Interactions with Single-stranded and Double-stranded DNA-binding
Factors and Alternative Promoter Conformation upon Transcriptional
Activation of the Htf9-a/RanBP1 and
Htf9-c Genes*
Gigliola
Di Matteo
§,
Massimiliano
Salerno
¶,
Giulia
Guarguaglini
,
Barbara
Di Fiore
,
Franco
Palitti**, and
Patrizia
Lavia

From the
CNR Centre of Evolutionary Genetics, c/o
Department of Genetics and Molecular Biology and the ** Department of
Biochemical Sciences, University "La Sapienza,"
Rome 00185, Italy
 |
ABSTRACT |
The murine Htf9-a/RanBP1 and
Htf9-c genes are divergently transcribed from a
shared TATA-less promoter. Transcription of both genes is initiated on
complementary DNA strands and is controlled by cell
cycle-dependent mechanisms. The bidirectional promoter harbors a genomic footprint flanking the major transcription start site
of both genes. Transient promoter assays showed that the footprinted
element is important for transcription of both genes. Protein-binding
experiments and antibody assays indicated that members of the retinoid
X receptor family interact with the double-stranded site. In addition,
distinct factors interact with single DNA strands of the element.
Double-stranded binding factors were highly expressed in liver cells,
in which neither gene is transcribed, while single-stranded binding
proteins were abundant in cycling cells, in which transcription of both
genes is efficient. In vivo S1 analysis of the promoter depicted an S1-sensitive organization in cells in which transcription of both genes is active; S1 sensitivity was not detected in conditions of transcriptional repression. Thus, the same element is a target for
either retinoid X receptor factors, or for single-stranded binding
proteins, and form distinct complexes in different cellular conditions
depending on the DNA conformation in the binding site.
 |
INTRODUCTION |
The murine Htf9 locus was isolated by virtue of
its association with a CG-rich genomic sequence (1) and mapped to mouse chromosome 16 (2). The locus contains two transcriptional units, Htf9-a and Htf9-c, that are
transcribed with opposite polarity from complementary DNA strands (see
map in Fig. 1A). The lower strand gene, called
Htf9-a, encodes Ran-binding protein 1 (RanBP1)1 (3, 4), an
interacting partner of the Ran GTPase, which is thought to cooperate in
control of several processes regulated by the Ran network and including
DNA replication, cell cycle progression, mitotic entry and exit,
nuclear structure, and nucleo/cytoplasmic transport (reviewed in Refs.
5 and 6). The Htf9-c gene codes for a protein sharing
extensive homologies with nucleic acid-modifying enzymes, including
bacterial tRNA methyltransferases and yeast DNA endo-exonucleases (7);
the evolutionary conservation of the protein sequence in the nucleic
acid-interacting domain suggests that the murine
Htf9-c gene product may also be implicated in nucleic
acid metabolism and/or modifications.
Both Htf9 divergently transcribed genes are expressed
at low levels in many tissues and cell types (1, 8) and are
up-regulated in proliferating cells: transcription is activated at the
G1/S transition of the cell cycle and peaks in S phase,
while being repressed in quiescent tissues and growth-arrested cells
(7, 9). Promoter deletion and site-directed mutagenesis assays have
revealed that cell cycle expression of both divergently transcribed genes is controlled by separate E2F and Sp1 promoter elements active in
each orientation (7).
Both the RanBP1 and Htf9-c transcriptional
units are initiated in a genomic sequence that has typical features of
the CG-rich class of mammalian promoters, which are mainly, although
not exclusively, associated with housekeeping genes. An intriguing
feature shared by most promoters of the housekeeping class is the
absence of a TATA box, whose function in transcriptional initiation is
well established for all three classes of RNA polymerase (reviewed in
Refs. 10-12). In most TATA-less promoters the DNA region surrounding the major transcriptional start site(s) plays a crucial role in directing initiation (see Refs. 11 and 13 for reviews). Like many
CG-rich promoters, the Htf9 bidirectional promoter
harbors no TATA box on either DNA strand; RNA transcription is
initiated at multiple sites on each strand; transcription start 1 (TS-1 in Fig. 1B) is the most frequently used start site on both
DNA strands (1). Deletion mapping analysis (7, 14) restricted the
promoter region required for basal transcription to a 74-bp long region
active in both orientations. Within that region, genomic footprinting
experiments depicted a site protected by unidentified factor(s),
located 20 bp apart from a functional Sp1-binding site and flanking
TS-1 (15).
The present study was undertaken to characterize the
Htf9 footprinted element (HFE) adjacent to TS-1. In
an initial set of experiments, we found that double-stranded HFE
interacts with factors belonging to the family of retinoid X receptors
(RXRs). In addition, distinct factors recognize each single HFE strand. The distinct DNA binding activities depicted with different forms of
the HFE element might have indicated that the Htf9
initiation region was organized either in the double-stranded or in the
single-stranded conformation in different cellular conditions. Since
both Htf9-associated genes are transcribed in a cell
cycle-dependent manner, we sought to determine whether the
interaction of factors with HFE varies in relation to proliferation and
transcription. Our findings show that the double-stranded nucleoprotein
complex is efficiently assembled with extracts from liver cells, in
which transcription from the Htf9 locus is extremely
low or absent, while single stranded DNA binding activities are
abundant in cycling NIH/3T3 fibroblasts, in which both
Htf9-associated genes are actively transcribed. In
addition, the region surrounding HFE acquires an S1-sensitive conformation in vivo in cycling, but not in growth-arrested,
NIH/3T3 cells, where transcriptional repression occurs. Therefore, the Htf9 promoter appears to assume alternative
conformations in relation to transcription. These data suggest that
factors of the RXR family and the newly identified single-stranded
binding proteins participate to transcriptional control of the
Htf9-associated genes by interacting with alternative
forms of the HFE element in different cellular conditions.
 |
EXPERIMENTAL PROCEDURES |
Cell Cultures--
Murine NIH/3T3 embryo fibroblasts (ATCC CRL
1658) were grown in Dulbecco's modified Eagle's medium supplemented
with 10% (v/v) fetal calf serum under 5% CO2 at 37 °C.
Asynchronously cycling cells were collected from 60-70% confluent
cultures. Proliferation arrest was induced by culturing the cells in
medium containing 0.5% fetal calf serum for at least 48 h. To
obtain S phase-enriched cultures, cell cycle reentry was induced after
starvation by adding 15% fetal calf serum and harvesting 15 h
after stimulation. Growth arrest and cell cycle progression were
monitored by flow cytometry as described (9). Cell samples were
analyzed in a FACStar Plus cytofluorimeter using the WinMDI software
(10.000 events/sample).
Protein Extract Preparation and Fractionation--
Protein
extracts were prepared from both NIH/3T3 fibroblast cultures and from
livers from 3-4-week-old black C57 mice after nuclei isolation as
described previously (8). All buffers and solutions contained 1 mM phenylmethylsulfonyl fluoride, 0.5 mM dithiothreitol, and 50 µg/ml antipain, leupeptin, chymostatin, and
pepstatin A. For chromatographic fractionation, NIH/3T3 nuclei were
isolated by centrifugation through a 0.8 M sucrose cushion overlaid by 0.3 M sucrose solution. Proteins were extracted
in 0.42 M KCl buffer and sequentially precipitated;
pelleted proteins from the 30% ammonium sulfate precipitation were
redissolved in D buffer (20 mM Hepes, pH 7.9, 20 mM KCl, 2 mM MgCl2, 0.2 mM EDTA, 20% glycerol), dialyzed, and again precipitated
using 60% ammonium sulfate. Proteins in the supernatant were loaded
onto a phosphocellulose column and subjected to chromatography.
Recovered fractions were extensively dialyzed against D buffer,
subjected to chromatography through DEAE 52 cellulose, and finally
recovered by step elution using KCl.
