BPV-1 nucleotides 1618 (ACC) in this sequence were changed to TAG to destroy the 5' portion of E2BS12. Samples were incubated for 30 min at 25 °C and then electrophoresed by 8% PAGE in 0·5x TrisborateEDTA (TBE) buffer (pH 7·5). Protein quantification was carried out using a Molecular Dynamics PhosphorImager.
Yeast two-hybrid assay.
Yeast transformations and liquid -galactosidase assays were performed as described by McShan & Wilson (2000)
. Alternatively, competent Saccharomyces cerevisiae strain SFY526 was co-transformed with pGBT9E1, pGBT9E11311 or pGBT9E1315605 and pGAD424E2 using the lithiumacetate method. Three separate clones from each transformation experiment were isolated and assayed for
-galactosidase activity using chloramphenicol red
-galactopyranoside as the substrate. Yeast co-transformants with E1N versus the E2 subdomains were made sequentially by first transforming yeast strain SFY526 with the pGBT9E11311 plasmid DNA. Competent pGBT9E11311 SFY526 cells were then transformed with DNA from either pGAD424E2TAD or pGAD424E2DBD. All E1N mutant co-transformants were made by first making competent pGAD424E2-, pGAD424E2TAD- and pGAD424E2DBD-transformed SFY526 cells. These cells were then re-transformed with pGBT9E11311 mutant plasmids. Fold stimulation was calculated from the
-galactosidase activities of individual co-transformants using the equation [(pGBT9X+pGAD424Y)-(pGBT9+pGAD424Y)]/(pGBT9X+pGAD424), where X is the E1 protein and Y is the E2 protein.
GST pulldown assay.
GSTE2, GSTE2TAD, GSTE2DBD or GST alone (10 µg) was incubated with 30 µl glutathioneSepharose beads in 500 µl of binding buffer (10 mM TrisHCl, pH 7·4, 50 mM NaCl, 2% BSA). Samples were incubated for 5 h at 4 °C and then centrifuged at 500 g for 4 min. In vitro translated 35S-labelled E1, E11311 or E1315605 was added to the beads in 500 µl binding buffer with 1% BSA and rocked for 20 h at 4 °C. Beads were washed first with TSA (10 mM TrisHCl, pH 8·0, 140 mM NaCl, 0·025% NaN3) and then three times with TSA containing 0·1% Triton X-100. Further washing steps were carried out with TSA containing 0·2% Triton X-100, then with TSA alone and finally with 50 mM TrisHCl, pH 6·8, containing 1 mM PMSF. After resuspension in each wash buffer, samples were vortexed for 5 s and incubated on ice for 23 min before centrifuging at 500 g. After washing, pellets were resuspended in 25 µl 2x SDS sample buffer and heated at 75 °C for 4 min. Half of each sample was electrophoresed by 10% SDSPAGE. Gels were then dried and visualized using a Molecular Dynamics PhosphorImager.
Transient replication assays.
pCGEagE1 (1 µg) or mutant pCGEagE1 (1 µg), and pCGE2 (0·1 µg) and either pBOR (1 µg; containing the ori) or pUC (1 µg; not containing the ori) were mixed with 12 µl PLUS reagent (Gibco) and incubated at room temperature for 20 min in 250 µl of Hams media with non-essential amino acids. LipofectAMINE reagent (Gibco) (12 µl) was mixed with 250 µl of Hams media containing non-essential amino acids and incubated for 10 min. LipofectAMINE solution was then added to the DNA/PLUS reagent mixture and incubated for an additional 20 min. Samples were added directly onto CHO cells, which had been seeded at a density of 1·4x106 cells/ml 1224 h earlier. Transfection was allowed to proceed for 3 h at 37 °C, after which the cells were trypsinized, split into three 60 mm plates and incubated for an additional 48, 72 or 96 h. DNA from each transfection experiment was harvested, digested with DpnI/HindIII and analysed by Southern blotting, as described previously (McShan & Wilson, 1997 ).
