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[Cancer Research 62, 3315-3321, June 1, 2002]
© 2002 American Association for Cancer Research

Tumor Biology

Human Papillomavirus E6-induced Degradation of E6TP1 Is Mediated by E6AP Ubiquitin Ligase1

Qingshen Gao2, Ajay Kumar2, Latika Singh, Jon M. Huibregtse, Sylvie Beaudenon, Seetha Srinivasan, David E. Wazer, Hamid Band and Vimla Band3

Division of Radiation and Cancer Biology, Department of Radiation Oncology, New England Medical Center [Q. G., A. K., L. S., S. S., D. E. W., V. B.], and Department of Biochemistry, Tufts University School of Medicine [V. B.], Boston, Massachusetts 02111; Institute for Cellular and Molecular Biology, Section of Molecular Genetics and Microbiology, University of Texas at Austin, Austin, Texas 78712-1095 [J. M. H., S. B.]; and Laboratory of Lymphocyte Biology, Division of Rheumatology, Immunology and Allergy, Brigham & Women’s Hospital, Harvard Medical School, Boston, Massachusetts 02111 [H. B.]


   ABSTRACT
Top
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
High-risk human papilloma viruses are known to be associated with cervical cancers. We have reported previously that the high-risk human papillomavirus (HPV) E6 oncoprotein interacts with E6TP1, a novel Rap GTPase-activating protein (RapGAP). Similar to p53 tumor suppressor protein, the high-risk HPV E6 oncoproteins target E6TP1 for degradation. The HPV16 E6-induced degradation of E6TP1 strongly correlates with its ability to immortalize human mammary epithelial cells. In this study, we used treatment with a proteasome inhibitor MG132, analysis in CHO-ts20 cells with a thermolabile ubiquitin-activating enzyme, and direct detection of ubiquitin-modified E6TP1 to demonstrate that E6TP1 is targeted for degradation by the ubiquitin-proteasome pathway both in the presence and in the absence of E6. Using deletion mutants of E6TP1, we mapped the region required and sufficient for E6 binding to COOH-terminal 40 amino acid residues and showed this region to be necessary for E6-dependent degradation of E6TP1. Furthermore, the E6-binding region of E6TP1 complexes with E6AP via E6. Importantly, the purified E6AP enhanced the ubiquitination and degradation of E6TP1 in the presence of E6 in vitro. Additionally, the expression of a dominant-negative E6AP mutant (C833A) in cells inhibited the E6-induced degradation of E6TP1. These findings demonstrate that the E6-induced decrease in the levels of E6TP1 protein involves the E6AP-mediated ubiquitination followed by proteasome-dependent degradation.


   INTRODUCTION
Top
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Studies with viral oncogenes that induce dominant cellular transformation have led to the identification and elucidation of a number of cellular pathways that are involved in human cancer (1, 2, 3, 4, 5, 6, 7, 8) . One such group of tumor viruses directly implicated in the pathogenesis of human cancer is the high-risk of HPVs,4 which are thought to be causally linked to the development of >90% cases of cervical cancer (9 , 10) . In vitro studies have defined two HPV oncogenes, E6 and E7, and these genes are always expressed in HPV-associated carcinomas and cell lines derived from them (11 , 12) .

The ability of HPV E6 and/or E7 oncogenes to induce the immortalization of human epithelial cells in vitro has provided an invaluable tool to identify the mechanism by which these oncogenes function (13, 14, 15, 16, 17) . E6 and E7 are thought to abrogate the cell growth control by inactivating key cellular regulatory proteins (5, 6, 7, 8) . For example, E7 proteins bind and inactivate the function of the retinoblastoma tumor suppressor protein (RB; Refs. 5 and 8 ) by targeting it for degradation via the ubiquitin pathway (17 , 18) . Similarly, E6 protein was shown to interact with and enhance the degradation of another tumor suppressor protein, p53 (6 , 7 , 16 , 17) , which plays an important role in cell cycle control and apoptosis in response to DNA damage (19 , 20) . Although E6-induced loss of p53 is an important element of E6-induced cellular transformation, recent studies have identified a number of additional cellular targets of E6 that may also play an important role. These included E6BP (E6 binding protein; Ref. 21 ), paxillin (22) , hDlg (the human homologue of the Drosophila discs large tumor suppressor protein; Refs. 23 and 24 ), Mcm7 (minichromosome maintenance protein 7; Refs. 25 and 26 ), IRF-3 (IFN regulatory factor-3; Ref. 27 ), myc (28) , Bak (Bcl-2 homologous antagonist/killer; Refs. 29 and 30 ), E6TP1 (E6 targeting protein 1; Ref. 31 ), CBP/p300 (32 , 33) , Tyk2 (protein-tyrosine kinase 2; Ref. 34 ), hScrib [the human homologue of the Drosophila Scribble (Vartul) tumor suppressor protein; Ref. 35 ], PKN (a novel protein kinase with a catalytic domain homologous to that of the protein kinase C; Ref. 36 ), MUPP1 (multi-PDZ-domain protein 1; Ref. 37 ), MAGI-1 [one Membrane-associated guanylate kinase (MAGUK) protein; Ref. 38 ), and Gps2 (G-protein pathway suppressor 2; Ref. 39 ). In addition to these E6-interacting proteins, E6 is also reported to induce the increase of the telomerase activity (40, 41, 42, 43) . Telomerase is normally at very low level in vivo and becomes activated during tumorigenesis (44) . The telomerase activity is dramatically increased in E6 immortalized cells (40, 41, 42, 43) . Understanding the role of these E6 targets in E6-induced oncogenesis is an area of intense research in many laboratories included ours.

