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J Virol, January 1998, p. 708-716, Vol. 72, No. 1
0022-538X/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Novel Binding Sites for Regulatory Factors in the Human
Papillomavirus Type 18 Enhancer and Promoter Identified by In
Vivo Footprinting
Paula H.
Bednarek,
Betty J.
Lee,
Sanjay
Gandhi,
Edward
Lee, and
Benette
Phillips*
Departments of Obstetrics and Gynecology and
Cell and Molecular Biology, Northwestern University Medical School,
Chicago, Illinois 60611
Received 16 July 1997/Accepted 25 September 1997
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ABSTRACT |
The E6 and E7 genes of human papillomaviruses (HPVs) associated
with anogenital cancers are largely responsible for the oncogenic activity of these viruses, and regulation of these genes has been intensively studied. Transcription of the E6 and E7 genes is controlled by the viral upstream regulatory region (URR). We have used in vivo
footprinting to examine the occupancy by regulatory factors of the HPV
type 18 (HPV18) URR enhancer and promoter in the cervical carcinoma
cell lines HeLa and C4-II. While corroborating occupancy in vivo of all
of the elements previously implicated in the transcriptional control of
the HPV18 E6 and E7 genes by in vitro DNase I footprinting, gel
retardation assays, and transfection studies, we also detect occupancy
in vivo of several enhancer and promoter sequences which have not been
previously identified as HPV18 URR regulatory elements. Our data
suggest that the HPV18 enhancer and promoter are more densely occupied
by DNA-binding proteins than previously thought and raise the
possibility that additional, possibly novel factors contribute to
transcription of the HPV18 early genes.
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INTRODUCTION |
Most cervical cancers develop
subsequent to infection with human papillomavirus types 16 and 18 (HPV16 and -18), or less commonly other high-risk HPV types, and
contain transcriptionally active copies of the viral DNA
(56). In all HPV18-associated cervical carcinomas so far
examined, and in most but not all HPV16-linked malignancies, viral DNA
is integrated into the host genome (5, 11, 45). Viral
sequences which encode the E6 and E7 proteins and an 800-bp viral
upstream regulatory region (URR) which controls transcription of these
viral genes are invariably intact in the integrated viral genome,
although the integration event usually interrupts sequences necessary
for expression of the E2 protein, the only viral product thought to
regulate HPV transcription (13, 42, 44). The E6 and E7 genes
encode proteins which interact with and inactivate the tumor
suppressors p53 and Rb, respectively (19, 41), and
expression of these viral proteins is thought to be crucial to the
initiation and progression of cervical tumors (27, 28, 33, 52,
54).
Recognition of the key role of these viral proteins in cervical cancer
has stimulated intense efforts to understand how E6/E7 gene expression
is regulated in cervical cells. Although posttranscriptional events are
clearly important (26), the rate of viral transcription controlled by the URR is a major determinant of E6 and E7 levels. Considerable effort has therefore been made to delineate which HPV16
and HPV18 URR sequences are important for transcriptional regulation of
the E6 and E7 oncogenes and to identify the cellular factors which
interact with these sequence elements. Attention has largely been
focused on the 400 to 500 bp of sequence which are immediately upstream
of the E6/E7 transcription start site, since transfection studies have
indicated that deleting the remaining sequences of the 800-bp URR
diminishes transcription only slightly (20, 47).
Transfection studies have further revealed that a region 230 bp long in
HPV18 and 400 bp long in HPV16, termed the constitutive enhancer, is
critical for efficient transcription, is preferentially active in
epithelial cells, and will function with heterologous promoters
(10, 16, 46). In both HPV16 and HPV18, DNase I footprinting
studies reveal the constitutive enhancer to be densely occupied by
cellular regulatory factors, and gel retardation assays suggest that
many of these are ubiquitous factors which regulate a wide variety of
cellular genes (7, 22, 51). The remaining URR sequences
downstream of the enhancer constitute the proximal promoter, which
contains binding sites for Sp1 and other cellular factors, e.g., YY1,
but which by itself drives only very low levels of viral transcription
(21, 47). With a few notable exceptions, many of the same
cellular factors appear to interact with both the HPV18 and HPV16
enhancer and promoter; however, the cognate sites for these factors are
arranged quite differently in the regulatory regions of the two
viruses.
Although invaluable tools in the analysis of regulatory regions, in
vitro DNase I footprinting and gel retardation assays utilize naked DNA
fragments and cellular extracts and therefore may detect DNA-protein
interactions which are not physiologically relevant or may fail to
detect all interactions which are occurring in vivo. Complementation of
in vitro binding assays with in vivo footprinting studies is therefore
becoming increasingly common in promoter analysis. Such studies allow
one to detect occupancy of regulatory regions in the unperturbed cell:
chromatin structure is intact, the localization and availability of
DNA-binding proteins are not perturbed, and the interactions of these
proteins with other protein partners are not disrupted by extract
preparation (15, 23, 34, 50).
While data from several independent studies agree as to the identity of
certain of the factors which recognize sites in the HPV16 and HPV18
URRs, e.g., AP1 and Oct-1 (7, 20, 36, 49), and the
functional importance of certain sites, there is less of a consensus
regarding other sites, e.g., putative NF1 sites (6, 8). To
verify that all sequence elements so far identified by in vitro binding
assays and transfection studies in HPV18 are occupied in vivo and to
look for additional sites of factor interaction in the HPV18 enhancer
and promoter which may have escaped detection, we have performed in
vivo footprinting in the cervical carcinoma cell lines HeLa and C4-II,
both of which contain integrated, transcriptionally active copies of
HPV18. While corroborating occupancy in vivo of most of the elements
previously implicated in the transcriptional control of the HPV18 E6
and E7 genes, our results also reveal occupancy in vivo of several
enhancer and promoter sequences which have not been previously
identified as HPV18 URR regulatory elements. Our data thus suggest that
the enhancer and promoter regions of HPV18 are more densely occupied by
DNA-binding proteins than previously thought and raise the possibility
that additional, possibly novel factors contribute to transcription of
the HPV18 early genes.
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MATERIALS AND METHODS |
Amplification and sequencing of the HPV18 enhancer and
promoter.
Genomic DNA (100 ng) from HeLa and C4-II cells was
amplified with primers 18E6 (5'-AGGGTCGCCGTGTTGGATCCT-3';
nucleotides [nt] 138 to 118), and primer 1 of bottom-strand
footprinting set I (nt 7363 to 7386; sequence listed below). PCR
conditions were 40 cycles of 94°C for 1 min, 56°C for 1 min, and
72°C for 1 min. Amplimers (632 bp) were isolated from a 0.9% NuSieve
GTG agarose gel (FMC) by melting, phenol extraction, and ethanol
precipitation and were sequenced by using an Amplicycle sequencing kit
(Perkin-Elmer). Sequencing primers (18E6, primer 1 of bottom-strand
footprinting set I, and primer 2 of bottom strand footprinting set III)
were end labeled with [
-33P]ATP.
In vivo footprinting.