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
E2 is unable to rescue the HR1 and HR3 E1DBD mutants to wild-type levels
The above results, in combination with previous reports (Benson & Howley, 1995 ; Leng et al., 1997
; Moscufo et al., 1999
; Thorner et al., 1993
), confirm that E2 interacts with the N-terminal region of E1, a region that includes the functional E1DBD (Chen & Stenlund, 1998
; Enemark et al., 2000
; Leng et al., 1997
). Consistent with these direct interaction results is the observation that origin binding of an isolated wild-type E1DBD polypeptide is cooperatively enhanced by full-length E2 in an EMSA (Chen & Stenlund, 1998
). Although indirect, the EMSA results probably indicate that the E1DBD region itself makes contact with E2. To begin to define the E1DBD residues critical for this interaction, we examined substitution mutants in three E1DBD hydrophilic domains, designated HR1, HR2 and HR3 (Fig. 3A
) (Gonzalez et al., 2000
). The E1DBD proteins were tested for cooperative binding with E2 in the EMSA using substrate B (BPV-1 nucleotides 792629), which contains the authentic BPV-1 18 bp E1BE and the 12 bp E2BS12. Wild-type GSTE1DBD protein, in the absence of E2, formed two discrete complexes on the DNA (Fig. 3B
, lanes 3 and 17). E2 alone formed a predominant proteinDNA complex that migrated faster than the E1DNA complexes (Fig. 3B
, lanes 2 and 16). Combination of E2 with the wild-type E1DBD protein resulted in a new E1DBDE2DNA complex that migrated to a position in-between the two E1DNA complexes (Fig. 3B
, lanes 4 and 18). In the absence of E2, all mutations in the E1DBD, with the exception of the K157A mutant, decreased E1DNA binding to some degree. As reported previously (Gonzalez et al., 2000
), origin binding by HR1 and HR3 mutants (K183A, K186A, T187A and K241A) was less than 10% of that seen for the wild-type (Fig. 3B
, lanes 7, 9, 11 and 21; Fig. 3C
), while non-HR mutants bound between 45 and 130% of wild-type (Fig. 3B
, lanes 5, 19, 23 and 25; Fig. 3C
). In the presence of E2, all the E1DBD proteins with mutations outside of HR1 and HR3 were rescued at levels similar to those seen for the wild-type (Fig. 3B
, lanes 14, 20, 24 and 26; Fig. 3C
). In contrast, no protein with a mutation falling within the two conserved hydrophilic domains was comparably rescued (Fig. 3B
, lanes 8, 10, 12 and 22; Fig. 3C
), with the exception a nonconserved amino acid mutant, T188A (Fig. 3B
, lane 14; Fig. 3C
). These results indicate that the interaction of full-length E2 with the E1DBD is able to compensate for reduced DNA binding by the E1DBD non-HR1 or -HR3 mutants and restore wild-type levels of E1DBDE2DNA complex formation. In contrast, the E1DBD mutants with amino acid changes at conserved residues in HR1 and HR3, which are likely to be involved in direct DNA contact, were only partially rescued by E2. These data support and extend previous work by Thorner et al. (1993)
, which demonstrated that E2 was unable to rescue DNA binding by the HR3 mutants and a single mutation in HR1.
|
|
E2E1 cooperative binding is not sufficient for in vivo replication
To relate the biochemical properties of the HR1 and HR3 domains to biological activity, the E1DBD mutations were transferred to full-length E1 to evaluate replication capacity using an in vivo triple plasmid transient replication assay (Fig. 5). All mutants were able to support replication of pBOR at levels comparable to those seen for wild-type E1, except for the T187A, K241A and K279A mutants. Surprisingly, the K186A mutant replicated quite well, even though its cooperative E2 binding in vitro was only marginally better than the T187A mutant (Fig. 3
). Conversely, the K279A mutant exhibited wild-type origin binding and E2 cooperativity, yet was completely defective for in vivo replication. Consequently, it appears that while cooperative E1E2 binding is required for in vivo replication capacity, it is not necessarily sufficient. All replication defective mutants were assayed by Western blot and found to express E1 at least as well as the wild-type, ensuring that the lack of replicative ability was not due to a lack of protein expression (data not shown).
|
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Our present study defined further the E1NE2 interaction by demonstrating that the N-terminal E1 domain is sufficient for interaction with the E2TAD as well as the E2DBD. Our E1NE2TAD results correlate well with an earlier study by Benson & Howley (1995) , which showed an E1 N-terminal domain interaction with E2191. However, several previous studies using immunoprecipitation or GST pulldown assays were unable to detect either a full-length E1E2DBD interaction or an E1NE2DBD interaction (Benson & Howley, 1995
; Mohr et al., 1990
; Moscufo et al., 1999
). The only previous reports of an E1NE2DBD interaction have been in assays using a DNA probe containing either the E1BE or the E1BE and the E2BS (Chen & Stenlund, 1998
, 2000
; Gillitzer et al., 2000
). Due to the apparently weak nature of this interaction, it may be that it is often difficult to detect E1E2DBD complexes in the absence of DNA. Nevertheless, we have now confirmed this interaction directly in both yeast two-hybrid and GST pulldown assays. Therefore, our combined results clarify some of the previous discrepancies and define three distinct E1E2 interactions: E1CE2, E1NE2TAD and E1NE2DBD.
