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Journal of Virology, November 2002, p. 11350-11358, Vol. 76, No. 22
0022-538X/02/$04.00+0     DOI: 10.1128/JVI.76.22.11350-11358.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.

Regulation of Bovine Papillomavirus Replicative Helicase E1 by the Ubiquitin-Proteasome Pathway

Marie-Helene Malcles, Nathalie Cueille,{dagger} Francisca Mechali, Olivier Coux, and Catherine Bonne-Andrea*

Centre de Recherches de Biochimie Macromoléculaire, CNRS, IFR 24, 34 293 Montpellier Cedex 5, France

Received 1 May 2002/ Accepted 13 August 2002


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ABSTRACT
 
Papillomaviruses maintain their genomes in a relatively constant copy number as stable extrachromosomal plasmids in the nuclei of dividing host cells. The viral initiator of replication, E1, is not detected in papillomavirus-infected cells. Here, we present evidence that E1 encoded by bovine papillomavirus type 1 is an unstable protein that is degraded through the ubiquitin-proteasome pathway. In a cell-free system derived from Xenopus egg extracts, E1 degradation is regulated by both cyclin E/Cdk2 binding and E1 replication activity. Free E1 is readily ubiquitinated and degraded by the proteasome, while it becomes resistant to this degradation pathway when bound to cyclin E/Cdk2 complexes before the start of DNA synthesis. This stabilization is reversed in a process involving E1-dependent replication activity. In transiently transfected cells, E1 is also polyubiquitinated and accumulates when proteasome activity is inhibited. Thus, the establishment and maintenance of a stable number of papillomavirus genomes in latently infected cells are in part a function of regulated ubiquitin-mediated degradation of E1.


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INTRODUCTION
 
Papillomaviruses are small DNA tumor viruses which have the capacity to establish a persistent infection in mammalian epithelial cells. The complete papillomavirus replication cycle is tightly linked to the differentiation state of the epithelial cells. According to a current model, a persistent infection is established when the virus enters a proliferating cell of the basal epithelial cell layer. Following the establishment stage, in which the circular viral genome has to be amplified, viral persistence is set up through the maintenance of a constant low copy number of extrachromosomal viral genomes in the nuclei of dividing host cells. The productive stage of infection begins only as the basal infected cells enter the terminal differentiation pathway and migrate to the upper levels of the stratified epithelium. Papillomavirus genomes are then amplified to allow the formation of new viral particles (reviewed in references 9 and 27). The genomes of papillomavirus can also be stably maintained as high-copy-number extrachromosomal elements in certain cell lines, in which the viral genomes replicate in synchrony with cellular DNA (17, 28). How the switch occurs from stable replication in the undifferentiated basal cells to unregulated genome amplification in differentiated cells is not known, but it is likely to involve a complex interplay between viral proteins and host cell factors.

These viruses exhibit a complete dependence on host cell functions for their replication (37, 39, 58), except for the virally encoded helicase, E1, capable of assembling an initiation complex on the viral origin of replication (56, 57). E1 has origin-specific DNA-binding activity (4, 22) and ATP-dependent DNA helicase activity (50, 51) and associates with the DNA polymerase {alpha}-primase (7, 11, 42) as well as with a key regulator of S-phase entry, cyclin E-Cdk2 (13, 36). While E1 is the only viral protein required to orchestrate the replication of papillomavirus DNA plasmids in different cell-free systems (6, 8, 13), a second viral protein is needed in vivo, the regulator of transcription E2 (10, 56). In addition to its auxiliary role in the formation of the replication initiation complex with E1 at the viral origin, E2 mediates the attachment of viral genomes to cellular chromosomes to ensure efficient segregation to daughter nuclei in mitosis (24, 30, 46, 52).

While little is known about how papillomavirus DNA replication is regulated, the finding that E1 is as efficient as the simian virus 40 T antigen lytic initiator in vitro (6) suggests that the papillomavirus replication protein E1 is a critical component in maintenance replication. The study of E1 in papillomavirus-infected cells has, however, been hindered by the fact that this protein is present at exceedingly low levels.

We have recently established a cell-free system derived from Xenopus egg extracts that catalyzes the E1-dependent replication of plasmid DNA containing the bovine papillomavirus type 1 (BPV1) origin of replication (13). In this system, translation of synthetic E1 mRNA leads to the accumulation of very low levels of radiolabeled E1. Using this novel approach, we determined that the viral initiator interacts functionally with cyclin E-Cdk2, as no E1-dependent replication was observed in cyclin E-Cdk2-depleted extracts, while it was restored by addition of E1-cyclin E-Cdk2 complexes.

In this study, we investigated the regulation of the viral initiator of DNA replication. We show that the lack of DNA replication in cyclin E-Cdk2-depleted Xenopus interphase egg extracts is due to the degradation of the viral initiator by the ubiquitin-proteasome pathway. We show that E1 is stabilized when associated with cyclin E-Cdk2 and that this protection is lost after replication. Polyubiquitinated E1 species can also be detected in extracts of human 293 cells synchronized in S phase, in correlation with replication function. Furthermore, although detection of E1 ubiquitination and degradation is more difficult when E1 is expressed alone than when E1 is expressed with the coactivator E2 and BPV origin-expressing plasmids, we show that the E1 steady-state level is also regulated by the ubiquitin-proteasome pathway in transfected primate cells. Together, these data suggest that the regulated proteolysis of E1 is likely a mechanism involved in the control of viral genome replication within latently infected cells.


