From the Consiglio Nazionale delle Ricerche Centre of
Evolutionary Genetics, c/o University La Sapienza, Via degli
Apuli 4, 00185 Rome, Italy and the
Division of Molecular
Carcinogenesis, The Netherlands Cancer Institute, Plesmanlaan 121, 1066 CX Amsterdam, The Netherlands
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ABSTRACT |
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The gene encoding Ran-binding protein 1 (RanBP1)
is transcribed in a cell cycle-dependent manner. The
RanBP1 promoter contains two binding sites for E2F factors,
named E2F-c, located proximal to the transcription start, and E2F-b,
falling in a more distal promoter region. We have now induced
site-directed mutagenesis in both sites. We have found that the distal
E2F-b site, together with a neighboring Sp1 element, actively controls
up-regulation of transcription in S phase. The proximal E2F-c site
plays no apparent role in cycling cells yet is required for
transcriptional repression upon growth arrest. Protein binding studies
suggest that each E2F site mediates specific interactions with
individual E2F family members. In addition, transient expression assays
with mutagenized promoter constructs indicate that the functional role of each site is also dependent on its position relative to other regulatory elements in the promoter context. Thus, the two E2F sites
play opposite genetic functions and control RanBP1
transcription through distinct molecular mechanisms.
The murine Htf9-a/RanBP1 gene encodes
Ran-binding protein 1 (RanBP1)1 (1), a major
partner of the Ran GTPase (2, 3), which cooperates with members of the
Ran signaling network in control of a variety of functions, including
DNA replication, mitotic entry and exit, chromatin condensation, and
nucleocytoplasmic transport (reviewed in Refs. 4 and 5). The
RanBP1 gene, unlike other members of the network, is
expressed in a cell cycle-dependent manner:
RanBP1 transcription is activated at the G1/S
transition and peaks in S cells (6, 7). Up-regulation of
RanBP1 transcription during S phase is an important
functional requirement for the Ran network activity: we have previously
shown that replacing the RanBP1 endogenous promoter with
cell cycle-independent regulatory sequences yielded deregulated
production of the RanBP1 protein and caused abnormalities in further
cell cycle progression, including inhibition or delay in S phase,
impaired mitotic exit, and failure of chromatin decondensation at the
mitosis-to-interphase transition (8).
In previous work, we identified two RanBP1 promoter regions
that control separate aspects of RanBP1 transcription (see
map in Fig. 1A). Basal transcription requires a proximal
region encompassing the major transcription start site, TS-1 (9). That
region harbors two prominent genomic footprints (10), termed Sp1.2,
acting as a bona fide Sp1-binding site (11), and Htf9
footprinted element (HFE), a basal control element that can interact
either with retinoid X receptor family members in quiescent cells or
with single-stranded DNA-binding proteins in cycling cells; the latter
determines the assumption of an active conformation around the
transcription start site (9). The HFE is flanked by a recognition site
for E2F/DP factors, termed E2F-c, which is not footprinted in
proliferating cells (10). G1/S up-regulation of
RanBP1 transcription is controlled by a region located from
E2F factors control expression of many cell cycle genes through
responsive promoter elements that confer either positive or negative
control; precise temporal regulation of these genes by E2F is a
prerequisite for ordered cell cycle progression. The E2F DNA binding
activity is shared by heterodimeric complexes, in which both subunits
are encoded by gene families: one heterodimeric component is
synthesized from one of five related E2F-encoding genes and dimerizes
with one of three related DP dimerization partner proteins. Control of
transcription by E2F/DP dimers is subjected to various levels of
complexity (see Refs. 12-14 for reviews). First, E2F-encoding genes
are differentially induced during the cell cycle: E2F-1, -2, and -3 appear in mid- or late G1, whereas E2F-4 and -5 are
expressed relatively constantly. Second, transcription of cell cycle
genes is controlled by the molecular balance established at any given
time between E2F/DP complexes and repressor pocket proteins.
Transactivation by E2F/DP heterodimers can be differentially
antagonized in the interaction with members of the pocket protein
family: E2F-1, E2F-2, and E2F-3 preferentially interact with the pRb
retinoblastoma gene product (15), whereas E2F-4 preferentially
associates with p107 and p130, and E2F-5 with p130 only (15-18);
however, pRb can also interact with E2F-4 (15). The loss of particular
pocket proteins affects distinct sets of genes (19), thus pinpointing
specific roles for particular E2F/DP/pocket complexes in
transcriptional control. These interactions are themselves temporally
regulated: the p130 protein is essentially active in G0
cells (Refs. 18 and 20; reviewed in Refs. 14 and 21); pRb and p107 are
instead transcriptionally induced during G1 and inactivated
by phosphorylation as cells approach S phase (reviewed in Refs. 14 and
22). The interactions of activating complexes with target genes is also
dependent upon regulated nuclear transport of particular E2F (23-26)
and DP (27) members, which determines which complexes will assemble in
the nucleus (28). Finally, a further level of control resides in the
promoter structure, within which E2F/DP/pocket complexes may productively interact with positive (29, 30) or negative (31, 32)
factors; recent experiments with synthetic reporter constructs indicate
that the repressing or activating function of E2F elements depends in
part on their position relative to neighboring regulatory sequences
(33, 34).
