Distinct Roles of Two Binding Sites for the Bovine Papillomavirus

JOURNAL OF VIROLOGY,0022-538X/98/$04.0010

July 1998, p. 5735–5744 Vol. 72, No. 7

Copyright © 1998, American Society for Microbiology. All Rights Reserved.

Distinct Roles of Two Binding Sites for the Bovine Papillomavirus(BPV) E2 Transactivator on BPV DNA Replication

THOMAS G. GILLETTE† AND JAMES A. BOROWIEC*

Department of Biochemistry and Kaplan Comprehensive Cancer Center, New York University Medical Center,New York, New York 10016

Received 4 December 1997/Accepted 31 March 1998

The modulation of DNA replication by transcription factors was examined by using bovine papillomavirustype 1 (BPV). BPV replication in vivo requires two viral proteins: E1, an origin-binding protein, and E2, atranscriptional transactivator. In the origin, E1 interacts with a central region flanked by two binding sites forE2 (BS11 and BS12), of which only BS12 has been reported to be essential for replication in vivo. Usingchemical interference and electrophoretic mobility shift assays, we found that the binding of E2 to each sitestimulates the formation of distinct E1-origin complexes. A high-mobility C1 complex is formed by usingcritical E2 contacts to BS12 and E1 contacts to the dyad symmetry element. In contrast, interaction of E2 withthe BS11 element on the other origin flank promotes the formation of the lower-mobility C3 complex. C3 is anovel species that resembles C2, a previously identified complex that is replication active and formed by E1alone. The binding of E1 greatly differs in the C1 and C3 complexes, with E1 in the C1 complex limited to theorigin dyad symmetry region and E1 in the C3 complex encompassing the region from the proximal edge ofBS11 through the distal edge of BS12. We found that the presence of both E2-binding sites is necessary forwild-type replication activity in vivo, as well as for maximal production of the C3 complex. These results showthat in the normal viral context, BS11 and BS12 play separate but synergetic roles in the initiation of viral DNAreplication that are dependent on their location within the origin. Our data suggest a model in which thebinding of E2 to each site sequentially stimulates the formation of distinct E1-origin complexes, leading to thereplication-competent complex.

Transcription factors have recently been recognized to playimportant modulatory roles during the initiation of eukaryoticDNA replication. In most cases, these factors act not by reg-ulating neighboring transcription units but, rather, by directlyinteracting with proteins bound to an origin of replication orwith the DNA itself (43). The mechanisms by which transcrip-tion factors regulate DNA replication have been most clearlydefined by using viral systems such as bovine papillomavirus(BPV) (29), adenovirus (19, 30), and simian virus 40 (SV40)(7), although chromosomal examples exist as well (see, e.g.,reference 11). In these systems, transcription factors actthrough various means, including the recruitment of replica-tion proteins to the origin, the modulation of the activity ofbound replication proteins, and the disruption of the localnucleosome structure, allowing replication factors access to theDNA.

In the BPV model system, in vivo replication of DNA con-taining the BPV origin requires two viral proteins designatedE1 and E2 (41), although only E1 is essential in vitro (36, 45).E1 is a DNA helicase (36, 46) and DNA-binding protein (29)that recognizes a dyad symmetry element within the viral originof replication (16, 42, 45). Origin binding by E1 is cooperativewith E2 (24, 32, 37, 45), and this cooperativity is mediated byphysical interaction between E1 and E2 (1–3, 22, 29). In thepresence of a single-stranded-DNA-binding protein such ashuman replication protein A (hRPA), E1 unwinds the DNA

outward from the origin, with its DNA helicase functioning inthe 39359 direction (36, 46).

The region encompassing the origin contains a dyad symme-try element adjacent to an AT-rich domain. This central regionis flanked by two binding sites for E2 termed BS11 and BS12,adjacent to the AT-rich and dyad elements, respectively. Mu-tational analysis to determine the minimal origin sequence hasindicated that the central region and BS12 are required forreplication in vivo (42). Consistent with the need only for E1 tosupport replication in vitro, a smaller region lacking most ofBS12 can suffice when cell-free systems are used (20). The twoviral proteins form various complexes over the origin region asdetected by electrophoretic mobility shift assays. E1 in theabsence of E2 forms a relatively slowly migrating complex (C2in the terminology of reference 23), using critical contactswithin the dyad element (16, 17, 23, 34). The binding of E1causes the ATP-dependent induction of structural changes tothe viral origin (13). On an origin containing the central regionand BS12, the addition of E2 to E1 can give rise to two distinctcomplexes: a fast-migrating complex containing a lower oligo-meric form of E1 (C1 in the terminology of reference 23; asimilar complex was observed by the Stenlund laboratory [32,33]), and an apparent C2 complex (lacking E2 [23, 32–34]). Byusing an origin that contained both E2-binding sites, it wasshown that an increase in the ratio of E1 to E2 caused anextension of the E1 footprint from the dyad region into theAT-rich region and stimulated distortion of the origin (13).The C1 and C2 complexes have different functional properties;the C2 complex is competent for replication, while C1 is inac-tive (23; see also reference 34).

We have used chemical interference and transient-replica-tion assays to examine the role of E2 and E2-binding sites inviral replication. We find that each flanking E2-binding siteplays distinct and important roles during the initiation of BPV

* Corresponding author. Mailing address: Department of Biochem-istry and Kaplan Comprehensive Cancer Center, New York UniversityMedical Center, 550 First Ave., New York, NY 10016. Phone: (212)263-8453. Fax: (212) 263-8166. E-mail: [email protected].

