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[Cancer Research 60, 4289-4298, August 1, 2000]
© 2000 American Association for Cancer Research

Virology

Human Papillomavirus Type 16 E6 and E7 Proteins Inhibit Differentiation-dependent Expression of Transforming Growth Factor-ß2 in Cervical Keratinocytes

Matthias Nees, Joel M. Geoghegan, Peter Munson, Vinayakumar Prabhu, Yun Liu, Elliot Androphy and Craig D. Woodworth1

Laboratory of Cellular Carcinogenesis and Tumor Promotion, National Cancer Institute [M. N., J. M. G., C. D. W.], Center for Information Technology [P. M., V. P.], NIH, Bethesda, Maryland, 20892, and Department of Dermatology, New England Medical Center and Tufts University School of Medicine, Boston, Massachusetts 02111 [Y. L., E. A.]


   ABSTRACT
Top
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Infection with high-risk human papillomaviruses (HPVs) represents a major risk factor for the development of cervical cancer. The HPV-16 E6 and E7 proteins are highly expressed in differentiating keratinocytes, where they inactivate the p53 and retinoblastoma (pRb) proteins, two important transcriptional regulators. We have used cDNA expression arrays to identify global alterations in gene expression induced by E6 and E7 in differentiating cultures of human cervical keratinocytes. We show that E6 and E7 decrease expression of TGF-ß2 mRNA and alter expression of multiple TGF-ß-responsive genes involved in cell cycle regulation, apoptosis, and tissue remodeling. E6 and E7 inhibited expression of TGF-ß2 RNA 7-fold (relative effectiveness, E6/E7 > E6 > E7 > control) and decreased secretion of biologically active TGF-ß2 by 70–80% (reduced from 70 to 10 pg/106cells/24 h). Down-regulation occurred through p53- and pRb-dependent pathways. In contrast, E6 and E7 did not alter expression of TGF-ß1 and TGF-ß3. Down-regulation of TGF-ß2 was biologically relevant because the addition of recombinant cytokine (10–200 pg/ml) to E6/E7-expressing cells restored expression of TGF-ß-responsive genes, inhibited growth of keratinocytes, and decreased immortalization by E6 and E7. These results suggest that TGF-ß2- and TGF-ß-responsive genes are important targets for the HPV-16 E6 and E7 oncoproteins in differentiating cervical keratinocytes.


   INTRODUCTION
Top
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The TGF2 -ß family of growth factors are important regulators of gene expression during embryogenesis, differentiation, and wound healing. Members of this family include TGF-ßs 1–5, bone morphogenetic proteins, growth/differentiation factors, and inhibins/activins. Human keratinocytes express three TGF-ß isoforms (1, 2, 3) , which have multiple effects on gene expression and cellular differentiation. TGF-ßs stimulate tissue remodeling, block cell cycle progression of epithelial cells, and inhibit the immune response (reviewed in Ref. 1 ). Regulation of TGF-ß activity is complex and occurs at multiple levels, including regulation of transcription, secretion of the latent inactive precursor, and cleavage of the latent complex to form biologically active TGF-ß (reviewed in Ref. 2 ). TGF-ßs activate a binary system of membrane receptors that signal via serine threonine kinase activity (3 , 4) . They function as tumor suppressors in animal models of carcinogenesis, and loss of one or both alleles stimulates malignant progression (5) . Loss or mutation of TGF-ß receptors also occurs during human carcinogenesis (4) , and most carcinoma cell lines are resistant to growth inhibition by TGF-ß in vitro (6) .

HPVs are small DNA tumor viruses that replicate in differentiating epithelial cells of the epidermis and anogenital tract. The E6 and E7 viral genes are expressed at low levels in proliferating basal cells, but transcription is activated as cells enter the terminal differentiation pathway. The E6 and E7 proteins alter cell differentiation (7 , 8) , reactivate host DNA synthesis (9) , and stimulate cell cycle progression (10) . This allows the virus to use host DNA synthetic enzymes to replicate the viral genome (reviewed in Ref. 11 ). Although many HPV types induce benign warts and papillomas, infection with certain high-risk types (HPV-16, HPV-18, HPV-31, and HPV-45) is a major risk factor for the development of cervical cancer (12) . The E6 and E7 genes are retained and expressed in most cervical carcinomas, and continued expression is required to maintain the malignant phenotype (13) .

An early step in HPV-associated carcinogenesis is perturbation of cellular gene expression by the HPV E6 and E7 oncoproteins. E6 binds to several cellular proteins including E6BP (14) and E6AP, a protein-ligase of the ubiquitin pathway of proteolysis (15) . E6/E6AP complexes target the p53 tumor suppressor protein for rapid degradation by the proteasome (15) . p53 integrates responses to genotoxic stress and DNA damage with cell cycle control and apoptosis. Therefore, loss of p53 function leads to increased genetic instability. p53 is a transcriptional activator, and numerous p53-responsive genes have been identified (16) . The HPV-16 E7 protein binds pRb and members of the pRb family (17) . Interaction occurs primarily with the hypophosphorylated form of pRb, causing release of active E2F transcription factors, which in turn stimulates expression of multiple genes involved in cell cycle progression. The E7 protein also binds and alters the function of AP-1 transcription factors, which regulate cellular gene expression (18 , 19) . E6 and E7 exert overlapping effects on cell cycle control, and in combination, they efficiently immortalize human keratinocytes (20, 21, 22, 23, 24) .

The p53 and pRb proteins are important transcriptional regulators. Thus, inactivation of their function by E6 and E7 is likely to alter keratinocyte gene expression significantly. We used cDNA array technology to identify global changes in gene expression in differentiating cultures of primary human cervical keratinocytes infected with retroviruses encoding HPV-16 E6 and E7. cDNA microarrays allow high-throughput, parallel expression analysis of thousands of cellular genes in a single experiment (25 , 26) . Our results showed that E6 and E7 influenced expression of a large number of cellular genes, including TGF-ß2. TGF-ß2 was induced during keratinocyte differentiation, and both E6 and E7 blocked this induction and altered expression of multiple TGF-ß-regulated genes. We describe inhibition of expression and release of biologically active TGF-ß2 from cervical keratinocytes by E6 and E7 and identify the biological consequences of TGF-ß2 inhibition.