The mechanisms whereby E6 alters the function of its cellular targets are of obvious interest. One mechanism is provided by E6-p300 interaction in which binding of E6 to a functionally important domain or in its vicinity induces loss of function (32 , 33) . A second mechanism is the ability of E6 to activate cellular protein function. An example of this mechanism is provided by E6-dependent recruitment of the ubiquitin ligase E6AP to crucial cellular proteins, such as p53, leading to their ubiquitination and subsequent degradation (45 , 46) . A related, but probably distinct, mechanism accounts for E6-induced degradation of other targets such as Mcm7 and bak (25 , 29) . In contrast to p53, which does not directly bind to E6, these other targets interact with E6 directly. Recent studies suggest that E6-induced ubiquitination of these targets also involves E6AP, although some evidence has been presented for the involvement of non-E6AP ubiquitin ligase (47) .

We have recently identified a novel GAP of the Rap family small GTPase, E6TP1, as a direct E6-binding protein (31) . The interaction with E6TP1 was observed with cancer-associated high-risk HPV E6 protein but not with benign lesion-associated low-risk HPV E6 protein (31) . Furthermore, an essentially perfect correlation between the ability of E6 mutants to bind to E6TP1 and their ability to immortalize human mammary epithelial cells was observed (42) . These observations suggest a potentially important role for E6TP1 in E6-induced oncogenesis. E6TP1, by virtue of its GAP activity, is expected to reduce the levels of GTP-bound Rap protein. Activated Rap has been indicated in sustained activation of mitogen-activated protein kinase upon growth factor stimulation of cells, and constitutively active Rap can transform fibroblasts (48 , 49) . In this context, our recent studies have shown E6TP1 to be a novel RapGAP.5 Given the fact that E6-E6TP1 interaction indicates a potential role for an important small G-protein signaling pathway in E6-induced oncogenesis, understanding the mechanism by which E6 alters E6TP1 function is of considerable interest. Here, we show that E6TP1 is targeted for degradation by the ubiquitin-proteasome pathway both in the absence and in the presence of E6. Additionally, we demonstrate that E6TP1 binds to E6AP in the presence of E6. Significantly, the purified E6AP can enhance the ubiquitination and degradation of E6TP1 in vitro in the presence of E6. Furthermore, overexpression of wild-type E6AP enhances, whereas a dominant-negative mutant of E6AP (C833A; Ref. 50 ) inhibits E6-induced degradation of E6TP1. These findings demonstrate that E6 targets E6TP1 for ubiquitination through E6AP and subsequent degradation by the proteasome pathway.


   MATERIALS AND METHODS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Chemicals and Plasmid Constructs.
Proteasome inhibitor MG132 (carbobenzoxy-L-leucyl-L-leucyl-L-leucinal) was purchased from Calbiochem (La Jolla, CA). The plasmid pMT-(HA-ubiquitin)8 encoding HA-tagged ubiquitin was kindly provided by Dr. Dirk Bohmann, European Molecular Biology Laboratory, Germany (51) . pGEX2TK-E6TP1–1590-1783 (previously referred as pGEX2TK-E6TP1-C-194), encoding the GST fusion protein for E6TP1 fragment 1590–1783, has been described (31) . pCR3.1-16E6 was constructed by subcloning 16E6 from pSG5-16E6 into pCR3.1 vector. pGEX-E6AP, pSP-E6AP, pCMV4-E6AP, and pCMV4-E6AP C833A were kindly provided by Dr. Peter Howley (Harvard Medical School, Boston, MA). pEF-myc-16E6 was kindly provided by Dr. M. Ishibashi (Aichi Cancer Center, Nagoya, Japan; Ref. 24 ). GFP construct and anti-GFP antibodies were purchased from Clontech (Palo Alto, CA).