Gel Shift Assays--
Oligonucleotides were synthesized at
Genenco Service (Department of Genetics and Molecular Biology,
University "La Sapienza," Rome), except for the RXR-binding
oligonucleotide (DR-1 configuration) and its mutagenized version
(MDR-1), which were from Santa Cruz Biotechnology. Oligonucleotide
sequences are indicated in the text. Gel shift reactions were set up
with 20-100 pg of [
32P]ATP-labeled oligonucleotide,
5-7 µg of NIH/3T3 cell or liver extract, 1 µg of
poly(dI-dC)·poly(dI-dC), 25 mM Hepes (pH 7.6), 10%
glycerol, 1 mM dithiothreitol, 0.5 mM EDTA, and
50 mM KCl for 15 min on ice. Slighlty different conditions
were used for ssG binding: 1 µg of poly(dI-dC)·poly(dI-dC), 500 ng
of nonspecific single-stranded DNA, 20 mM Hepes, pH 7.9, 10% glycerol, 1 mM dithiothreitol, 1 mM EDTA,
0.1 mM EGTA, 60 mM KCl and 0.1% Nonidet-P 40. In certain experiments, sodium deoxycholate (0.8% final concentration)
was added either to the protein mixture before addition of the probe, or to the complete reaction, and incubated for 10 min on ice; incubations were terminated by adding Nonidet-P 40 (1.5% final concentration). Where required reactions were preincubated with competitor DNA for 10 min on ice before adding the probe. Supershift reactions were set up as above, except that 2 µl (2 µg) of anti-RXR or anti-retinoic acid receptors (RAR) antibody were added to the binding mixtures and further incubated for 45 min at room temperature. Anti-RXR and anti-RAR specificities are described below.
Western Blot Assays--
Nuclear protein extracts from cycling
and growth-arrested NIH/3T3 cells, as well as from liver fractions,
were electrophoresed through 10% SDS-polyacrylamide gel,
electroblotted in 48 mM Tris-HCl, pH 8.3, 39 mM
glycine, 0.037% SDS, and 20% methanol on nitrocellulose membranes and
analyzed by Western blotting using either anti-RXR or anti-RAR
antibodies (1:1000 dilution). The anti-RXR (
N 197, Santa Cruz
sc-774) reacts with all murine RXR subtypes (RXR
, RXR
, and
RXR
), but not with RAR factors; the anti-RAR (M-454, Santa Cruz
sc-773) reacts with all RAR subtypes (
1,
2,
1,
2,
1,
and
2) but not with RXR factors. Membranes were
incubated with horseradish peroxidase-conjugated secondary antibody
(1:20.000 dilution), and bands were revealed following the enhanced
chemiluminescence protocol (ECL reagents, Amersham Corp.).
Plasmid Constructs--
The pEA-A and pEA-C constructs were
generated by inserting the EarI-AluI fragment
from the Htf9 promoter (X05830 sequence) in both
orientations upstream of the chloramphenicol acetyltransferase (CAT)
sequence; the pES-A and pES-C constructs were generated by inserting
the EarI-SmaI fragment from the
Htf9 promoter in both orientations upstream of the
CAT sequence. HFE-mutated constructs carried an oligonucleotide
identical to the EarI-AluI fragment, except that
HFE was mutated to
5
-GAACTCTGATCTCTGGTCTCAGC-3
and 5
-GCTGAGACCAGAGATCAGAGTTC-3
on complementary strands (C
T and G
A mutations are
underlined). The fragment was cloned in both orientations upstream of
the CAT sequence, yielding the pEA-mA and pEA-mC constructs
respectively, or ligated to the AluI-SmaI fragment from the Htf9 promoter to yield the pES-mA
and pES-mC constructs. In an independent series of constructs, the HFE
oligonucleotide was ligated in multiple copies downstream or upstream
of the pA10.CAT2 promoter, which carries two copies of the SV40 21-bp
repeat (including Sp1 sites) and a TATA box (16).
Transfections--
NIH/3T3 cells were routinely diluted 1:6 the
day before transfection. Typical experiments were carried out using
5 × 106 cells and a mixture composed of DOTAP
liposome reagent (Boehringer Mannheim), 5 µg of CAT reporter DNA, and
1 µg of pCMV-lacZ plasmid. The medium was changed 6 h after
lipofection, and cells were harvested 36-48 h later. In all
experiments, mock-transfected cultures were harvested and analyzed by
flow cytometry as a control to verify that cells were actively
proliferating. Promoter strengths were quantified by immunoenzymatic
staining of the CAT protein (CAT enzyme-linked immunosorbent assay kit,
Boehringer Mannheim) and normalized relative to the amount of
synthesized
-galactosidase from the co-transfected plasmid
(
-galactosidase enzyme-linked immunosorbent assay, Boehringer
Mannheim). pSV0 or pCAT promoterless vectors (Promega) were used as
negative controls. Four to eight transfections experiments were carried
out for each construct.
Northern Blots--
NIH/3T3 cultures were growth-arrested or
synchronized in S phase as described above. Total RNA was extracted
following the guanidine-acid phenol method, electrophoresed, stained
with ethidium bromide to visualize the 28 and 18 S ribosomal bands, and
transferred to GeneScreen membranes as reported in detail elsewhere
(9). Probes used for Northern hybridizations were gel-purified
fragments corresponding to the Htf9-c,
Htf9-a/RanBP1, and glyceraldehyde-3-phosphate dehydrogenase coding sequences.
S1 Analysis of Plasmids and Nuclear Chromatin--
Plasmid DNAs
were prepared using Qiagen columns, which yield approximately 75% of
all DNA molecules in the supercoiled form. Plasmids were subjected to
S1 digestion in S1 buffer (3 mM ZnCl2, 30 mM sodium acetate, pH 4.5, 30 mM NaCl, and 0.2 mM EDTA) for 30 min at 37 °C. Nuclei from
growth-arrested or S phase NIH/3T3 cells were resuspended in buffer A
(homogenization buffer without sucrose) and digested with increasing
amounts of S1 nuclease (Amersham) in S1 buffer as above, except that
300 mM NaCl was used. Digestions were stopped by adding 10 mM EDTA, 10 mM Tris-HCl, pH 7.6, 0.1% SDS, and
0.1 mg/ml proteinase K at 37 °C for 5 h. Genomic DNA was
repeatedly extracted with phenol and phenol-chloroform, precipitated, resuspended, digested with EcoRI, electrophoresed, and
blotted with conventional methods. Terminal probes for indirect
end-labeling experiments were purified from the pL9.2 subclone, which
contains the Htf9 bidirectional promoter (X05830
sequence); probes were prepared by double digestion either with
EcoRI and HindIII, yielding a 315-bp probe from
the 3
region, or with EcoRI and EcoRV, yielding a 485-bp probe from the 5
region of the Htf9 insert,
eluted from preparative gels, uniformly labeled, hybridized to the
filter-bound DNA, and washed using standard procedures.
 |
RESULTS |
Mutations within the Footprinted Site Flanking TS-1 Impair Promoter
Activity--
In a previous characterization of the
Htf9 locus (shown in Fig.
1A), we mapped the full-length
bidirectional promoter to a 273-bp fragment (14), which was
subsequently found to contain both basal and cell cycle control
elements in each orientation. Deletion of the elements responsible for
cell cycle control of each gene yielded a 74-bp fragment, called EA to
design the EarI-AluI restriction ends (see Fig.
1B), which directed basal transcription in both directions
(7). Genomic footprinting identified functional sites of protein
interaction in the fragment (15). Among those, a footprint (framed in
Fig. 1B) exactly starts at the nucleotide adjacent to the
major start site of transcription (TS-1 in Fig. 1B) used for
initiation of both the Htf9-a/RanBP1 and
Htf9-c mRNA transcripts. We set out to establish
whether the sequence defined by the footprint, called
Htf9 footprinted element (HFE), identified a
functional element.