   MATERIALS AND METHODS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Culture.
Primary cultures of human ectocervical keratinocytes were established from fresh cervical tissue obtained after hysterectomy because of fibroids or endometriosis (20) . Keratinocytes were maintained in serum-free MCDB153-LB medium (Life Technologies, Gaithersburg, MD), and cultures were sampled for medium, RNA, or protein extraction when 95% confluent. In some experiments, cells were induced to undergo terminal differentiation by removal of growth factors (epidermal growth factor, bovine pituitary extract, insulin, hydrocortisone, triiodothyronine, and transferrin) for 24 h, followed by addition of 1.4 mM calcium for an additional 24 h. Primary cultures of human ectocervical keratinocytes were infected with high titer retroviruses expressing the neomycin resistance gene only (pLXSN) or the E6 and E7 oncogenes of the high-risk HPV type 16 and low-risk HPV 6b (27) . Retroviruses producing mutant E6 proteins with single amino acid exchanges (F2V and F125V) have been described (28) . A retrovirus producer line expressing a dominant-negative p53 miniprotein was established in the PA317 packaging cell line, using a recombinant vector kindly provided by Moshe Oren (29) . After infection, keratinocytes were selected in 200 µg/ml G418 for 2–4 days and were subcultured at least once prior to extraction of protein or RNA. Selected primary cultures were analyzed by RT-PCR for expression of E6 or E7 genes of several high-risk HPV types and found to be negative.

Cell Proliferation Assays.
Primary keratinocytes were added to 100-mm culture dishes at a density of 0.5–1.0 x 105 cells/dish. After 24 h, cells were treated with various concentrations of recombinant human TGF-ß2 or TGF-ß1 (R&D Systems, Minneapolis, MN). Fresh medium with recombinant cytokine was added every other day, and cells were counted after 8 days using a Coulter counter. All proliferation experiments were repeated three times in duplicate dishes.

Cell Immortalization Assay.
Keratinocytes expressing HPV-16 E6/E7 were seeded on 150-mm plastic dishes at a density of 0.5–1.0 x 104 cells/dish. Cultures at higher passage numbers that showed evidence of crisis were seeded at densities of 2.5–5.0 x 104 cells/dish. After 24–48 h, cells were treated with various concentrations of recombinant human TGF-ß2 (50, 100, 200, and 1000 pg/ml medium) every second day, and TGF-ß2 treatment was continued until the first rapidly and continuously growing colonies appeared (usually after 10–14 days). For quantification of immortalization, colonies were allowed to grow to a diameter of ~1 cm in the absence of TGF-ß2. Selected colonies were picked and cultured for three more passages to confirm that they were immortal.

RNA Isolation and Purification.
RNA was extracted from monolayer cultures that were growing or that were induced to differentiate as described above. For RNA extraction, cells were washed with PBS and lysed with Trizol (Life Technologies), and DNA was removed by digestion with RNase-free DNaseI (Life Technologies) or DNase RQ1 (Promega, Inc., Palo Alto, CA.). DNase, proteins, and other contaminants were removed by two extractions with buffer-saturated phenol/chloroform and precipitation with 1.5 volumes of isopropanol. The integrity of RNA was analyzed by electrophoresis, and samples were stored at -80°C.

cDNA Synthesis and cDNA Array Hybridization.
The human and human Cancer cDNA arrays on nylon membranes were purchased from Clontech, Inc. (Palo Alto, CA; ATLAS cDNA arrays). The protocol for cDNA synthesis was modified from the manufacturer’s recommendations. Briefly, 32P-labeled cDNAs were synthesized from DNase-treated total RNA. We used total RNA because polyadenylated RNA often resulted in severe background and variability in signal intensity from experiment to experiment. Five to 10 µg of RNA in a volume of 5 µl were denatured in the presence of 1 µl of first-strand primer mix at 70°C for 5 min in a thermocycler (GeneAmp PCR System 9700; PE Applied Biosystems, Foster City, CA). RNA was incubated for 5 min at 48°C to allow primer annealing. Reverse transcription was performed using 5 µl of [{alpha}-32P]dATP (3000 Ci/mmol; Amersham Life Science, Cleveland, OH), 50 µM dNTP mix lacking dATP, 1 µl 100 mM DTT, and 200 units Superscript MMLV Reverse Transcriptase (Life Technologies) in a total volume of 20 µl. The reaction was incubated at 48°C for 60 min, and an additional 200 units of Superscript were added after the first 30 min. Non-incorporated radioactive nucleotides were removed by spin column purification. Approximately 15–30 x 106 cpm were incorporated in the final product. Radioactive cDNA probes were denatured for 20 min at 68°C by adding one-tenth volume of 1 M NaOH, 10 mM EDTA, neutralized with 1 M sodium phosphate buffer (pH 7.0), and 7.5 µg of Cot-1 DNA and 5 µg poly(A)-DNA were added. Membranes were prehybridized at 68°C for 6–8 h. Hybridization was performed overnight at 68°C in a total volume of 7.5–10 ml of ExpressHyb hybridization solution and a final probe concentration of ~2.5–5 x 106 cpm/ml. Membranes were washed three times at 68°C with 2x SSC, 1.0% SDS, and twice with 0.2x SSC, 0.5% SDS for 10 min each, sealed in plastic pouches, and exposed to phosphor imager screen cassettes. Screens were exposed for 1, 3, and 10 days and analyzed using the Storm Phosphor Imager System 860 (Molecular Dynamics, Sunnyvale, CA). Computer analysis and normalization of array hybridization data were performed using the P-SCAN software.