† Present address: Department of Pathology, University of TexasSouthwestern Medical Center, Dallas, TX 75235.

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DNA replication. E2 binding to BS12 serves to recruit E1 tothe origin. In contrast, the interaction of E2 with BS11 stabi-lizes the binding of E1 across the central origin and BS12regions, yielding a novel complex that we term C3. We proposethat in this final initiation complex, E1 recognizes the origin ina structure similar to that formed by the SV40 T antigen on itscognate origin, using the central palindromic element to pro-duce a complex with twofold dyad symmetry encompassing theAT-rich, dyad, and BS12 regions.

MATERIALS AND METHODS

E1 and E2 proteins. The GST-E1 (4) and E2 (22) proteins were overexpressedin Sf9 insect cells by using recombinant baculovirus. The E2 protein was purifiedas described by Seo et al. (37). E1 protein was purified by a modified procedureof that described by Bonne-Andrea et al. (4). Infected cells were thawed in 5volumes of hypotonic buffer (20 mM Tris-HCl [pH 8.0], 5 mM KCl, 1 mM MgCl2,1 mM dithiothreitol [DTT], 0.1 mM phenylmethylsulfonyl fluoride, proteinaseinhibitors [0.05 mM EGTA, 20 mg of aprotinin per liter, 20 mg of leupeptin perliter, 10 mg of antipain per liter]) and lysed by Dounce homogenization with 20strokes with a type B pestle. The lysate was centrifuged for 15 min at 10,000 rpmin a Beckman SS-34 rotor, and the pelleted nuclei washed with 20 mM Tris-HCl(pH 8.0)–10% (wt/vol) sucrose–1 mM EDTA. Nuclei were resuspended in NRbuffer (20 mM Tris-HCl [pH 8.0], 50 mM MgSO4, 500 mM NaCl, 0.5% [vol/vol]Nonidet P-40, 5 mM DTT, 0.1 mM phenylmethylsulfonyl fluoride, proteinaseinhibitors) and incubated on ice for 30 min. The nuclei were pelleted as above,and the supernatant was mixed with glutathione-Sepharose beads (previouslyequilibrated in NR buffer) and nutated for 1 h at 4°C. The beads were washedwith 50 bead volumes of NR buffer: three washes with NR buffer containing 1 MNaCl, and three washes with XPa cleavage buffer (50 mM Tris-HCl [pH 8.0], 10mM MgSO4, 100 mM NaCl, 10% [vol/vol] glycerol, 1 mM CaCl2, 5 mM DTT).E1 was then cleaved from the beads by incubation with biotinylated XPa (Boehr-inger Mannheim) for 4 h at 4°C. The beads were briefly centrifuged, and thesupernatant containing the liberated E1 was removed. Streptavidin beads(Boehringer Mannheim) were added to remove the biotinylated XPa.

BPV constructs. The BPV DNA containing the mutated origin was generatedby PCR with the Stratagene Quick Change kit. The following oligonucleotideswere used for mutagenesis (mutated bases underlined): DBS12, 59-GTTGTTAACAATAATCACGTTCTCACGTACTTTTCAAGCGGGAAAAAATAGCC(top primer) and 59-GGCTATTTTTTCCCGCTTGAAAAGTACGTGAGAACGTGATTATTGTTAACAAC (bottom primer); and DBS11, 59-GCAGCATTATATTTTAAGCTCGTTCAAACGTACAAGTAAAGACTATGTATTTTTTCC (top primer) and 59 GGAAAAAATACATAGTCTTTACTTGTACGTTTGAACGAGCTTAAAATATAATGCTGC (bottom primer). The top strand isdefined as that containing the run of T’s between BPV positions 7925 and 7930.The template plasmid used to prepare the DBS12 and DBS11 mutants was pXS(in which the BPV XbaI-SmaI fragment from nucleotides [nt] 6132 to 945, wasinserted into a vector derived from pML-1 [21]). The template plasmid for theDBS12DBS11 double mutant was pXSDBS12. Mutations were verified by se-quencing. To generate the full-length viral DNA containing the mutated origins,the pXS plasmids were digested with MluI and MunI and the origin-containingfragment was inserted into the MluI-MunI site of pSS3 (28).

BPV origin-containing DNA fragments. The BPV origin-containing DNAfragments (;120 bp) were generated by PCR amplification of pKSO (45),pXSDBS12, or pXSDBS11 (to prepare the wild-type, DBS12, or DBS11 origin,respectively). One of the two origin-flanking primers was 59-32P labeled with T4polynucleotide kinase (Boehringer Mannheim) to a specific activity of approxi-mately 1 3 106 to 2 3 106 cpm/pmol.

Electrophoretic mobility shift assays. To prepare E1-origin and E1-E2-origincomplexes, reaction mixtures (30 ml) containing 25 mM potassium phosphate(pH 7.5), 0.1 M potassium glutamate, 7 mM MgCl2, 1 mM EDTA, 0.5 mM DTT,4 mM ATP, 10% glycerol, 300 ng of pBluescript KS1 (as a nonspecific compet-itor), 200 fmol of the origin-containing fragment, and E1 alone or E1 and E2 (asindicated) were incubated for 15 min at 37°C. Glutaraldehyde (final concentra-tion, 0.1%) was added, and the reaction mixtures were incubated for an addi-tional 5 min. The resulting complexes were separated by electrophoresis througha native 5% polyacrylamide gel (acrylamide/bisacrylamide ratio, 29:1) and visu-alized by autoradiography.