View larger version (30K):
[in this window]
[in a new window]
|
Fig. 1.
The Htf9 bidirectional
promoter. A, map of the murine Htf9 locus,
showing the 5 ends of the Htf9-a/RanBP1 and
Htf9-c genes, the major transcription start site of
both genes (vertical arrows), coding sequences (filled
boxes), untranslated regions (empty boxes), and introns
(lines). The start codon in each open reading frame is
marked and the orientation of transcription is arrowed. B,
sequence of the Htf9 promoter in the
RanBP1 orientation. Footprinted sites are
underlined; identified footprinting factors are indicated
above the sequence. The HFE footprint is framed. The
EarI, SmaI, and AluI sites used in
promoter reporter construction are indicated. C, promoter
regions assayed in transient experiments; dotted lines
represent deleted regions. Mutations within HFE are represented by the
crossed box. The graph on the right shows the relativity
activity of constructs; values were calculated in pg of CAT enzyme/100
µg of protein extract and normalized relative to the amount of
-galactosidase from a cotransfected plasmid. Mean values and S.D.
were calculated from four to eight experiments for each construct. pSV0
is the empty vector.
|
|
An oligonucleotide corresponding to HFE was saturated with mutations
(see "Experimental Procedures") and assayed in either orientation,
both in the context of the 74-bp basal promoter (pEA series), and
within a larger promoter fragment (180 bp) harboring activator binding
sites (pES series); assayed constructs in transient expression
experiments in NIH/3T3 cells are shown in Fig. 1C. In the
orientation of the Htf9-C gene, HFE inactivation
significantly reduced the basal activity of the pEA-C promoter, whose
strength decreased by 50%; a milder effect (25% reduction) was
recorded in the 180-bp pES-C construct. In the RanBP1
orientation, HFE mutation lowered activity of the 180-bp pES-A promoter
by approximately 50%; it was difficult to determine whether the
reduction was significant in the pEA-A basal promoter, which worked
per se with very low efficiency. Thus, HFE contributes to
promoter activity in both orientations, although the effect of
mutations is sensed differently in the context of different promoter
architectures.
To assess whether HFE acted as an activator-binding element, the
pA10.CAT2 vector was used, which contains a minimal promoter composed
of two Sp1-binding sites and a TATA box (16). If HFE functioned as a
classical modular promoter element, multimerized copies should activate
transcription from pA10.CAT2. No difference was instead detected in the
efficiency of chimeric promoter constructs bearing one to four HFE
copies upstream or downstream of the TATA box compared with that of
pA10.CAT2 alone (data not shown). These results suggest that HFE does
not identify a strong activator-binding element, yet its integrity in
the Htf9 promoter is required for full activity in
both orientations in NIH/3T3 cells.
Sequence Requirements for Protein Binding to HFE--
As an
initial step to characterize the factors conferring the protection
flanking TS-1, an oligonucleotide was synthesized from the footprinted
HFE window and incubated with NIH/3T3 cell extracts. A discrete
nucleoprotein complex was detected by gel-shift assays (Fig.
2B, lanes 1,
11, and 14). Since the HFE location near the
transcription start is similar to that of certain initiators, competition assays were designed with oligonucleotides characterized for their ability to initiate transcription in TATA-less promoters, including the adenovirus Inr initiator element and binding
sites for the YY1, USF, and Sp1 factors (see Weis and Reinberg (13) and
references therein). A canonical TATA box was also assayed, since
transcription initiation involves direct or indirect interaction of the
TBP factor with several TATA-less promoters (17-19). None of the
tested sequences interfered with HFE complex assembly; the complex was
also insensitive to the addition of anti-TFIID antibodies and failed to
interact with purified recombinant TBP protein (data not shown). Thus,
the HFE complex does neither include TBP nor characterized
initiator-binding proteins.

View larger version (48K):
[in this window]
[in a new window]
|
Fig. 2.
The HFE site. A, HFE sequence and
mutagenized versions; only the strand corresponding to the
RanBP1 orientation is shown. Repeats in the wild-type site
(WT) are underlined, base substitutions are
indicated by asterisks. B, gel shift assays of wild-type HFE
with 5 µg of NIH/3T3 extract (lanes 1, 11, and 14) and increasing amounts of wild-type (lanes 2 and 3) or mutated (lanes 4-10) oligonucleotides
(50- and 100-fold excess of competitor were used in each set of
reactions), after protein extract preincubation with sodium
deoxycholate (NaDOC) (lane 12), and after
NaDOC addition to the assembled complex (lane 13).
|
|
The HFE sequence (5
-GGGTCAGGGGTCAGGG-3
) harbors a tandem repeat in
several combinations, GGTCAGG, GGGTCA, or TCAGGG. To define the
sequence requirement for protein binding, mutated HFE versions were
used as competitors (shown in Fig. 2A). M0 was mutated throughout HFE and used in transient expression experiments (Fig. 1C); M1 and M2 were, respectively, mutagenized in the distal
(leftward) or proximal (rightward) repeat relative to TS-1; M3 carried
mutations in the central region of the oligonucleotide which affected
both repeats. The results in Fig. 2B show that M3, although
carrying only three base substitutions, like M1 and M2, was unable to
compete for protein binding to wild-type HFE. In contrast, both M1 and M2, each of which retained one repeat, partly competed with HFE. These
experiments indicate that the GGGTCAGGGGTCAGGG sequence represents the
optimal binding site; the low efficiency of the competition by M1 and
M2 shows that their affinity for HFE-binding factor(s) was not
completely abolished yet was significantly reduced compared with the
wild-type site. These observations suggest that the HFE complex may be
stabilized by interactions among proteins binding to adjacent repeats.
Indeed, preincubation of protein extracts with sodium deoxycholate,
which disrupts weak protein-protein interactions, prevented the complex
assembly (Fig. 2B, lane 12); furthermore,
deoxycholate addition to the DNA/protein binding reaction disrupted the
assembled complex (Fig. 2B, lane 13). The deoxycholate effect was specific because it did not affect the interaction of Sp1, which binds as a monomer, with its DNA target site
(data not shown).
HFE Is Recognized by Members of the RXR Family--
The repeated
structure and sequence of HFE are similar to those contained in gene
promoters regulated by retinoic acid, known as retinoic acid response
elements, that are composed of a direct repetition of the PuGGTCA motif
and are recognized by members of two large families of transcription
factors, RARs and RXRs. Searching transcription factor data bases with
the HFE sequence depicted the highest degree of raw homology with the
H2RII element in the major histocompatibility complex class I gene
promoter (Fig. 3A), a target
of the RXR
subtype of retinoid X receptors (20). Unlabeled H2RII
interfered with the HFE complex assembly (Fig. 3A,
lanes 6-9) almost as effectively as the homologous
competitor (Fig. 3, A, lanes 2-5, and
C). Reciprocal experiments using labeled H2RII as the probe
showed that homologous competitions were highly effective (Fig.
3A, lanes 10-14), while competition by unlabeled HFE required a 50-fold excess to achieve 50% inhibition and plateaued thereafter (Fig. 3, A, lanes 15-18, and
B). The finding that H2RII was an effective competitor of
HFE, while HFE only partially competed for factors binding to the H2RII
probe, suggests that H2RII and HFE are bound by both common and
specific partners. We next used a canonical RXR-binding oligonucleotide
in which PuGGTCA repeats are separated by one-nucleotide spacer (DR-1
configuration), as in HFE; extensive characterization of DR-1 had shown
it to be the preferred target sequence of RXR factors (21).
Heterologous competitions for factors between DR-1 and HFE were now
complete (Fig. 3D); thus, the binding properties of HFE are
indistinguishable from those of the DR-1 oligonucleotide. Together,
these results implicate RXR factors in binding to HFE.