RNA from keratinocytes infected with vector retroviruses was compared with RNA from cells infected with one or both viral oncogenes of HPV-16. Cells from three to five different individuals were pooled to reduce biological variation between samples. For both the Normal Human and Human Cancer cDNA arrays, experiments were repeated three times for each oncogene combination (total, 24 array hybridizations). Comparisons between growing and differentiating cells, or between untreated cultures and cultures treated with TGF-ß2, were each performed twice using independent samples (total, 4 hybridizations). Differential gene expression data were considered significant if the average factor of differential mRNA expression exceeded 2.0.

RT-PCR.
Fifty to 100 µg of total RNA were purified using lithium chloride precipitation. An equal volume of 5 M LiCl, 0.1 M Tris-HCl (pH 7.4) was added to samples, and RNA was precipitated overnight at 4°C, centrifuged at 15000 rpm, washed with 70% ethanol, dried by evaporation, and resuspended in diethyl pyrocarbonate-treated water. For cDNA synthesis, 5–10 µg of purified total RNA were reverse transcribed for 1 h at 42°C in a volume of 50 µl, containing 500 µM individual dNTPs; 10 µM DTT, 1.25 µM oligo-dT primer (T16–18), 20–40 units RNasin (Promega), and 75 units Superscript Reverse Transcriptase II (Life Technologies) in 1x RT buffer. RT reactions were diluted 5–10-fold, and for each PCR, one-tenth (or 2 µl) of diluted RT-mix was included. Amplifications were performed in a volume of 20 µl in 96-well plates using a GeneAmp 9700 thermocycler, in the presence of 50 µM dNTPs, 0.5 µM of each oligonucleotide primer, 1.75 mM magnesium chloride, and 1 unit of AmpliTaq Gold (PE Applied Biosystems). To avoid saturation or plateau effect of amplification, PCR was limited to a total of 20–25 cycles. Each reaction was performed twice using independent reverse transcription reactions to confirm reproducibility. For direct quantification of PCR products, 1.0–2.0 µCi of radioactive [{alpha}-32P]dCTP was incorporated, and PCR products were separated on 6% denaturing polyacrylamide/50% urea gels. DNA was detected and quantified by autoradiography or using a STORM Phosphor Imager (Molecular Dynamics, Inc.). Oligonucleotide primers were designed using the GCG (Genetics Computer Group, Madison, Wisconsin) Program Package. Table 1Citation summarizes all primer combinations used for RT-PCR analysis. Northern blot analyses were performed as described (30) .


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  		Table 1 Primer combinations used for RT-PCRa

 
ELISA Analysis of Cell-associated and Secreted TGF-ß1 and TGF-ß2 Proteins.
For analyses of cell-associated TGF-ßs, cells were lysed in buffer containing 150 mM NaCl, 50 mM Tris-HCl (pH 7.5), 1.5 mM MgCl2, 1% glycerol, 1% Triton X-100, 5 mM EGTA, and protease inhibitors AEBSF, leupeptin, and aprotinin (100 µg/ml each). Protein samples were diluted to a concentration of 0.5 µg protein/µl, acid-activated by incubation with 0.2 volume of 1 M HCl for 10 min at room temperature and neutralized with 0.2 volume of 1 M HEPES/1.2 M NaOH. For ELISA analysis, protein samples and TGF-ß2 standards were diluted in lysis buffer. For analyses of secreted TGF-ßs, cell conditioned medium (5 ml/10 cm dish) was supplemented with 10 mM EDTA and protease inhibitors including [4-(2 aminoethyl)-benzenesulfonyl fluoride hydrochloride (AEBSF, 1 mM), aprotinin, pepstatin A, and leupeptin (10 µg/ml each), and centrifuged for 5 min at 5000 x g to remove cell debris. Samples were frozen at -80°C in polypropylene tubes and analyzed within 2 weeks. After sampling media, cells were counted for normalization of cytokine secretion using a Coulter counter. Alternatively, cells were lysed, and total protein concentration was determined by the BCA protein assay (Pierce, Rockland, IL), and protein data were used for normalization of ELISA. Samples and standards were added to wells in duplicate (200 µl/well) and incubated overnight at 4°C. Plates were washed five times, and the protocol continued according to the manufacturer’s recommendations (Quantikine sandwich ELISA; R & D Systems).

Immunoblotting.
Western blot analysis was performed as described (10) using polyclonal antibodies to HPV-16 E7 (Zymed, San Francisco, CA) and p53 (DO-1; Santa Cruz Biotechnology, Santa Cruz, CA).


   RESULTS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
HPV-16 E6 and E7 Down-Regulate TGF-ß2 and TGF-ß-responsive Genes in Differentiating Cervical Keratinocytes.
We have used cDNA arrays to examine changes in the overall pattern of gene expression in human cervical keratinocytes after infection with retroviruses encoding HPV-16 E6 and E7 genes. A large number of genes were differentially expressed in E6/E7-infected cells in comparison with vector controls. Approximately 50 cellular genes were up- or down-regulated >2-fold (~5% of all genes on arrays). These genes were clustered into functional groups, revealing important differences in global gene expression induced by HPV16 E6 and E7 (Table 2)Citation . Data in column 2 (E6/E7) represent the average of three independent experiments and were highly reproducible. Growth factors and cytokines were a prime target for HPV oncogenes, including insulin like growth factor-1 receptor and heparin-binding epidermal growth factor. E6 and E7 also decreased expression of genes involved in tissue remodeling and wound healing including the MMPs 2, 3, 7, 9, and 10–13, fibronectin, and PAI-1 and PAI-2. Interestingly, E6 and E7 altered expression of multiple apoptosis-related genes known to decrease susceptibility to programmed cell death including survivin (inhibitor of apoptosis hIAP4). As expected, E6 and E7 also stimulated expression of multiple genes that regulate cell cycle progression including cyclins A and B, cyclin-dependent kinases 1, 2, and 3, and DNA synthetic enzymes as described previously (31) . Expression of DNA repair enzymes including RAD 50, RAD 51, and RAD 52 were also induced by HPV.