The NH2-terminal myc epitope in full length of E6TP1 was incorporated using the PCR and cloned in pCMV vector. The pGEX4T1-E6TP1 mutant constructs encoding GST fusion proteins for E6TP1 fragments 1759–1783, 1744–1783, 1716–1783, 1704–1783, 1704–1715, 1704–1743, 1704–1758, 1716–1758, 1716–1743, and 1744–1758 were generated using the PCR. All of the constructs were sequenced to ensure that they lack PCR-generated mutations.

Transfection of 293T Cells, Proteasome Inhibitor Treatment, and Western Blotting.
293T cells (1 x 106 per 100-mm dish) were transfected with 2 µg of pCMV-E6TP1 with and without 6 µg of pEF-myc-16E6 using the calcium phosphate method (52) . Each dish was cotransfected with 500 ng of GFP to control for transfection efficiency. Total amount of DNA was kept constant by adding empty vector. After 40 h of transfection, the cells were treated with MG132 (final concentration, 50 µM) for 4 h. Cell lysates were then prepared in sample buffer [50 mM Tris (pH 6.8), 2% SDS, and 10% glycerol], resolved by SDS-PAGE, and subjected to Western blotting using a rabbit anti-E6TP1 serum (42) or anti-myc 9E10 or anti-GFP antibody (Clontech). Signals were then detected using the enhanced chemiluminescence (ECL) method, as suggested by manufacturer (Amersham Biosciences, Inc., Piscataway, NJ). For assessment of E6TP1 ubiquitination, a plasmid construct encoding HA-tagged Ub (HA-Ub) was cotransfected together with the indicated plasmids. Forty h after transfection, the cells were mock-treated or treated with 50 µM MG132 for 4 h and extracted in RIPA buffer [100 mM Tris (pH 8.0), 100 mM NaCl, 1% NP40, 1% deoxycholic acid, and 0.1% SDS]. Myc-E6TP1 was immunoprecipitated with anti-myc 9E10 antibody and probed with anti-HA antibody 12CA5 to detect the ubiquitinated E6TP1. The same blot was reprobed with anti-myc antibody to detect E6TP1.

Expression of E6TP1 in CHO-ts20 Cells.
CHO-ts20 cells (5 x 105 ; Ref. 53 ) grown at 30°C were transfected with 2 µg of pCMV-myc-E6TP1 or pCMV-FLAG-p53 with or without 6 µg of pCR3.1-HPV16 E6 using the calcium phosphate method. The total amount of DNA was kept constant by adding vector DNA. Forty h after transfection, one set of transfectants was maintained at 30°C, and another set was switched to 42°C for 8 h. The cell lysates were prepared and subjected to Western blotting using the anti-myc 9E10 or anti-FLAG antibodies (Sigma, St. Louis, MO) to assess the levels of transfected E6TP1 or p53 protein, respectively.

In Vitro Binding between E6 and E6TP1 or Its Mutants.
HPV16 E6 protein was generated by in vitro translation from pCR3.1–16E6. Translation reactions were carried out in the presence of [35S]cysteine (NEN, Boston, MA) in a wheat germ-based coupled transcription/translation system (TNT wheat germ system; Promega Corp., Madison, WI), according to the supplier’s recommendations. Equal aliquots of 35S-labeled E6 translation reaction were allowed to bind to 1 µg of GST or GST fusion proteins of E6TP1 or its mutants noncovalently bound to glutathione-Sepharose beads in 300 µl of lysis buffer [100 mM Tris (pH 8.0), 100 mM NaCl, and 0.5% NP40]. After 2 h incubation at 4°C, the bound 35S-labeled E6 was resolved by SDS-PAGE and visualized by fluorography.

In Vitro Binding between E6AP and E6TP1.
35S-labeled E6AP was generated by in vitro translation in the presence of [35S]methionine (DuPont NEN), as described above for E6. Equal aliquots of 35S-labeled E6AP translation reaction were incubated with 1 µg of GST or GST fusion of E6TP1 residues 1590–1783 noncovalently bound to glutathione-Sepharose beads in 300 µl of lysis buffer in the presence or absence of 35S-labeled in vitro translated E6 (see above). After 2 h of incubation at 4°C, the bound 35S-labeled E6AP was resolved by SDS- PAGE and visualized by fluorography. The gel was stained with Coomassie Brilliant Blue for the expression of GST-fusion proteins.