Using a series of substitution mutations in the E1DBD, we were able to evaluate the contribution of this region to E2TAD and E2DBD interaction and replication function (summarized in Table 1). Mutants with mutations at conserved residues in HR1 (K183, K186 and T187) and HR3 (K241), as well as at residue K267, were unable to bind to the E2TAD. From the E1DBD crystal structure (Enemark et al., 2000
), these five amino acids form a confluent accessible surface that could be available for interaction with the E2TAD. However, it does not appear that the interaction defined by these mutations is required for viral genome replication, as three of the five mutants are wild-type for DNA replication activity. Also, it seems unlikely that the E2TAD could be interacting with this E1 region to facilitate replication, since this region is apparently involved in direct contact with origin DNA (Enemark et al., 2000
). Using a different set of mutants, the same approximate region of the E1DBD was shown recently to be required for mediating E1 modulation of E2 transcriptional activity (Parker et al., 2000
). Since the E1 transcriptional effect requires the E1BS (Parker et al., 2000
), there would be no conflict between DNA binding and E2TAD binding in this context. Taken together, our results and those of Parker et al. (2000)
strongly suggest that the E1DBDE2TAD interaction is relevant for transcription, rather than playing a role in replication. Therefore, the E1E2TAD requirement for replication (Berg & Stenlund, 1997
; Chen & Stenlund, 1998
) is probably mediated through sequences downstream of the E1DBD in the C-terminal region of E1.
|
The explanation for the replication defects of the K241A and K279A mutants is less clear. The K241A mutant interacted with the E2DBD and exhibited more cooperative origin binding activity in vitro than the K186A mutant; total inability to form the origin initiation complex seems unlikely. However, this mutation could clearly affect some more subtle aspect of initiation complex assembly or could affect post-assembly processes, such as DNA unwinding (Gillette et al., 1994 ) and interaction with host cell proteins. Likewise, the K279A mutant was not impaired in any tested function, yet was unable to replicate. Based on a hydropathy plot of the E1DBD (Gonzalez et al., 2000
), amino acid 279 falls within a 12 amino acid hydrophobic sequence (aa 272283) (Fig. 3A
). Another mutation in this hydrophobic region, L275A, has the same phenotype and is functional for origin recognition and E2 cooperative binding, but is extremely impaired for replication (Thorner et al., 1993
). From the crystal structure (Enemark et al., 2000
), this hydrophobic region contains the
-helix 5 and, consistent with the wild-type DNA binding properties of the K279A mutant, does not appear to contribute directly to the DNA contact region formed by HR1 and HR3. Consequently, while their precise role in replication function remains obscure, mutants L275A and K279A appear to define a new functional subregion for the E1DBD that is critical for replication but does not directly involve origin recognition activity.
![]() |
Acknowledgments |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Berg, M. & Stenlund, A. (1997). Functional interactions between papillomavirus E1 and E2 proteins. Journal of Virology 71, 3853-3863.[Abstract]
Blitz, I. L. & Laimins, L. A. (1991). The 68-kilodalton E1 protein of bovine papillomavirus is a DNA binding phosphoprotein which associates with the E2 transcriptional activator in vitro. Journal of Virology 65, 649-656.[Medline]
Bonne-Andrea, C., Santucci, S., Clertant, P. & Tillier, F. (1995). Bovine papillomavirus E1 protein binds specifically DNA polymerase but not replication protein A. Journal of Virology 69, 2341-2350.[Abstract]
Bonne-Andrea, C., Tillier, F., McShan, G. D., Wilson, V. G. & Clertant, P. (1997). Bovine papillomavirus type 1 DNA replication: the transcriptional activator E2 acts in vitro as a specificity factor. Journal of Virology 71, 6805-6815.[Abstract]
Chen, G. & Stenlund, A. (1998). Characterization of the DNA binding domain of the bovine papillomavirus replication initiator E1. Journal of Virology 72, 2567-2576.
Chen, G. & Stenlund, A. (2000). Two patches of amino acids on the E2 DNA-binding domain define the surface for interaction with E1. Journal of Virology 74, 1506-1512.