In this work, we have sought to establish the role of the E2F sites in
cell cycle-regulated transcription of the RanBP1 gene. We
have induced site-directed mutagenesis of both E2F sites to assess
their individual contribution to cell cycle regulation of
RanBP1 promoter activity. We have found that the distal
E2F-b, together with the neighboring Sp1.3 site, controls up-regulation of transcription at the G1/S boundary. In contrast, the
proximal E2F-c site is dispensable for expression in cycling cells, yet is absolutely required for transcriptional repression in G0
cells. Thus, both E2F sites independently contribute to cell
cycle-regulated activity of the RanBP1 promoter and mediate
genetically distinct control mechanisms.
Cell Cultures and FACS Analysis--
Murine NIH/3T3 fibroblast
cultures (ATCC CRL 1658) were grown in Dulbecco's modified Eagle's
medium supplemented with 10% (v/v) fetal calf serum (FCS) under 5%
(v/v) CO2 at 37 °C. Cell samples to be analyzed were
harvested in phosphate-buffered saline, fixed in acetone:methanol
solution (1:5 v/v), and incubated with RNase (10 µg/ml) for 5 min at
0 °C. Propidium iodide (0.5 µg/ml) was added and the DNA content
of cell samples was measured in a FACStar Plus cytofluorometer (Beckton
Dickinson). In experiments designed to monitor S phase, cells were
incubated with 45 µM bromodeoxyuridine for 30 min before
harvesting. Harvested cells were incubated in 1 N HCl for
45 min, neutralized, incubated with anti-bromodeoxyuridine antibody (Ig
G clone BU5.1, Ylem) for 30 min and then with a fluorescein-conjugated secondary anti-IgG antibody (Ylem), and finally subjected to
biparametric FACS analysis for simultaneous determination of the DNA
content and of bromodeoxyuridine incorporation using the WinMDI
software (10,000 events/sample).
Expression Constructs--
The pA10 and pE1 clones carry the
SV40 minimal promoter and a chimeric derivative containing the E2F-b
site, respectively (for details, see Ref. 6), upstream of the
chloramphenicol acetyltransferase (CAT) reporter sequence. The pTS-A
clone carries the wild-type RanBP1 promoter upstream of the
CAT gene (11). Derivative clones were synthesized by site-directed
mutagenesis, by either ligating synthetic oligonucleotides mutated in
the E2F-b and/or Sp1.3 sites and terminating with a Sau96 end to the
Sau96 unique site of the RanBP1 promoter
(GenBankTM accession number X05830), or using the
Quick-Change site-directed mutagenesis kit (Stratagene). Resulting
clones carried the following mutations (underlined): pmE, mutation in
the E2F-b site (sequence TTTGGCGGGA mutated to TTTACTCAGA);
pmS, mutation in the Sp1.3 site (sequence GGGGCGGGC mutated to
GAGATGGGC); pmES, simultaneous mutations of
both the E2F-b and Sp1.3 sites; pGi, mutation in the proximal E2F-c
site (sequence TTTCCCGCCGC mutated to
TTTACTCACGC); pQl, double E2F-b and E2F-c site
mutations; pmBB was derived from pmE and hence carries the mutated
version of the distal E2F-b site, whereas the proximal E2F-c site
(TTTCCCGCCGC) was replaced by a wild-type E2F-b site (TTTGGCGGGA) by
site-directed mutagenesis; pBB was similarly derived from pTS-A and
carries site E2F-b in both the distal and the proximal locations.
Effector constructs were synthesized by cloning the coding sequences
for the E2F-1, E2F-4, DP1, pRb, p130, and p107 proteins under the
control of the cytomegalovirus promoter/enhancer region in a
pBluescript vector. A construct carrying the lac Z gene
under the cytomegalovirus promoter was used to control the efficiency
of transfection.