Subconfluent HeLa and C4-II cells,
growing on 15-cm-diameter dishes in Dulbecco modified Eagle medium
supplemented with 10% fetal bovine serum, were exposed to dimethyl
sulfate (DMS) for 2 min at room temperature as follows: spent medium
was removed, DMS was suspended in 10 ml of this medium to 0.2% final
concentration by vortexing, and the mixture was added back to the
cells. The reaction was stopped by aspiration of the DMS-containing
medium and immediate addition of ice-cold phosphate-buffered saline. Cells were scraped from the plates and lysed to isolate nuclei. Genomic
DNA was isolated from the lysed nuclei (4), digested with
HindIII, cleaved with piperidine, and used for
footprinting. Footprinting was also carried out on DNA which was
isolated from C4-II and HeLa cells, purified free of protein (naked
DNA), cleaved with HindIII, and treated with DMS in
vitro (40 µg of naked DNA was exposed to 0.25% DMS for 1 min).
Footprinting was carried out on multiple batches of cells and naked DNA
independently exposed to DMS to ensure reproducibility of results.
Footprinting analyses were performed by using a ligation-mediated PCR
method (32). Footprinting of the top and bottom strands of
the HPV18 enhancer and promoter was performed with eight sets of
primers, three primers per set. For footprinting of the bottom strand,
the primers in set I were primer 1 (5'-GCTTGTTGGGCTATATATTGTCCT-3'), primer 2 (5'-CTGCACACCTTACAGCATCCATTTTATCCTACA-3'), and primer 3 (5'-CTGCACACCTTACAGCATCCATTTTATCCTACAATCCTC-3'). The primers in set II were primer 1 (5'-ATACAGTACGCTGGCACTATTG-3'),
primer 2 (5'-GGGCACTGCTCCTACATATTTTGAACCATTG-3'), and
primer 3 (5'-GGGCACTGCTCCTACATATTTTGAACCATTGGCG-3'). The
primers in set III were primer 1 (5'-CTACATATTTTGAACCATTGGCG-3'), primer 2 (5'-ACCTGGTATTAGTCATTTTCCTGTCCAGG-3'), and
primer 3 (5'-ACCTGGTATTAGTCATTTTCCTGTCCAGGTGCG-3'). The
primers in set IV were primer 1 (5'-TCCCTATGTAATAAAACTGCTTTTAGG-3'), primer 2 (5'-GCTAATTGCATACTTGGCTTGTACAACTACTTTCAT-3'), and primer 3 (5'-GCTAATTGCATACTTGGCTTGTACAACTACTTTCATGTCC-3'. For
footprinting of the top strand, the primers in set I were primer 1 (5'-GCCTAAAAGCAGTTTTATTACATAGGG-3'), primer 2 (5'-GGGAGTGGATATAGTTATGCAAGCAATTGTTGT), and primer 3 (5'-GGGAGTGGATATAGTTATGCAAGCAATTGTTGTAGC-3'). The primers in
set II were primer 1 (5'-CTTAGTCATATTATAGTTCATGTTAAGG-3'),
primer 2 (5'-GACAGAATGTTGGACATGAAAGTAGTTGTACAA), and
primer 3 (5'-GACAGAATGTTGGACATGAAAGTAGTTGTACAAGCC-3'). The
primers in set III were primer 1 (5'-GAAAAGTATAGTATGTGCTGCC-3'), primer 2 (5'-CCCAACCTATTTCGGTTGCATAAACTATGTAT-3'), and
primer 3 (5'-CCCAACCTATTTCGGTTGCATAAACTATGTAT-3'). The
primers in set IV were primer 1 (5'-AAGTGTTCAGTTCCGTGCACA-3'),
primer 2 (5'-GTGTTGGATCCTCAAAGCGCGCCA-3'), and primer
3 (5'-CCTCAAAGCGCGCCATGGTATTGTGGTGTG-3'). The positions of
the 3' ends of the eight primers 3 used for footprinting are indicated
by arrowheads in Fig. 1.
Primer 1 was annealed to piperidine-treated DNA and extended with
Sequenase, primer 2 was used in conjunction with a linker
primer for
amplification, and primer 3 was phosphorylated and
used to label the
amplified fragments. Amplification conditions
were 19 cycles of 1 min
at 94°C, 2 min of annealing at a temperature
which varied with each
primer 2, and 3 min at 76°C. To label the
fragments, we performed one
cycle of 2 min at 94°C, 2 min of annealing
at a temperature which
varied with each primer 3, and 10 min at
76°C. Labeled fragments were
electrophoresed on 6% polyacrylamide-8
M urea sequencing gels, which
were dried and exposed to film.
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RESULTS |
In vivo footprinting of the HPV18 URR enhancer.
C4-II cervical
carcinoma cells contain a single integrated and transcriptionally
active copy of the HPV18 genome (44), and because all
regulatory factor-DNA interactions detected are likely to be relevant
to transcription, it was our original intent to use C4-II cells
exclusively in this study. However, we obtained much better sensitivity
and nearly identical results with HeLa cells, which contain 10 to 50 integrated copies of HPV18 DNA (44), and we therefore show
results from both lines interchangeably. We confirmed that the C4-II
and HeLa stocks maintained in our laboratory actively transcribe the
E6/E7 gene by transcription run-on analysis (data not shown).
Eight sets of primers were used to examine factor occupancy in the
enhancer and promoter (Fig.
1). To
facilitate the design
of these primer sets, as well as the matching of
DMS reactivity
patterns to sequence, we amplified URR nt 7363 to 138 from each
cell line and sequenced these amplimers directly. When
compared
to the sequence of the prototype HPV18 genome cloned from a
cervical
biopsy specimen (
9 [accession no.
X05015]), seven base pair
changes were found in the HPV18 DNA
integrated in the C4-II cell
line and six changes were found in the
HPV18 sequences integrated
in HeLa cells, with five of these changes
common to both cell
lines (Table
1). An
independent sequencing of the HPV18 DNA cloned
from the biopsy specimen
(
48) and sequencing of HPV18 DNA isolated
from HeLa cells by
using an enhancer trap strategy (
46) revealed
very similar
sets of changes.
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FIG. 1.
Sequence of the HPV18 enhancer (nt 7510 to 7740) and
promoter (nt 7741 to 105), showing proposed binding sites for
transcription factors, as suggested from previous studies. The
arrowheads denote the 3' ends of primer 3 from each primer set; DMS
reactivity patterns on the opposite strand can be visualized starting
approximately 30 nt downstream of the 3' end of primer 3.
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TABLE 1.
Deviations from the depositeda
enhancer and promoter sequences in HPV18 sequences in HeLa and
C4-II cells
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To identify sequences in the HPV18 enhancer and promoter to which
factors were binding and potentially regulating transcription
in vivo,
both HeLa and C4-II cells were treated briefly with 0.2%
DMS, and the
DMS reactivity patterns in DNA isolated from these
cells were compared
to those of DNA isolated from the same cell
lines, purified free of
proteins, and then exposed to DMS in vitro.
Differences in the
susceptibility of guanines to methylation by
DMS in DNA exposed in vivo
and in vitro are taken to indicate
the presence of bound regulatory
factors (
4). Reactivity to
DMS is not perturbed by the
association of DNA with histone or
nonhistone components of chromatin
(
30). Adenines are also reactive
with DMS, although much
less so than guanines (
40). We have
noticed, however, that
adenines which are part of a purine-rich
stretch are fairly reactive
with DMS. The HPV18 URR enhancer and
promoter contain many short A-rich
stretches, and these stretches
are frequently hyperreactive in DNA from
DMS-treated cells, the
basis for which is presently unclear. Therefore,
in this study,
only adenines which are particularly hyperreactive to
DMS are
considered to mark factor binding sites.