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  		Table 2 Genes differentially regulated by E6 and E7 cultured cervical keratinocytes

 
E6 and E7 Differentially Regulate TGF-ß2, TGF-ß-responsive Genes, and Differentiation-induced Genes.
One of the most striking observations was that many of the E6/E7-regulated genes that we identified have been shown previously to be TGF-ß inducible. Because E6 and E7 strongly reduced expression of TGF-ß2, we examined whether other E6/E7-responsive genes were also regulated by TGF-ß in cultured cervical keratinocytes. For these experiments, normal keratinocytes were treated with recombinant TGF-ß2 (1.0 ng/ml for 24 h), and cDNA arrays were used to compare patterns of gene expression in untreated and treated cells. TGF-ß2 had an inverse effect on gene expression compared with the HPV-16 oncogenes in two independent experiments (Table 2Citation ; TGF-ß2). Many genes that were down-regulated by HPV-16 E6/E7 were also identified as transcriptionally regulated by TGF-ß2.

The cDNA array experiments showed that effects of E6 and E7 on cellular gene expression were most pronounced under conditions promoting keratinocyte differentiation (removal of growth factors and addition of 1.4 mM calcium). E6 and E7 induced similar changes in proliferating cells, although the magnitude of changes were smaller (data not shown). This suggested that E6 and E7 might alter keratinocyte gene expression in part by delaying or blocking differentiation. Therefore, cDNA arrays were used to compare cellular gene expression in proliferating keratinocytes versus cells that were induced to undergo differentiation for 24 h. A large number of genes (~15% on arrays) showed altered mRNA expression during the process of differentiation (Table 2)Citation . Differentiation was confirmed by decreased expression of keratins 5, 14, and 6 (data not shown), which are down-regulated during terminal differentiation in vivo. Most genes that were down-regulated by E6/E7 were up-regulated during terminal differentiation, and one of the most strongly induced was TGF-ß2. Differentiation-dependent induction was also observed for many TGF-ß-responsive genes such as MMP 7, 10–12, Fas/APO-1, and Fas ligand. These results suggest that E6 and E7 may alter expression of TGF-ß and TGF-ß-inducible genes indirectly through their ability to block or delay keratinocyte differentiation.

Confirmation of Differential Gene Expression by Northern Analysis and RT-PCR.
Fig. 1ACitation shows one sector of a typical cDNA array comparing gene expression in vector-infected controls (left) and E6/E7 infected keratinocytes (right). This sector contained TGF-ßs 1, 2, and 3 as well as multiple cytokines and growth factors, and it showed the most striking patterns of differential gene expression. Differences in the relative signal intensity of each TGF-ß isoform in response to E6, E7, or E6/E7 are shown in Fig. 1BCitation . Expression of TGF-ß2 mRNA was down-regulated 5–7-fold in keratinocytes expressing E6/E7 and 2–3-fold in cells expressing E6 or E7 individually (Fig. 1BCitation ). Expression of other TGF-ß isoforms including TGF-ßs 1 and 3 were not significantly changed by E6, E7, or E6/E7. Down-regulation of TGF-ß2 mRNA was confirmed using Northern blot hybridization (Fig. 1CCitation ), semiquantitative RT-PCR (Fig. 1DCitation ), and RNase protection (data not shown). All three methods detected similar levels of down-regulation of TGF-ß2 RNA. However, no differences in TGF-ß1 or TGF-ß3 expression were detected. Interestingly, retroviral expression of the low risk HPV-6b E6 and E7 proteins decreased TGF-ß2 RNA expression with much lower efficiency, and effects were not always reproducible (Fig. 1DCitation ).



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  		Fig. 1. HPV-16 E6 and E7 oncoproteins down-regulate expression of TGF-ß2 RNA in cultured cervical keratinocytes. A, sector F of the Human ATLAS cDNA expression array showing differential expression of TGF-ß1 and TGF-ß2 RNA (box) in vector control and HPV-16 E6/E7 expressing cells. B, summary of array hybridization data showing effects of specific HPV proteins on expression of TGF-ßs 1, 2, and 3. Data show signal intensity relative to the vector control (PLXSN) and represent the means from three independent array hybridizations; bars, SD. C, Northern blot analysis of TGF-ß2 expression in primary cultures of cervical keratinocytes infected with retroviruses encoding HPV oncogenes and cells immortalized by HPV-16 E6/E7. D, RT-PCR analysis of TGF-ß1 and 2 expression in keratinocytes infected with HPV-16 or HPV-6b (low-risk) E6 and E7 genes. NC, a control in which the reverse transcriptase was heat inactivated prior to the reaction.

 
Genes that were up- or down-regulated by E6/E7 in cDNA arrays were examined by semiquantitative RT-PCR to confirm differential expression and to assess whether changes were attributable to E6, E7, or E6/E7 (for primer pairs, see Table 1Citation ). Fig. 2Citation shows representative RT-PCR analyses for several of these genes including the MMPs 2, 3, and 10, cyclins A and B, PAI-1 (a well-established TGF-ß-inducible gene), TRAIL, and GAPDH. These experiments confirmed that each gene was differentially regulated by E6/E7 and indicated that regulation was generally more dramatic in response to E6/E7 than to E6 or E7 alone. The results also confirmed that many but not all of the E6/E7 regulated genes were TGF-ß responsive. Importantly, addition of recombinant TGF-ß2 to infected cultures did not alter expression of E6 or E6/E7 mRNAs, which in retrovirus-infected cells are regulated by the mouse mammary tumor virus long terminal repeat (see below). Western blot analyses confirmed that retrovirus-infected keratinocytes actually expressed the HPV-16 E7 protein (Fig. 2BCitation ). Although the E6 protein was not detected with several commercially available antibodies, E6 or E6/E7-infected cells had dramatically reduced levels of p53, a known consequence of E6 expression.