In Vitro Ubiquitination Assay.
Recombinant baculovirus expressing HPV16 E6 was produced using the BaculoGold system (PharMingen) in High5 cells (Invitrogen), and the E6 protein was isolated and purified as described previously (35) . Baculovirus E6AP protein and human E1 Ub-activating enzyme were expressed and purified from insect cells, and AtUbc8 E2-Ub conjugating enzyme was expressed in Escherichia coli (35) .

In vitro translation of E6TP1 was performed in reticulocyte lysate system (TNT system; Promega) in the presence of [35S]methionine. Ubiquitination assay mixtures contained 4 µl of translation reaction, 2 mM ATP, and 2 µg of Ub (Sigma) in 35 µl of 25 mM Tris (pH 7.5), 50 mM NaCl, 2 mM MgCl2, 50 µM DTT, with purified E1 and E2 with or without 1 µg of 16E6 and/or E6AP and/or E6TP1. Reaction mixtures were incubated for 45 min at room temperature, resolved by SDS-PAGE, and visualized by fluorography.

In Vivo E6-induced E6TP1 Degradation in the Presence of E6AP.
To assess the effect of E6AP or its mutant on E6TP1 degradation in vivo, 2 µg of pCMV-E6TP1 were cotransfected into 1 x 106 293T cells together with 2 µg of pCMV4 constructs encoding wild-type E6AP or 2 µg of its dominant-negative mutant (C833A) with or without 4 µg of pEF-myc-16E6. Each dish was also cotransfected with 500 ng of GFP for transfection efficiency control. The total amount of DNA was held constant by adding vector DNA. After 48 h, cell lysates were prepared in sample buffer, and E6TP1 was detected by Western blot using an anti-E6TP1 antiserum. HPV16 E6 or E6AP proteins were detected using anti-myc or anti-HA 12CA5 antibodies. GFP was detected using anti-GFP antibody.


   RESULTS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Proteasome Inhibitor Treatment Stabilizes E6TP1 Both in the Presence and Absence of HPV16 E6.
To assess whether the stability of E6TP1 is regulated by proteasomal degradation, we examined the effect of the proteasome inhibitor MG132 on E6TP1 levels in the presence and absence of E6. 293T cells were transfected either with E6TP1 alone or together with HPV16 E6, followed by treatment with MG132 (final concentration, 50 µM) for 4 h, and cell lysate was analyzed for the levels of E6TP1 protein by Western blotting. As we have observed previously (31) , cotransfection with E6 induced a marked decrease in E6TP1 protein levels (Fig. 1Citation , upper panel, compare Lanes 1 and 3). Notably, MG132 treatment led to a substantial increase in E6TP1 levels as compared with untreated cells both in the presence (upper panel, Lanes 3 and 4) and in the absence (upper panel, Lanes 1 and 2) of HPV16 E6 expression. As expected, 16 E6 expression was detected in Lanes 3 and 4 (middle panel). These results indicate that E6TP1 levels are regulated by the Ub-proteasome pathway both basally and when degradation is induced by E6 protein.



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  		Fig. 1. Levels of E6TP1 protein upon proteasome inhibitor treatment. 293T cells (1 x 106) were cotransfected with 2.0 µg each of pCMV construct encoding E6TP1 with or without 6.0 µg of pEF-myc-HPV16 E6. Each dish was also cotransfected with 500 ng of GFP for transfection efficiency control. The total amount of DNA/dish was kept constant by adding vector DNA. After 40 h of transfection, cells were either mock-treated or treated with MG132 (final concentration, 50 µM) for 4 h. Cell lysates were then analyzed for E6TP1 or E6 proteins using anti-E6TP1 antiserum or anti-myc immunoblotting, respectively. The same blot hybridized with anti-GFP antibody served as a transfection efficiency control. M.W., molecular weight.