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MATERIALS AND METHODS
 
Proteins and DNA plasmids. BPV1 E1 protein, Xenopus cyclin E, and Xenopus Cdk2 were prepared by translation of mRNA-encoding constructs in RNase-treated translation extracts from Xenopus eggs in the presence of [35S]methionine as described previously (13). The E1 mutant in which the three Cdk phosphorylation sites were changed to alanines (E1-AAA) was obtained by using a transformer site-directed mutagenesis kit from Clontech and then confirmed by dideoxy sequencing. BPV1 E1 protein was purified from recombinant baculovirus-infected Sf9 cells (7). 35S-labeled BPV1 E2 was prepared by coupled transcription-translation reactions in a rabbit reticulocyte lysate (Promega).

Glutathione S-transferase (GST)-tagged human Cdk2, GST-tagged human Cdk2K33R (47), GST-ubiquitin (49), 6His-tagged ubiquitin, and 6His-tagged K48R ubiquitin (1, 53) were expressed in Escherichia coli BL21 and purified by affinity chromatography on glutathione-Sepharose or nickel-agarose. The GST fusion protein of ubiquitin contains a protein kinase A site between the GST and the ubiquitin portion of the fusion protein to allow specific labeling (49). Methylated ubiquitin was prepared by the method of Hershko and Heller (20). The green fluorescent protein (GFP)-E1 construct was generated by inserting wild-type E1 into the BamHI site of pEGFP-C2 (Clontech, Palo Alto, Calif.). The BPV1 E1 expression vectors pCGEag, pCGE2, and pSKori+, containing a 160-bp origin-bearing BPV1 DNA fragment (nucleotides 7855 to 81) in the pBluescript SK+ vector, have been described previously (8, 57). pCDNA3-6HisUb and pCDNA3-HAUb were provided by M. Piechaczyk, Institut de Genetique Moleculaire de Montpellier, Montpellier, France.

Preparation of soluble extracts and in vitro DNA replication assays. Crude Xenopus interphase egg extracts for translation assays and membrane-free interphase egg cytosol used for E1-dependent replication assays were prepared essentially as described previously (13). Replication assays were carried out by first mixing 18 µl of membrane-free egg cytosol supplemented with an ATP-regenerating system and with 2 µl of radiolabeled E1 protein for 30 min at 25°C before the addition of 300 ng of either pSKori+ or pSKori-. Reaction mixtures were incubated at 25°C, and samples were taken for both protein and DNA analysis. Samples to be analyzed for DNA synthesis were supplemented with 2 µCi of [32P]dCTP.

The 293 cell extracts for cell-free replication were prepared as described previously (32). Cells were synchronized in early S phase by a double block in culture medium containing 2.5 mM thymidine. The conditions for cell-free replication in the presence of 2 µCi of [32P]dCTP have been described (7). Reaction mixtures (50 µl) containing 300 ng of pSKori DNA, 100 ng of E1 protein, and 200 µg of 293 cell cytoplasmic extract were incubated at 37°C for 60 min. Aphidicolin was added at a final concentration of 40 µg/ml in both cell-free systems. Samples for DNA analysis were deproteinized with proteinase K (500 µg/ml) and 0.5% sodium dodecyl sulfate (SDS) for 60 min at 37°C, followed by two phenol-chloroform extractions, and analyzed on agarose gels.

In vitro E1 degradation and ubiquitination assays. For the degradation assays, Xenopus egg extracts were depleted with anti-Cdk2 antibody as previously described and supplemented with an ATP-regenerating system (13). NLVS (4-hydroxy-5-iodo-3-nitrophenylacetyl-Leu-Leu-Leu-vinylsulfone) (Calbiochem), an irreversible inhibitor of the three proteolytic activities of the proteasome 26S (5), was added to extracts before addition of 1 to 2 µl of radiolabeled E1 protein. Ubiquitination assays in crude Xenopus egg extracts were carried out by supplementing egg extracts with ubiquitin aldehyde (4 µM; Sigma) (21) and ATP{gamma}S [adenosine 5'-O-(3-thiotriphosphate); 4 mM] in the presence or absence of either hexahistidine-ubiquitin (6His-ubiquitin), methylated ubiquitin, or K48R ubiquitin at a final concentration of 1.25 mg/ml, before addition of 2 µl of radiolabeled E1 protein.

Reaction mixtures were incubated at 28°C for 30 to 60 min, and 3-µl aliquots were analyzed by SDS-polyacrylamide gel electrophoresis (PAGE) followed by autoradiography. The conjugation of ubiquitin to E1 for replication in 293 cell extracts was carried out by supplementing reaction mixtures with 1 µg of 32P-labeled GST-ubiquitin and ubiquitin aldehyde (4 µM). After 30 min at 37°C, E1 was immunoprecipitated with polyclonal anti-C-terminal E1 antibody. Immunoprecipitates were subjected to polyacrylamide gel electrophoresis, and radioactively labeled bands were visualized by autoradiography.

Cell culture and DNA transfection. COS-7 and 293 cells were grown in Dulbecco's modified Eagle's medium with 10% fetal bovine serum. To inhibit protein synthesis, cells were cultured in the presence of 10 µg of cycloheximide per ml for 4 h. After this, the cells started to die. Inhibition of the proteasome was performed by culturing cells for up to 18 h in the presence of 5 µM MG132 (Calbiochem). A stock solution of MG132 was made in dimethyl sulfoxide and used at a 1:2,000 dilution.

Cells were 80% confluent when trypsinized for electroporation (56). Each electroporation mixture contained 10 µg of pEGFP-E1, 5 µg of pCGE2, 3 µg of supercoiled pSKori, and 50 µg of carrier DNA. Electroporation was performed with 6.6 million cells in 250 µl of growth medium containing 10% fetal bovine serum and 5 mM BES buffer (N,N-bis[2-hydroxyethyl]-2-aminoethanesulfonic acid, pH 7.2) at 960 µF and 170 V for 293 cells and 180 V for COS-7 cells, with a Bio-Rad Gene Pulser with a capacitance extender.