Before a region was designated as a putative site for factor
interaction, several criteria had to be met. Changes in methylation
sensitivities were not considered significant unless they were
consistently seen in multiple footprinting assays which used DNA
exposed to DMS in independently performed experiments. Binding
of a
regulatory factor was generally ascribed only to those regions
in which
the relative densities of several bands in a set differed
between DNA
treated in vitro and in vivo. There was often considerable
overlap in
the sequences which could be visualized with different
primer sets, and
when this overlap encompassed regions of differences
in the reactivity
patterns between in vivo- and in vitro-exposed
DNA, similar changes in
the band patterns had to be seen with
both primer sets.
In vivo footprinting of the distal region of the enhancer.
Regulatory proteins which have been proposed to bind to the HPV18
enhancer include the ubiquitous transcription factors NF1 (6,
17), Oct-1 (6, 29), AP1 (14, 49), and YY1
(1), as well as a C/EBP 
-YY1 complex (3) and
KRF-1 (29), a factor which has not yet been characterized
(Fig. 1). We performed in vivo footprinting of the enhancer starting
from the distal end. Within the distal portion of the enhancer are two
pairs of TTGGC half-sites, one pair at nt 7513 to 7526, where neither
half-site is a perfect match to the consensus, and the other at nt 7569 to 7586, where both half-sites are TTGGC. These pairs of half-sites have been proposed to be binding sites for the factor NF1
(17). The sequence TGACTAA, which deviates in
only one position from a consensus AP1 site, is found just downstream
at nt 7608 to 7614.
Using bottom primer set I, which allowed us to visualize nt 7490 to
7625, we saw only a few prominent differences in the DMS
reactivity
patterns of DNA from DMS-treated cells and naked DNA
exposed to DMS in
vitro (Fig.
2A). However, there were
other,
less prominent changes in reactivity which were seen with
absolute
consistency and are likely to signify factor occupancy. Just
upstream
of the distal pair of sequence-aberrant TTGGC
half-sites, G-7496
was hyperreactive to DMS in vivo, while G-7501
and G-7506 were
hyporeactive; sequences comprising these guanines have
not previously
been proposed to constitute a binding site. Within and
bordering
the TTGGC half-sites, three guanines showed diminished
reactivity
in vivo: G-7511 and G-7519, which were weakly protected from
DMS
in vivo, and G-7525, which was strongly protected. These G's all
flank or lie in within sequences corresponding to the distal pair
of
TTGGC half-sites. Between G-7525 and the end of the region
visualized
with this primer set, only two other bottom-strand
G's showed clear
and reproducible differences in reactivity to
DMS. G-7588, just
downstream of the proximal pair of TTGGC repeats,
was clearly
hyperreactive in vivo, while G-7599 showed a less
pronounced
hyperreactivity. We saw no change in reactivity to
DMS of G-7613 in the
AP1 site in assays using this set of primers.
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FIG. 2.
In vivo footprinting of the distal portion of the HPV18
enhancer (nt 7450 to 7625), using top and bottom primer sets I and DNA
from HeLa cells. The left lane in each gel shows the methylation
pattern of protein-free DNA exposed to DMS in vitro; the right lane
shows the methylation pattern of DNA isolated from DMS-treated cells.
Open circles represent G's or A's which were hyporeactive to DMS in
DNA treated in vivo; closed circles represent G's or A's which were
hyperreactive to DMS in DNA treated in vivo. Regions corresponding to
proposed factor binding sites are denoted by vertical lines.
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Footprinting of top-strand sequences (nt 7450 to 7595) of the distal
region of the enhancer consistently showed differences
in the
methylation patterns in sequences comprising both pairs
of putative NFI
sites (Fig.
2B). Within the proximal pair of TTGGC
half-sites, three
protected G's, G-7587, G-7576, and G-7572, and
one hypersensitive G,
G-7574, were seen in DNA exposed to DMS
in vivo, and a marked
hypersensitivity at A-7565 was seen just
upstream. In the top-strand
sequences of the distal pair of imperfect
TTGGC half-sites, A-7528 was
strikingly hyperreactive to DMS in
vivo, G-7515 was also hyperreactive,
and G-7516 was hyporeactive.
Taken together, our top- and bottom-strand
footprinting data clearly
suggest that sequences which include the two
pairs of TTGGC half-sites
are occupied in vivo, although the footprints
for the distal and
proximal pairs are quite distinct. It is also
possible that sequences
just upstream of the distal pair of half-sites
are occupied by
factors, although in this region changes were only seen
in bottom-strand
sequences.
In vivo footprinting of the central region of the enhancer.
Top-strand primer set II, which was used to footprint the central
region of the enhancer, also allowed us to visualize reactivity patterns in the proximal and distal TTGGC repeats, although with less
resolution, since these regions were now far from the primers, i.e.,
near the top of the gel. Nevertheless, this primer set yielded very
similar DMS reactivity patterns in top-strand sequences comprising the
two pairs of TTGGC repeats (Fig. 3B).
With this primer set, top strand reactivity patterns covering sequences
between nt 7590 and 7700 could also be visualized, and within these
sequences we saw many differences between DNA exposed to DMS in vitro
and in vivo (Fig. 3B), suggesting that virtually the entire central portion of the enhancer is occupied by DNA-binding proteins. Clear protection from DMS of G-7611 in the AP1 site was apparent in DNA from
DMS-treated cells, and two guanines just upstream (G-7604 and G-7605)
were also protected. Just downstream of the AP1 site, we observed
protection of G-7630 and G-7632 in a region which does not comprise a
known HPV18 enhancer binding site. Sequences between nt 7644 and 7651 match in six of eight positions a consensus Oct-1 site and appear to
bind Oct-1 proteins in a nuclear extract (29); immediately
downstream and possibly overlapping the Oct-1 site are sequences (nt
7641 to 7675) first identified by DNase I protection studies and
subsequently shown to interact with a keratinocyte specific factor,
KFR-1 (29). Two G's within the putative Oct-1 site, G-7644
and G-7648, and G-7671, at the 3' end of the sequences proposed to
interact with KRF-1, were all protected in vivo (Fig. 3B).
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FIG. 3.
In vivo footprinting of the central portion of the HPV18
enhancer (nt 7510 to 7760), using top and bottom primer sets II and DNA
from C4-II cells. The left lane in each gel shows the methylation
pattern of protein-free DNA exposed to DMS in vitro; the right lane
shows the methylation pattern of DNA isolated from DMS-treated cells.
Open circles represent G's or A's which were hyporeactive to DMS in
DNA treated in vivo; closed circles represent G's or A's which were
hyperreactive to DMS in DNA treated in vivo. Regions corresponding to
proposed factor binding sites are denoted by vertical lines.
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At the proximal end of the central enhancer, additional hyporeactive
G's were seen: G-7682 was clearly protected from DMS
in vivo, as were
G-7690 and G-7702. Although several A's upstream
of G-7682 were
hyperreactive in vivo, A-7678 was especially so.