Chow, L. T. & Broker, T. R. (1994). Papillomavirus DNA replication. Intervirology 37, 150-158.[Medline]
Enemark, E. J., Chen, G., Vaughn, D. E., Stenlund, A. & Joshua-Tor, L. (2000). Crystal structure of the DNA binding domain of the replication initiation protein E1 from papillomavirus. Molecular Cell 6, 149-158.[Medline]
Ferran, M. C. & McBride, A. A. (1998). Transient viral DNA replication and repression of viral transcription are supported by the C-terminal domain of the bovine papillomavirus type 1 E1 protein. Journal of Virology 72, 796-801.
Fouts, E. T., Yu, X., Egelman, E. H. & Botchan, M. R. (1999). Biochemical and electron microscopic image analysis of the hexameric E1 helicase. Journal of Biological Chemistry 274, 4447-4458.
Gillette, T. G. & Borowiec, J. A. (1998). Distinct roles of two binding sites for the bovine papillomavirus (BPV) E2 transactivator on BPV DNA replication. Journal of Virology 72, 5735-5744.
Gillette, T. G., Lusky, M. & Borowiec, J. A. (1994). Induction of structural changes in the bovine papillomavirus type 1 origin of replication by the viral E1 and E2 proteins. Proceedings of the National Academy of Sciences, USA 91, 8846-8850.[Abstract]
Gillitzer, E., Chen, G. & Stenlund, A. (2000). Separate domains in E1 and E2 proteins serve architectural and productive roles for cooperative DNA binding. EMBO Journal 19, 3069-3079.
Giri, I. & Yaniv, M. (1988). Structural and mutational analysis of E2 trans-activating proteins of papillomaviruses reveals three distinct functional domains. EMBO Journal 7, 2823-2829.[Abstract]
Gonzalez, A., Bazaldua-Hernandez, C., West, M., Woytek, K. & Wilson, V. G. (2000). Identification of a short, hydrophilic amino acid sequence critical for origin recognition by the bovine papillomavirus E1 protein. Journal of Virology 74, 245-253.
Han, Y. F., Loo, Y. M., Militello, K. T. & Melendy, T. (1999). Interactions of the papovavirus DNA replication initiator proteins, bovine papillomavirus type 1 E1 and simian virus 40 large T antigen, with human replication protein A. Journal of Virology 73, 4899-4907.
Haugen, T. H., Turek, L. P., Mercurio, F. M., Cripe, T. P., Olson, B. J., Anderson, R. D., Seidl, D., Karin, M. & Schiller, J. (1988). Sequence-specific and general transcriptional activation by the bovine papillomavirus-1 E2 trans-activator require an N-terminal amphipathic helix-containing E2 domain. EMBO Journal 7, 4245-4253.[Abstract]
Holt, S. E., Schuller, G. & Wilson, V. G. (1994). DNA binding specificity of the bovine papillomavirus E1 protein is determined by sequences contained within an 18-base-pair inverted repeat element at the origin of replication. Journal of Virology 68, 1094-1102.[Abstract]
Le Moal, M. A., Yaniv, M. & Thierry, F. (1994). The bovine papillomavirus type 1 (BPV1) replication protein E1 modulates transcription activation by interacting with BPV1 E2. Journal of Virology 68, 1085-1093.[Abstract]
Leng, X., Ludesmeyers, J. H. & Wilson, V. G. (1997). Isolation of an amino-terminal region of bovine papillomavirus type 1 E1 protein that retains origin binding and E2 interaction capacity. Journal of Virology 71, 848-852.[Abstract]
Lusky, M. & Fontane, E. (1991). Formation of the complex of bovine papillomavirus E1 and E2 proteins is modulated by E2 phosphorylation and depends upon sequences within the carboxyl terminus of E1. Proceedings of the National Academy of Sciences, USA 88, 6363-6367.[Abstract]
Lusky, M., Hurwitz, J. & Seo, Y.-S. (1994). The bovine papillomavirus E2 protein modulates the assembly of but is not stably maintained in a replication-competent multimeric E1-replication origin complex. Proceedings of the National Academy of Sciences, USA 91, 8895-8899.[Abstract]
McBride, A. A., Byrne, J. C. & Howley, P. M. (1989). E2 polypeptides encoded by bovine papillomavirus type 1 form dimers through the common carboxyl-terminal domain: transactivation is mediated by the conserved amino-terminal domain. Proceedings of the National Academy of Sciences, USA 86, 510-514.[Abstract]
McBride, A. A., Romanczuk, H. & Howley, P. M. (1991). The papillomavirus E2 regulatory proteins. Journal of Biological Chemistry 266, 18411-18414.