Interference assays. Chemical modification of the origin-containing DNAfragment was performed as described previously (35). After the separation of theprotein-DNA complexes by a gel retardation assay (see above), gel slices con-taining the complexes were excised, crushed, and soaked overnight in gel elutionbuffer (0.5 M ammonium acetate, 0.1% sodium dodecyl sulfate, 1 mM EDTA).The eluted DNA was precipitated with ethanol and then resuspended in TE (10mM Tris-HCl [pH 8.0], 1 mM EDTA). The DNA was extracted with phenol-chloroform (1:1, vol/vol) and precipitated with ethanol. The modified DNA wasthen cleaved as described previously (35). Assignation of interfering or stimula-tory modifications was determined by careful comparison of the results of mul-tiple independent interference experiments.

Transient-replication assays. Transient-replication assays were performed asdescribed by Mendoza et al. (28). The BPV genomic DNA was released from thevector by digestion with BamHI. Linearized DNA was purified by phenol-chlo-roform (1:1, vol/vol) extraction followed by ethanol precipitation and resuspen-sion in TE. C127 cells growing in the log phase were trypsinized, pelleted,washed, and resuspended in Dulbecco’s modified Eagle’s medium (DMEM)–5mM N,N-bis(2-hydroxyethyl)-2-aminoethanesulfonic acid (BES) (pH 7.2) at aconcentration of 2 3 107 cells/ml. The cell suspension (0.25 ml) was mixed with2 mg of input DNA, 0.5 mg of pSS3 linearized with SalI (containing the wild-typeviral DNA not released from the vector, to serve as an internal standard), and 50mg of sheared salmon sperm DNA and transferred to a electroporation cuvette(0.4-cm gap; Bio-Rad). Electroporation was performed at 270 V and 960 mF ina Bio-Rad gene pulser. The cell material was then transferred to 10 ml ofDMEM containing 10% fetal bovine serum, and 1 ml was added to each 10-cmplate containing 9 ml of DMEM with 10% fetal bovine serum. For each timepoint, low-molecular-weight DNA was extracted from two plates of cells by themethod of Hirt (15). The isolated viral DNA was restricted with MunI to lin-earize the viral genome and DpnI to remove unreplicated DNA. DNA wasdetected by Southern blot analysis with nick-translated pSS3 as a probe. Thereplication activity in vivo was quantitated by excision of the bands and countingin a scintillation counter.

RESULTS

Key origin contacts used by E1 and E2 identified by inter-ference assays. Three distinct complexes can be detected whenthe BPV E1 protein alone or E1 and E2 proteins are incubatedwith DNA fragments containing the wild-type origin (the cen-tral region flanked by BS11 and BS12) in an electrophoreticmobility shift assay (Fig. 1). In the presence of E1 alone, arelatively slow-migrating C2 complex is detected (lane 1) (23,34). When both E1 and E2 are added, a novel complex (that wedefine as C3) that migrates slightly more slowly than C2 isgenerated, in addition to the quickly migrating C1 complex(lane 2 [23; see also references 32 and 33]).

To characterize these different complexes and explore theirfunction in viral DNA replication, we determined the criticalcontacts on DNA used by the E1 and E2 proteins to form eachcomplex. These contacts were determined by an interferenceassay. In this approach, 59-32P-labeled DNA fragments con-taining the BPV origin were modified at the base or phosphate

FIG. 1. Complexes formed on the wild-type BPV origin by E1 and E2. E1alone (100 ng) (lane 1) or E1 (100 ng) plus E2 (60 ng) (lane 2) were incubatedfor 15 min at 37°C with a 32P-labeled DNA fragment containing the wild-typeorigin. The complexes were cross-linked by treatment with glutaraldehyde, sep-arated by electrophoresis through a native 5% polyacrylamide gel, and visualizedby autoradiography. The locations of the C1, C2, and C3 complexes are indi-cated.

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positions to an extent of less than one ‘hit’ per molecule. Themodified substrate was incubated with E1 or with E1 and E2,and the resulting protein-DNA complexes were separated bynondenaturing gel electrophoresis. The free and bound DNAfragments were excised and cleaved at the modified sites in achemical cleavage reaction. The modification pattern was thendetermined by subjecting the cleavage products to denaturinggel electrophoresis. Bands corresponding to modifications thatinterfere with complex formation are underrepresented in thebound fraction and overrepresented in the free fraction, com-pared to the initial substrate.

Differential requirements for E2 binding in the C1 and C3complexes. Key protein contacts with purines and pyrimidineswere examined by the “missing-contact” approach of Brunelleand Schlief (6). Purine contacts were determined by using aDNA fragment modified with formic acid, leading to partial

depurination (Fig. 2) (6, 27). Conversely, protein contacts withpyrimidines were examined by using a DNA fragment treatedwith hydrazine, a reagent that causes destruction of the pyrim-idine base (Fig. 3) (6, 27). Each reagent leaves the sugar-phosphate backbone intact. The effect of each modificationwas examined on a DNA fragment that was 59-32P labeled oneither the top (Fig. 2A and 3A) or bottom (Fig. 2B and 3B)strand.