Sequences between nt
7691 and 7700, CACATATTTT, are identical
to a site in HPV16
which was previously shown to bind TEF-1 (
24),
a factor
which was initially identified as a simian virus 40 enhancer-binding
protein (
12). Our data thus show changes in a potential
TEF-1
binding site but also indicate occupancy of sequences immediately
upstream. In DNase I footprinting assays carried out by Nakshatri
et
al. (
35), nt 7673 to 7704 were protected from DNase I when
C33A nuclear extracts were used, although not when extracts of
four
other cell lines, including HeLa, were used. Two other in
vitro
footprinting studies failed to detect protection of this
region
(
14,
17).
Footprinting of bottom-strand sequences (nt 7600 to 7760) in the
central region of the enhancer with primer set II confirmed
that this
portion of the enhancer is densely occupied by regulatory
factors.
Sequences corresponding to the AP1 site were seen at
higher resolution
with this primer set, and a weak but reproducible
protection of G-7613
within this site was seen (Fig.
3A). Downstream
of the AP1 site, in the
same region where protections of G-7630
and G-7632 were seen with
top-strand primer set II, bottom-strand
G-7633, G-7636, and G-7639 were
weakly protected from DMS in vivo.
In bottom-strand sequences
comprising the Oct-1 and KRF-1 sites,
there were very clear and
reproducible differences in the methylation
patterns. These include
protected G's at positions 7645, 7649,
7654, 7661, and 7663, as well
as a very hypersensitive A at position
7643. Furthermore, in the 75 nt
of bottom-strand sequences corresponding
to bands in the top third of
the gel, numerous and prominent differences
in the reactivity patterns
between DNA exposed to DMS in vitro
and in vivo were apparent (Fig.
3A). G-7691, in the putative TEF-1
site, and G-7683, upstream of this
site, were protected. Four
adenines between these G's and three
stretches of adenine repeats
downstream of G-7693 were clearly
hypersensitive to methylation
in vivo. Sequences downstream of this
A-rich stretch, corresponding
to bands near the top of the gel, include
a site for a proposed
C/EBP
-YY1 complex, termed the switch region
(nt 7710 to 7718
[
2,
3]) and a putative Oct-1/NF1
binding site (nt 7720 to
7735 [
6]). Within these
sequences, e.g., at G-7726, G-7730,
and G-7735, and in sequences
immediately downstream, e.g., G-7747,
there were numerous differences
in susceptibility to methylation
between DNA exposed to DMS in vitro
and in vivo (Fig.
3A).
In vivo footprinting of the proximal region of the enhancer.
Bands near the top of the gel shown in Fig. 3A were better resolved
when footprinting was performed with bottom primer set III. This primer
set allowed visualization of DMS reactivity patterns between G-7654 in
the KRF-I site and G-7784, 45 nt downstream of the enhancer/promoter
boundary. Much of this sequence overlapped with that seen with bottom
primer set II, and within the region of overlap, the band patterns for
DNA exposed to DMS in vitro and in vivo in assays using these two
primer sets were virtually identical (compare Fig. 3A and
4A). The single G in the switch region
site, G-7713, was not visible as a band in C4-II cells, for reasons
unknown, but was visible and clearly protected from methylation in HeLa
cells (data not shown). G-7719, just upstream of the Oct-1 site, showed
clear protection in vivo with primer set III, and protections of
G-7730, between the Oct-1 and NF1 sites, and G-7735, within the TTGGC
half-site, were very obvious as well (Fig. 4A). The marked differences
in the bottom-strand methylation patterns in sequences comprising this
TTGGC half-site, which borders an Oct-1 site, are in contrast to the
minor differences seen for the proximal pair of TTGGC half-sites. One
possible explanation for this difference is that binding of Oct-1 to
its cognate site enhances occupancy of the adjacent TTGGC site. For an
almost identical Oct-1/NF1 motif in HPV16, data obtained from gel shift
assays suggested cooperative binding of Oct-1 and NF1 (36).
Immediately downstream of the Oct-1/NF1 site, protections of G-7741,
G-7744, and G-7747 were also very apparent with this primer set (Fig. 4A).
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FIG. 4.
In vivo footprinting of the proximal portion of the
HPV18 enhancer (nt 7605 to 7785), using bottom primer set III and DNA
from C4-II cells and top primer set III and DNA from HeLa cells. The
left lane in each gel shows the methylation pattern of protein-free DNA
exposed to DMS in vitro; the right lane shows the methylation pattern
of DNA isolated from DMS-treated cells. Open circles represent G's
which were hyporeactive to DMS in DNA treated in vivo; closed circles
represent G's which were hyperreactive to DMS in DNA treated in vivo.
Regions corresponding to proposed factor binding sites are denoted by
vertical lines.
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Using primer set III, we were able to examine top-strand sequences in a
region which included the proximal part of the enhancer
and the distal
part of the promoter. Near the top of the gel,
where there was overlap
between the sequences visualized with
top-strand primer sets II and
III, changes in the DMS reactivity
patterns were very similar to those
seen in Fig.
3 (compare Fig.
3B and
4B). With this primer set, we also
detected protection
from DMS in vivo of G-7706, just downstream of the
putative TEF-1
binding site, as well as protections of G-7718 in the
switch region,
G-7725 and G-7734 in the Oct-1/NF1 site, and G-7738,
G-7754, and
G-7766 in the distal end of the promoter (Fig.
4B). The
data obtained
with top and bottom primer sets III strengthen the notion
that
enhancer sequences between the AP1 site at 7610 and the
enhancer-promoter
boundary at nt 7740 are almost fully occupied by
factors. Prominent
changes in DMS reactivity patterns were also seen in
both the
top and bottom strands of sequences just downstream of the
enhancer-promoter
boundary, a region not previously proposed to be a
factor binding
site.
In vivo footprinting of the promoter.
The HPV18 promoter,
which encompasses nt 7738 to 105, contains an AP1 site at its distal
end, an Sp1 site at its proximal end, and, between these two sites, a
putative glucocorticoid response element (GRE) and a YY1 binding site
(Fig. 1). The promoter also contains three binding sites for the viral
E2 protein, although neither HeLa nor C4-II cells are thought to
express E2, due to disruption of its coding region during integration
of the viral DNA. Primer sets III allowed us to examine the distal 30 to 40 bp of the promoter (Fig. 4), and we used primer sets IV to
inspect the DMS reactivity patterns of the rest of the promoter.
Using bottom primer set IV, we were able to visualize bottom-strand
sequences starting at nt 7770 (just upstream of the promoter
AP1 site)
and extending well into the coding region (Fig.
5A).
Because in approximately one-half of
the viral copies in HeLa
cells, integration has disrupted the URR
between the enhancer-promoter
junction and the TATA box (
42,
46), the bottom strand of the
promoter could be footprinted in
its entirety only in C4-II cells.
Several G's hyporeactive to DMS in
vivo were seen in the lower
half of the gel: these included G-7795 and
G-7800, within and
flanking the AP1 site, G-7805 and G-7809, downstream
of the AP1
site, and G-7820 and G-7823, within and flanking the E2
binding
site at nt 7822 to 7833 (Fig.