 
Stabilization of E6TP1 in CHO-ts20 Cells at Nonpermissive Temperature.
Accumulation of E6TP1 protein upon MG132 treatment indicated the role of ubiquitination in regulating E6TP1 turnover. To demonstrate the role of the Ub pathway, we assessed the level of myc-tagged E6TP1 or p53 (used as a positive control), expressed in the presence or absence of E6, in CHO-ts20 cells with a thermolabile Ub-activating enzyme (E1), the first enzyme of Ub machinery (53) . One set of transfectants was maintained at permissive temperature (30°C), and another set was switched to a nonpermissive temperature (42°C) for the last 8 h before cell lysis. Cell lysates were prepared 48 h after transfection, and the levels of E6TP1 protein were assessed by anti-myc immunoblotting. The levels of p53 protein were assessed using anti-FLAG antibody. As seen in Fig. 2A,Citation the level of E6TP1 was reduced (~3-fold) in cells coexpressing E6 compared with cells expressing E6TP1 in the absence of E6, when cells were maintained at 30°C (compare Fig. 2Citation A, Lanes 1 and 3). Notably, the level of E6TP1 increased (~2-fold) when cells were incubated for 8 h at 42°C; this was observed in cells expressing E6TP1 alone (compare Lanes 1 and 2) as well as those coexpressing E6 (compare Lanes 3 and 4). As expected, the levels of p53 protein decreased in E6-expressing cells at 30°C (~3-fold), and stabilization of p53 (~3-fold) was observed at 42°C (Fig. 2BCitation , Lanes 3 and 4). These results support an important role of the Ub machinery in targeting E6TP1 for proteasomal degradation.



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  		Fig. 2. Levels of E6TP1 protein in the absence of Ub-activating enzyme E1. CHO cells (5 x 105; containing temperature-sensitive Ubiquitin-activating enzyme E1) were cotransfected with 2.0 µg of myc-E6TP1 or FLAG-p53 with or without 6.0 µg of pCR3.1–16 E6. The total amount of DNA/dish was kept constant by adding vector DNA. Forty h after transfection, one set of transfectants was transferred to 30°C, and another set of transfectants was transferred to 42°C and incubated for an additional 8 h. Cell lysates were subjected to Western blotting using anti-myc (9E10) or anti-FLAG antibodies to examine the levels of E6TP1 or p53 protein, respectively. The expression was quantitated by densitometry, and the data are presented as fold change compared with the levels of proteins at 30°C. M.W., molecular weight.

 
Ubiquitination of E6TP1 in Vivo.
The above experiments strongly supported the role of the Ub-proteasome pathway in controlling the levels of E6TP1 and its destabilization by E6. To directly examine whether E6TP1 is ubiquitinated in vivo, we coexpressed myc-E6TP1 with HA-tagged ubiquitin, which markedly facilitates the detection of ubiquitinated protein (51) . To further enhance the detection of ubiquitinated E6TP1, 293T cells transfected with myc-tagged E6TP1 and HA-tagged Ub with or without E6 were either mock-treated or treated with MG132 (final concentration, 50 µM) for 4 h before cell lysis. Cell lysates were subjected to anti-myc immunoprecipitation, and immunoprecipitates were blotted with anti-HA antibody 12CA5. As shown in Fig. 3Citation , ubiquitinated E6TP1 was readily observed when both E6TP1 and HA-tagged Ub were cotransfected into 293T cells in the absence of HPV16 E6 (Fig. 3Citation , Lane 3). Notably, a much reduced level of ubiquitinated E6TP1 was observed when E6 was coexpressed, coinciding with a markedly reduced E6TP1 protein level attributable to E6-induced degradation of E6TP1 (Fig. 3Citation , Lane 5). Importantly, treatment with MG132 led to a substantial accumulation of the ubiquitinated E6TP1 protein both in the absence and presence of E6 (Fig. 3Citation , Lanes 4 and 6). These results demonstrate that E6TP1 is ubiquitinated in vivo both basally and when destabilized by interaction with E6.



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  		Fig. 3. Ubiquitination of E6TP1 in the presence or absence of HPV16 E6. 293T cells (1 x 106) were cotransfected with pCMV-myc-E6TP1, pCR3.1–16E6, and HA-ubiquitin individually or in combination, as indicated. The total amount of DNA/dish was held constant by adding vector DNA. Forty h after transfection, cells were either mock-treated or treated with (final concentration, 50 µM) MG132 for 4 h and harvested in lysis buffer. Myc-E6TP1 was immunoprecipitated (IP) with anti-Myc antibody 9E10, fractionated on SDS-PAGE, and immunoblotted (IB) with anti-HA antibody 12CA5 to detect ubiquitinated E6TP1. The same blot was reprobed with anti-Myc antibody 9E10 to detect levels of E6TP1 protein.

 
Determination of the Minimal E6-binding Domain of E6TP1.
Our previous analysis showed that the COOH-terminal 194 amino acids of E6TP1 (residue 1590–1783) were sufficient for E6 binding (31) . To further narrow down the E6-binding domain on E6TP1, we generated 10 mutants with deletion within the previously characterized 194 amino acid region. The E6TP1 mutants were expressed as fusion proteins with GST and used in in vitro binding experiments with in vitro translated HPV16 E6. As shown in Fig. 4Citation , E6TP1–1704-1783, 1716–1783, and 1744–1783 bound HPV16 E6 similar to E6TP1. The E6TP1 mutant 1590–1783, shown previously to be competent at E6 binding (31) , clearly showed binding in these experiments. The remaining seven mutants were unable to bind E6. Notably, although the COOH-terminal 40 amino acids of E6TP1 (1744–1783) bound to HPV16 E6 efficiently, the COOH-terminal 25 amino acids (1759–1783) were unable to bind. These results defined the COOH-terminal 40 amino acids of E6TP1 as the minimal E6 binding domain. The doublet of E6 protein is attributable to the presence of two alternate methionine initiation codons (amino acids 1 and 8) of E6.