McShan, G. D. & Wilson, V. G. (1997). Reconstitution of a functional bovine papillomavirus type 1 origin of replication reveals a modular tripartite replicon with an essential AT-rich element. Virology 237, 198-208.[Medline]
McShan, G. & Wilson, V. G. (2000). Contribution of bovine papillomavirus type 1 E1 protein residue 48 to replication function. Journal of General Virology 81, 1995-2004.
Mohr, I. J., Clark, R., Sun, S., Androphy, E. J., MacPherson, P. & Botchan, M. R. (1990). Targeting the E1 replication protein to the papillomavirus origin of replication by complex formation with the E2 transactivator. Science 250, 1694-1699.[Medline]
Moscufo, N., Sverdrup, F., Breiding, D. E. & Androphy, E. J. (1999). Two distinct regions of the BPV-1 E1 replication protein interact with the activation domain of E2. Virus Research 65, 141-154.[Medline]
Park, P., Copeland, W., Yang, L., Wang, T., Botchan, M. R. & Mohr, I. J. (1994). The cellular DNA polymerase -primase is required for papillomavirus DNA replication and associates with the viral E1 helicase. Proceedings of the National Academy of Sciences, USA 91, 8700-8704.[Abstract]
Parker, L. M., Harris, S., Gossen, M. & Botchan, M. R. (2000). The bovine papillomavirus E2 transactivator is stimulated by the E1 initiator through the E2 activation domain. Virology 270, 430-443.[Medline]
Rangasamy, D. & Wilson, V. G. (2000). Bovine papillomavirus E1 protein is sumoylated by the host cell Ubc9 protein. Journal of Biological Chemistry 275, 30487-30495.
Sanders, C. M. & Stenlund, A. (1998). Recruitment and loading of the E1 initiator protein: an ATP-dependent process catalysed by a transcription factor. EMBO Journal 17, 7044-7055.
Sanders, C. M. & Stenlund, A. (2000). Transcription factor-dependent loading of the E1 initiator reveals modular assembly of the papillomavirus origin melting complex. Journal of Biological Chemistry 275, 3522-3534.
Sarafi, T. R. & McBride, A. A. (1995). Domains of the BPV-1 E1 replication protein required for origin-specific DNA binding and interaction with the E2 transactivator. Virology 211, 385-396.[Medline]
Sedman, J. & Stenlund, A. (1995). Co-operative interaction between the initiator E1 and the transcriptional activator E2 is required for replicator specific DNA replication of bovine papillomavirus in vivo and in vitro. EMBO Journal 14, 6218-6228.[Abstract]
Sedman, J. & Stenlund, A. (1996). The initiator protein E1 binds to the bovine papillomavirus origin of replication as a trimeric ring-like structure. EMBO Journal 15, 5085-5092.[Abstract]
Sedman, J. & Stenlund, A. (1998). The papillomavirus E1 protein forms a DNA-dependent hexameric complex with ATPase and DNA helicase activities. Journal of Virology 72, 6893-6897.
Seo, Y. S., Müller, F., Lusky, M. & Hurwitz, J. (1993). Bovine papillomavirus (BPV)-encoded E1 protein contains multiple activities required for BPV DNA replication. Proceedings of the National Academy of Sciences, USA 90, 702-706.[Abstract]
Spalholz, B. A., Yang, Y. C. & Howley, P. M. (1985). Transactivation of a bovine papillomavirus transcriptional regulatory element by the E2 gene product. Cell 42, 183-191.[Medline]
Thorner, L. K., Lim, D. A. & Botchan, M. R. (1993). DNA-binding domain of bovine papillomavirus type 1 E1 helicase: structural and functional aspects. Journal of Virology 67, 6000-6014.[Abstract]
Ustav, M. & Stenlund, A. (1991). Transient replication of BPV-1 requires two viral polypeptides encoded by the E1 and E2 open reading frames. EMBO Journal 10, 449-457.[Abstract]
Wilson, V. G. & Ludes-Meyers, J. (1991). A bovine papillomavirus E1-related protein binds specifically to bovine papillomavirus DNA. Journal of Virology 65, 5314-5322.[Medline]
Yang, L., Mohr, I., Fouts, E., Lim, D. A., Nohaile, M. & Botchan, M. (1993). The E1 protein of bovine papillomavirus 1 is an ATP-dependent DNA helicase. Proceedings of the National Academy of Sciences, USA 90, 5086-5090.[Abstract]
Yasugi, T., Benson, J. D., Sakai, H., Vidal, M. & Howley, P. M. (1997). Mapping and characterization of the interaction domains of human papillomavirus type 16 E1 and E2 proteins. Journal of Virology 71, 891-899.[Abstract]
Received 21 March 2001;
accepted 1 June 2001.