5A). We note with interest that
in a previously reported DNase I footprinting study of the HPV18
promoter, protection of two regions, nt 7781 to 7803 and 7809
to 7827, was observed (
14). Just downstream of the E2 site are
sequences (nt 7839 to 7853) which comprise a proposed GRE and
which
have been shown to mediate weak hormonal responsiveness
(
6,
31). Sequences between nt 7841 and 7850 also match the
consensus
for a TEF-1 binding site, 5'-N(G/A)CAT(T/A)(T/C)(T/C)(T/A)-3',
however (
25). Within and flanking these sequences we
saw four
weakly but consistently protected G's at positions 7838, 7843,
7847, and 7852 (Fig.
5A). Immediately downstream of these
sequences
is a stretch of 40 nt, encompassing both the YY1 and Sp1
sites,
which is very guanine poor. Bands corresponding to G-8, which
lies within the proposed YY1 site, and G-15 were always extremely
faint
and are barely visible in Fig.
5. However, we have seen
protection of
G-8 in vivo in several other independently performed
footprinting
assays of this region. Abutting the 3' end of the
Sp1 site (nt 35 to
40) is a pair of closely spaced E2 binding
sites, nt 42 to 53 and 57 to
68, and we consistently observed
hyporeactivity in vivo of G-43, just
3' to the Sp1 site, and hyperreactivity
of G-54, which lies between the
two E2 sites. Viral sequences
encoding the E2 protein are not intact in
C4-II cells (
42);
therefore, if the changes in DMS
reactivity seen at this pair
of E2 binding sites as well as at nt 7822 to 7833 signify factor
binding, such a factor must be of cellular
origin.
View larger version (36K):
[in this window]
[in a new window]
|
FIG. 5.
In vivo footprinting of the HPV18 promoter (nt 7730 to
105), using top and bottom primer sets IV and DNA from C4-II cells. The
left lane in each gel shows the methylation pattern of protein-free DNA
exposed to DMS in vitro; the right lane shows the methylation pattern
of DNA isolated from DMS-treated cells. Open circles represent G's
which were hyporeactive to DMS in DNA treated in vivo; closed circles
represent G's which were hyperreactive to DMS in DNA treated in vivo.
Regions corresponding to proposed factor binding sites are denoted by
vertical lines.
|
|
Footprinting performed with top-strand primer set IV allowed us to
examine all but the two most proximal top strand G's in
these same E2
sites, and these guanines showed equivalent reactivity
to DMS in vivo
and in vitro (Fig.
5B). Immediately upstream of
the pair of E2 sites,
clear protections in vivo of all four G's
(at positions 35, 36, 37, and 39) in a putative Sp1 site were
evident. These sequences have been
shown by gel retardation assays
to bind Sp1 and to be required for
HPV18 URR promoter activity
(
21). G-20 and G-23 were also
protected; these G's are just
5' to an upstream TATA consensus element
which in the HPV18 promoter
does not mediate transcription initiation
but rather comprises
part of the origin of replication in the intact
viral genome (
38).
A cluster of bands whose relative
densities consistently differed
in the in vitro and in vivo lanes
corresponded to nt 7825 to 7840
(Fig.
5B). G-7840, although faint in
the lane representing DNA
exposed to DMS in vitro, was fainter in the
in vivo lane, G-7831
and G-7832 were weakly protected in vivo, and
G-7825 was hyperreactive
in vivo (Fig.
5B). Considered along with the
altered reactivities
seen in bottom-strand G's in this region (Fig.
5A), the altered
reactivities of these top-strand G's further support
the notion
that a factor or factors constitutively occupy sequences
between
nt 7830 and 7850. The decreased reactivities of G-7802 and
G-7804
seen in vivo (Fig.
5B) may be due to in vivo occupancy of the
AP1 site at nt 7792 to 7798; however, changes in reactivities
of G's
at positions 7805 and 7809 in the bottom strand were also
seen (Fig.
5A), and nt 7807 to 7815 deviate in only one position
from a TEF-1
consensus site. We did not detect changes in reactivities
of G's in
the top strand of the AP1 site, similar to our failure
to detect
changes in the bottom strand of the distal AP1 site.
Except for G's in
the Sp1 site, most of the guanines in the promoter
showed only small
differences in reactivity to DMS in vitro and
in vivo. Nonetheless, our
data raise the possibility that in the
promoter also, there are
heretofore unrecognized binding sites
for regulatory factors.
| |
DISCUSSION |
Previous work using in vitro binding assays and/or transfection
studies have identified multiple cis-acting elements in the HPV18 URR enhancer and promoter. In vivo footprinting analyses performed in this study confirm that all of these elements are occupied
in vivo in both HeLa and C4-II cells. Importantly, these in vivo
analyses have also identified several additional candidate factor
binding sites; in both HeLa and C4-II cells, much of the proximal 300 bp of the URR exhibits changes in DMS reactivity in vivo suggestive of
factor occupancy (Fig. 6).
View larger version (81K):
[in this window]
[in a new window]
|
FIG. 6.
Summary of in vivo footprinting results. Guanines which
were hyporeactive to DMS in DNA isolated from DMS-treated cells are
indicated by open circles; guanines which were hyperreactive to DMS in
DNA isolated from DMS-treated cells are indicated by closed circles.
Several adenines which were particularly hyperreactive to DMS are also
indicated by closed circles.
|
|
One of the candidate sites identified in this study, nt 7675 to 7706, includes 10 bp which are a perfect match to a TEF-1 binding site
previously identified in the HPV16 enhancer (24). Importantly, a BLAST search using nt 7675 to 7706 as a query revealed very close matches in 17 other HPV types, including many which are
oncogenic for anogenital tissue, e.g., HPV16, -31, -33, -35, and -39. When used in an electrophoretic mobility shift assay, an
oligonucleotide corresponding to nt 7675 to 7706 forms a
sequence-specific complex with proteins in whole-cell extracts of HeLa,
C4-II, and C33A (HPV-negative cervical carcinoma) cells
(43). Studies to characterize the factors which interact
with this region and to assess its contribution to HPV18 URR-driven
transcription are in progress.
Interestingly, changes in DMS reactivity patterns were seen at two
other candidate TEF-1 binding sites in the promoter. One of these
regions, nt 7639 to 7653, has also been identified as a putative GRE
(6, 31). Mutation of this site, while abolishing glucocorticoid responsiveness of a transfected URR reporter construct, also increased its basal expression (6). Such an effect
would be consistent with our footprinting data, which suggest basal occupancy of these sequences in vivo. It will be interesting to determine whether in C4-II cells, which exhibit increased HPV18 transcription after treatment with glucocorticoids (53),
concomitant changes in DMS reactivity patterns encompassing nt 7639 to
7653 are seen. The other candidate TEF-1 binding site, nt 7807 to 7815, located just downstream of the promoter AP1 site, has not been previously identified as a regulatory element. While we are very interested in the possibility that TEF-1 interacts with multiple sites
in the HPV18 URR, the highly degenerate nature of the TEF-1 binding
site mandates caution in predicting that a particular nucleotide
sequence will bind TEF-1 with high affinity.
We hope to verify that one or more factors can recognize other HPV18
URR sequence stretches in which multiple G's exhibit altered
reactivity to DMS in vivo, strongly suggestive of factor occupancy.