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  		Fig. 2. Confirmation of altered gene expression by RT-PCR (A) and Western analysis (B). A, RT-PCR analyses showing differential regulation of TGF-ß2 RNA by HPV-16 E6 and E7. Keratinocytes were infected with retroviruses encoding vector, E6, E7, or E6/E7 and were maintained for 24 h in the absence or presence of TGF-ß2 (1 ng/ml). NC, negative control with RNA but heat-inactivated reverse transcriptase. TRAIL, TNF-related apoptosis inducing ligand/Apo-2 ligand; GAPDH served as a control for loading. B, Western analysis of HPV-16 E7 and p53 protein in retrovirus-infected keratinocytes.

 
HPV Oncogenes Down-Regulate Expression of Cell-associated TGF-ß2 Protein.
Consistent with the mRNA data, cervical keratinocytes expressing HPV-16 E6 and/or E7 produced significantly less cell-associated TGF-ß2 protein (Fig. 3ACitation ). As observed for mRNA expression, the combination of HPV-16 E6 and E7 was significantly more efficient at reducing TGF-ß2 than either oncogene alone. No differences were detected for TGF-ß1 protein expression (data not shown). Interestingly, localization and expression patterns for TGF-ß1 and TGF-ß2 proteins were strikingly different. About 33% of the cell-associated TGF-ß2 protein was localized in the extracellular matrix and was removed by a brief trypsin digest prior to cell extraction. The remainder was intracellular and resistant to trypsin treatment. The ratio of matrix-associated to intracellular TGF-ß2 was increased in keratinocytes expressing E7 or E6/E7 (Fig. 3ACitation ). In contrast, TGF-ß1 protein was almost exclusively intracellular and not associated with the extracellular matrix; trypsin digest did not reduce the amount of cellular TGF-ß1 detected by ELISA (Fig. 3BCitation ). This observation might be related to differences in activation of TGF-ß1 and TGF-ß2 observed in vitro.



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  		Fig. 3. HPV-16 E6 and E7 proteins down-regulate levels of cell-associated TGF-ß2. A, keratinocytes were infected with retroviruses encoding E6 and/or E7 genes and levels of intracellular ({square}) and matrix-associated () TGF-ß2 were determined by ELISA. B, levels of intracellular and matrix-associated TGF-ß1. C, relative levels of total cell-associated TGF-ß2 in keratinocytes infected with retroviruses encoding vector-only (PLXSN); HPV-16 E6 wild type (wt); a HPV-16 E6 mutants that binds E6BP but does not degrade p53 (E6mutF2V), or efficiently degrades p53 (E6mutF125V); a dominant-negative p53 mini protein (p53 DD); HPV-16 E7 and 16 E6/E7. Values represent the means of three individual experiments; bars, SD.

 
Down-Regulation of Cell-associated TGF-ß2 Protein by HPV-16 E6 Was Dependent on Binding and Inactivation of p53.
We examined cell-associated TGF-ß2 after infection with retroviruses encoding several mutant HPV-16 E6 proteins that differed in their ability to bind p53 or other cellular proteins (28) . One mutant, which was deficient in p53 degradation in vitro but bound to E6BP (E6mut F2V), failed to decrease intracellular TGF-ß2 protein (Fig. 3CCitation ). One E6 mutant that retained the ability to degrade p53 (E6mut F125V) decreased levels of intracellular TGF-ß2 as efficiently as wild-type HPV-16 E6. To further investigate the importance of p53, we constructed retroviruses encoding a dominant-negative p53 miniprotein (29) . Expression of this dominant-negative mutant decreased intracellular TGF-ß2 as strongly as wild-type E6.

HPV-16 E6 and E7 Decrease Secretion of Biologically Active TGF-ß2.
Conditioned medium from keratinocytes infected with E6, E7, E6/E7, or vector-only viruses (pLXSN) was used for detection of biologically active TGF-ß1 and TGF-ß2. Medium was subsequently acid-treated to activate and detect latent TGF-ß1 and TGF-ß2 because the ELISA only detects the mature, active cytokine. Secretion of biologically active TGF-ß2 was strongly induced in differentiating keratinocytes relative to growing cells (Fig. 4ACitation ). E6 and E7 down-regulated secretion of biologically active TGF-ß2 in both rapidly growing and differentiating cultures. Differentiating keratinocytes secreted an average of 112 pg of total TGF-ß2/106 cells/24 h. Most of the secreted protein was biologically active TGF-ß2 (Fig. 4BCitation ). In differentiating cells, secretion of active TGF-ß2 was significantly reduced by HPV-16 E6 and E7 oncogenes. The combination of E6 and E7 was most efficient (7-fold reduction), suggesting a cooperative interaction. E6 and E7 also decreased secretion of latent TGF-ß2 (Fig. 4BCitation ). In contrast to differentiating cultures, growing keratinocytes activated only 15% of the secreted TGF-ß2 protein, and low levels of biologically active TGF-ß2 were detected (10–24 pg/106 cells/24 h; data not shown). In most experiments with growing cells, expression of E6 and E7 resulted in a complete loss of active TGF-ß2 secretion detectable by ELISA. In contrast, cervical keratinocytes secreted TGF-ß1 exclusively as the latent, inactive form and E6 and E7 did not alter secretion (data not shown).