View larger version (45K):
[in this window]
[in a new window]
|
Fig. 3.
HFE competes with RXR-binding sites.
A, HFE and major histocompatibility complex (MHC)-H2RII
sequences are shown at the top; single repeats are
framed. Below, competition assays of HFE with 5 µg of NIH/3T3 cell extract (lane 1) and increasing amounts
of unlabeled HFE (lanes 2-5) or H2RII site (lanes
6-9), and reciprocal assays using the H2RII probe (lane
10) with increasing amounts of unlabeled H2RII (lanes
11-14) or HFE (lanes 15-18) DNA. B, the
graph shows the ratio of complexed/total input DNA calculated from
microdensitometric scanning of H2RII gel shifts in the presence of the
indicated amounts of homologous or heterologous competitor.
C, complexed/total input DNA ratios calculated after microdensitometric scanning of HFE gel-shifts in the presence of the
indicated amounts of homologous or heterologous competitor. D, top, HFE and DR-1 sequences; repeats are
framed. Below, competitive inhibition of both the HFE
(lane 1) and DR-1 (lane 6) complexes formed with
NIH/3T3 extracts (5 µg) by preincubation with a 50-fold excess of
unlabeled HFE (lanes 2 and 7) or DR-1
(lanes 3 and 8) sites, but not with mutagenized
versions of either HFE (M3, lanes 4 and 9) or
DR-1 (MDR1, lanes 5 and 10).
|
|
Distinct RXR-containing Complexes Are Assembled with Extracts from
Liver and NIH/3T3 Fibroblasts--
Since in previous work the
strongest HFE in vitro footprints were detected using mouse
liver extracts (8), the study of HFE-binding factors was further
pursued using such extracts. We found that the HFE complex with liver
nuclear extracts had higher abundance, and lower electrophoretic
mobility, than seen using NIH/3T3 extracts (Fig.
4, compare lanes 1 and
7). Addition of an anti-RXR antibody recognizing all three
RXR subtypes (
,
, and
) to the HFE reaction with liver
extracts supershifted the complex in a manner comparable to that seen
with DR-1 (Fig. 4, lanes 2 and 5). An antibody
recognizing all members (
1,
2,
1,
2,
1, and
2) of the RAR family did neither affect the HFE nor the
DR-1 complex (Fig. 4, lanes 3 and 6). When
immunoassays were carried out using the same amount of NIH/3T3 nuclear
extract, no clear supershift was detected with either HFE or DR-1
probes; however, the amount of assembled complex with both probes
decreased with the addition of both anti-RXR (Fig. 4, lanes
8, 9, 13, and 14) and anti-RAR
(Fig. 4, lanes 10, 11, 15, and
16) antibodies, suggesting that component(s) of both the RXR
and RAR families interacted with HFE and DR-1 in NIH/3T3 cells. Thus,
the differences in abundance, mobility, and antibody reactivity
indicate that specific complexes are formed by both probes depending on
the extract source.

View larger version (105K):
[in this window]
[in a new window]
|
Fig. 4.
Double-stranded HFE forms distinct complexes
in different cell types. The HFE (lane 1) and DR-1
(lane 4) complexes formed with liver extracts are
supershifted (arrow) by anti-RXR (lanes 2 and
5) but not anti-RAR (lanes 3 and 6)
antibodies; neither HFE (lane 7) nor DR-1 (lane
12) complexes formed with NIH/3T3 extracts are visibly
supershifted, yet decrease, with increasing amounts of either anti-RXR
(lanes 8, 9, 13, and 14) or
anti-RAR (lanes 10, 11, 15, and
16) antibodies. Reactions were set up using 105
cpm of labeled probes and 6 µg of nuclear extract from either source.
|
|
The results in Fig. 4 suggest that the RXR supershift depicted with
liver extracts reflected the higher abundance of these factors in
liver, compared with NIH/3T3, nuclei. To further investigate that
possibility, liver extracts were fractionated through sequential phosphocellulose and DEAE 52 chromatography (Fig.
5A). All RXR subtypes (
,
, and
) recognized by the anti-RXR antibody were eluted with, and
were abundantly expressed in, fractions that were positive for HFE
binding (compare Fig. 5, B and C, upper panel). In NIH/3T3 extracts, only one subtype, whose
electrophoretic mobility was compatible with that expected for RXR
(22), was detected; the relative abundance of the reacted protein was
significantly lower than that of the corresponding liver protein, and
in cycling NIH/3T3 cells was less abundant than in growth-arrested
cultures (Fig. 5C, upper panel, lanes
8 and 9). In NIH/3T3 extracts we also detected RAR
factor(s) that had not been depicted in liver cells (Fig.
5C, lower panel). In summary, HFE is bound by RXR factors which associate with different partners and assemble specific complexes with different extract types. Abundant complexes are formed
with liver factors, among which RXRs are highly expressed. RXR-containing complexes have lower abundance in NIH/3T3 fibroblasts and include members of the RAR family. In retrospect, the cell type-specific reactivity in supershift assays reflects the different distribution and relative abundance of retinoid receptors depicted in
Western blot assays.

View larger version (28K):
[in this window]
[in a new window]
|
Fig. 5.
Cofractionation of the HFE-binding activity
and RXR factors from liver extracts. A, schematic of the
procedure used for fractionation of the HFE-binding activity from liver
extracts. AS, ammonium sulfate. B, gel-shift
assays of HFE with eluted fractions (indicated above each lane). 4 µg
of protein from nuclear extract (NE) and phosphocellulose
fractions (PC), and 2 µg of DEAE 52 (DE)
fractions were used. The addition of homologous competitor DNA is
indicated by +. C, Western blot assays of protein from liver
fractions and from growth-arrested or cycling NIH/3T3 cells with
anti-RXR (top panel) and anti-RAR (lower panel)
antibodies.
|
|
Distinct Proteins Bind to Each Strand of HFE in NIH/3T3
Fibroblasts--
Since the region surrounding the transcription start
of both Htf9 genes can be expected to open up during
transcription and expose single-stranded DNA templates to
transcriptional complexes, we wondered whether single DNA strands
interacted with proteins. The G-rich and C-rich strands of the HFE
oligonucleotide were separately incubated with NIH/3T3 extracts and
their ability to form nucleoprotein complexes was assessed in gel-shift
assays. All forms of the site (i.e. G-rich, C-rich, and
double-stranded) assembled discrete complexes (Fig.
6A). The complex formed with the C-rich strand was designed ssC (single-stranded C-rich
DNA-containing complex). The G-rich strand formed two predominant
complexes, ssG1 and ssG2 (single-stranded G-rich DNA-containing
complexes 1 and 2) and two fainter complexes of lower mobility. The
specificity of the complexes was assessed in competition experiments.
The G-rich strand was firstly examined: the assays in Fig.
6B (lanes 1-10) show that none of the competitor
DNAs inhibited the faint complexes, which therefore reflect nonspecific
associations. Both ssG1 and ssG2 were instead inhibited by
preincubation with homologous (lanes 3 and 4),
but not with double-stranded (lanes 8-10), DNA. The
complementary strand partially competed for protein(s) (lanes 5-7); however, the excess of C-rich DNA required to inhibit ssG1 and ssG2 formation was at least 5-fold higher than that used in the
homologous competition (compare lane 3 to 5, and
4 to 6). Competition experiments were then set up
between the C-rich strand and homologous, reverse complementary or
double-stranded DNA: ssC assembly was inhibited by preincubation with
homologous DNA (lanes 13 and 14), while remaining
unaffected in the presence of double-stranded (lanes 18-20)
HFE. Precompetition with G-rich DNA affected the formation of ssC in a
dose-dependent manner (lanes 15-17); however,
the inhibition remained partial even with the largest competitor excess
(200-fold). To further establish whether double- and single-stranded
complexes were independently assembled, HFE and the C-rich strand were
simultaneously assayed. As shown in Fig. 6C, double-stranded
competitor DNA selectively inhibited the HFE complex (lanes
3-5), while the C-rich strand competed for only ssC protein(s)
(lanes 6-8), thus confirming the specificity of the binding
events. Conclusive evidence that the factors binding to HFE and to each
DNA strand are chemically distinct was obtained from NIH/3T3 extract
fractionation. While HFE-binding activities were eluted in the 0.5 M KCl fraction, ssG and ssC proteins fractionated with the
phosphocellulose flow-through (data not shown). Thus, distinct proteins
bind to HFE and interact either with each DNA strand, or with the
double-stranded form, of the element.