Because these sequences were not protected in any of three previously
performed DNase footprinting assays, we anticipate that demonstrating
binding in vitro may require manipulations of the assay conditions or
fractionation of the extracts to enrich for the putative binding
protein. Establishing an in vitro system in which factors will
recognize the site under investigation is a prerequisite for
identifying the factor, especially where the site does not resemble
known factor binding sites. Furthermore, the ability to determine which
bases are critical for binding allows one to more easily assess the
consequences on expression of mutating the candidate
cis-acting element.
Enhancer sequences previously shown to bind factors in vitro by DNase I
footprinting assays and proposed to be a binding site for a novel
keratinocyte specific factor, KFR-1 (29), exhibited one of
the most pronounced footprints seen in our analyses. Mutation of this
site in the context of the 230-bp HPV18 enhancer fragment diminished
enhancer activity by 80% in HeLa cells (26) and in primary
human keratinocytes (6). Coupled with these previous results, the observation that the KRF-1 binding site is clearly occupied in vivo underscores the importance of characterizing the
factor which recognizes it.
One of the more controversial aspects of HPV18 (and HPV16)
transcriptional regulation has been the role of TTGGC sites,
which are present in multiple copies and which have been proposed to bind NF1 (although its consensus motif is a partial palindrome TTGGCTN3AGCCAA). Only one of three DNase I
footprinting studies detected protections of the distal and proximal
pairs of TTGGC half-sites as well as the TTGGC at the enhancer-promoter
boundary (17). The second study reported a protection at the
proximal pair only (35), and the third study did not report
protections at any of the TTGGC motifs (14). Binding of NF1
to oligomers corresponding to either pair of half-sites or to the
Oct-1/TTGGC motif in the HPV18 enhancer could be demonstrated only when
purified NF1 was used (6), probably reflecting the
demonstrated low affinity of NF1 for nonpalindromic sites. Within the
context of the intact URR, mutating all three NF1 sites so as to
abolish binding of NF1 did not substantially reduce transcription;
however, in the context of the intact enhancer (nt 7510 to 7739) or a
distal enhancer fragment which extended through the AP1 site (nt 7510 to 7625), deleting the distal 73 bp so as to remove both the distal and
proximal pairs of TTGGC half-sites substantially reduced enhancer activity (6). To explain this discrepancy, it was suggested that the deleted sequences contained binding sites for additional factors. Although in vivo footprinting provides no insight into the
nature of the factor which occupies the five TTGGC or TTGGC-like sites
in the URR enhancer, it does suggest that all of them are occupied in
vivo. Equally important, our data do not reveal occupancy of sites
other than the TTGGC and AP1 motifs within the region from nt 7510 to
7625.
Our data demonstrating in vivo occupancy of the switch region,
sequences which have been shown to be necessary for the YY1 site in the
promoter to mediate activation rather than repression of transcription,
are in accord with those obtained in a previous study (2).
In this earlier study, which focused on the switch region, a single set
of top- and bottom-strand primers was used to confirm in vivo occupancy
of this site in both HeLa and C4-I cells (derived from the same tumor
as C4-II cells), but these primers also allowed visualization of DMS
reactivity patterns of flanking sequences as well. Consistent with our
results, changes in reactivity to DMS were also seen in sequences just
upstream of the switch region (nt 7682 to 7706) as well as in sequences downstream of the putative Oct-1/NF1 site (nt 7741 to 7757). However, our data also suggest occupancy of some sequences not detected in the
previous analysis, which may reflect our use of lower DMS concentrations or multiple primer sets which allowed us to view the
reactivity patterns at higher resolution.
Data generated from the application of in vivo footprinting to HPV18
transcriptional regulation suggest that the number and variety of
trans-acting factors which interact with the enhancer and
promoter are even greater than was previously thought. Occupancy of the
additional sites identified in our footprinting analyses awaits
confirmation by in vitro binding assays and mutagenesis studies. We
cannot rule out the possibility that causes other than factor
occupancy, e.g., distortions of the helix due to binding of factors to
adjacent sites, are responsible for some of the changes in DMS
reactivity that we observed. That packaging of DNA into chromatin does
not affect the reactivity of guanines to DMS has been thoroughly
documented, however: both we and others have footprinted genes which
are transcriptionally silent and inaccessible to transcription factors
and have seen identical DMS reactivity patterns in vitro and in vivo
(18, 37).
While the currently accepted model depicts the URR as occupied in large
part by well-characterized and commonly utilized transcription factors,
regulatory proteins whose features and target genes are unknown or less
well characterized may in fact play a major role in controlling
transcription of the HPV18 early genes or in regulating other aspects
of the viral life cycle. If further studies confirm this hypothesis,
the next challenge will be to identify these proteins and elucidate
their roles in HPV18 regulation.
| |
ACKNOWLEDGMENT |
This work was supported by the Evelyn Dyba Women's Health
Fellowship award to B.P.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Departments of
Obstetrics and Gynecology and Cell and Molecular Biology, Northwestern University Medical School, 303 E. Chicago Ave., Chicago, IL 60611-3008. Phone: (312) 503-7883. Fax: (312) 908-8773. E-mail:
schallma{at}casbah.acns.nwu.edu.
| |
REFERENCES |
| 1.
|
Bauknecht, T.,
P. Angel,
H.-D. Royer, and H. zur Hausen.
1992.
Identification of a negative regulatory domain in the human papillomavirus type 18 promoter: interaction with the transcriptional repressor YY1.
EMBO J.
11:4607-4617[Medline].
|
| 2.
|
Bauknecht, T.,
F. Jundt,
I. Herr,
T. Oehler,
H. Delius,
Y. Shi,
P. Angel, and H. zur Hausen.
1995.
A switch region determines the cell-type-specific positive or negative action of YY1 on the activity of the human papillomavirus type 18 promoter.
J. Virol.
69:1-12[Abstract].
|
| 3.
|
Bauknecht, T.,
R. H. See, and Y. Shi.
1996.
A novel C/EBP B-YY1 complex controls the cell-type-specific activity of the human papillomavirus type 18 upstream regulatory region.
J. Virol.
70:7695-7705[Abstract].
|
| 4.
|
Becker, P. B., and G. Schutz.
1988.
Genomic footprinting, p. 1-19. In
J. K. Setlow (ed.), Genetic engineering, principles and methods, vol. 10.
Plenum Press, New York, N.Y.
|
| 5.
|
Berumen, J.,
E. R. Unger,
L. Casas, and P. Figueroa.
1995.
Amplification of human papillomavirus types 16 and 18 in invasive cervical cancer.
Hum. Pathol.
26:676-681[Medline].
|
| 6.
|
Butz, K., and F. Hoppe-Seyler.
1993.
Transcriptional control of human papillomavirus (HPV) oncogene expression: composition of the HPV type 18 upstream regulatory region.
J. Virol.
67:6476-6486[Abstract/Free Full Text].
|
| 7.
|
Chong, T.,
W.-K. Chan, and H.-U. Bernard.
1990.
Transcriptional activation of human papillomavirus 16 by nuclear factor 1, AP1, steroid receptors and a possibly novel transcription factor, PVF: a model for the composition of genital papillomavirus enhancers.
Nucleic Acids Res.