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  		Fig. 4. HPV16 E6 binding to various mutants of E6TP1. Upper panel, deletion mutants of E6TP1 used in binding assays. The GAP domain, PDZ domain, and leucine zipper domains are indicated. In vitro binding of E6 with various E6TP1 mutants was carried out by coincubating 35S-labeled E6 protein, generated by in vitro translation in wheat germ lysate, and various mutants of E6TP1 fused to GST. GST alone was used as a negative control. Bound E6 proteins were resolved by SDS-PAGE and visualized by fluorography. The input lane contains a 10% aliquot of E6 protein used in binding reactions. L, leucine; E, glutamate. +/-, binding/nonbinding.

 
E6-dependent Binding of E6AP to E6TP1.
High-risk HPV E6 proteins are known to target p53 (6 , 45 , 46) , hScrib (35) , Mcm7 (25) , and Bak (29) for degradation via the HECT domain-containing Ub ligase E6AP. In the presence of E6, these proteins associate with E6AP and get targeted for degradation by the Ub-proteasome pathway. To define the role of E6AP in E6-induced degradation, we wanted to first examine whether E6AP can bind to E6TP1. For this purpose, GST or GST-E6TP1 (1590–1783) was incubated with in vitro translated 35S-labeled E6AP either in the presence or in the absence of in vitro translated HPV16 E6. Notably, E6TP1 was unable to bind to E6AP in the absence of E6 (Fig. 5Citation , upper panel, Lane 3); however, easily detectable binding was observed in the presence of E6 (Fig. 5Citation , upper panel, Lane 4). Protein staining (lower panel) showed an equal amount of fusion proteins used in each binding reaction. These results showed that E6AP could be recruited to E6TP1 in the presence of E6.



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  		Fig. 5. In vitro binding of E6AP and E6TP1. HPV16 E6 and E6AP proteins were generated by in vitro translation in the presence of [35S]cysteine or methionine (NEN) using a wheat germ-based coupled transcription/translation system (Promega). The 35S-labeled in vitro-translated E6AP protein was incubated with 1 µg of GST or GST E6TP1 (1590–1783) fusion proteins noncovalently bound to glutathione-Sepharose beads in 300 µl of lysis buffer in the presence (Lane 4) or absence (Lane 3) of E6 for 2 h at 4°C, and bound 35S-labeled proteins were resolved by SDS- PAGE and visualized by fluorography (upper panel). The same gel was stained with Coomassie Brilliant Blue for the expression of GST-fusion proteins (lower panel). M.W., molecular weight.

 
E6 Induces E6AP-dependent Ubiquitination and Degradation of E6TP1 in Vitro.
To determine whether E6 can induce E6AP-dependent ubiquitination and degradation of E6TP1 directly in vitro, rabbit reticulocyte lysate-translated E6TP1 was incubated with purified human E1 and AtUbc8 E2 enzyme that supports E6AP function, with or without E6 and E6AP proteins. Although low levels of E6TP1 ubiquitination were seen without E6 and E6AP (Fig. 6Citation , compare Lane 1 and 2), the addition of E6 or E6AP further stimulated appearance of high molecular weight forms of E6TP1 (Lanes 3 and 4) together with reduction in the intensity of unmodified E6TP1 band. The addition of E6 and E6AP together further enhanced the accumulation of ubiquitinated higher molecular weight species of E6TP1 and the disappearance of the unmodified form (Lanes 5 and 6). Taken together, these results indicate that E6TP1 is a direct target for E6AP-mediated ubiquitination.



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  		Fig. 6. In vitro ubiquitination of E6TP1 in the presence of E6 and E6AP. In vitro translated 35S-labeled E6TP1 was mixed with purified E1, E2, Ub, and ATP in the presence or absence of purified E6 and/or E6AP, as indicated. The ubiquitination reactions were carried out as described in "Materials and Methods," and 35S-labeled E6TP1 was resolved by SDS- PAGE and visualized by fluorography. M.W., molecular weight.