View larger version (39K):
[in this window]
[in a new window]
|
Fig. 6.
Distinct factors bind the double-stranded and
single-stranded HFE oligonucleotides. A, gel shift assays
with NIH/3T3 extracts and the G-rich (lane 2), C-rich
(lane 4), or double-stranded (lane 6) probes;
lanes 1, 3, and 5, probes alone.
B, competition assays between single-stranded
oligonucleotides. Lanes 1-10, gel shift of the G-rich
strand alone (lane 1), after incubation with NIH/3T3
extracts (lane 2), and in the presence of 50- and 100-fold molar excess of homologous strand (lanes 3 and
4), 50-, 100-, and 200-fold excess of C-rich strand
(lanes 5-7), and 50-, 100-, and 200-fold double-stranded
HFE (lanes 8-10). Lanes 11-20, C-rich probe
alone (lane 11), after incubation with NIH/3T3 extracts (lane 12), or in the presence of 50- and 100-fold excess of
homologous strand (lanes 13 and 14), 50-, 100-, and 200-fold excess of G-rich strand (lanes 15-17) and 50-, 100-, and 200-fold double-stranded HFE (lanes 18-20).
C, the C-rich and double-stranded HFE probes were mixed
(lane 1) and incubated with NIH/3T3 extracts (lane 2), in the presence of 50-, 100-, and 200-fold double-stranded (lanes 3-5), C-rich (lanes 6-8), or G-rich
(lanes 9-11) DNA.
|
|
Several single-stranded binding activities have been characterized in
the last few years, some of which preferentially recognize C-rich or
G-rich DNA. To establish whether protein binding to single DNA strands
was sequence-specific or was driven by the base composition, binding
reactions were challenged with single-stranded oligonucleotides of
comparable C or G content yet unrelated in sequence; the sequence of
used competitors is shown in Table I. No
tested sequence, even of similar base composition to that of the C-rich
probe, inhibited ssC (Fig.
7A). Factors involved in ssG1
and ssG2 did interact with the MG1, MG2, and MG3 derivatives of the
wild-type site (Fig. 7B, lanes 2-4), suggesting
that the factors recognized closely related sequences to the G-rich
strand of HFE, yet did not bind to the heavily mutagenized LI24
version, or to unrelated G-rich oligonucleotides (Fig. 7B,
lanes 7-12). Thus, single-stranded binding proteins
interact with their cognate sites in a substantially sequence-specific
manner.

View larger version (28K):
[in this window]
[in a new window]
|
Fig. 7.
Specificity of single-stranded binding
activities. A, gel shifts of the C-rich probe with NIH/3T3
extracts (lanes 1 and 13), in the presence of a
50- and 100-fold excess of unlabeled probe (lanes 2 and
3) or 100-fold excess of the indicated competitors (lanes 4-12). B, gel shifts of the G-rich probe
with NIH/3T3 extracts (lanes 1 and 13), in the
presence of 50- and 100-fold excess of unlabeled probe (lanes
5 and 6), or with a 100-fold excess of the indicated
competitors (lanes 2-4 and 7-12). Competitor
sequences are shown in Table I.
|
|
Different Forms of the HFE Show Differential Interactions with
Protein Factors in Different Cell Types--
We next asked whether the
distribution of single-stranded DNA-binding factors varied in different
cells, as observed for RXR and RAR binding to double-stranded HFE. Gel
shift assays were carried out with probes corresponding to C-rich,
G-rich, and double-stranded HFE using nuclear extracts from liver, or
from cycling and growth-arrested NIH/3T3 cells. Consistent with the
cell-type distribution of RXR factors seen in Western experiments, the
double-stranded HFE complex was more abundant in growth-arrested,
compared with cycling, NIH/3T3 cells (Fig.
8, lanes 2 and 3),
and highest in liver (Fig. 8, lane 1). G-rich DNA-binding
factors showed the reverse pattern, being either more abundant or more
active, in cycling compared with quiescent NIH/3T3 cells, and
undetectable in liver (Fig. 8, lanes 7-9). The ssC complex
was comparatively more widespread, although somewhat less abundant in
liver compared with both arrested and proliferating NIH/3T3 cells (Fig.
8, lanes 4-6). Together, the results suggest that different
cell types are equipped with distinct sets of factors capable of
binding either double-stranded HFE or single DNA strands.
Single-stranded complexes most efficiently form with protein extracts
from proliferating, i.e. actively transcribing, NIH/3T3
fibroblasts, whereas factors interacting with double-stranded HFE were
most abundantly expressed in liver.

View larger version (59K):
[in this window]
[in a new window]
|
Fig. 8.
Distribution of factors binding different HFE
forms. Double-stranded HFE (lanes 1-3), C-rich
(lanes 4-6), and G-rich (lanes 7-9)
oligonucleotides were incubated with protein extracts (4 µg) from
liver (lanes 1, 4, and 7),
growth-arrested (lanes 2, 5, and 8)
and cycling (lanes 3, 6, and 9)
NIH/3T3 cells.
|
|
Plasmids Containing the HFE Sequence Are Sensitive to
S1--
Since single-stranded DNA-binding activities were depicted in
cell cultures in which the Htf9 locus is
transcriptionally active (9), it was important to assess whether
single-stranded structures actually formed in the
Htf9 promoter. Computer analysis revealed that HFE
fell within one potential stem-and-loop structure; theoretical estimates of the free energy associated with the formation of the
single-stranded loop give
G =
21.7 kcal/mole. We
wondered whether these theoretical sequence features gave rise to the
formation of unusual DNA structures in the Htf9
initiation region. Supercoiled plasmids carrying either the full-length
(pTS-A, 273 bp), or the minimal (pEA-A, 74 bp), Htf9
promoter were digested with S1 nuclease and subsequently restricted
with PstI to separate the Htf9 sequences from the plasmid replication origin (Fig.
9A), which might contain unwound DNA. Results in Fig. 9B show that both the pTS-A and
pEA-A constructs contained S1-sensitive sequences that were not present in the vector. Hybridization with labeled HFE oligonucleotide confirmed
that the region containing HFE was sensitive to S1 cleavage (Fig.
9C). To directly ascertain whether HFE in particular
represented a DNA target for S1 cleavage, pA10-derived clones carrying
one or three HFE copies (Fig. 9D) were subjected to
digestion using increasing S1 nuclease doses. While the pA10 vector was
inefficiently cleaved by S1 (Fig. 9E, lanes
1-4), plasmids containing one or more HFE copies were sensitive
to S1 in a dose-dependent manner. Microdensitometry
quantification of the ratio of supercoiled to linear DNA revealed that
the extent of cleavage by S1 increased with the number of HFE copies.
Thus, the HFE sequence confers S1 sensitivity to supercoiled
plasmids.

View larger version (35K):
[in this window]
[in a new window]
|
Fig. 9.