18:465-470[Abstract/Free Full Text].
|
| 8.
|
Chong, T.,
D. Apt,
B. Gloss,
M. Isa, and H.-U. Bernard.
1991.
The enhancer of human papillomavirus type 16: binding sites for the ubiquitous transcription factors Oct-1, NFA, TEF-2, NF1, and AP1 participate in epithelial cell-specific transcription.
J. Virol.
65:5933-5943[Abstract/Free Full Text].
|
| 9.
|
Cole, S. T., and O. Danos.
1987.
Nucleotide sequence and comparative analysis of the human papillomavirus type 18 genome.
J. Mol. Biol.
193:599-608[Medline].
|
| 10.
|
Cripe, T. M.,
T. H. Haugen,
J. P. Turk,
F. Tabatabai,
P. G. Schmid III,
M. Durst,
L. Gissman,
A. Roman, and L. B. Turek.
1987.
Transcriptional regulation of the human papillomavirus-16 E6-E7 promoter by a keratinocyte-dependent enhancer, and by viral E2 trans-activator and repressor gene products: implications for cervical carcinogenesis.
EMBO J.
6:3745-3753[Medline].
|
| 11.
|
Cullen, A. P.,
R. Reid,
M. Campion, and A. T. Lorincz.
1991.
Analysis of the physical state of different human papillomavirus DNAs in intraepithelial and invasive cervical neoplasm.
J. Virol.
65:606-612[Abstract/Free Full Text].
|
| 12.
|
Davidson, I.,
J. H. Xiao,
R. Rosales,
A. Staub, and P. Chambron.
1988.
The HeLa cell protein TEF-1 binds specifically and cooperatively to two SV40 enhancer motifs of unrelated sequences.
Cell
54:931-942[Medline].
|
| 13.
|
El Awady, M. K.,
J. B. Kaplan,
S. J. O'Brien, and R. D. Burk.
1987.
Molecular analysis of integrated human papillomavirus 16 sequences in the cervical cancer cell line SiHa.
Virology
159:389-398[Medline].
|
| 14.
|
Garcia-Carranca, A.,
F. Thierry, and M. Yaniv.
1988.
Interplay of viral and cellular proteins along the long control region of human papillomavirus type 18.
J. Virol.
62:4321-4330[Abstract/Free Full Text].
|
| 15.
|
Garrity, P. A.,
D. Chen,
E. V. Rothenberg, and B. J. Wold.
1994.
Interleukin-2 transcription is regulated in vivo at the level of coordinated binding of both constitutive and regulated factors.
Mol. Cell. Biol.
14:2159-2169[Abstract/Free Full Text].
|
| 16.
|
Gloss, B.,
H. U. Bernard,
K. Seedorf, and G. Klock.
1987.
The upstream regulatory region of the human papilloma virus-16 contains an E2 protein-independent enhancer which is specific for cervical carcinoma cells and regulated by glucocorticoid hormones.
EMBO J.
6:3735-3743[Medline].
|
| 17.
|
Gloss, B.,
M. Yeo-Gloss,
M. Meisterernst,
L. Rogge,
E. L. Winnacker, and H.-U. Bernard.
1989.
Clusters of nuclear factor I binding sites identify enhancers of several papillomaviruses but alone are not sufficient for enhancer function.
Nucleic Acids Res.
17:3519-3533[Abstract/Free Full Text].
|
| 18.
|
Gorzowski, J. J.,
C. A. Eckerley,
R. G. Halgren,
A. B. Mangurten, and B. Phillips.
1995.
Methylation-associated transcriptional silencing of the MHC-linked hsp70 genes in mouse cell lines.
J. Biol. Chem.
270:26940-26949[Abstract/Free Full Text].
|
| 19.
|
Heck, D. V.,
C. L. Yee,
P. M. Howley, and K. Munger.
1992.
Efficiency of binding the retinoblastoma protein correlates with the transforming capacity of the E7 oncoproteins of the human papillomaviruses.
Proc. Natl. Acad. Sci. USA
89:4442-4446[Abstract/Free Full Text].
|
| 20.
|
Hoppe-Seyler, F.,
K. Butz, and H. zur Hausen.
1991.
Repression of the human papillomavirus type 18 enhancer by the cellular transcription factor Oct-1.
J. Virol.
65:5613-5618[Abstract/Free Full Text].
|
| 21.
|
Hoppe-Seyler, F., and K. Butz.
1992.
Activation of human papillomavirus type 18 E6-E7 oncogene expression by transcription factor SpI.
Nucleic Acids Res.
20:6701-6706[Abstract/Free Full Text].
|
| 22.
|
Hoppe-Seyler, F., and K. Butz.
1994.
Cellular control of human papillomavirus oncogene transcription.
Mol. Carcinog.
10:134-141[Medline].
|
| 23.
|
Hornstra, I. K., and T. P. Yang.
1993.
In vivo footprinting and genomic sequencing by ligation-mediated PCR.
Anal. Biochem.
213:179-193[Medline].
|
| 24.
|
Ishiji, T.,
M. J. Lace,
S. Parkkinen,
R. D. Anderson,
T. H. Haugen,
T. P. Cripe,
J. H. Xiao,
I. Davidson,
P. Chambron, and L. P. Turek.
1992.
Transcriptional enhancer factor (TEF)-1 and its cell-specific co-activator activate human papillomavirus-16 E6 and E7 oncogene transcription in keratinocytes and cervical carcinoma cells.
EMBO J.
11:2271-2281[Medline].
|
| 25.
|
Jacquemin, P.,
J. J. Hwang,
J. A. Martial,
P. Dolle, and I. Davidson.
1996.
A novel family of developmentally regulated mammalian transcription factors containing the TEA/ATTS DNA binding domain.
J. Biol. Chem.
271:21775-21785[Abstract/Free Full Text].
|
| 26.
|
Jeon, S., and P. F. Lambert.
1995.
Integration of human papillomavirus type 16 DNA into the human genome leads to increased stability of E6 and E7 mRNAs: implications for cervical carcinogenesis.
Proc. Natl. Acad. Sci. USA
92:1654-1658[Abstract/Free Full Text].
|
| 27.
|
Kaur, P.,
J. K. McDougall, and R. Cone.
1989.
Immortalization of primary human epithelial cells by cloned cervical carcinoma DNA containing human papillomavirus type 16 E6/E7 open reading frames.
J. Gen. Virol.
70:1261-1266[Abstract/Free Full Text].
|
| 28.
|
Liu, Z.,
J. Ghai,
R. S. Ostrow, and A. J. Faras.
1995.
The expression levels of the human papillomavirus type 16 E7 correlate with its transforming potential.
Virology
207:260-270[Medline].
|
| 29.
| Mack, D. H., and L. A. Laimins. A
keratinocyte-specific transcription factor, KRF-1, interacts with AP-1
to activate expression of human papillomavirus type 18 in squamous
epithelial cells. Proc. Natl. Acad. Sci. USA 88:9102-9106.
|
| 30.
|
McGhee, J. D., and G. Felsenfeld.
1979.
Reaction of nucleosome DNA with dimethyl sulfate.
Proc. Natl. Acad. Sci. USA
76:2133-2137[Abstract/Free Full Text].
|
| 31.
|
Medina-Martinez, O.,
N. Morales-Peza,
M. Yaniv,
A. Garcia-Carranca, and F. Thierry.
1996.