Western blotting and antibodies. Transfected cells were harvested from a 35-mm plate in 100 µl of RIPA buffer (0.5% NP-40, 0.5% Tween 20, 0.5% deoxycholic acid, 0.15 M NaCl, 10 mM KCl, 20 mM Tris-HCl [pH 7.5], and 1 mM EDTA) and boiled with 1% SDS, 48 h after electroporation. Then 50 µg of protein was loaded onto SDS-PAGE gels. Western blotting was performed by standard procedures with 5,000-fold dilutions of anti-GFP (Torrey Pines Biolabs) or monoclonal antihemagglutinin (HA) antibody (generous gift of A. Debant, Centre de Recherches de Biochimie Macromoleculaire, Montpellier, France).

Transient-replication assays in transfected cells. Transfected 293 cells used in transient-replication assays were plated onto 100-mm-diameter dishes, and samples were taken at 24-h intervals on days 2 to 4. DNA plasmids were isolated as described before (31), and DNA samples were digested with a mix of SmaI and DpnI. The samples were run on 1% agarose gels and blotted to nitrocellulose under standard conditions for Southern blot analysis. High-specific-activity probes were generated by random priming of pSKori+ DNA with a kit from Amersham (Rediprime), and the blots were visualized and quantitated with a PhosphorImager.

In vivo ubiquitination assays. Asynchronously growing COS-7 cells were transfected with 10 µg of pEGFP-E1, 5 µg of pCDNA3-6HisUb, or 5 µg of pCDNA3-HAUb and with or without 5 µg of pCGE2 and 3 µg of pSKori. Control transfections used the same amount of empty vectors. Cells were cultured in the presence of 5 µM MG132 overnight and lysed after 48 h. For the ubiquitination assay with HA-ubiquitin, cells were trypsinized, pelleted, and lysed in 50 µl of RIPA buffer containing MG132 and ubiquitin aldehyde to block isopeptidase activity. For the ubiquitination assay with 6His-ubiquitin, cells from three 100-mm-diameter dishes were lysed in 5 ml of 6 M guanidine-HCl-0.1 M Na2HPO4/NaH2PO4-0.01 M Tris-HCl (pH 8.0). 6His-ubiquitin conjugates in the extracts were purified on nickel-agarose.


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RESULTS
 
E1 is protected from degradation within cyclin E/Cdk2 complexes. Coincubation of E1, cyclin E, and Cdk2 translated separately in egg extracts depleted of Cdk2 led to the generation of a tight complex that was coimmunoprecipitated by either anti-Cdk2 or anti-E1 antibodies, as previously described (13). The relative level of E1 was similar in both immunoprecipitates, suggesting that radiolabeled E1 molecules are detected only when associated with cyclin E-Cdk2 (Fig. 1A). To determine whether cyclin E/Cdk2 protects E1 from degradation, E1 was first mixed with increasing amounts of Xenopus cyclin E and Cdk2 and then added to Xenopus Cdk2-depleted interphase egg extract (5). While E1 alone was mostly degraded (Fig. 1B, lane 2), increasing E1 levels were detectable upon addition of cyclin E/Cdk2 (19 and 28% protection in lanes 3 and 4, respectively).



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  		FIG. 1. E1 is unstable in Xenopus egg extract in the absence of cyclin E/Cdk2. (A) BPV1 E1 (E1), Xenopus cyclin E (CycE), and Xenopus Cdk2 (Cdk2), translated separately in Cdk2-depleted Xenopus egg extracts in the presence of [35S]methionine, were mixed together, and immunoprecipitations (IP) were performed with anti-Cdk2 antibodies (K2) or with an anti-C-terminal E1 polyclonal antibody (E1). Immunoprecipitates were analyzed by SDS-PAGE and autoradiographed. (B) 35S-labeled E1 translated in Xenopus Cdk2-depleted extracts was incubated in the absence (lane 2) or presence of increasing amounts of labeled Xenopus cyclin E and Xenopus Cdk2 (lanes 3 and 4) obtained by translation in Xenopus Cdk2-depleted extracts. After 30 min of incubation at 25°C, equal amounts of each mixture were added to fresh Cdk2-depleted egg cytosol and incubated for an additional 30 min. E1 was recovered by immunoprecipitation with an anti-C-terminal E1 polyclonal antibody and analyzed by SDS-PAGE and autoradiography. Input E1, lane 1. (C) Purified complexes reconstituted with 35S-labeled E1, 35S-labeled Xenopus cyclin E, and GST-Cdk2 (Cdk2) or GST-Cdk2K33R (Cdk2 K33R) were added to egg extract and incubated for the indicated times at 25°C. Samples were analyzed by SDS-PAGE and autoradiography. (D) 35S-labeled E1 translated in undepleted Xenopus egg extracts was added to a fresh membrane-free interphase egg extract and incubated for the indicated times. After 60 min of incubation, a sample of extract was incubated with beads coupled to the cyclin/Cdk-associated protein Suc1 (suc1). The behavior of radiolabeled E1 was analyzed by SDS-PAGE and quantitated as a percentage of that at 0 min.

E1 is phosphorylated within E1/cyclin E/Cdk2 complexes but is not released from the kinase after phosphorylation (13, 36). To test whether the protective effect of cyclin E-Cdk2 is a direct consequence of binding or phosphorylation of E1, we examined the stability of E1 within complexes reconstituted with cyclin E and either catalytically inactive or active recombinant GST-Cdk2. In interphase egg extracts, E1 levels remained constant in complexes formed with both forms of Cdk2, indicating that the kinase activity had no effect on the stability of E1. The presence of cyclin E-Cdk2 in the E1 immunoprecipitates, as well as the protective effect observed with catalytically inactive kinase complexes, strongly suggests that E1 is stabilized as a consequence of its interaction with cyclin E-Cdk2.