Comparing the cleavage pattern of the DNA within the C2complex (Fig. 2 and 3, lanes 3) to unbound DNA (Fig. 2 and 3,lanes 2) reveals numerous top- and bottom-strand bases withinthe dyad region whose modification inhibited C2 complex for-mation (i.e., whose intensity was lower in lane 3 than in lane 2).Inhibition of complex formation was caused by modification ofall top-strand bases from nt 7940 to 16 and bottom-strandbases from nt 7943 to 14 (compiled below in Fig. 5C), althoughthe loss of bases encompassing positions 7940 to 10 was ob-

FIG. 2. Interference of the formation of E1- and E1-E2-origin complexes bypartial origin depurination. Duplex DNA fragments, 59-32P labeled on either thetop (A) or bottom (B) strand, were subjected to partial depurination by treat-ment with formic acid. The DNA fragments were then incubated with E1 alone(300 ng) (lanes 2 and 3) or E1 (75 ng) plus E2 (60 ng) (lanes 4 to 6). Eachreaction mixture was then subjected to native gel electrophoresis to isolate theC2 complex (lane 3), or the C1 (lane 5) and C3 (lane 6) complexes from thecorresponding free (unbound) DNA (lanes 2 and 4, respectively). After separa-tion, the free DNA pools, the DNA molecules within each complex, and theinitial origin substrate DNA (lane 1) were isolated and chemically cleaved at thesites of modification. The cleavage products were separated by electrophoresisthrough a denaturing 8% polyacrylamide gel and visualized by autoradiography.

FIG. 3. Interference of the formation of E1- and E1-E2-origin complexes bypartial origin depyrimidation. Duplex DNA fragments, 59-32P labeled on eitherthe top (A) or bottom (B) strand, were subjected to partial depyrimidation bytreatment with hydrazine. The DNA fragments were then incubated with E1alone (300 ng) (lanes 2 and 3) or E1 (75 ng) plus E2 (60 ng) (lanes 4 to 6). Eachreaction mixture was then subjected to native gel electrophoresis to isolate theC2 complex (lane 3) or the C1 (lane 5) and C3 (lane 6) complexes from thecorresponding free (unbound) DNA (lanes 2 and 4, respectively). After separa-tion, the free DNA pools, the DNA molecules within each complex, and theinitial origin substrate DNA (lane 1) were isolated and chemically cleaved at thesites of modification. The cleavage products were separated by electrophoresisthrough a denaturing 8% polyacrylamide gel and visualized by autoradiography.

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served to have a greater effect on complex formation. Outsideof this region, the loss of two top-strand purines within theBS12 element at nt 19 and 22 and three bottom-strand purineswithin the AT-rich element (nt 7925 to 7927) were also seen tohave significant effects on C2 complex formation (Fig. 2, lanes3). Although the effects of modification of these purines weremodest, they were consistently observed in our depurinationstudies. Thus, we conclude that E1 binding in the C2 complexis stabilized primarily by contacts with the dyad element, al-though the interaction of E1 with bases in BS12 and the AT-rich element play a supporting role.

In the presence of E2, removal of most purine (Fig. 2) orpyrimidine (Fig. 3) bases within the dyad region affected bothC1 (lanes 5) and C3 (lanes 6) complex formation (comparewith lanes 4). Bases within BS12 also affected the formation ofboth complexes but in a differential manner, as described be-low. Modification of bases in the dyad region were observed tohave a smaller effect on C1 and C3 formation than on C2formation (compare lanes 5 and 6 with lanes 3). This resultsuggests that E2-mediated stabilization of E1 within the C1and C3 complexes causes E1 binding to be less dependent oncontacts with the dyad element (see also references 32 and 34).

Comparing the C1 and C3 complexes, we observed cleardifferences in the effect of modification of each flanking E2binding site. Formation of the C1 complex was dependentupon contacts within BS12, as indicated by the effect of mod-ification of top-strand bases at nt 15 to 18 and 24 to 27 and ofbottom-strand bases at nt 15 to 18, 21, and 24 to 26. These dataare in agreement with previous results showing the importanceof E2 binding to BS12 for the formation of the C1 and similarfast-migrating complexes (24, 32, 34). Concerning the C3 com-plex, modification of BS12 bases needed for C1 complex for-mation had relatively modest effects on C3 complex formation.In contrast, modification of the E2 BS11 had more severeconsequences. Top-strand bases from nt 7897 to 7910 andbottom-strand bases from nt 7896 to 7900 and nt 7904 to 7909were found to be critical for C3 complex formation. We notedthat when the interference pattern of DNA from the C3 com-plex was compared to that of unbound DNA, the intensity ofcertain bands in the BS11 region was reduced .90% in den-sitometric analysis (data not shown). Because base modifica-tion in this region had no effect on C2 complex formation,these data demonstrate that the recovered C3 complex was notcontaminated by significant levels (.10%) of C2, which couldpotentially complicate the analysis of our interference results.In summary, because we have previously shown that E2 bindsthe BS11 region when incubated with E1 (13), our data indi-cate that E2 binding to BS11 is critical for C3 complex forma-tion.

It was observed that modifications within BS11 stimulate theformation of the C1 complex, seen as an increase in the inten-sity of bands within the BS11 region compared to those for theunbound and substrate DNA (compare lanes 1, 4, and 5 in Fig.2 and 3). The use of DNA substrates modified with dimethylsulfate for methylation interference also indicates that modi-fication of BS11 stimulates C1 complex formation (data notshown). These data argue that the C3 complex can occur in apathway that uses the C1 complex as an intermediate.

Overlap of AT-region phosphate contacts with site of pri-mary origin distortion. The contacts made by E1 and E2 to thesugar-phosphate backbone were examined (Fig. 4). The DNAsubstrate was ethylated at phosphate positions with N-nitroso-N-ethylurea, generating phosphotriesters (38). Similar to thatobserved for the missing-base assays, E1 interaction with thebackbone of the dyad region was critical for C2 complex for-mation (Fig. 4, lanes 3; compiled in Fig. 5C).