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  		Fig. 4. HPV-16 E6 and E7 proteins decrease secretion of biologically active and latent TGF-ß2. A, detection of biologically active TGF-ß2 by ELISA analysis of culture medium from growing ({square}) or differentiating HPV-infected keratinocytes (). B, detection of biologically active ({square}) and latent () TGF-ß2 by ELISA of medium from differentiating cultures of HPV-infected keratinocytes. C, relative levels of biologically active TGF-ß2 secreted by differentiating keratinocytes infected with retroviral constructs. HPV-6b E6 (6b E6), HPV-16 E6 (16 E6), p53 dominant-negative mutant (p53 DD), p53 sense, HPV-6b E7 (6b E7), HPV-16 E7 mutant that fails to bind pRb (16E7mut p24gly; 16 E7 mut), HPV-16 E7 (16 E7), transcription factor E2F-1, HPV-16 E7/E7, and keratinocytes infected with vector approaching replicative senescence. Bars, SD.

 
Treatment with all-trans retinoic acid stimulates secretion of biologically active TGF-ß2 from keratinocytes (32) . We found that HPV-16 E6/E7 genes strongly decreased release of active TGF-ß2 in response to retinoic acid. Retinoic acid exposure induced growing cultures of vector-infected cells to secrete 30 pg of active TGF-ß2/106 cells/24 h, compared with only 2 pg of TGF-ß2 released by E6/E7- expressing cells under the same conditions (data not shown). Thus, E6 and E7 inhibit secretion of TGF-ß2 induced by either cell differentiation or retinoic acid.

Down-Regulation of Biologically Active TGF-ß2 Secretion Involves the pRb and p53 Tumor Suppressor Genes.
Down-regulation of TGF-ß2 secretion was dependent on the ability of the E6 or E7 proteins to bind and inactivate the p53 and pRb proteins, respectively. A dominant-negative p53 miniprotein (p53 DD) decreased secretion of active TGF-ß2 as efficiently as wild-type HPV-16 E6 (Fig. 4CCitation ). In contrast, overexpression of wild-type p53 resulted in a 2-fold induction of TGF-ß2 secretion. A similar induction was observed in senescent keratinocytes (in vitro passages >4). These data point to a functional role for the p53 tumor suppressor gene in regulation of TGF-ß2 expression. The low-risk E6 and E7 proteins of HPV-6b, which do not degrade or inactivate the p53 and pRb proteins, also failed to alter secretion of biologically active TGF-ß2 (Fig. 4CCitation ). HPV-6b E6 and E7 also did not alter expression of cellular genes regulated by TGF-ß (data not shown). A mutant HPV-16 E7 protein unable to bind the retinoblastoma protein [E7-p24gly (33) ] did not diminish secretion of biologically active TGF-ß2. Interestingly, infection of keratinocytes with a retrovirus overexpressing the E2F-1 transcription factor, which signals downstream of pRb, was not sufficient to decrease TGF-ß2 secretion.

TGF-ß2 Inhibits Growth of E6/E7-expressing Keratinocytes.
TGF-ßs strongly inhibit growth of keratinocytes. We found that keratinocytes expressing HPV-16 E6 and E7 retained sensitivity to growth inhibition by recombinant TGF-ß2 in three independent experiments (Fig. 5Citation ). In fact, cells expressing E6 and E7 were slightly more sensitive to TGF-ß2 than primary cultures of normal human keratinocytes. Addition of recombinant TGF-ß2 at concentrations produced endogenously by differentiating keratinocytes (10–200 pg/ml) inhibited growth of E6/E7-expressing cells by 25–75%. Cells expressing HPV-16 E6/E7 were inhibited 50% by 100 pg/ml TGF-ß2 compared with uninfected keratinocytes, which were inhibited 50% by 200 pg/ml (Fig. 5Citation ). E6/E7 positive cells grew faster than control cells, but this difference was lost after physiological doses of TGF-ß2 were added to the culture medium. An HPV-immortalized cell line examined at low passage retained sensitivity to growth inhibition by TGF-ß2. Cells with reduced growth potential were particularly sensitive to low concentrations of TGF-ß2. Keratinocytes infected with HPV-16 E6/E7 and approaching crisis because of prolonged culture in vitro (Fig. 5Citation ) were growth inhibited 50% by 5–10 pg/ml of TGF-ß2. Interestingly, E6/E7 expressing cells in crisis or senescing cultures of normal keratinocytes up-regulated expression of TGF-ß2 2-fold (Fig. 4CCitation ).



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  		Fig. 5. Recombinant TGF-ß2 inhibits growth of cervical keratinocytes expressing HPV-16 E6 and E7 genes. Cervical keratinocytes were infected with retroviruses encoding HPV-16 E6/E7 or vector (normal), cells were maintained in culture for three passages (rapidly growing) or five passages (approaching senescence) and treated with various concentrations of TGF-ß2 for 8 days. The shaded region corresponds to concentrations of active TGF-ß2 secreted by differentiating keratinocytes in culture as determined by ELISA. Bars, SD.

 
TGF-ß2 Inhibits Immortalization of Keratinocytes by HPV E6/E7.
HPV-16 E6/E7-infected cells were seeded at low density in 150-mm dishes and treated with TGF-ß2 concentrations similar to those secreted endogenously by cultures of differentiating keratinocytes (10–200 pg/ml). The ability of infected keratinocytes to grow continuously and form immortal colonies was significantly decreased (P < 0.05, t test) by 100–200 pg/ml TGF-ß2 (Fig. 6ACitation ). Growth inhibition and severe crisis preceded formation of colonies arising from cells that survived crisis. Colonies of small, rapidly growing cells were picked and subcultured to confirm that they were immortal. Of 20 colonies selected, 18 grew for three more in vitro passages and were established as immortal cell lines. Fig. 6BCitation summarizes data from six independent experiments.