In undepleted egg extracts, a significant proportion of E1 molecules were degraded within 30 min (Fig. 1D). The E1 molecules that remained stable over the incubation time bound to a Cdk affinity matrix, suc1-agarose, indicating that this fraction of E1 molecules was associated with endogenous cyclin E/Cdk2 complexes. These results indicate that all the E1 molecules in stoichiometric excess relative to the available cyclin E/Cdk2 complexes are potentially unstable, while those bound to endogenous cyclin E/Cdk2 complexes become protected from degradation.

Protection of E1 bound to cyclin E/Cdk2 is reversed as a consequence of replication. The effect of BPV DNA replication on the stability of E1 was then monitored. Membrane-free interphase egg extracts supplemented with E1 translated in egg extracts catalyzed the efficient replication of plasmid DNA molecules containing the BPV origin of replication (Fig. 2A, lane 1). The reaction was almost completely inhibited by aphidicolin, which induces the accumulation of early-replicating intermediates by blocking DNA polymerase {alpha} activity (Fig. 2A, lane 2). As shown previously, the replication was dependent on the presence of a BPV origin of replication, since very little DNA synthesis was observed with a template lacking BPV sequences (Fig. 2A, lane 3).



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  		FIG. 2. E1 protection is reversed by replication. (A) 35S-labeled E1 was added to a membrane-free interphase egg extract supplemented or not with aphidicolin (Aphi) before addition of pSKori+ or pSKori-. Samples to be analyzed for DNA synthesis were supplemented with [32P]dCTP. After a 30-min incubation at 25°C, DNA products from pSKori+ (lanes 1), pSKori+ and aphidicolin (lanes 2), and pSKori- (lanes 3) extracts were purified, subjected to agarose gel electrophoresis, and detected by incorporation of [32P]dCTP. FI and RI designate the migration positions of supercoiled monomer circles of pSKori+ and pSKori- markers and replicative intermediates, respectively. (B) Samples from the same reactions were taken at intervals to monitor E1 behavior and analyzed by SDS-PAGE and autoradiography. (C) Complementation with BPV1 E2. 35S-labeled E1 was incubated in a membrane-free interphase egg extract in the absence (lane 1) or presence (lane 2) of 35S-labeled E2 before addition of pSKori+ and [32P]dCTP. The replication was allowed to proceed for 10 min, and the DNA products were analyzed by agarose gel electrophoresis. (D) The levels of E1 and E2 before addition of pSKori+ (lane 1) and after 30 min of incubation with pSKori+ (lane 2) were analyzed by SDS-PAGE and autoradiography.

As shown in Fig. 1D, E1 molecules not bound to cyclin E/Cdk2 were rapidly degraded after addition to membrane-free interphase egg cytosol. To specifically follow the behavior of E1 molecules stabilized by association with cyclin E/Cdk2 complexes, E1 was therefore incubated for 30 min in membrane-free interphase egg extract prior to the addition of DNA template. As shown in Fig. 2B, the fraction of E1 molecules stably bound to cyclin E/Cdk2 was degraded 20 to 30 min after addition of the BPV origin-containing plasmids. This was not observed in the reaction carried out in the presence of aphidicolin or after addition of a plasmid without a BPV origin of replication (Fig. 2B), demonstrating that E1 protection is reversed by replication.

While BPV E2 is dispensable in this cell-free system, we examined whether it could affect E1 stability during the in vitro replication process. In vitro-translated and -labeled E2 protein stimulated E1-dependent replication, as demonstrated by the enhanced level of late replicative intermediates at an early time point (Fig. 2C). While E2 levels remained unchanged, the E1 levels were diminished at the end of the reaction (Fig. 2D). Thus, the successful execution of E1-directed replication events causes E1 degradation, a phenomenon that is unaffected by the presence of E2.

E1 degradation in egg extract occurs by the ubiquitin-proteasome pathway. Analysis of E1 for the in vitro replication process revealed the transient appearance of high-molecular-weight E1 species in parallel with the gradual loss of E1 monomers (Fig. 3A). This behavior is characteristic of proteins that are modified by polyubiquitination for degradation by the proteasome. To test this in the case of E1, we identified E1-ubiquitin conjugates by supplementing the egg extracts with different modified forms of ubiquitin as well as by inhibiting the activity of cellular isopeptidases, which cleave ubiquitin molecules from protein substrates. As shown in Fig. 3B, addition of hexahistidine-tagged ubiquitin led to a notable increase in very large E1 conjugates (Fig. 3B, lane 3). This ubiquitin chain elongation was inhibited by incorporation of methylated ubiquitin, which terminates the elongation of ubiquitin chains and led to the appearance of apparently monoubiquitinated E1 (Fig. 3B, lane 4). The addition of K48R ubiquitin decreased the overall level of ubiquitin conjugates (Fig. 3B, lane 5).