S1 sensitivity in HFE-containing
plasmids. A, maps of the pSV0-CAT vector and of
Htf9-derived promoter constructs, respectively,
carrying the basal (pEA-CAT, 74 bp) and the full-length (pTSA-CAT, 273 bp) promoter region. The pBR322 origin of replication (ORI)
is marked. P, PstI; H,
HindIII. In the series of constructs, the promoter insert
(cloned in the HindIII site) is oriented in the direction of
RanBP1 transcription relative to the CAT gene. B, ethidium-bromide staining of constructs subjected to
digestion with S1 nuclease (lanes 2, 4, and
6) or to mock digestion (indicated by a dash (-) in
lanes 1, 3, and 5), and subsequently
restricted with PstI to separate the replication origin from
the Htf9 promoter region. Lane 7, DNA
digested with EcoRI and HindIII. C,
autoradiograph of the blot from the gel in A after
hybridization with labeled HFE oligonucleotide. D, chimeric
promoter constructs carrying no (pA10), one
(pA10G), or three (pA10G9) HFE copies upstream of
the pA10 minimal promoter. The arrow marks the promoter
insertion site. E, ethidium bromide-stained gel showing 1 µg of DNA from each construct after digestion with 0, 0.5, 1, and 2.5 units of S1.
|
|
The Genomic Region Surrounding HFE Is Sensitive to S1 during
Transcription in Vivo--
We finally assessed whether S1-sensitive
structures identified in HFE-containing clones were maintained in the
higher order organization of genomic DNA. NIH/3T3 cell cultures were
either brought to proliferation arrest by serum withdrawal for 48 h, or stimulated to reenter the cycle by adding fresh serum. Flow cytometric analysis confirmed that cells collected prior to serum stimulation were arrested in the G0/G1 state,
while cells collected 15 h after serum refeeding were traversing S
phase (Fig. 10A). Neither
Htf9-associated gene was transcribed in
growth-arrested cells, while transcription was active in S phase cells
(Fig. 10B), confirming our previous data on cell cycle
control of the Htf9 promoter (7). Nuclei were
isolated from both growth-arrested and S phase NIH/3T3 cultures to
assess the Htf9 promoter sensitivity to S1 in
transcribing and nontranscribing cells. Nuclei were digested with
increasing amounts of S1 nuclease; DNA was extracted, restricted with
EcoRI and hybridized with a Htf9-derived
fragment that was flush with one EcoRI restriction end (map
in Fig. 10C). These experiments depicted a restricted region
(approximately 100 bp) that was sensitive to S1 cleavage in S phase
NIH/3T3 cells (Fig. 10C, lanes 5-7), but not in
NIH/3T3 cultures brought to quiescence (Fig. 10C,
lanes 1-3). Indirect end-labeling revealed that the
S1-sensitive region in S phase cells encompassed the 74-bp region
sufficient for basal transcription as functionally defined in promoter
assays (see Fig. 1).

View larger version (26K):
[in this window]
[in a new window]
|
Fig. 10.
S1 sensitivity of the HFE-containing region
in S phase and in growth-arrested cells. A, flow cytometry
analysis of cell cultures maintained in low serum for 48 h
(top) and after 15 h of serum restimulation
(below). The DNA content of cells is indicated on the
x axis, cell numbers are indicated on the y axis.
B, total RNA (15 µg) was extracted from growth-arrested (G0, lane 1) or S phase (lane
2) NIH/3T3 cultures, stained with ethidium bromide to assess the
integrity of the 28S and 18S ribosomal bands (top panels),
blotted, and hybridized with the indicated probes. C, nuclei
from growth-arrested (G0) and S phase NIH/3T3 cells
were digested with the following S1 units/µg DNA: 1, lanes
1 and 5; 5, lanes 2 and 6; 10, lanes 3 and 7. DNA was extracted, restricted with
EcoRI, and hybridized with a fragment flush with one
EcoRI end (shown on the map). Lane 5, EcoRI-HindIII marker fragments (M).
The full-length restriction fragment is indicated by a thick
arrowhead; fragments derived from internal S1 cleavage (indicated
by thin arrowheads) were detected in S phase, but not in
G0, NIH/3T3 nuclei. The region of S1 cleavage
(arrowheads) encompasses the minimal promoter region (box).
|
|
 |
DISCUSSION |
The HFE Near TS-1 Is Required for Promoter Activity--
A 74-bp
promoter region carries sufficient information to direct transcription
of both the RanBP1 and Htf9-c genes. We
have presently began to characterize the HFE element, originally
identified by genomic footprinting, which flanks the TS-1 start site
used for initiation of both genes. Promoter sites showing interactions with factors in vivo often identify regulatory elements. HFE
is indeed a functional element in both orientations, as mutational inactivation reduces promoter strength in each direction. However, HFE
inactivation is sensed differently depending on the orientation and
arrangement of neighboring regulatory sequences in the promoter context; HFE inactivation does mildly impair the pES-C promoter, while
severely affecting activity of both the pES-A and pEA-C constructs. The
effectiveness of regulatory sequences depends, at least in part, on
their position in the promoter context. It is possible that regulatory
elements included between the pEA-C and pES-C promoter boundaries
participate in the assembly of transcriptional complexes and relieve in
part the effect of HFE inactivation. Thus, the functional promoter
mapping results suggest that the HFE site is involved in mediating
interactions among various regulatory elements in each promoter
orientation.
RXR Factors Bind the Double-stranded HFE--
In the search for
factor(s) protecting HFE, we have identified distinct activities which
bind different HFE forms. The double-stranded site is recognized by
complexes containing retinoid X receptors. These complexes showed cell
type-specific differences in both their relative abundance and
interacting partners. NIH/3T3 extracts yielded a complex that was
similarly reacted by both anti-RXR and anti-RAR antibodies, while liver
extracts formed a complex that was specifically reacted by anti-RXR but
not anti-RAR antibodies. Thus, RXRs interacting with HFE associate with
specific partners in different cells. These findings were paralleled by
the distribution of retinoid receptors: in liver nuclei, all three RXR
subtypes were abundant and were recovered in chromatographic fractions positive for HFE binding. In NIH/3T3 cells, RAR factors were also depicted, while only one RXR subtype was above the level of detection in Western experiments.
Sites in a DR-1 configuration, such as HFE, can be bound by RXR
homodimers or heterodimers involving different partners (reviewed in
Refs. 23-25). The DR-1 oligonucleotide used here for control was
previously characterized as a preferred target site of RXR homodimers
(21). An identical sequence to HFE was also included in an
oligonucleotide, designed R7, characterized as a high affinity site for
RXR
homodimer binding (26). RXRs are versatile factors that can act
as transcriptional activators, be transcriptionally silent although
remaining engaged in the interaction with target DNA sites, or
contribute to repression in differentiated tissues, as shown for
several proliferation-associated genes (27-29), in a
ligand-dependent manner (see Minucci and Ozato (30) and
references therein). Immune reactions of both the HFE complexes and of
fractions enriched in HFE-binding activity indicate that RXR family
members interact with the HFE site in liver. Since transcription from the Htf9 locus is low or absent in liver cells,
unless regeneration is induced by surgical hepatectomy (9), while the
binding of RXR complexes is highest compared with other cell types, the
interaction of RXR factors with HFE appears to correlate with
transcriptional inactivity in liver nuclei.
Western immunoblotting and supershift experiments with NIH/3T3
fibroblast extracts suggest that the RXR subtype(s) in these cells
rather heterodimerize with RAR factors while binding HFE. HFE-binding
complexes in NIH/3T3 cells have lower abundance than in liver cells,
and in cycling NIH/3T3 cells are less abundant than in growth-arrested
cultures which cease transcription of the
Htf9-associated genes. We previously noticed that
undifferentiated embryo stem cells were the only cell type in which HFE
was not footprinted in vivo; embryo stem cell extracts also
lacked double-stranded HFE binding activity (15). Since both
Htf9 genes are transcribed in both embryo stem and F9
undifferentiated cells (31), it appears in retrospect that RXR binding
is dispensable for expression of the Htf9 genes
during embryonal proliferation. Thus, the binding of RXR-containing
complexes to HFE is up-regulated during differentiation and is
inversely related to Htf9 promoter activity.