Transfections--
Cells were passaged at approximately 1 × 106 cells/25-cm2 flask the day before
transfection; on the following day, a mixture containing DOTAP reagent
(Boehringer Mannheim) and DNA was added; 4 µg of CAT and 1 µg of
Protein Extracts and Western Blot Assays--
Extracts were
prepared either from whole cells or after nuclei isolation as described
in Ref. 23 with minor modifications. Briefly, cells were resuspended in
two packed cell volumes of hypotonic buffer (10 mM Hepes,
10 mM NaCl, 3 mM MgCl2, 0.05%
Nonidet P-40, 1 mM EDTA, and 1 mM EGTA), with
freshly added aprotinin, leupeptin, and pepstatin A (1 µg/ml each), 1 mM sodium orthovanadate, and 1 mM sodium
fluoride, incubated 30 min on ice and disrupted by repeated pottering
while microscopically monitoring the incorporation of trypan blue
(0.25% solution). The homogenate was centrifuged at 500 × g for 20 min at 4 °C; the supernatant containing the cytoplasmic fraction was concentrated and adjusted to 150 mM NaCl (final concentration). The nuclear pellet was
washed in five packed cell volumes of hypotonic buffer, resuspended in
radioimmune precipitation assay buffer (50 mM Tris, pH 8.0, 150 mM NaCl, 1% Nonidet P-40, 0.5% sodium deoxycholate, 1 mM EDTA, and 1 mM EGTA) containing protease and
phosphatase inhibitors as above, and lysed on ice. Gel electophoresis
in SDS-polyacrylamide and electroblotting on polyvinylidene difluoride
membranes were carried out as described elsewhere (9). Kaleidoscope
molecular weight markers were from Bio-Rad. Membranes were blocked in
5% (w/v) low-fat milk in TBST buffer (10 mM Tris, pH 8.0, 150 mM NaCl, 0.1% Tween 20) at 4 °C overnight and then
incubated for 2 h at room temperature with the following primary
antibodies in 5% milk/TBST: anti-E2F-4 (C-20, Santa Cruz
Biotechnology), anti-p27 (F-8, Santa Cruz Biotechnology), anti- Gel Shift Assays--
Protein extracts from NIH/3T3 cultures
were prepared essentially as described in Ref. 35. All buffers and
solutions contained 1 µg/ml each aprotinin, leupeptin, and pepstatin
A; 4 mM sodium orthovanadate; and 4 mM sodium fluoride. The following oligonucleotides and
their reverse complementary strands were used: Sp1-3,
5'-AATTCCGGCCCGCCCCGCGCTTG-3'; E2F-b,
5'-GCATCGCCGCGGGCGTTTTGGCGGGAAGCGC-3'; E2F-c,
5'-AATTCGCGTTTCCCGCCGCTG-3'; TATA, 5'-GCAGAGCATATAAGGTGAGGTAGGA-3'. All
gel shift experiments were routinely performed using at least two
independent extract preparations. Binding reactions with E2F
oligonucleotides were set up as in Ref. 36 with 20-100 pg
of [ Northern Blot Experiments in E2F-overexpressing Cell
Lines--
Cell lines were all based on NIH/3T3 fibroblasts.
Retroviral infections were used to establish stable E2F overexpressing
cell lines. Expression constructs were based on the pBabe-puro vector harboring hemagglutinin-tagged versions of either E2F-1, E2F-2, E2F-3,
E2F-4, or E2F-5 (see also Ref. 37). Cultures were grown in Dulbecco's
modified Eagle's medium supplemented with 10% (v/v) newborn calf
serum, starved by washing the cells twice with phosphate buffered
saline and culturing them for 46 h in medium containing low serum,
i.e. 0.25% (v/v) FCS, and restimulated for 15 h by applying high serum again. RNA was extracted as described in Ref. 38;
15-20 µg of total RNA/lane were used for Northern blot analyses. The
probe was generated by polymerase chain reaction amplification of the
p19.6 plasmid containing the RanBP1 cDNA
(GenBankTM accession X6045) and subsequent purification of
the radiolabeled product on a Sephadex-G50 spin column. A
glyceraldehyde-3-phosphate dehydrogenase cDNA probe was also used
for control. Radioactive hybridization signals were both
autoradiographed and quantified on a phosphorimager (Fuji).
Identification of RanBP1 Promoter Elements Required for Promoter
Activity in Asynchronously Cycling Cells--
Previous deletion
mapping analysis established that a 273-base pair fragment from the
RanBP1 promoter, schematically shown in Fig.
