Department of Molecular Genetics and Biochemistry, University of Pittsburgh School of Medicine, Pittsburgh, PA 15261, USA
Correspondence
Saleem Khan
khan{at}pitt.edu
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ABSTRACT |
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Introduction |
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The E1 proteins of PVs have ATPase, helicase, origin binding and unwinding activities and it is assumed that they play a similar role during replication of different PVs (Jenkins et al., 1996; Mansky et al., 1997
; Muller et al., 1997
; Rocque et al., 2000
; Santucci et al., 1995
; Sedman & Stenlund, 1998
; Seo et al., 1993a
, b
; Stenlund, 1996
; Thorner et al., 1993
; Wilson & Ludes-Myers, 1991
; Yang et al., 1993
). Biochemical studies with the full-length E1 proteins of BPV-1 and HPVs 6b, 11, 16, 31b and 33 have shown that either they do not bind to the ori, or they bind very weakly and with low specificity (Bream et al., 1993
; Chen & Stenlund, 2001
; Dixon et al., 2000
; Leng et al., 1997
; Liu et al., 1995
, 1998
; Masterson et al., 1998
; Muller & Sapp, 1996
; Muller et al., 1997
; Rocque et al., 2000
; Sanders & Stenlund, 2000
). However, in the presence of the viral E2 protein, ori binding by E1 becomes more efficient and specific (Berg & Stenlund, 1997
; Mohr et al., 1990
; Sarafi & McBride, 1995
; Woytek et al., 2001
). Thus, the detection of full-length E1ori complexes by electrophoretic mobility-shift assays (EMSAs) requires the use of cross-linking agents, which is consistent with the weak DNA binding activity of E1 (Chen & Stenlund, 1998
, 2001
; Gonzalez et al., 2000
; Liu et al., 1995
, 1998
). On the other hand, truncated derivatives of the BPV-1 E1 protein containing its DNA binding domain bind to the ori stably, and this binding can be detected by EMSA in the absence of any cross-linking agents (Berg & Stenlund, 1997
; Chen & Stenlund, 1998
, 2001
; Gonzalez et al., 2000
; Leng et al., 1997
; Liu et al., 1995
, 1998
). The binding of the full-length E1 proteins of BPV-1 and some HPVs to specific sequences within the ori has also been demonstrated by footprint analysis (Chen & Stenlund, 1998
, 2001
; Frattini & Laimins, 1994a
, b
; Gilette & Boroweic, 1998
; Holt et al., 1994
; Muller et al., 1997
; Sanders & Stenlund, 2000
). The following model has emerged from a number of recent studies. The E2 protein targets E1 to the ori, resulting in the formation of an E1E2ori complex. In the case of BPV-1, this complex is subsequently converted to a stable E1ori complex, which lacks the E2 protein, has a larger size and contains a higher multimeric form of E1 (Berg & Stenlund, 1997
; Gilette & Boroweic, 1998
; Lusky et al., 1994
; Sedman & Stenlund, 1995
; Stenlund, 1996
). This multimeric E1ori complex is postulated to be the substrate for the initiation of BPV-1 replication. However, in the case of HPV-11, the E1E2ori complex does not appear to be a precursor for an E1ori complex and it is possible that the E1E2ori complex may be competent for initiation (Chao et al., 1999
).
Previous studies in our laboratory have shown that the HPV-1 E1 protein is sufficient for the transient replication of ori plasmids, suggesting that a stable E1ori complex is formed that is competent for the initiation of replication. To find biochemical support for this model, we have purified the HPV-1 E1 and E2 proteins as fusion proteins, with the FLAG peptide as an epitope tag. EMSAs showed that the E1 protein bound to the ori region containing the E1BS in the absence of any cross-linking agents. In vitro-translated, native E1 protein also bound to the ori. In the presence of both the E1 and E2 proteins, an E1E2ori complex was also observed. The E1 protein was shown to have a DNA-independent ATPase activity with a Km value that was comparable with that of the SV40 large T antigen. Our results suggest that the E1 protein of HPV-1 (and possibly other related HPVs) may be capable of supporting in vivo replication of ori plasmids in the absence of E2 due to a stable interaction with the HPV-1 origin.
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Methods |
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Expression and purification of the E1 and E2 proteins.