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  		FIG. 3. E1 is degraded by the ubiquitin-proteasome pathway. (A) 35S-labeled E1 was added to a membrane-free interphase egg extract 30 min before addition of pSKori+. Aliquots were taken at intervals and analyzed by SDS-PAGE and autoradiography. The autoradiogram shows the appearance of high-molecular-weight labeled E1 species (lanes 2 and 3). (B) E1 is ubiquitinated in egg cytosol. Radiolabeled E1 was incubated in a membrane-free interphase egg treated with ubiquitin aldehyde and supplemented with either 6His-ubiquitin (lane 3), methylated ubiquitin (lane 4), or K48R ubiquitin (lane 5). No supplement was used in lane 2, and input was used in lane 1. The formation of large radiolabeled ubiquitin-E1 (Ub-E1) conjugates (lane 3) was blocked by addition of methylated ubiquitin (lane 4) and decreased in the presence of K48R ubiquitin (lane 5). (C) Radiolabeled E1 protein was added to Cdk2-depleted membrane-free interphase egg supplemented with NLVS at a final concentration of 50, 100, or 200 µM (lanes 3 to 5, respectively) or dimethyl sulfoxide (DMSO) as a control (lane 2). TO, input, lane 1. Quantitation of the fluctuation of E1 levels is shown on the lower panel. (D) Radiolabeled wild-type E1 (lanes 1 to 3) or E1-AAA (lanes 4 to 6) was added in a membrane-free interphase egg supplemented with ubiquitin aldehyde and either 6His-ubiquitin (lanes 2 and 5), methylated ubiquitin (lanes 3 and 6), or no ubiquitin complement (lanes 1 and 4). Aliquots of reaction mixtures were subjected to SDS-PAGE, and radiolabeled E1 species were visualized by autoradiography (upper panel). A shorter exposure is shown on the bottom panel to emphasize the accumulation of monoubiquitinated E1 species in the presence of methylated ubiquitin, indicated by an asterisk on the right.

We then tested the effects of a potent proteasome inhibitor (NLVS) on E1 degradation in membrane-free interphase egg extracts. As shown in Fig. 3C, NLVS stabilized radiolabeled E1 molecules translated in the absence of cyclin E-Cdk2 in egg extracts. This is consistent with the fact that the covalent attachment of ubiquitin to E1 targets the protein for degradation by the 26S proteasome (12). In addition, a mutant in which the three cyclin/Cdk phosphorylation sites were mutated to alanines (E1-AAA) was ubiquitinated with the same efficiency as the wild-type E1 protein (Fig. 3D). Restriction to monoubiquitination by the addition of methylated ubiquitin yielded similar amounts of monoubiquitinated E1 molecules in both the wild-type E1 and E1-AAA reactions, suggesting that the targeted degradation of E1 is independent of its phosphorylation status.

E1 ubiquitination in S-phase extracts from human 293 cells depends on replication. Given that replication caused E1 degradation in Xenopus egg extracts, we tested for a similar process in mammalian cell extracts with cytoplasmic fractions from human 293 cells. These extracts replicated the BPV origin-containing plasmid in the presence of E1 protein produced from baculovirus vectors (Fig. 4A, lane 2). Interestingly, the purified protein was phosphorylated and tightly associated with a kinase activity from insect cells (5). Replication and cell cycle factors are much less concentrated in this replication system than the Xenopus extracts, necessitating 100-fold more E1. We were unable to detect ubiquitinated E1 species on immunoblots following in vitro replication into the cytosol from asynchronously growing 293 cells.



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  		FIG. 4. E1 is ubiquitinated in a replication-dependent manner in a cell-free system from S-phase 293 cell extracts. (A) A total of 100 ng of E1 produced from baculovirus vector in Sf9 cells was added to reaction mixtures containing a cytoplasmic extract prepared from 293 cells synchronized in S phase by a double block with thymidine, followed by release into the culture medium for 1 h. Reaction mixtures were supplemented with aphidicolin (lane 1) or dimethyl sulfoxide (lane 2) before addition of pSKori+. Half of each reaction mixture was supplemented with [32P]dCTP and used in the BPV1 origin replication assay. Radiolabeled DNA products were purified after 60 min of incubation at 37°C and subjected to agarose gel electrophoresis. The fastest-migrating supercoiled form I (FI) and early and late replication intermediates (RI) are marked. (B) Half of the reaction mixtures described in A containing aphidicolin (lane 2) or dimethyl sulfoxide (lane 3) were used to monitor E1 ubiquitination (Ub-E1) for the replication assay. As a control, a reaction mixture lacking pSKori+ was used (lane 1). Reactions were supplemented with 32P-radiolabeled GST-ubiquitin and ubiquitin aldehyde and incubated for 30 min at 37°C. E1 was recovered by immunoprecipitation (IP), and radioactively labeled bands were visualized by autoradiography.

Cytosols were then prepared from 293 cells synchronized in S phase, and 32P-radiolabeled GST-ubiquitin was added together with ubiquitin aldehyde, which blocks isopeptidases and enhances the detection of ubiquitin conjugates. In the absence of DNA, E1 conjugates were not detected (Fig. 4B, lane 1). However, under conditions in which DNA synthesis was initiated by adding BPV plasmids, high-molecular-weight radiolabeled E1 conjugates were present in E1 immunoprecipitates after 30 min (Fig. 4B, lane 3) that were reduced in aphidicolin-treated cytosol (Fig. 4B, lane 2). These data suggest that some E1 molecules are also ubiquitinated in a replication-dependent manner in extracts of mammalian cells synchronized in S phase.

E1 is degraded by the proteasome in living cells. These results indicated that E1 is an unstable protein in cell-free systems. To test this conclusion in vivo, we used an expression vector encoding wild-type E1 with the green fluorescent protein attached to the N terminus. This vector allows us to force E1 expression and readily detect it, since we were unable to detect E1 in a BPV-transformed C127 line. The intrinsic replication capacity of the GFP-E1 recombinant protein was first tested in vitro. As shown in Fig. 5A, the GFP-E1 protein generated similar amounts of early and late replicative intermediates in the cell-free system and thus displayed an activity equivalent to that of the nontagged E1 protein. We also did not observe any significant effect of the fusion to GFP on E1 interactions with other proteins or its stability in the cell-free system (data not shown).