1A, carries all the
information required for control of transcription (11). Two sites match
the E2F consensus: site E2F-c (positions The E2F Sites Exert Independent Roles in Cell Cycle-regulated
RanBP1 Transcription--
RanBP1transcription is up-regulated after
15 h of cell cycle entry, i.e. when cells progress
beyond the G1/S transition (6, 7). To assess the role of
particular promoter elements in up-regulation of transcription, the
constructs shown in Fig. 1A were transfected in cell
cultures that were serum-starved and either maintained in conditions of
growth arrest or restimulated and harvested 15 h after release of
the proliferation block; cell cycle arrest and progression through S
phase were assessed by FACS analysis (Fig.
2A). We found that the
activity of the wild-type pTS-A promoter construct was up-regulated
after 15 h of restimulation (Fig. 2B) and ranged from
20 to 30% above the recorded level in asynchronously growing cells.
None of the mutated promoters in the distal E2F-b or Sp1.3 sites (pmE,
pmS, pmES, and pQl) reached the activity level of the wild-type
construct in S phase: thus, G1/S up-regulation of
transcription requires the distal sites and cannot be sustained by the
E2F-c site alone, as also seen in asynchronously proliferating cells
(Fig. 1B). However, inactivation of site E2F-c (pGi
construct) was not neutral, but resulted in failure of
G0-associated repression of transcription. Thus, formally
distinct functions are exerted by each E2F site: E2F-c mediates
transcriptional repression in arrested cells, whereas the neighboring
E2F-b and Sp1.3 sites are both required for G1/S activation
of transcription. To further verify that site E2F-b acted as an
activating promoter element, a 60-base pair long fragment from the
RanBP1 promoter retaining site E2F-b but not site Sp1.3 (6),
was cloned upstream of the SV40 minimal promoter, formed by two copies
of the 21-base pair repeat and a TATA box (39); as shown in Fig.
3A, in the pE1 chimeric
derivative, site E2F-b is again immediately 5' of a functional Sp1
site, yet is now inserted in a TATA-dependent context.
These experiments (Fig. 3B) show that site E2F-b confers
up-regulation to the SV40-derived promoter in an S
phase-dependent manner.
The Distal E2F-b Site Is Responsive to Exogenously Expressed E2F
Factors--
The finding that each E2F site contributes to cell
cycle-regulated RanBP1 transcription through formally
distinct mechanisms suggests that they respond differently to E2F
factors. Co-transfection experiments were designed to assess the
responsiveness of the wild-type RanBP1 promoter and mutated
derivatives to exogenously expressed E2F factors. These experiments
were carried out in growth-arrested cultures, in which the basal
activity of the RanBP1 promoter is low. Both the E2F-1 and
E2F-4 members of the family were chosen for this analysis: E2F-1
carries its own nuclear localization signal and is efficiently
transported to the nucleus; in contrast, E2F-4 has no nuclear
localization signal and is only transported to the nucleus when
complexed with other proteins (23, 25). In our experiments, E2F-4 was
used in combination with a construct expressing the DP1 dimerization partner.
Various molar ratios of E2F and DP expression constructs to CAT
reporters were preliminarily assayed; FACS analysis revealed that
transfected cells using high ratios of E2F-1, alone or with DP1, to
reporter construct underwent significant apoptosis in response to
a threshold level of E2F-1, as indicated by the appearance of a
hypodiploid cell population of high cellular density (data not shown).
In our experiments, transfection of 0.5-1 µg of E2F-1 construct
alone or of 1 µg of both E2F-4 and DP1 expression constructs per
106 cells induced a low level or no apoptosis and no
significant change in the FACS profile of transfected compared with
control cultures. Western blot experiments were carried out with
extracts from transfected and control cultures: although the endogenous E2F-1 and E2F-4 factors were found to be expressed at different basal
levels, with E2F-1 being virtually undetectable in G0
cells, both plasmids yielded overexpression of both E2F members in
transfected cells (data not shown). Under these conditions, repression
of the wild-type RanBP1 promoter (pTS-A construct) during
growth arrest was fully relieved (Fig.
4). The E2F-4/DP1 combination restored
promoter activity to a comparable level to that measured in S phase
cells, and E2F-1 up-regulated it above S phase levels; transfection of
E2F-4 had instead a minor effect in the absence of a co-transfected DP
partner (data not shown).
The relief of repression in G0 cells was dependent on the
integrity of the distal sites, because the pmE construct failed to
respond to exogenous E2F factors. The pmS construct also showed a very
low level of basal transcription, which was only modestly increased by
exogenous E2F factors; thus, Sp1.3 site disruption impaired the
responsiveness of the neighboring E2F-b site to exogenous E2F factors.