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  		Fig. 6. Recombinant TGF-ß2 inhibits immortalization of cervical keratinocytes by HPV-16 E6/E7. A, cultures of E6/E7-infected keratinocytes that were treated with TGF-ß2 for 10–14 days and subsequently stained with 0.01% crystal violet to visualize colonies. B, relative colony-forming ability of keratinocytes expressing HPV-16 E6/E7 after treatment with various concentrations of TGF-ß2 for 10–14 days. Values represent the means of six independent experiments; bars, SD.

 

   DISCUSSION
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
HPV-16 E6 and E7 Proteins Inhibit Differentiation-dependent Expression of TGF-ß2 and TGF-ß-responsive Genes.
HPV infection of the cervix is characterized by high-level expression of the E6 and E7 oncoproteins in suprabasal keratinocytes undergoing squamous differentiation. We used cDNA expression arrays to identify cellular genes that are up- or down-regulated by the HPV-16 E6 and E7 in differentiating cultures of human cervical keratinocytes. Over 100 cellular genes or ~7% of genes on the array were either increased or decreased in expression by >2-fold. These genes were classified into several functional groups including growth factors/cytokines, apoptosis-related genes, genes involved in tissue remodeling, and cell cycle genes. One of the genes that was strongly down-regulated by E6/E7 was TGF-ß2. Expression of TGF-ß2 is increased during epidermal differentiation (34) , and our array experiments confirmed that TGF-ß2 RNA was up-regulated 7-fold in differentiating cervical keratinocytes. We also demonstrated that many of the genes that were perturbed by E6 and E7 in differentiating keratinocytes were TGF-ß-responsive genes and/or genes regulated during keratinocyte differentiation. Thus, our results suggest that E6 and E7 indirectly alter expression of diverse cellular genes by decreasing expression of TGF-ß2. Our results also suggest that differential expression of TGF-ß2 may represent a consequence of the ability of E6 and E7 to interfere with aspects of keratinocyte differentiation. However, we also observed down-regulation of TGF-ß2 expression in growing cells that did not differentiate, and E6 and E7 also blocked up-regulation of TGF-ß2 expression by all-trans retinoic acid. Thus, down-regulation of TGF-ß2 expression by E6 and E7 is not restricted to differentiating cells.

HPV-16 E6 Down-Regulates TGF-ß2 Production by a p53-dependent Pathway.
The HPV-16 E6 oncoprotein binds to several cellular proteins that have been implicated in cell cycle regulation or gene expression (14 , 15) . In particular, E6 proteins of high-risk HPV types bind to the p53 tumor suppressor protein and target it for degradation by a ubiquitin-mediated pathway (15) . We used HPV-16 E6 mutants that differed in their ability to bind and degrade p53 to demonstrate a direct correlation between inactivation of p53 and decreased TGF-ß2 protein production. In particular, the E6-F2V mutant that interacts with E6BP but fails to degrade p53 (28) did not decrease levels of TGF-ß2. The low-risk HPV-6b E6 protein, which does not efficiently degrade p53, also failed to decrease TGF-ß2. We confirmed this association by showing that a dominant-negative p53 mutant inhibited TGF-ß2 production as effectively as the wild-type HPV-16 E6 protein. Furthermore, retroviral expression of a wild-type p53 gene in cervical keratinocytes increased TGF-ß2 protein levels. We also showed that TGF-ß2 secretion was induced 2-fold in cells undergoing replicative senescence, a process that involves activation of p53 and p53-responsive genes (16) . Our data indicate that TGF-ß2 is a p53-responsive gene. However, the TGF-ß2 promoter does not contain p53 response elements, suggesting that regulation might be indirect. Of more general interest, our results infer that p53 inactivation by mutations, which occur frequently in premalignant lesions and carcinomas, might also block production of biologically active TGF-ß2. This might confer a selective growth advantage on cells with mutated p53 and contribute to malignant progression.

Interactions between E6/E7, p53, and TGF-ß in Keratinocyte Differentiation.
Differentiation in HPV-16-infected keratinocytes and premalignant lesions is altered and partially uncoupled from proliferation (10 , 35) . In particular, HPV-16 E6 has been shown to impair serum- and calcium-induced keratinocyte differentiation (7 , 9) . This function correlates with the ability of E6 mutants to degrade p53, although additional functions of E6 might also be involved (7 , 9) . The biochemical activities of p53 and p53-mediated genes like p21-waf1 are increased during keratinocyte differentiation (9 , 37) , and overexpression of wild-type p53 in keratinocytes causes premature differentiation and cell flattening (38) . In turn, TGF-ß isoforms control stability and phosphorylation patterns of p53 protein and activate p53-dependent expression of p21/waf-1 and GADD45 (39) . p53-null keratinocytes show impaired responsiveness to negative growth regulators, including TGF-ß (40) .

The expression of HPV-16 oncoproteins and TGF-ß2 protein coincide in the differentiating, suprabasal layers of HPV-infected cervical epithelia (9 , 34) . Expression of mitotic regulatory genes (e.g., cyclins A and B and cyclin-dependent kinases) are elevated in HPV-infected keratinocytes (31) but repressed in growth-inhibited and/or differentiating keratinocytes. These genes are also repressed in response to p53-mediated growth arrest or by TGF-ß. Therefore, HPV-mediated inhibition of TGF-ß2 expression, which supports epithelial differentiation (41) , may contribute to growth stimulation and defective differentiation observed in HPV-infected keratinocytes. TGF-ß has also been reported to induce replicative senescence in normal keratinocytes (42) and apoptosis in HPV-immortalized keratinocytes (43) . Inhibition of p53- and TGFß- responsive proapoptotic genes (Fas/APO-1, FasL, TNF-receptor, TRAIL, and others) or induction of protective genes (survivin or inhibitor of apoptosis 4) may help to explain the partial inhibition of differentiation-related cell death by HPV-16 E6 (8) .