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  		FIG. 5. Functional GFP-E1 proteins are degraded by the proteasome in vivo. (A) In vitro-translated and 35S-radiolabeled GFP, E1, and GFP-E1 were analyzed by SDS-PAGE and autoradiography (left panel). Membrane-free interphase egg extracts were supplemented with either GFP (lane 1), E1 (lane 2), or GFP-E1 (lane 3) before addition of pSKori+ and [32P]dCTP. After a 30-min incubation at 25°C, DNA products were purified, subjected to agarose gel electrophoresis, and detected by incorporation of [32P]dCTP. FI and RI designate the migration positions of supercoiled monomer circles of pSKori+ and replicative intermediates, respectively (right panel). (B) Transient replication of pSKori+ in the presence of E1 and E2 (lanes 1), GFP and E2 (lanes 2), and GFP-E1 and E2 (lanes 3) expression vectors. Low-molecular-weight DNA was extracted from 293 cells at 48, 72, and 96 h after transfection, digested with SmaI and DpnI, and analyzed by Southern blotting. Filters were probed with radiolabeled pSKori+ plasmid. pOri indicates the band generated after digestion of pSKori+ with SmaI. (C) 293 cells were transfected with GFP-E1 expression vector, and proteins from total cell lysate were resolved on an SDS gel and analyzed by Western blotting with an antibody directed against the GFP (lane 2). Cell lysate from untransfected cells is shown in lane 1 as a control. The presence of GFP-E1 is indicated (left panel). GFP-E1 expression plasmid was transfected into asynchronously growing 293 cells. Cells were treated with cycloheximide (CHX), and the level of GFP-E1 was evaluated at the indicated time (hours) by Western blotting with the GFP antibody (right panel). Only relevant parts of the blot are shown. 293 cells transfected with GFP-E1 and E2 expression vectors and pSKori+ plasmid were treated with cycloheximide as above in the presence of either MG132 (+) or dimethyl sulfoxide (-). Cells were harvested, and lysates were prepared for Western blotting with an antibody to GFP.

We then tested the ability of GFP-E1 to support BPV origin-dependent replication in vivo in the presence of BPV E2. In transiently transfected 293 cells, GFP-E1 yielded a level of replication comparable to that of the wild-type nontagged E1 protein at different times after cotransfection (Fig. 5B, compare lanes 3 with lanes 1). In the presence of GFP and E2, no replication was observed (Fig. 5B, lanes 2). Thus, E1 monomers carrying a GFP tag at the amino terminus are able to form a functional hexameric helicase on the BPV origin without affecting replication levels either in vitro or in vivo.

Since we do not have a suitable E1-specific antibody to analyze its levels by Western blotting, we used a GFP antibody that can detect low levels of GFP-E1 (Fig. 5C). To measure E1 steady-state levels in 293 cells transfected with the GFP-E1 expression vector, cells were treated with cycloheximide to block de novo protein synthesis, and extracts were prepared at various times thereafter. Anti-GFP immunoblots showed no apparent changes in E1 levels during the 4-h treatment.

Since degradation of E1 was only detectable in the presence of E2 and BPV plasmids, i.e., conditions that allow viral replication, 293 cells were cotransfected with the GFP-E1 and E2 expression vectors as well as the BPV origin-containing plasmid and then cultured in the presence of cycloheximide. One transfection was cultured in the presence of the proteasome inhibitor MG132, while the other was cultured with dimethyl sulfoxide, the solvent for MG132. In contrast to the apparent stability of E1 when expressed alone in 293 cells, significant degradation of E1 was observed in the presence of E2 and BPV plasmids, with an estimated half-life of less than 2 h. In addition, MG132 stabilized E1 over the 4 h of cycloheximide treatment, confirming the proteasome-dependent turnover of E1 under these conditions.

E1 is polyubiquitinated in vivo. To investigate whether E1 ubiquitination can be detected in vivo, we cotransfected COS-7 cells with the GFP-wild-type E1 expression vector and a hexahistidine-tagged ubiquitin (6His-ubiquitin) expression vector either with or without the E2 expression vector and the BPV origin-containing plasmid. Cell extracts were prepared and analyzed by immunoblotting with the antibody to GFP (Fig. 6A, lanes 1 to 3). An aliquot was also bound to nickel affinity resin prior to immunoblotting (Fig. 6A, lanes 4 to 6). A ladder of slower-migrating E1 species was specifically detected in samples from total cells as well as in the 6His-ubiquitin-conjugated protein purified fraction (Fig. 6A, lanes 1 and 4). However, cotransfection of the E2 and BPV plasmids with the GFP-E1 plasmid strongly increased the amount of polyubiquitinated E1 (Fig. 6A, lanes 2 and 5).



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  		FIG. 6. In vivo polyubiquitination of E1. (A) COS-7 cells were transiently cotransfected with pEGFP-E1 and pCDNA3-6HisUb in the absence (lanes 1 and 4) or presence (lanes 2 and 5) of pCGE2 and pSKori+. COS-7 cells transfected with pCDNA3-6HisUb were used as a control (lanes 3 and 6). After 48 h, cells were harvested and lysed. An aliquot corresponding to 1% of each cell extract was kept for input analysis (lanes 1 to 3). The ubiquitinated proteins were purified from the extracts on nickel-agarose (lanes 4 to 6). Purified proteins were subjected to SDS-PAGE and analyzed by immunoblotting with anti-GFP antibody. Polyubiquitinated GFP-E1 proteins are detected as a smear of high-molecular-weight species (lanes 1, 2, 4, and 5). 6xHisUb, 6His-ubiquitin. (B) COS-7 cells were transfected with the indicated plasmids. After 48 h, cells were lysed in the presence of MG132 and ubiquitin aldehyde. Cell lysates were immunoprecipitated with anti-E1 C-terminal (Cter) antibody or with preimmune antiserum ({alpha}-PI), and the Western blot was probed with anti-HA. Ubiquitin-E1 (Ub-E1) conjugates were detected in cells transfected with pEGFP-E1, pCGE2, pSKori+, and pCDNA3-HAUb. 6xHAUb, HA-ubiquitin. Sizes are shown in kilodaltons.