A single element mediates glucocorticoid hormone response of HPV18 with no functional interactions with AP1 or hbrm.
Virology
217:392-396[Medline].
|
| 32.
|
Mueller, P. R., and B. Wold.
1989.
In vivo footprinting of a muscle specific enhancer by ligation mediated PCR.
Science
246:780-785[Abstract/Free Full Text].
|
| 33.
|
Munger, K.,
W. C. Phelps,
V. Bubb,
P. M. Howley, and R. Schlegel.
1989.
The E6 and E7 genes of the human papillomavirus type 16 together are necessary and sufficient for transformation of primary human keratinocytes.
J. Virol.
63:4417-4421[Abstract/Free Full Text].
|
| 34.
|
Murphy, S. P.,
J. J. Gorzowski,
K. D. Sarge, and B. Phillips.
1994.
Characterization of constitutive HSF2 DNA binding activity in mouse embryonal carcinoma cells.
Mol. Cell. Biol.
14:5309-5317[Abstract/Free Full Text].
|
| 35.
|
Nakshatri, H.,
M. M. Pater, and A. Pater.
1990.
Ubiquitous and cell-type-specific protein interactions with human papillomavirus type 16 and type 18 enhancers.
Virology
178:92-103[Medline].
|
| 36.
|
O'Connor, M., and H.-U. Bernard.
1995.
Oct-1 activates the epithelial-specific enhancer of human papillomavirus type 16 via a synergistic interaction with NF1 at a conserved composite regulatory element.
Virology
207:77-88[Medline].
|
| 37.
|
Pfeifer, G. P.,
R. L. Tanguay,
S. D. Steigerwald, and A. D. Riggs.
1990.
In vivo footprint and methylation analysis by PCR-aided genomic sequencing: comparison of active and inactive X chromosomal DNA at CpG island and promoter of human PGK-1.
Genes Dev.
4:1277-1287[Abstract/Free Full Text].
|
| 38.
|
Remm, M.,
R. Brain, and J. R. Jenkins.
1992.
The E2 binding sites determine the efficiency of replication for the origin of human papillomavirus type 18.
Nucleic Acids Res.
20:6015-6021[Abstract/Free Full Text].
|
| 39.
|
Romanczuk, H.,
L. L. Villa,
R. Schlegel, and P. M. Howley.
1991.
The viral transcriptional regulatory region upstream of the E6 and E7 genes is a major determinant of the differential immortalization activities of human papillomavirus types 16 and 18.
J. Virol.
65:2739-2744[Abstract/Free Full Text].
|
| 40.
|
Saluz, H. P., and J. P. Jost.
1993.
Approaches to characterize protein-DNA interactions in vivo.
Crit. Rev. Eukaryotic Gene Expression
3:1-29[Medline].
|
| 41.
|
Scheffner, M.,
B. A. Werness,
J. M. Huibregtse,
A. J. Levine, and P. M. Howley.
1990.
The E6 oncoprotein encoded by human papillomavirus types 16 and 18 promotes the degradation of p53.
Cell
63:1129-1136[Medline].
|
| 42.
|
Schneider-Gadicke, A., and E. Schwarz.
1986.
Different human cervical carcinoma cell lines show similar transcription patterns of human papillomavirus type 18 early genes.
EMBO J.
5:2285-2292[Medline].
|
| 43.
| Schrock, A., and B. Phillips. Unpublished data.
|
| 44.
|
Schwarz, E.,
U. K. Freese,
L. Gissmann,
W. Mayer,
B. Roggenbuck,
A. Stremlau, and H. zur Hausen.
1985.
Structure and transcription of human papillomavirus sequences in cervical carcinoma cells.
Nature
314:111-114[Medline].
|
| 45.
|
Stoler, M. H.,
C. R. Rhodes,
A. Whitbeck,
S. M. Wolinsky,
L. T. Chow, and T. R. Broker.
1992.
Human papillomavirus type 16 and 18 gene expression in cervical neoplasias.
Hum. Pathol.
23:117-128[Medline].
|
| 46.
|
Swift, F. V.,
K. Bhat,
H. B. Younghusband, and H. Hamada.
1987.
Characterization of a cell type-specific enhancer found in the human papilloma virus type 18 genome.
EMBO J.
6:1339-1344[Medline].
|
| 47.
|
Thierry, F.,
A. Garcia-Carranca, and M. Yaniv.
1987.
Elements that control the transcription of genital papillomavirus type 18.
Cancer Cells
5:23-32.
|
| 48.
|
Thierry, F.,
J. M. Heard, and M. Yaniv.
1987.
Characterization of a transcriptional promoter of human papillomavirus 18 and modulation of its expression by simian virus 40 and adenovirus early antigens.
J. Virol.
61:134-142[Abstract/Free Full Text].
|
| 49.
|
Thierry, F.,
G. Spyrou,
M. Yaniv, and P. Howley.
1992.
Two AP1 sites binding junB are essential for human papillomavirus type 18 transcription in keratinocytes.
J. Virol.
66:3740-3748[Abstract/Free Full Text].
|
| 50.
|
Tommasi, S., and G. P. Pfeifer.
1995.
In vivo structure of the human cdc2 promoter: release of a p130-E2F-4 complex from sequences immediately upstream of the transcription initiation site coincides with induction of cdc2 expression.
Mol. Cell. Biol.
15:6901-6913[Abstract].
|
| 51.
|
Turek, L. P.
1994.
The structure, function, and regulation of papillomavirus genes in infection and cervical cancer.
Adv. Virus Res.
44:305-356[Medline].
|
| 52.
|
von Knebel Doeberitz, M.,
T. Oltersdorf,
E. Schwartz, and L. Gissmann.
1988.
Correlation of modified human papilloma virus early gene expression with altered growth properties in C4-1 cervical carcinoma cells.
Cancer Res.
48:3780-3786[Abstract/Free Full Text].
|
| 53.
|
von Knebel Doeberitz, M.,
T. Bauknecht,
D. Bartsch, and H. zur Hausen.
1991.
Influence of chromosomal integration on glucocorticoid-regulated transcription of growth-stimulating papillomavirus genes E6 and E7 in cervical carcinoma cells.
Proc. Natl. Acad. Sci. USA
88:1411-1415[Abstract/Free Full Text].
|
| 54.
|
von Knebel Doeberitz, M.,
C. Rittmuller,
F. Aengeneyndt,
P. Jansen-Durr, and D. Spitkovsky.
1994.
Reversible repression of papillomavirus oncogene expression in cervical carcinoma cells: consequences for the phenotype and E6-p53 and E7-pRB interactions.
J. Virol.
68:2811-2821[Abstract/Free Full Text].
|
| 55.
|
Xiao, J. H.,
I. Davidson,
H. Matthes,
J. M. Garnier, and P. Chambon.
1991.
Cloning, expression, and transcriptional properties of the human enhancer factor TEF-1.
Cell
65:551-568[Medline].
|
| 56.
|
zur Hausen, H.
1991.
Human papillomavirus in the pathogenesis of anogenital cancer.
Virology
184:9-13[Medline].
|
J Virol, January 1998, p. 708-716, Vol. 72, No. 1
0022-538X/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