Bacmid DNAs were transfected into Sf-21 cells using Cellfectin (Life Technologies) and expression of the FLAGE1 and FLAGE2 proteins was confirmed by Western blot analysis, as described below. To isolate the recombinant baculovirus expressing the E1 and E2 proteins, supernatant from cells grown for 48 h after transfection with bacmid DNAs was collected. The titre of the viral stock was determined by a plaque assay, as recommended by the supplier. Various conditions were tested for optimal expression of the E1 and E2 proteins by isolating protein lysates from infected cells, followed by SDS-PAGE and Western blotting using the anti-FLAG M2 monoclonal antibodies. The HPV-1 FLAGE1 and FLAGE2 proteins were optimally expressed at 24 and 72 h, respectively. The optimum m.o.i. was 1 for E1 and 2 for the E2 protein. Large-scale cultures of insect cells expressing the HPV-1 E1 and E2 proteins were obtained from the National Cell Culture Center (Minneapolis, MN, USA). For purification of the FLAGE1 protein, the cell pellet was thawed and suspended in buffer A [50 mM Tris/HCl, pH 8·0, 1 mM EDTA, 1 mM DTT, 0·15 M NaCl, 10 % glycerol and Complete (Boehringer Mannhein) protease inhibitor (1 tablet in 50 ml extraction buffer); the tablet contains EDTA, aprotinin, leupeptin and Pefabloc SC]. The cells were lysed by sonication at a continuous cycle for about 8 min with 30 s intervals after each 2 min. The supernatant containing the E1 protein was collected by centrifugation (at least 3 h at 37 000 r.p.m. in an SW41 rotor), and the protein was precipitated by the addition of ammonium sulfate to a final concentration of 40 %. At this cut-off range, most of the E1 protein was precipitated. The precipitated proteins were resuspended in buffer A and loaded directly on to a DEAESepharose column (pre-equilibrated with buffer A) to remove the DNA associated with the proteins. The column was washed with two bed volumes of buffer A. The wash fractions containing unbound proteins free of DNA were pooled. The A280/A260 ratio was close to 1 for the E1 protein preparations. Western dot blots were performed to identify the fractions containing the E1 protein. Fractions containing E1 were pooled and dialysed for at least 2 h at 4 °C with one change of buffer. The pooled fraction was centrifuged to remove any debris and loaded on to 23 ml of an affinity column containing anti-FLAG M2 monoclonal antibody coupled to Sepharose, according to the manufacturer's protocol (Sigma). The effluent was passed through the column at least three times to enhance retention of the E1 protein on the column. The column was then washed several times with buffer B (as buffer A but containing 1 M NaCl in place of 0·15 M) to remove proteins that were bound non-specifically to the resin. The FLAGE1 protein was then eluted with buffer A containing 10 mM FLAG peptide, and 0·5 ml fractions were collected. Aliquots of these fractions were analysed by SDS-PAGE to identify the fractions containing the E1 protein. Western blot analysis was used to confirm the presence of the E1 protein in these fractions. The fractions containing the purified E1 protein were pooled, concentrated and dialysed in buffer A without the protease inhibitor. The FLAGE1 protein was stored in small aliquots at -80 °C. The procedure for the purification of the FLAGE2 protein has been described elsewhere (Van Horn et al., 2001). The MBPE2 fusion gene was constructed by PCR amplification of the E2 cDNA using primers with EcoRI ends followed by ligation into the pMALC2 expression vector (New England Biolabs). The MBPE2 fusion protein was expressed in E. coli by IPTG induction,as suggested by the manufacturer. Induced cells from a 500 ml culture were resuspended in 15 ml of buffer C (50 mM Tris/HCl, pH 8·0, 1 mM EDTA, 1 mM DTT, 100 mM NaCl and 10 % glycerol) and lysed by treatment with lysozyme (1 mg ml-1) for 30 min at 4 °C. The cell lysate was subjected to two freeze/thaw cycles and sonication at a continuous cycle twice for 90 s bursts. The cell debris was removed by centrifugation for 20 min at 15 000 r.p.m. in an SS34 rotor. The supernatant was collected and passed through an affinity column containing amyloseSepharose resin. The column was washed with two column volumes of buffer C and the MBPE2 protein was eluted with buffer C containing 15 mM maltose. The fractions were checked for the presence of the MBPE2 protein by SDS-PAGE on a 10 % gel followed by staining with Coomassie brilliant blue. The amount of E1 and E2 protein in purified fractions was estimated using the Bio-Rad protein assay kit, with BSA as the standard.