Specific Factors Bind Each DNA Strand of the HFE--
One
unexpected HFE feature is that specific activities present in NIH/3T3
cells bind to each DNA strand. That finding prompted us to investigate
whether single-stranded structures actually formed in the
Htf9 promoter. Theoretical energy estimates support the possibility that the Htf9 promoter forms a
stem-and-loop structure, and the identification of S1-sensitive sites
in HFE-containing plasmids suggests that the formation of
single-stranded structures represent an inherent feature of the
Htf9 initiation region. Many CG-rich, TATA-less
promoters potentially form single-stranded loops in the region
surrounding the transcription start sites (32). Such looping structures
have been suggested to serve as structural landmarks and facilitate the
recognition of start sites by basal factors in complex genomes, where
TATA box recognition by linear sequence scanning would be inefficient
compared with smaller genomes (32).
In the Htf9 bidirectional promoter each DNA strand
must serve as a transcriptional template. In vivo
sensitivity to S1 was detected in NIH/3T3 cells in which the
Htf9 promoter was active, but not in conditions in
which transcription was repressed. The link between proliferation,
promoter activity, S1 sensitivity, and expression or activity of the
single-stranded binding proteins, suggest that single-stranded proteins
identified in this work exert a positive role in transcription. A
growing number of single-stranded DNA-binding proteins have now been
found to interact with regulatory elements and influence transcription
by altering the DNA topology or conformation (33-37). Extensive
studies of the c-myc promoter (37-40) are suggestive of a
model in which the cellular conditions control the interaction of
double-stranded or single-stranded DNA-binding factors to specific
promoter elements; the binding of proteins to single DNA strands is
thought to induce structural transitions that are sensed as
transcriptional signals and mediate c-myc response to cell
cycle regulators, growth factors, and other inducing stimuli. On a
similar line, two studies of hormone-inducible promoters have shown
that the binding of double-stranded and single-stranded binding
activities is hormonally regulated and independently mediate control of
basal transcription and hormone induction (41, 42). From these studies
it appears that the transcriptional response of one same element to
inducing or repressing stimuli can be mediated by factors influencing
the promoter structural organization and either allow, or prevent,
productive interactions among neighboring regulatory factors in
different cellular conditions.
It is increasingly clear that RXR complexes, particularly with RARs,
can recruit corepressors that cooperate in maintaining a
transcriptionally inactive structural organization (43). Ligand addition can destabilize the interaction with corepressors, thereby triggering chromatin remodeling (reviewed in Ref. 44). Various possibilities might be envisaged concerning HFE. RXR complexes might
normally occupy HFE and set the TS-1 region in the double-stranded form. In the presence of cell cycle-related stimuli, RXRs could be
displaced from the DNA, either directly by the single-stranded binding
proteins, or by general factors with helicase activity (12). The
single-stranded binding proteins may then facilitate the assembly of
initiation complexes on each strand, and/or contribute to looping
structures in which activator-binding sites productively interact with
the transcription start site. Alternatively, the inherent features of
the Htf9 initiation region might favor the single-stranded conformation, which may represent a natural target for
single-stranded binding proteins setting the promoter in a transcriptionally competent state. In cells in which neither
Htf9-associated gene is to be expressed, RXR
complexes might catalyze the formation of a double-stranded structure.
RXR complexes might not necessarily determine repression in all cells
and may exert a negative or a positive function depending on their
interactions with positive or negative co-factors (45), the presence of
ligands, and the occupancy of adjacent promoter sites by
transcriptional repressors or activators.
We have previously shown that cell cycle-dependent
transcription of both Htf9-associated genes is
mediated by target elements for both the E2F and Sp1 families of
activators, while being repressed by the pRb retinoblastoma protein and
its relative p107 (7, 9). The present data show that transcription is
also associated with S1 sensitivity and with the interaction of
specific proteins with single DNA strands near TS-1, while absence of
transcription is associated with loss of S1 sensitivity and increased
binding of RXR complexes to the double-stranded HFE. The pRb protein
interferes with single-stranded DNA-binding by the Pur
protein (46).
Thus, the Htf9 promoter may not only be controlled by
the antagonism between retinoblastoma-related factors and E2F
activators, but may also involve regulated interactions between
single-stranded binding proteins and each DNA strand in the region of
initiation, or assumption of the double-stranded conformation
associated with the binding of retinoid receptors.
 |
ACKNOWLEDGEMENTS |
We are grateful to E. Cundari for flow
cytometry, to C. Pittoggi for many contributions to this work, to F. D'Ottavio for technical assistance, to G. Bonelli for photography, and
to our colleagues D. Carotti, V. Colantuoni, A. Farsetti, and R. Strom for helpful comments and suggestions for this manuscript.
 |
FOOTNOTES |
*
This work was supported by grants from EEC (contract
BMH4-C796-1529), CNR (Progetto Strategico Cell Cycle and Apoptosis), Associazione Italiana per la Ricerca sul Cancro (AIRC), and Fondazione Cenci-Bolognetti.The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement" in
accordance with 18 U.S.C. Section
1734 solely to indicate this fact.
§
Supported by a fellowship from the Associazione Italiana per la
Ricerca sul Cancro (AIRC).
¶
Present address: Dept. of Surgical Pathology, "Tor
Vergata" University, 00133 Rome, Italy.
Supported by a fellowship from the MURST Ministry of Public
Education.

To whom correspondence should be addressed. Tel.: 39-6 445 7528; Fax: 39-6 445 7529; E-mail:lavia{at}axrma.uniroma1.it.
1
The abbreviations used are: RanBP1, Ran-binding
protein 1; TS-1, transcription start 1; HFE, Htf9
footprinted element; CAT, chloramphenicol acetyltransferase; RXR,
retinoid X receptor; RAR, retinoic acid receptor; ssG1, single-stranded
G-rich DNA-containing complex 1; ssG2, single-stranded G-rich
DNA-containing complex 2; ssC, single-stranded C-rich DNA-containing
complex; bp, base pair(s).
 |
REFERENCES |
-
Lavia, P.,
Macleod, D.,
and Bird, A.
(1987)
EMBO J.
6,
2773-2779[Abstract]
-
Bressan, A.,
Somma, M. P.,
Lewis, J.,
Santolamazza, C.,
Copeland, N.,
Gilbert, D.,
Jenkins, N.,
and Lavia, P.
(1991)
Gene (Amst.)
103,
201-209[CrossRef][Medline]
[Order article via Infotrieve]
-
Coutavas, E.,
Ren, M.,
Oppenheim, J.,
D'Eustachio, P.,
and Rush, M.
(1993)
Nature
366,
585-587[CrossRef][Medline]
[Order article via Infotrieve]
-
Bischoff, F. R.,
Krebber, H.,
Smirnova, E.,
Dong, W.,
and Ponstingl, H.
(1995)
EMBO J.
14,
705-715[Abstract]
-
Rush, M. G.,
Drivas, G.,
and D'Eustachio, P.
(1996)
Bioessays
18,
103-112[Medline]
[Order article via Infotrieve]
-
Sazer, S.
(1996)
Trends Cell Biol.
6,
81-85 [CrossRef]
-
Guarguaglini, G.,
Battistoni, A.,
Pittoggi, C.,
Di Matteo, G.,
Di Fiore, B.,
Lavia, P.
(1997)
Biochem. J.
325,
277-286[Medline]
[Order article via Infotrieve]
-
Somma, M. P.,
Gambino, I.,
and Lavia, P.
(1991)
Nucleic Acids Res.