Compared to the C2 complex, C1 had a smaller number ofphosphate contacts in the dyad region (Fig. 4, lanes 5 [top-strand phosphates at nt 7940 to 7942] and lanes 3 and 4 [bot-tom-strand phosphates at nt 7947 and 9 to 11]). E1 also ap-peared to utilize top-strand phosphate contacts at nt 12 to 15,located between the dyad element and BS12, because theywere seen to play a role in C2 complex formation. Thesecontacts, when plotted on a helix map, appear on one face ofthe helix (data not shown). C1 complex formation also utilizedsix phosphate contacts within the BS12 (top-strand phosphatesat nt 23 to 25; bottom-strand phosphates at nt 18 to 20). TheC3 complex had a similar pattern of contacts in the dyad regionto that observed for C2 (Fig. 4, lanes 6). The most notablefeature of the C3 complex is the deleterious effect of BS11modification (top-strand phosphates at nt 7896, 7897, 7905,7906, and 7909; bottom-strand phosphates at nt 7898 to 7901,7909, and 7910). Thus, as was seen for the base interferencestudies, the C1 and C3 complexes had differential require-ments for the flanking E2-binding sites.

FIG. 4. Interference of the formation of E1- and E1-E2-origin complexes byethylation of the origin phosphates. Duplex DNA fragments, 59-32P labeled oneither the top (A) or bottom (B) strand, were ethylated on a small fraction ofDNA phosphates. The DNA fragments were then incubated with E1 alone (300ng) (lanes 2 and 3) or E1 (75 ng) plus E2 (60 ng) (lanes 4 to 6). Each reactionmixture was then subjected to native gel electrophoresis to isolate the C2 com-plex (lane 3) or the C1 (lane 5) and C3 (lane 6) complexes from the correspond-ing free (unbound) DNA (lanes 2 and 4, respectively). After separation, the freeDNA pools, the DNA molecules within each complex, and the initial originsubstrate DNA (lane 1) were isolated and chemically cleaved at the sites ofmodification. The cleavage products were separated by electrophoresis througha denaturing 8% polyacrylamide gel and visualized by autoradiography. Phos-phates are numbered according to the base position on the 59 side.



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The interference data for the C1, C3, and C2 complexeswere compiled (Fig. 5A, B, and C, respectively). Included ineach compilation were the results of previous DNase I foot-printing and KMnO4 modification studies, the latter indicatingthe sites of ATP-dependent DNA distortion induced by E1(13). We noted that the patch of phosphate and purine con-tacts in the nt 7930 region for the C3 complex overlapped theprimary site of DNA distortion (centered at nt 7932). Whenthese phosphate and base contacts were mapped on a DNAhelix, they were found to be located on the same face of thehelix (Fig. 5D).

Each E2-binding site stimulates formation of a differentcomplex. Our data suggest that, contrary to published data(40), both BS11 and BS12 play key roles during the initiationof BPV DNA replication. We therefore examined complexformation on BPV origins in which one of the two E2-bindingsites was mutated to prevent E2 binding (Fig. 6). Complexformation was tested by a gel retardation assay in the presenceof low levels (6.25 to 25 ng) of E1 to more closely mimic theexpression levels in infected cells. Binding to the wild-type

origin, as well as to mutant origins lacking BS11 (DBS11) orBS12 (DBS12), was tested. In the absence of E2, E1 formed theC2 complex on the wild-type (lanes 1 to 3) and DBS11 (lanes7 to 9) origins, but only at the highest levels of E1 (25 ng). Incontrast, C2 formed to a lower degree on the DBS12 origin at25 ng of E1 (lanes 4 to 6). This result again indicates thatsequences within the BS12 element stabilize E1 binding to theBPV origin.

The presence of E2 moderately stimulated C3 complex for-mation on the wild-type origin and allowed significant C1 com-plex formation at all levels of E1 (Fig. 6, lanes 10 to 12). A C1complex was not detected on DBS12 (lanes 13 to 15), evenusing 10-fold-higher levels of E1 or E2 (data not shown). Inconstrast, C3 complex formation was observed on DBS12 using

FIG. 5. Compilation of depurination, depyrimidation, and phosphate ethyla-tion interference data for E1 and E2 binding to the BPV origin. Maps ofmodifications that interfere with C1 (A), C3 (B), and C2 (C) complex formationare shown. Phosphates whose modification strongly inhibits complex formationare indicated by solid triangles; weakly interfering phosphates are shown by opentriangles. Bases whose removal (i.e., by depurination and depyrimidation) re-duces complex formation are indicated by solid circles above (bottom strand) orbelow (top strand) the affected base. We also include the results from previousfootprinting analyses (13). The top- and bottom-strand regions protected fromDNase I cleavage by each complex are indicated by solid boxes above and belowthe sequence, respectively. Thymines hyperreactive to KMnO4 (a probe of DNAstructure) are indicated by ovals. The previous data for the C1 and C3 complexeswas taken from complexes formed at low (50 ng) and high (400 ng) E1 levels. (D)Helix map of phosphates and purines in the nt 7930 region whose modificationinterferes with C3 complex formation. Phosphates whose ethylation inhibitscomplex formation are indicated by solid circles, while critical purines are indi-cated by open circles. The top and bottom strands are indicated. The dashed linein the center of the map distinguishes the two helical sides.