To further demonstrate that these high-molecular-weight species contain ubiquitin, an immunoprecipitation-Western blot assay was used in which HA-tagged ubiquitin was cotransfected with the GFP-E1 plasmid into COS-7 cells, once again with or without the E2 and BPV origin-containing plasmids. Extracts were immunoprecipitated with an anti-E1 or control antibody, and the ligation of HA-tagged ubiquitin to E1 was evaluated by Western blotting with an antibody to the HA tag. As shown in Fig. 6B, a robust signal of polyubiquitinated E1 species was detected only when E2 and BPV origin-containing plasmids were present in COS-7 cells. These ubiquitination experiments, together with our inability to detect subtle changes in E1 levels without E2 and BPV origin-containing plasmids, indicate that conditions allowing viral replication facilitate the detection of ubiquitin-mediated E1 degradation within living cells.


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DISCUSSION
 
The results presented in this study provide new insights into how papillomavirus DNA replication is regulated and coordinated within the eukaryotic cell cycle. We show that the papillomavirus E1 protein is regulated by the ubiquitin-proteasome pathway both in vitro and in vivo, that E1 stability is in part regulated by cyclin E/CDk2, and that free E1 (E1 not bound to cyclin E/Cdk2) is an unstable protein whose turnover is controlled by the ubiquitin-proteasome pathway. In contrast, E1 bound to cyclin E/Cdk2 is stable. However, E1 stably bound to cyclin E/Cdk2 can be destabilized as a consequence of E1-dependent replication events.

Our detection of regulated, ubiquitin-mediated E1 proteolysis was greatly facilitated by the in vitro reconstitution of E1-dependent replication in Xenopus egg extracts (13). These egg extracts have been shown to represent a complete system for regulated degradation of a large number of factors, such as mitotic cyclins and Cdk inhibitor (16, 40). Our results show that a fraction of E1 expressed in low levels in these extracts is associated with cyclin E/Cdk2 and remains stable in the absence of viral replication. The protective effect is observed within catalytically inactive kinase complexes, suggesting that E1 is stabilized as a direct consequence of its interaction with cyclin E/Cdk2. However, newly translated E1 molecules also accumulated in cyclin E/Cdk2-depleted translation extracts, indicating that the cyclin E/Cdk2 complex is not the only cellular factor that stabilizes E1.

The protein translated in the absence of cyclin E/Cdk2 is nevertheless rapidly turned over upon its addition to fresh egg extract depleted of cyclin E/Cdk2. This suggests that newly synthesized E1 molecules might be bound to molecular chaperones prior to association with cyclin E/Cdk2. Human papillomavirus 11 E1 protein was previously found to bind to the human chaperone proteins Hsp70 and Hsp40 (35), but the direct effect of these proteins on E1 folding and stability remains to be investigated. We found that E1 was stabilized by complementing cyclin E/Cdk2-depleted extracts with cyclin E and Cdk2 produced by in vitro translation in egg extracts. This was not seen upon complementation with E2, and in contrast to cyclin E/Cdk2, E2 did not coimmunoprecipitate with E1 under conditions of low expression (data not shown).

Significant differences exist between the cell cycle of amphibian eggs and human somatic cells. For example, in Xenopus eggs, ~=90% of the Cdk2 is associated with cyclin E, and cyclin A is associated with Cdk1 (25), while in human cells, cyclin A is a primary partner of Cdk2 (48). Nonetheless, cyclin E/Cdk2 has been shown recently to be specifically required for active human papillomavirus replication in human cells (33). In our cell-free system, E1 is resistant to ubiquitin-mediated degradation within E1/cyclin E/Cdk2 complexes but becomes unstable following replication. These results led us to hypothesize that E1, cyclin E, and Cdk2 dissociate as a consequence of replication. In this scenario, E1 stability might therefore be coupled to the periodic accumulation of cyclin E at the G1/S-phase transition of the cell cycle (14, 26). Importantly, this notion correlates with the biology of papillomavirus. Specifically, if E1 is uniquely stable for the short time period when cyclin E/Cdk2 is present in the cell, this would explain why it cannot be detected in papillomavirus-transformed cells.

Once engaged in replication, E1 degradation can be blocked by aphidicolin, an inhibitor of the elongation phase of DNA replication. This indicates that replication elongation is required for E1 destruction. E1 is thought to provide the helicase activity at the replication fork (34). Our analysis of the DNA products obtained under conditions in which E1 is stabilized indicate that E1 degradation occurs after replication is complete (C. Bonne-Andrea, unpublished data).

Analysis of E1 steady-state levels in living cells revealed that E1 is also a substrate of the ubiquitin-proteasome pathway in vivo. However, E1 overexpression by transient transfection did not facilitate the detection of E1 ubiquitination and degradation in the absence of other viral components. This is in apparent contradiction to a previous study that reported a cell cycle-dependent fluctuation in the level of E1 protein. When E1 was expressed under the control of an inducible promoter, its levels were low in G1 phase, increased as the cells entered S phase, and remained high until G2/M (3). This observation would fit with the proposed protective effect of cyclin E/Cdk2 at each entry into S phase. Our data show that significant E1 turnover as well as the bulk of ubiquitin-mediated E1 degradation can be preferentially detected in the presence of E2 and BPV origin-containing plasmids that allow viral replication.