19,
4451-4458[Abstract]
-
Di Matteo, G.,
Fuschi, P.,
Zerfass, K.,
Moretti, S.,
Ricordy, R.,
Cenciarelli, C.,
Tripodi, M.,
Jansen-Durr, P.,
and Lavia, P.
(1995)
Cell Growth Differ.
6,
1213-1224[Abstract]
-
Hernandez, N.
(1993)
Genes Dev.
7,
1291-1308[CrossRef][Medline]
[Order article via Infotrieve]
-
Novina, C. D.,
and Roy, A. L.
(1996)
Trends Genet.
12,
351-355[CrossRef][Medline]
[Order article via Infotrieve]
-
Orphanides, G.,
Lagrange, T.,
and Reinberg, D.
(1996)
Genes Dev.
10,
2657-2683[CrossRef][Medline]
[Order article via Infotrieve]
-
Weis, L.,
and Reinberg, D.
(1992)
FASEB J.
6,
3300-3309[Abstract/Free Full Text]
-
Somma, M. P.,
Pisano, C.,
and Lavia, P.
(1991)
Nucleic Acids Res.
19,
2817-2824[Abstract]
-
Stapleton, G.,
Somma, M. P.,
and Lavia, P.
(1993)
Nucleic Acids Res.
21,
2465-2471[Abstract]
-
Laimins, L. A.,
Khoury, G.,
Gorman, C. M.,
Howard, B.,
Gruss, P.
(1982)
Proc. Natl. Acad. Sci. U. S. A.
79,
6453-6457[Abstract]
-
Wiley, S.,
Kraus, R. J.,
and Mertz, J. E.
(1992)
Proc. Natl. Acad. Sci. U. S. A.
89,
5814-5818[Abstract]
-
Zenzie-Gregory, B.,
Khachi, A.,
Garraway, I. P.,
Smale, S.
(1993)
Mol. Cell. Biol.
13,
3841-3849[Abstract]
-
Kaufman, J.,
and Smale, S.
(1994)
Genes Dev.
8,
821-829[Abstract]
-
Hamada, K.,
Gleason, S. L.,
Levi, B.-Z.,
Hirschfeld, S.,
Appella, E.,
and Ozato, K.
(1989)
Proc. Natl. Acad. Sci. U. S. A.
86,
8289-8293[Abstract]
-
Kliewer, S. A.,
Umesono, K.,
Heyman, R. A.,
Mangelsdorf, D. J.,
Dyck, J. A.,
Evans, R. M.
(1992)
Proc. Natl. Acad. Sci. U. S. A.
89,
1448-1452[Abstract]
-
Nagata, T.,
Kanno, Y.,
Ozato, K.,
and Taketo, M.
(1994)
Gene (Amst.)
142,
183-189[CrossRef][Medline]
[Order article via Infotrieve]
-
Mangelsdorf, D. J.,
and Evans, R. M.
(1995)
Cell
83,
841-850[Medline]
[Order article via Infotrieve]
-
Mangelsdorf, D. J.,
Thummel, C.,
Beato, M.,
Herrlich, P.,
Schutz, G.,
Umesono, K.,
Blumberg, B.,
Kastner, P.,
Mark, M.,
Chambon, P.,
and Evans, R. M.
(1995)
Cell
83,
835-839[Medline]
[Order article via Infotrieve]
-
Chambon, P.
(1996)
FASEB J.
10,
940-954[Abstract/Free Full Text]
-
Yang, Y-Z.,
Subauste, J. S.,
and Koenig, R.
(1995)
Endocrinology
136,
2896-2904[Abstract]
-
Hudson, L. G.,
Santon, J. B.,
Glass, C. K.,
Gill, G. N.
(1990)
Cell
62,
1165-1175[Medline]
[Order article via Infotrieve]
-
Bouterfa, K. L.,
Piedrafita, F. J.,
Doenecke, D.,
Pfahl, M.
(1995)
DNA Cell Biol.
14,
909-919[Medline]
[Order article via Infotrieve]
-
Soprano, D. R.,
Chen, L. X.,
Wu, S.,
Donigan, A. M.,
Borghaei, R. C.,
Soprano, K. J.
(1996)
Oncogene
12,
577-584[Medline]
[Order article via Infotrieve]
-
Minucci, S.,
and Ozato, K.
(1996)
Curr. Opin. Genet. Dev.
6,
567-574[CrossRef][Medline]
[Order article via Infotrieve]
-
Tyndall, C.,
Watt, F.,
Molloy, P. L.,
Vincent, P. C.,
Frommer, M.
(1992)
J. Mol. Biol.
226,
289-299[Medline]
[Order article via Infotrieve]
-
Ackerman, S.,
Minden, A. G.,
and Yeung, C. Y.
(1993)
Proc. Natl. Acad. Sci. U. S. A.
90,
11865-11869[Abstract]
-
Xu, G.,
and Goodridge, A.
(1996)
J. Biol. Chem.
271,
16008-16019[Abstract/Free Full Text]
-
Chen, S.,
Supakar, P. C.,
Vellanoveth, R. L.,
Song, C. S.,
Chatterjee, B.,
Roy, A. K.
(1997)
Mol. Endocrinol.
11,
3-15[Abstract/Free Full Text]
-
Giffin, W.,
and Hache', R. J. G.
(1995)
DNA Cell Biol.
14,
1025-1035[Medline]
[Order article via Infotrieve]
-
Santra, M.,
Danielson, K.,
and Iozzo, R.
(1994)
J. Biol. Chem.
269,
579-587[Abstract/Free Full Text]
-
Duncan, R.,
Bazar, R.,
Michelotti, G.,
Tomonaga, T.,
Krutzsch, H.,
Avigan, M.,
and Levens, D.
(1994)
Genes Dev.
8,
465-480[Abstract]
-
Bazar, l.,
Meighen, D.,
Harris, V.,
Duncan, R.,
Levens, D.,
and Avigan, M.
(1995)
J. Biol. Chem.
270,
8241-8248[Abstract/Free Full Text]
-
Tomonaga,
and Levens, D.
(1996)
Proc. Natl. Acad. Sci. U. S. A.
93,
5830-5835[Abstract/Free Full Text]
-
Michelotti, G. A.,
Michelotti, E. F.,
Pullner, A.,
Duncan, R. C.,
Eick, D.,
Levens, D.
(1996)
Mol. Cell. Biol.
16,
2656-2669[Abstract]
-
Shimura, H.,
Shimura, Y.,
Ohmori, M.,
Ikuyama, S.,
and Kohn, L. D.
(1995)
Mol. Endocrinol.
9,
527-539[Abstract]
-
Saito, H.,
and Oka, T.
(1996)
J. Biol. Chem.
271,
8911-8918[Abstract/Free Full Text]
-
Nagy, L.,
Kao, H.-Y.,
Chakravarti, D.,
Lin, R.,
Hassug, C. A.,
Ayer, D. E.,
Schreiber, S. L.,
Evans, R. M.
(1997)
Cell
89,
373-380[Medline]
[Order article via Infotrieve]
-
Schulman, I. G.,
Li, C.,
Schwabe, J. W. R.,
Evans, R. M.
(1997)
Genes Dev.
11,
299-308[Abstract]
-
Perlmann, T.,
and Evans, R. M.
(1997)
Cell
90,
391-397[Medline]
[Order article via Infotrieve]
-
Johnson, E. M.,
Chen, P.-L.,
Krachmarov, C. P.,
Barr, S. M.,
Kanovsky, M.,
Ma, Z.-W.,
Lee, W.-H.
(1995)
J. Biol. Chem.
270,
24352-24360[Abstract/Free Full Text]
-
Brunel, F.,
Zakin, M. M.,
Buc, H.,
and Buckle, M.
(1996)
Nucleic Acids Res.
24,
1608-1615[Abstract/Free Full Text]
Copyright © 1998 by The American Society for Biochemistry and Molecular Biology, Inc.