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25 ng of E1. On DBS11, C1 complex formation was seen at alevel similar to that on the wild-type origin, while C3 formationwas not observed. In summary, BS12 was required for C1complex formation, BS11 was necessary for formation of theC3 complex, and both BS11 and BS12 were required for max-imal stimulation of C3 complex formation.

Both E2-binding sites are required for wild-type levels ofreplication in vivo. The requirement for the E2-binding sites inviral DNA replication was tested in vivo using a transientreplication assay. To reproduce the levels of E1 and E2 occur-ring during viral infection, mutant origins were constructed inthe context of viral DNA. A plasmid containing the BPV ge-nome (pSS3 [28]; designated the wild type) was mutated toremove one or both of the E2-binding sites from the origin(designated DBS11, DBS12, and DBS11DBS12, lacking BS11,BS12, and both elements, respectively). As a negative control,an origin was used with a 15-bp insertion in the dyad symmetryelement (designated LI 15C), previously shown to inactivatethe viral origin (28). Each of these constructs was linearizedwith BamHI to liberate the viral genome from the vector, andthe linearized DNA was transfected into C127 cells by electro-poration. The test plasmids were cotransfected with wild-typeviral DNA, not released from the vector, which served as aninternal control. After 72 and 96 h, the viral DNA was isolatedand linearized and the unreplicated DNA was destroyed bydigestion with DpnI.

The DBS11DBS12 and LI 15C viral constructs failed to rep-licate (Fig. 7). At each time point, the DBS11 and DBS12origins were defective compared to the wild type, since DBS11and DBS12 replicated to 30 and 20% of the wild-type level,respectively. Similar effects on replication were observed whenthese experiments were repeated by varying the ratio of thewild-type-to-mutant origin template (1:10 and 1:1; data notshown), indicating that the inhibition is not a result of alteredexpression of E1 and E2. The presence of both E2-bindingsites therefore resulted in a synergistic response of viral DNAreplication. The inability of the mutant origins to replicate atwild-type levels indicates that both flanking E2-binding sitesare important for BPV replication under physiological levels ofE1 and E2 expression.

DISCUSSION

Our data indicate that the BPV E2 transactivator enablesviral replication by facilitating discrete steps during formationof the viral initiation complex. These data lead to a model inwhich the initiation complex is formed in a pathway that entailsthe sequential interaction of E2 with two binding sites flankingthe central origin region, each stabilizing a distinct E1-origincomplex. In the first step, E2 binding to the BS12 element isrequired to form a replication-incompetent complex whichcontains a low oligomeric form of E1 bound to the dyad sym-metry element within the central origin region. Additional E1monomers bind and extend this complex outward, forming areplication-active species, in a step in which E2 binding toBS11 becomes paramount and E2 bound to BS12 is displaced.

A previous study concluded that only one of the two E2-binding sites (BS12) flanking the central origin region wasrequired to support efficient viral DNA replication in vivo (40).In contrast, we found that both BS11 and BS12 play distinctroles of similar importance, indicated by the comparable re-duction of transient replication caused by mutation of either ofthese two sites. The likely cause for this difference is that Ustavet al. (40) overexpressed E1 and E2 while our replicationassays expressed E1 and E2 in the context of the viral genome.Since E1 is detected at extremely low levels in infected cells(31, 39), our data suggest that E1 overexpression inadequatelyreproduces replication conditions during infection.

The initiation of viral replication appears to involve theformation of the C1 recognition complex, although this com-plex per se is not active in replication. Stenlund and colleagueshave made compelling arguments that a similar fast-migratingcomplex is critical by virtue of increasing the specificity of E1for origin sequences (32, 33). Indeed we found that at lowlevels of E1, the C1 complex recruits E1 more effectively to theorigin than does the C3 complex. These workers also foundthat a single E2-binding site engineered at different positionswithin the dyad-proximal flank of the origin can support com-plex formation by E1 and E2 (32). Using origins that containBS11 but lack BS12, we were unable to observe a rapidlymigrating complex similar to C1, even at very high levels of E1or E2 (data not shown). In that the E1 recognition element hasgeneral twofold symmetry (see, e.g., reference 16), it seemsunlikely that C1-like complexes can form only by using anE2-binding site located on the BS12 side of the origin. Twonon-mutually exclusive causes appear more reasonable. First,the distance between E2 bound to BS11 and E1 bound to thedyad element may be too great to allow stable complex forma-tion. Second, since the transition of C1 to C3 correlates with anextension of the E1 footprint into the AT-rich region (13),physical interaction between E2 binding to BS11 and the ad-jacent E1 may greatly favor the formation of C3 compared toa complex containing E1 bound only to the dyad region.

The C1 and C3 complexes are distinctly different by variouscriteria. Most obvious is the relative importance of the BS11and BS12 elements for C3 and C1 complex formation, respec-tively, but other notable differences exist. Formation of C3requires a larger number of phosphate contacts, particularly inthe dyad region, supporting the hypothesis that E1 is in ahigher oligomeric state in the C3 (and C2) complex comparedto that in C1 and similar complexes (23, 33). C3 formation alsoutilizes contacts with purines in the AT-rich region (this work)and, from our previous DNA-probing analysis of the C3 com-plex in solution (13), results in protection of the minimal coreorigin sequence from nuclease attack. In contrast, E1 and E2 inthe C1 complex had significant interactions only with the righthalf of the origin including the dyad region. Finally, our prior

FIG. 6. Effect of E2 binding-site mutation on complex formation by E1 andE2. DNA fragments (32P labeled) containing the wild-type (WT) DBS12, orDBS11 mutant origin were incubated with increasing levels of E1 (6.25, 12.5, and25 ng) in the absence or presence of E2 (15 ng; as indicated). Complexes werecross-linked with glutaraldehyde, separated by electrophoresis through a native5% polyacrylamide gel, and visualized by autoradiography. The locations of theC1, C2, and C3 complexes are indicated. Note that the amounts of E1 are lessthan that used in the experiment in Fig. 1 (100 ng), accounting for the reducedlevel of C3 complex on the wild-type origin (lanes 10 to 12) in this experiment.