Determination of the ATPase activity of the E1 protein.
ATP hydrolysis was measured in a 50 µl reaction mixture containing 50 mM Tris/HCl, pH 8·0, 10 mM MgCl2, 1 mM DTT and 0·0310 µM [-32P]ATP (3000 Ci mmol-1). After incubation at 37 °C for 60 min, the reaction was stopped by addition of EDTA to a final concentration of 100 mM. An aliquot of the reaction was spotted on to a polyethyleneiminecellulose thin-layer plate, which was developed with 0·5 M potassium phosphate buffer (pH 3·5). After TLC, the plates were dried and subjected to autoradiography for 30 min at 25 °C. The products were quantified using a Phosphorimager (Molecular Dynamics).
In vitro synthesis of the E1 protein.
The pSG5 expression vector containing the HPV-1 E1 gene downstream of the T7 RNA polymerase promoter (Gopalakrishnan & Khan, 1994) was used for the in vitro synthesis of native, unfused E1 protein using the TNT Quick Coupled Transcription/Translation System (Promega). Reactions were carried out according to the manufacturer's protocol with some modifications. Briefly, 20 µl rabbit reticulocyte lysate (TNT Master Mix) was mixed with 1 µg DNA and 1 µl [35S]methionine (1175 Ci mmol-1; ICN) in a total volume of 25 µl. Reactions were incubated at 30 °C for 90120 min. The samples were run on 10 % polyacrylamide gels (29·2 : 0·8, acrylamide : bis-acrylamide). Before drying, gels were soaked in 30 % methanol and 10 % acetic acid for 15 min. The gels were soaked for an additional 15 min in the Enhancer solution (NEN). The gels were rinsed three times with water, placed on a piece of Whatman paper, dried for 2 h at 90 °C and subjected to autoradiography. Non-radioactive E1 protein was synthesized as above except that the labelled methionine was replaced with 1 µl 1 mM cold methionine supplied with the TNT kit.
DNA binding experiments.
The 171 bp optimal HPV-1 ori (ori171) containing one putative E1BS, one high-affinity E2BS and one low-affinity E2BS was isolated by digesting the pori171 plasmid (Gopalakrishnan & Khan, 1994) with HindIII and BamHI. A 60 bp oligonucleotide containing the E1BS and the surrounding AT-rich region (ori60) was chemically synthesized (Life Technologies). A 244 bp origin mutant fragment was isolated by digesting the pori312 plasmid with HindIII and HpaI (Gopalakrishnan & Khan, 1994
). This fragment contains HPV-1 nt 75937815/1-3 and disrupts the putative E1BS at the HpaI site. An unrelated 166 bp fragment containing an AT-rich region that includes the origin of replication of plasmid pT181 (Koepsel et al., 1986
) was used as a negative control. EMSAs were performed by labelling the ori DNAs using 50 µCi [
-32P]ATP (3000 Ci mmol-1) and T4 polynucleotide kinase (Sambrook et al., 1989
). DNA binding reactions contained 25 mM Tris/HCl, pH 8·0, 7 mM MgCl2, 1 mM DTT, 5 % glycerol (v/v), 45 or 200 ng poly(dI·dC), labelled DNA and FLAGE1 and/or FLAGE2 (or MBPE2) protein. Some EMSA reactions also included NP-40 at a final concentration of 0·1 % (v/v). The reactions were incubated at room temperature for 20 min and the DNAprotein complexes analysed by electrophoresis on 5·5 or 6 % native polyacrylamide or 1·2 % agarose gels (Sambrook et al., 1989
). For EMSA with native in vitro-synthesized E1, rabbit reticulocyte lysates containing in vitro-translated, non-radioactive E1 protein were incubated with the labelled DNA probes as above and subjected to electrophoresis on 1·2 % agarose gels.