HPV-16 E7 Protein Down-Regulates TGF-ß2 Production by a pRb-dependent Pathway.
The HPV-16 E7 protein binds to and inactivates pRb and related proteins p107 and p130 (17) . We have shown that two different E7 proteins that fail to bind pRb were not effective in decreasing secretion of TGF-ß2. Specifically, the low-risk HPV-6b E7 protein and an HPV-16 E7 mutant [E7-p24gly (33) ], which do not bind pRb, also do not perturb TGF-ß2 production. However, we found that retroviral expression of the E2F-1 transcription factor, which acts downstream of pRb, was not sufficient to decrease TGF-ß2 expression, suggesting that additional factors are involved. E2F-1 overexpression had strong effects on the growth of keratinocytes, which did not involve changes in TGF-ß2 expression. These data indicate that a functional pRb protein is necessary for regulation of TGF-ß2 protein expression. Interestingly, pRb has been reported to increase expression of the TGF-ß2 gene by binding to the transcription factor ATF-2, which in turn interacts with the promoter of the TGF-ß2 gene (44) . Furthermore, pRb binds to c-jun and alters activity of members of the AP-1 family of transcription factors, which serve an important role in epithelial differentiation (18 , 19) .

E6 and E7 Regulate TGF-ß2 Activity at the Transcriptional Level.
Regulation of TGF-ß activity is complex and may be mediated by changes in transcription, secretion, or activation of the latent peptide. Our results suggest that E6 and E7 decreased levels of biologically active TGF-ß2 mainly by inhibiting expression of TGF-ß2 RNA (5–7-fold). Levels of total cell-associated TGF-ß2, secretion of latent peptide, and levels of active TGF-ß2 generally decreased by a similar factor. We found significant differences between localization of the different TGF-ß isoforms. TGF-ß1 was almost exclusively intracellular, whereas 30–50% of TGF-ß2 was found in the extracellular matrix. TGF-ßs are activated in association with matrix proteins; therefore, the absence of TGF-ß1 from the extracellular matrix might explain in part why most TGF-ß1 remained in the latent state.

Inhibition of Keratinocyte Growth and Immortalization by TGF-ß2.
Previous studies have reported conflicting results regarding the sensitivity of E6/E7-expressing keratinocytes to TGF-ß growth inhibition (36 , 45) . Our results show that cervical keratinocytes expressing E6 and E7 were growth inhibited by low concentrations of recombinant TGF-ß2 (10–200 ng/ml), similar to levels secreted by differentiating keratinocytes in the absence of E6 and E7. Furthermore, the induction of TGF-ß-responsive genes by recombinant TGF-ß2 confirmed that the downstream signaling and response to TGF-ß were intact in cells expressing E6 and E7. These results suggest that inhibition of TGF-ß2 expression by HPV E6 and E7 is biologically significant. Recombinant TGF-ß2 at levels secreted by differentiating keratinocytes also significantly inhibited immortalization of primary cervical keratinocytes by HPV-16 E6/E7 genes. Senescent keratinocytes or E6/E7-expressing cells approaching crisis were strongly growth inhibited by recombinant TGF-ß2 at levels secreted by differentiating keratinocytes. Furthermore, expression and secretion of TGF-ß2 increased 2-fold (Fig. 4CCitation ) in these cells. Autocrine TGF-ß has been implicated in blocking growth and immortalization of keratinocytes in response to vitamin D3 or retinoic acid (46 , 47) . TGF-ß isoforms actively induce cellular senescence (42) and programmed cell death (43) , processes that directly precede and counteract immortalization of keratinocytes. Therefore, decreased secretion of TGF-ß2 in HPV-infected keratinocytes may result in loss of an autocrine or paracrine pathway that inhibits cellular immortalization in vitro.

Importance of TGF-ß2 in Cervical Cancer.
The role of TGF-ßs in epithelial carcinogenesis is complex. During multistage carcinogenesis in the skin, loss of the tumor suppressor function of TGF-ßs 1 and 2 accelerates malignant progression. Loss of TGF-ß expression in mouse skin papillomas is associated with suprabasal proliferation (48) , increased genomic instability (49) , and rapid malignant progression (5) . The loss of TGF-ß responsiveness through somatic mutations of TGF-ß receptor subunits I and II is common in many human cancers (reviewed in Ref. 4 ), including cervical carcinoma (50) . Loss or altered expression of TGF-ß1 has been described as an early event in cervical carcinogenesis and frequently observed in cervical intraepithelial neoplasia (51) . Frequent loss or decreased expression of TGF-ß2 also has been described in cervical dysplasia (52) . The latter observations are consistent with the idea that TGF-ßs function as tumor suppressors in cervical epithelia and also confirm that loss of TGF-ß expression in HPV-infected keratinocytes is not restricted to observations in cell culture. Thus, down-regulation of TGF-ß2 expression by HPV oncoproteins may represent a novel mechanism that contributes to growth stimulation, immortalization, and carcinogenesis.


   ACKNOWLEDGMENTS
 
We thank Darci A. Gaiotti and Eudora A. Jones for excellent technical assistance, Moshe Oren for the dominant-negative p53DD construct, Adam Glick for critical reading of the manuscript, and Christa Walter for editorial assistance.


   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 To whom requests for reprints should be addressed, Biology Department, Clarkson University, Potsdam, NY 13699. Phone: (315) 268-6641; Fax: (315) 268-6610; E-mail: woodworth{at}clarkson.edu Back

2 The abbreviations used are: TGF, transforming growth factor; HPV, human papillomavirus; pRb, retinoblastoma protein; GAPDH, glyceraldehyde phosphate dehydrogenase; E6BP, E6 binding protein; E6AP, E6-associated protein, RT-PCR, reverse transcription-PCR; MMP, matrix metalloproteinase; dNTP, deoxynucleotide triphosphate; PAI, plasminogen activator inhibitor. Back

Received 11/24/99. Accepted 5/30/00.


   REFERENCES
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 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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H. You, Y. Liu, M. J. Carey, C. L. Lowery, and P