As demonstrated in the cell-free system, E1 degradation may occur as a consequence of replication. However, given the level of E1 proteolysis detected in unsynchronized transfected cells, a major proportion of E1 molecules are likely degraded in a replication-independent manner. A recent study reported that E2 is necessary to recruit E1 in nuclear foci, an effect that was increased by the presence of an origin-containing plasmid (54). This observation raises the possibility that the targeting of E1 by the ubiquitin-proteasome pathway may be localized to specific subnuclear compartments. Testing this hypothesis will be the subject of further work.

On the other hand, in the cell-free system, E1 degradation was not affected by its viral partner E2. Like E1, E2 was recently shown to be an unstable protein that is degraded by the ubiquitin-proteasome pathway in vivo (2, 43). Our results show that, in contrast to E1, E2 levels remain constant during viral DNA replication in interphase egg cytosols, suggesting that E2 may be targeted to degradation either at a different stage of the cell cycle or by a different ubiquitin ligase.

It was recently reported that the level of human papillomavirus 31 E1 expression is regulated by both transcriptional and posttranscriptional control mechanisms during the early phases of the viral life cycle (23). E1 RNA levels were also found to be upregulated during the maximal amplification of viral genomes in raft cell cultures that support the complete human papillomavirus life cycle (41). Our data demonstrate for the first time that, in dividing cells, a major regulatory restriction on viral replication can also be imposed by regulated proteolysis of the viral initiator.

Variation in E1 protein levels is one of the mechanisms that govern relative amounts of viral DNA synthesis. The level of E1 in various cell stages may be the result of an equilibrium between expression and degradation. Replication control could therefore be abolished in terminally differentiated cells by increased levels of E1. There are several potential mechanisms by which increased steady-state levels of E1 might be generated. High E1 expression resulting from an increased number of genomes could override controls on E1 steady-state levels. This might correlate with the switch from the semiconservative theta structure mode of the human papillomavirus 16 viral genome replication to the rolling-circle mode of viral replication when cells enter the terminal differentiation pathway in the raft system (15). In addition, the ubiquitin-proteolytic system responsible for E1 degradation might be inactive in differentiated cells.

The E1 ubiquitination reaction yields very high molecular mass conjugates that are bona fide polyubiquitinated E1 species, since addition of methylated ubiquitin, an inhibitor of polyubiquitin chain formation, blocks the formation of these large conjugates. The efficiency of E1 ubiquitination is therefore very high in comparison to certain proteins such as cyclin B, which presents a much lower level of ubiquitin conjugation in standard in vitro ubiquitination assays (18, 19). The pattern of ubiquitin conjugation to E1 resembles that of the p53 tumor suppressor, generated in vitro by the cellular ubiquitin ligase E6-AP and the human papillomavirus 16 E6 protein (49). In this case, p53 was found to form a stable complex with E6 and E6-AP. In contrast, the binding of cyclin B to the activators of the anaphase-promoting complex/cyclosome (APC/C) ubiquitin ligase, which is responsible for its ubiquitination, was shown to be weak and difficult to detect (45). This may indicate that the interaction between E1 and the cellular ubiquitin-protein ligase that mediates its ubiquitination is rather strong.

The ubiquitin-dependent proteolytic pathway targets many key regulators of chromosomal DNA replication for intracellular degradation. In contrast to the viral initiation process, the mechanisms leading to the initiation of DNA replication on cellular chromatin depend on the sequential association of several initiation factors. The six proteins of the origin recognition complex recruit the Cdc6 and Cdt1 proteins, which, in turn, are required to recruit the proteins that form the minichromosome maintenance protein (MCM) complex, which presents the helicase activity (reviewed in references 29 and 55). The regulation of this sequence of events involves cyclin/Cdks, which help activate the firing of replication origins for the S phase of the cell cycle and block reinitiation of DNA replication within a single cell cycle. This leads to inactivation by degradation, nuclear exclusion, or inhibition of chromatin binding.

Two replication factors have recently been shown to be targeted for ubiquitin-mediated destruction in human cells by two different ubiquitin ligase complexes: the hOrc1p subunit of the origin recognition complex involves the ubiquitin ligase complex SCF (38), while Cdc6 ubiquitination requires the APC/C (44). Identification of the cellular ubiquitin ligase involved in the degradation of E1 will be critical to understanding the mechanisms by which papillomavirus DNA synthesis is differentially regulated and further investigating whether the viral protein E1 affects cell cycle regulatory pathways and contributes to the pathogenesis of papillomavirus infections.


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ACKNOWLEDGMENTS
 
We thank Bob Hipskind for critical reading of the manuscript and Jean-Claude Labbe, Thierry Lorca, and Marc Piechaczyk for helpful discussions. We gratefully acknowledge Guillaume Bossis for advice and assistance with the in vivo ubiquitination experiments.

M.-H.M. and N.C. were supported successively by a fellowship from the French MRT and a fellowship from the Association pour la Recherche sur le Cancer. This work was supported by a grant from the Association pour la Recherche sur le Cancer to C.B.-A. O.C. is supported by a grant from the Association pour la Recherche sur le Cancer and the ATIPE Program of the French CNRS.


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FOOTNOTES
 
* Corresponding author. Mailing address: Centre de Recherches de Biochimie Macromoléculaire, CNRS, IFR 24, 1919 Route de Mende, 34 293 Montpellier Cedex 5, France. Phone: 33 4 67 61 33 32. Fax: 33 4 67 52 15 59. E-mail: bonne-andrea{at}crbm.cnrs-mop.fr. Back

{dagger} Present address: Swiss Institute for Experimental Cancer Research, CH-1066 Epalinges/Lausanne, Switzerland. Back


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Journal of Virology, November 2002, p. 11350-11358, Vol. 76, No. 22
0022-538X/02/$04.00+0     DOI: 10.1128/JVI.76.22.11350-11358.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.



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