 
A Dominant-Negative E6AP Mutant Inhibits the E6-induced Degradation of E6TP1.
To further assess the role of E6AP in E6-induced E6TP1 degradation in vivo, we cotransfected 293T cells with E6TP1 and a well-characterized dominant-negative E6AP mutant C833A (50) or the wild-type E6AP, together with myc-tagged 16 E6. The levels of E6TP1 or E6 proteins were determined using anti-E6TP1 antiserum or anti-myc immunoblotting, respectively. As shown in Fig. 7Citation , cotransfection with HPV16 E6 reduced the levels of E6TP1 protein (upper panel, compare Lanes 2 and 3). Notably, the levels of E6TP1 were significantly reduced when wild-type E6AP was cotransfected with E6 (upper panel, compare Lanes 3 and 4). In contrast, cotransfection with E6AP-C833A mutant led to a marked inhibition of E6-induced decrease in the levels of E6TP1 protein when compared with E6 plus E6TP1 (upper panel, compare Lanes 3 and 5). Taken together, these results clearly demonstrate that E6AP mediates E6-induced degradation of E6TP1.



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  		Fig. 7. In vivo degradation of E6TP1 in the presence of wild-type or mutant E6AP. 293T cells (1 x 106) were cotransfected with pCMV constructs encoding E6TP1, HA-tagged E6AP, or E6AP mutant (C833A) with myc-tagged 16 E6. Each dish was cotransfected with GFP construct for transfection efficiency control. The total amount of DNA was kept constant by adding vector DNA. Forty-eight h later, the cell lysates were subjected to immunoblotting with anti-E6TP1 antiserum to examine the levels of E6TP1 (upper panel). The same blot hybridized with anti-HA antibody shows the presence of almost equal levels of E6AP in lanes where E6AP or its mutant was transfected (second panel). E6 expression was assessed by Western blotting using anti-myc antibody (third panel). Anti-GFP Western blotting (lower panel) controls for transfection efficiency. M.W., molecular weight.

 

   DISCUSSION
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
E6TP1 was identified previously as a direct HPV16 E6-binding protein using the yeast two-hybrid system (31) . Significantly, the high-risk but not the low-risk HPV E6 proteins bind to E6TP1 and target it for degradation (31) . High-risk HPV E6 proteins are known to induce the degradation of a number of their targets by promoting their ubiquitination and subsequent targeting to the proteasome (6 , 25 , 28 , 29 , 35 , 45, 46) . It was, therefore, reasonable to hypothesize that high-risk HPV E6 proteins induce the degradation of E6TP1 via the Ub pathway. To provide direct support for this hypothesis, we first demonstrated that E6-induced degradation of E6TP1 is inhibited by treating cells with a proteasome inhibitor MG132. A second line of evidence for Ub-dependent E6TP1 degradation by E6 was provided by our analysis of a CHO cell mutant CHO-ts20 (53) , which contains a temperature-sensitive defect in Ub-activating enzyme, E1. We observed that under our experimental conditions, E6 induced comparable degradation of E6TP1 as well as p53 (a known E6 target), and degradation of both proteins was significantly decreased when E1 was inactive. Finally, we showed that E6TP1 is ubiquitinated. The ubiquitinated E6TP1 was readily detected both in the presence or absence of HPV E6 protein, although less ubiquitinated E6TP1 was observed in the presence of HPV16 E6 because of the enhanced degradation. We observed a smear rather than clear-cut bands of ubiquitinated E6TP1. Notably, polyubiquitination of many cytosolic proteins, such as E6TP1, leads to smearing as polyubiquitination involves elongation of Ub chains through attachment of additional Ub units to lysine residues such as K48. Because each protein may have multiple Ub attachment sites, the various lengths of polyubiquitin chains on these sites result in smearing (54) . Given the HPV E6-induced ubiquitination and degradation of E6TP1 in our studies, it will be of obvious interest to determine whether high-risk HPV E6 oncoproteins reduce the steady-state levels of the endogenous E6TP1 in E6-immortalized cells and/or HPV-positive tumor cells. Our initial attempts to address this question have been unsuccessful because of the inability of the existing anti-E6TP1 antibodies to detect the endogenous E6TP1 (data not shown). Notably, MG132 treatment also inhibited the degradation of E6TP1, even in the absence of HPV E6, implicating that even normal turnover of E6TP1 protein involves Ub-proteasome pathway. Furthermore, stabilization of E6TP1 in the absence of E6 was noted in CHO-ts20 cells at 42°C, and E6TP1 was found to be ubiquitinated in the absence of E6. These results indicate that similar to other E6 targets such as p53, E6TP1 levels are normally regulated by the Ub-proteasome pathway.