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footprinting study of the C3 and C1 complexes in solutionindicated that the C3 complex was competent to induce dis-tortion in the origin structure while C1 was lacking in thisability (13).

In contrast to the dissimilarity between C3 and C1, theoverall disposition of E1 in the C2 and C3 complexes appearssimilar. The pattern of base and phosphate interference for theC2 and C3 complexes is alike (Fig. 5), as is the ability of thesecomplexes to distort origin structure (13). The mobility of theC3 complex by a gel retardation assay was only slightly reducedcompared to that of C2, reflective of the additional molecularweight provided by E2 binding to BS11, indicating that the E1oligomeric state in the two complexes is similar. Because of thegreat similarity of the C2 and C3 complexes, our data arguesthat C3, like C2, is replication competent. It is clear that E2binding to BS11 significantly stabilizes C3 at low E1 levels (Fig.6), indicating that the main role of E2 binding to BS11 is tostabilize this replication-active complex. The low expressionlevels of E1 in infected cells lead us to suggest that the normal

replication-active complex in BPV-infected cells is C3 ratherthan C2.

Various data indicate that the C3 complex forms by using afavored pathway with C1 as an intermediate. First, our mobilityshift assays show that formation of the C3 complex is stimu-lated by the presence of BS12, which is critical for C1 forma-tion. Conversely, the BS11 element does not stimulate forma-tion of the C1 complex. Second, modification of BS12phosphates which are important for C1 complex formation alsoinhibits C3 formation, although to a lesser degree. This obser-vation would be expected for a pathway in which C1 precedesC3. Third, modifications within BS11 that prevent C3 complexformation were found to stimulate the formation of C1. Inother words, preventing E2 from binding to BS11 led to anincrease in C1 complex formation. This result would also bepredicted by a model in which C1 converts to C3. These ob-servations strongly support the hypothesis that the C1 complexinitially forms and then is transformed into the C3 complex.

We note, however, that if C1 complex formation were es-

FIG. 7. Mutation of either BS11 or BS12 is deleterious for transient BPV DNA replication in vivo. (A) Schematic showing the origins that were tested for replicationactivity. From top to bottom, these origins are the wild-type origin, LI 15C (containing a 15-bp insertion in the dyad symmetry element which inactivates the origin [28]),DBS12 and DBS11 (lacking the BS12 and BS11 elements, respectively), and DBS12DBS11 (lacking both E2-binding sites). (B) The viral DNA molecules (2 mg) werereleased from the vector and transfected into murine C127 cells by electroporation. As a control, the test plasmids were cotransfected with wild-type viral DNA (0.5mg) not released from the vector. After 72 and 96 h, the viral DNA was isolated by the method of Hirt (15) and treated with MunI to linearize the viral genome andwith DpnI to digest unreplicated DNA. DNA was detected by Southern blot analysis with nick-translated pSS3 as a probe. (C) The replication activity in vivo wasquantitated by excising bands corresponding to linearized viral DNA and counting in a scintillation counter. Replication activity was normalized with respect to thereplication activity of the wild-type origin at 96 h.



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sential for the subsequent formation of the final initiationcomplex, BS12 modification would be expected to have similareffects on C1 and C3 complex formation, a result we did notobserve. We therefore suggest that replication-active com-plexes can form independent of C1, although this pathway isconditional on the stabilization of E1 binding to the origin bythe E2-BS11 complex. Similarly, we can conclude that BS11 isnot essential for replication if E2 binding to BS12 is possible.Thus, formation of an active viral replication complex canoccur by multiple pathways. Because the viral DNA lackingeither BS12 or BS11 replicates to levels 20 to 30% of that in thewild type, the pathway involving both the C1 and C3 complexesappears to be favored.

We found that removal of nearly any base in the dyad ele-ment is similarly deleterious for C1 and C3 complex formation.This is particularly surprising with respect to C1, since thephosphate contacts fall predominantly on one helical face(data not shown), a result observed previously for a C1-likecomplex (33). We find it doubtful that E1 could form impor-tant contacts with nearly every dyad base yet use primarily onehelical face of the DNA for binding. A more probable expla-nation derives from observations that DNA molecules contain-ing an abasic residue have structural perturbations (see, e.g.,reference 9). Loss of bases in the origin would therefore beexpected to alter both the conformation and dynamic proper-ties of the DNA, resulting in destabilization of the E1-origincomplex.