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Results |
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Discussion |
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Since the HPV-1 E1 protein alone is sufficient to support robust replication of ori plasmids (Gopalakrishnan & Khan, 1994; Gopalakrishnan et al., 1995
, 1999
; Van Horn et al., 2001
), we tested whether it was capable of stable interaction with the HPV-1 ori by EMSA. E1 was found to form a single DNAprotein complex in the presence of DNA fragments containing the putative E1BS (Fig. 3A, B
). Furthermore, in vitro-translated native HPV-1 E1 protein also bound to the HPV-1 ori (Fig. 4B
). In a similar experiment, in vitro-translated HPV-18 E1 protein did not bind to the HPV-18 origin (L. Sheahan and S. A. Khan, unpublished data). While an ori mutant fragment that is deleted for half of the E1BS also bound to the E1 protein, a non-specific DNA fragment did not stably bind to this protein (Fig. 3C, D
). These results suggest that sequences in addition to the putative E1BS may be required for E1 binding. Since in vivo replication in the presence of E1 alone is specific to the HPV-1 ori (Gopalakrishnan & Khan, 1994
), HPV-1 E1 must be able to recognize specifically the HPV-1 ori during initiation. The FLAGE2 protein also generated a single DNAprotein complex in the presence of the optimal ori (Fig. 5
). In the presence of both the E1 and E2 proteins, a novel, slower-migrating complex (E1-E2-CX) was also observed, presumably corresponding to an E1E2ori complex (Fig. 5A
). In the case of the BPV-1 E1 protein, it has been shown that the E2 protein stimulates binding of E1 to the ori, initially generating an E1E2ori complex, which is subsequently converted to an E1ori complex that contains a larger multimeric form of E1 and is competent for initiation (Berg & Stenlund, 1997
; Chen & Stenlund, 1998
, 2001
; Gilette & Boroweic, 1998
; Lusky et al., 1994
). We did not observe such an E1ori complex in the presence of E2, similar to the results obtained with the HPV-11 E1 protein (Chao et al., 1999
; Rocque et al., 2000
). It is possible that both the E1ori and the E1E2ori complexes that assemble in the presence of both the E1 and E2 proteins are capable of supporting the initiation of HPV-1 replication.
In EMSA, the E2 protein did not significantly stimulate the ori-binding activity of E1 or the multimeric state of E1 in E1ori complexes (Fig. 5 and data not shown). In the case of BPV-1 and HPV types 11, 16 and 31b, the E2 protein has a strong stimulatory effect on the ori binding activity of E1, and E2 also enhances the specificity of the E1ori interaction (Berg & Stenlund, 1997
; Chen & Stenlund, 1998
; Frattini & Laimins, 1994a
, b
; Gilette et al., 1994
; Masterson et al., 1998
; Seo et al., 1993a
, b
). Since HPV-1 E1 can stably interact with the ori in the absence of E2, it is possible that E2 may have a more limited role in targeting E1 to the ori in this case. However, it is known that the E2 protein stimulates E1-dependent replication of HPV-1 ori plasmids in vivo (Gopalakrishnan & Khan, 1994
; Gopalakrishnan et al., 1995
). Thus, it is possible that the E2 protein may have a more important role in the establishment of a nucleosome-free ori region, recruitment of host factors to the ori, etc. during HPV-1 replication.
Our observations suggest that the E1 protein of HPV-1 can stably interact with the ori. Recent studies have shown that the BPV-1 E1 protein recognizes the hexanucleotide sequence AACAAT, or its variants, which are present in multiple copies in the ori of BPV-1 as well as several other HPVs (Chen & Stenlund, 2001; Holt et al., 1994
). Interestingly, the HPV-1 ori has six such putative E1 binding sequences, more than in any other PV (Chen & Stenlund, 2001
). Thus, it is possible that the E1 protein of HPV-1 stably interacts with the ori due to the presence of a larger number of high-affinity E1 binding sequences. This prediction is consistent with the lack of an absolute requirement for the E2 protein or E2BSs in HPV-1 replication (Gopalakrishnan & Khan, 1994
). Since HPVs that infect cutaneous tissues replicate to much higher levels and produce higher levels of virions than the mucosotropic HPVs, it is possible that an efficient E1ori interaction in the case of HPV-1 (and similar HPV types) may play a critical role in the high-level replication of the virus in the productive phase during terminal differentiation of epithelial cells.
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ACKNOWLEDGEMENTS |
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Received 18 April 2002;
accepted 25 September 2002.