Previous studies have shown that degradation of several cellular targets of high-risk HPV E6 proteins, including p53, hScrib, Bak, myc, and Mcm7, involve the HECT domain-containing ubiquitin ligase E6AP (45 , 46 , 35 , 29 , 28 , 25) . Notably, p53 and myc use distinct ubiquitin ligase in the absence and presence of E6 (45 , 46 , 28) . The degradation of endogenous p53 is mediated by mdm2 (55, 56, 57) . Additionally, the degradation of myc in the absence of E6 involves E3{alpha} and E3-Fos (28) . Interestingly, two E6 targets, Mcm7 and Bak, also use E6AP for their normal turnover, even in the absence of E6 (25 , 29) . Our results strongly support a role for E6AP in E6-induced degradation of E6TP1. We defined the minimal E6-binding domain in E6TP1, which is essential for E6-induced E6TP1 degradation. Next, we showed that E6AP can form a complex with E6TP1 in the presence of E6. Furthermore, we showed that purified E6AP can enhance the ubiquitination and degradation of E6TP1 in vitro in the presence of E6. Because these experiments were done using rabbit reticulocyte lysates that contain E6AP (45) , we observed enhanced ubiquitination with E6 alone, even in the absence of exogenously added E6AP. Finally, we showed that a dominant-negative mutant of E6AP, shown previously to prevent E6-induced degradation of p53 when expressed in E6-expressing cells, markedly reversed the E6-induced degradation of E6TP1. These data demonstrate that E6-induced degradation of E6TP1 via the ubiquitination pathway is mediated by the ubiquitin ligase, E6AP.

Our studies reported here have also defined the COOH-terminal 40-amino acid region of E6TP1 as the minimal E6 binding domain. Notably, this sequence shows no significant homology with the defined E6 binding domains of other E6-binding proteins, such as the 18-amino acid E6-binding domain in E6AP (58) , the 13-amino acid sequence in E6BP (59) , the 13-amino acid LD1 sequence of paxillin (60) , or the L2G box in Mcm7 (25) . Previous studies have revealed three conserved motifs, including Lhx{phi}Ls (59) , E/D-L/I/F-L/V-G (61) , and the L2G box (S/TXXXLLG; Ref. 25 ). None of these motifs were found within the 40-amino acid E6-binding domain of E6TP1. Although the E6-binding domain of E6TP1 includes part of the leucine zipper (residue 1744–1758), this portion of the leucine zipper alone is not sufficient to bind E6 but is required for binding to E6. Further mutational and structural studies will be needed to further refine the E6-binding domain of E6TP1 and to define whether it has a secondary or tertiary structural similarity with other E6-binding motifs.

E6TP1 is a novel target of E6-induced degradation, and sequence homology predicts it to be a GAP toward Ras-related small G-protein of the Rap family. Indeed, our recent analyses have demonstrated that E6TP1 functions as a GAP toward Rap1 as well as Rap2.6 These findings suggest the role of Rap G-protein signaling cascade in the homeostasis of normal epithelial cells and in HPV-induced cancer.


   ACKNOWLEDGMENTS
 
We thank Drs. Peter Howley (Harvard Medical School, Boston, MA) for E6AP plasmids, Ger Strous (University Medical Center Utrecht, Utrecht, the Netherlands) for CHO-ts20 cell line, Dirk Bohmann (EMBO, Heidelberg, Germany) for the HA-ubiquitin expression construct, and M. Ishibashi (Aichi Cancer Center, Nagoya, Japan) for the pEF-myc-16E6 construct.


   FOOTNOTES
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported by NIH Grants CA81076 and CA70195 (to V. B.) and CA87986, CA75075, and CA76118 (to H. B.). A. K. is a recipient of a fellowship from Massachusetts Department of Public Health. Back

2 Co-first author. Back

3 To whom requests for reprints should be addressed, at Department of Radiation Oncology, New England Medical Center, 750 Washington Street, Boston, MA 02111. Phone: (617) 636-4776; Fax: (617) 636-6205; E-mail: vband{at}lifespan.org. Back

4 The abbreviations used are: HPV, human papillomavirus; GAP, GTPase activating protein; E6TP1, E6 targeted protein 1; HA, hemagglutinin antigen; Ub, ubiquitin; GFP, green fluorescent protein; GST, glutathione S-transferase; CHO, Chinese hamster ovary. Back

5 Unpublished data. Back

6 Q. Gao, L. Singh, A. Kumar, S. Srinivasan, and V. Band, unpublished data. Back

Received 12/ 3/01. Accepted 4/ 2/02.


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Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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