We previously observed that E1 binding to the origin in-duced ATP-dependent structural distortions within the AT-rich region and, to a lesser degree, within the dyad element andBS12 (Fig. 5B) (13). In the present study, we found six phos-phates and three bases in the nt 7930 region whose modifica-tion inhibited C3 complex formation. The base contacts wereimportant for both C2 and C3 formation, showing that E1 useslimited sequence recognition of the AT-rich region. The nt7930 region base and phosphate contacts, which fall on onehelical face of the DNA, overlap the primary site of DNAdistortion (Fig. 5B) (13), suggesting that these contacts areused to induce the structural transitions. These phosphate con-tacts are apparently more critical for C3 formation becausetheir modification did not noticeably disrupt C2 complex for-mation. E2 was shown previously to lower the amount of E1required to distort this region (13). These data suggest that E2bound to BS11 causes E1 to more closely approach the DNAin the nt 7930 region. This effect may be a more generalproperty of E1-E2 interactions, because ethylation of top-strand phosphates at nt 12 to 15 (adjacent to BS12) had agreater effect on C1 formation than on C2 formation. Theability of E2 to increase the interfering properties of phosphateethylation may be due to an E2-induced conformationalchange within E1 that alters the interaction of E1 with DNA.Since E2 is known to bend DNA fragments (14), a relatedeffect may be that E2-mediated DNA bending at each E2-binding site both heightens the deleterious effects of phosphatemodification between the dyad and each E2 binding site andfacilitates the E1-mediated DNA distortion in the nt 7930region.

Our previous DNase I footprinting analysis of the wild-typeviral origin showed that in the presence of E2, an increase inthe concentration of E1 caused an extension of protection intothe region between the dyad element and BS11 (13). Theboundary of the footprint at low E1 concentrations (corre-sponding to the C1 complex [Fig. 5A]) overlapped the regionof primary structural distortion. From this data, we suggestedthat increases in E1 concentration caused an additional lobe ofE1 to bind adjacent to that E1 situated over the dyad element

(13). Since we observed few key phosphate contacts in theAT-rich region other than those in the nt 7930 region, ourresults do not support the suggestion that two independentlobes of E1 bind to the dyad and AT-rich regions. Instead, theyindicate that E1 binding to the AT-rich region is an extensionof the E1 bound over the dyad element. This larger E1 struc-ture would therefore be responsible for the induction of struc-tural changes within the viral origin.

Clues to the mechanism of formation of the C3 complex byE1 and E2 can likely be obtained from comparison with theSV40 T antigen. E1 is homologous to T antigen, particularly intheir C-terminal regions (8), which, for T antigen, contain theATPase and other elements critical for DNA helicase activity(12). The two proteins have similar activities including theability to bind and unwind the origin in the presence of asingle-stranded-DNA-binding protein (10, 36, 44, 46). Eachprotein can bind the origin in both low and high oligomericforms by using a central palindromic structure (23, 25, 26, 32,33). In the presence of ATP, structural changes are induced oneach side of the central palindrome (5, 13). Analysis of theT-antigen–SV40 origin complex by scanning transmission elec-tron microscopy and DNA probing has shown that T antigen isbound as a double hexamer, with twofold symmetry around thecentral palindrome (26, 35).

The similar characteristics of E1 and T antigen, combinedwith the results of previous binding studies, lead to the follow-ing model of E1 binding to the BPV origin (Fig. 8). E1 initiallybinds the origin in a low oligomeric state, using symmetricalcontacts with the dyad symmetry element, and is stabilized byE2 bound to BS12 (13, 23, 32, 33) (see above). The binding ofadditional E1 monomers enlarges the complex, both towardthe proximal edge of the BS11 element and 10 bp beyond thedistal edge of the BS12 element, as observed in the DNase Ifootprint of the C2 complex (Fig. 5C). A key feature of thismodel is that the twofold symmetry around the central palin-drome is maintained. The rightward extension of the complexdisplaces the E2 bound to the BS12 element but is stabilized by(and stabilizes) E2 binding to BS11. Recent data by Berg andStenlund (2) has shown that E2 has two distinct domains thatcan interact with E1, the DNA-binding domain and the trans-activating domain, and our data indicates that each domainmay be differentially used in the C1 complex and the C3 com-

FIG. 8. Model of E1 and E2 binding to the BPV origin to form a replicationinitiation complex. See the text for details. The light zigzag lines indicate regionsof DNA distortion induced by E1 in the AT-rich and BS12 regions.



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plex. In the presence of ATP, this higher oligomeric C3 com-plex induces structural transitions, primarily within the nt 7930region, but also to significant levels within BS12. Similar to Tantigen, the final complex would have two helicase entities, onebound to the left half of the dyad element and the AT-richregion and the other bound to the right half of the dyadelement and the BS12 region. Since hRPA can denature DNAunder conditions supporting DNA replication (18), the inter-action of hRPA with the distorted DNA region(s) leads toorigin denaturation and unwinding of the DNA by the DNAhelicase activity of E1.

Although transcriptional transactivators have been found tomodulate DNA replication by a diversity of mechanisms, ourdata indicates that E2 is perhaps unique in its ability to usemultiple binding sites to sequentially activate viral replication.Other papillomaviruses have multiple E2-binding sites in ori-gin-proximal regions, and it would not be surprising to find thedifferential usage by E2 of these sites during the initiation ofreplication.

ACKNOWLEDGMENTS

We thank Philippe Clertant for his generous gift of the baculovirusGST-E1 construct and Mike Botchan for his kind gift of the pSS3 andBPV LI 15C plasmids. We thank Cristina Iftode, Natalia Smelkova,Jennifer Garner, and Yaron Daniely for constructive comments duringthe course of this project and for critical readings of the manuscript.

This research was supported by NIH grant CA62198, Kaplan CancerCenter Developmental Funding, and a Kaplan Cancer Center SupportCore Grant (NCI P30CA16087).

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Documents

Distinct Roles of Two Binding Sites for the Bovine Papillomavirus