Conversely, the pE1 construct, in which the E2F-b site flanks the SV40
Sp1 sites (see Fig. 3A), was up-regulated by exogenous E2F
factors in growth-arrested cells (data not shown). Finally, the pGi
construct, which is mutated in the proximal E2F-c site yet maintains
both functional E2F-b and Sp1.3 sites, was highly expressed in
G0 cells; the overall activity further increased above
(with exogenous E2F-1) or close to (with the E2F-4/DP combination) S
phase levels, further confirming that responsiveness to E2F factors was
essentially conferred by the distal site. Therefore, the data in Fig. 4
depict a specific responsiveness of the E2F-b site to exogenous E2F factors.
Regulation of RanBP1 Promoter Activity by Pocket Proteins--
We
next asked how the RanBP1 promoter would respond to pocket
proteins by cotransfecting wild-type or mutagenized promoters in the
presence of constructs expressing pRb, p107, or p130 proteins. We
firstly examined cells stimulated to cycle and harvested during S
phase, when RanBP1 promoter activity is highest. All three
pocket proteins clearly affected the wild-type promoter (pTS-A
subclone) and reduced its activity by 50-60% (Fig.
5A). However, repression by
exogenous pocket proteins (particularly p130) in S phase cells was less
effective than that induced by endogenous factors in G0
cells. Incomplete repression may reflect the partial inactivation of
pocket proteins by cyclin/kinase complexes in S phase cells. Indeed,
the dose of transfected pocket proteins used in our experiments did not
significantly alter the cycling profile of transfected, compared with
control, cultures, as indicated by FACS analysis (data not shown); in
those conditions, exogenous pocket proteins underwent S phase-specific
modifications similar to their endogenous counterpart. In particular,
Western blot experiments showed that p130, although overexpressed from
the plasmid vector in G0 cells, underwent degradation in S
phase cells to a similar extent than the endogenous protein (data not
shown). Indeed, overexpression of pocket proteins in G0
cultures effectively repressed the RanBP1 promoter (Fig.
5A). Mutagenized promoters in both distal sites (pmE, pmS,
and pmES constructs) showed a low basal activity, and it was difficult
to appreciate whether further repression by exogenous pocket proteins
was statistically significant. Interestingly, the pGi construct, the
basal activity of which was high in S phase and which might therefore
have been susceptible of undergoing significant variations in the
presence of repressing factors, did not show the 50-60% reduction
typical of the wild-type promoter and was essentially unaffected by
pocket proteins. Pocket proteins caused indeed some repression in
G0 cells, which was evidently mediated by the distal
region; however, pGi activity in the presence of all three pockets
remained substantially higher than that of the wild-type promoter in
the same conditions, indicating that integrity of site E2F-c is
required for effective repression by pocket proteins.
Quiescence-associated repression by pocket proteins is largely exerted
by interacting with E2F promoter elements through E2F factors,
particularly E2F-4, which represents the most abundant family member in
G0 cells (reviewed in Ref. 14). In order to rule out the
possibility that the results in Fig. 5A simply reflected a
nonspecific inhibition of transcription by pocket factors, E2F-4 and
DP1 were coexpressed with pocket-encoding constructs in cotransfection experiments with the RanBP1 promoter. In these experiments,
we analyzed both antagonistic partners of E2F-4, i.e. p107
and p130. As shown in Fig. 5B, both proteins counteracted
relief of G0 repression by E2F-4/DP1; p130, which normally
acts as the preferential E2F-4 partner in G0 cells
(reviewed in Refs. 12 and 14), actually antagonized E2F-4/DP1-mediated
activation in a dose-dependent manner and eventually
reduced RanBP1 promoter activity down to the low basal level
normally seen in G0 cells. Thus, the RanBP1 gene
is indeed a regulatory target of pocket proteins during growth arrest,
which act at least in part through the interaction with E2F-4.
The Regulatory Elements in the RanBP1 Promoter Show Cell
Cycle-regulated Interactions with DNA-binding Factors--
We next
examined the interactions established between DNA-binding factors and
genetically identified promoter elements. Preliminary experiments were
carried out to characterize site Sp1.3 using extracts from
asynchronously cycling cells, where the site showed in vivo
occupancy by protein factors (10). The highest proportion of the
assembled complex migrated with an electrophoretic mobility compatible
with that of Sp1 (Fig. 6A, lanes
1 and 7) was competitively inhibited by excess of
canonical Sp1 sites, including site Sp1.2 from the RanBP1
proximal promoter (lanes 2-3) and an SV40 Sp1 site
(lanes 4-6), and was supershifted by the addition of
anti-Sp1 antibody (lane 8), indicating that Sp1.3 is indeed
a bona fide Sp1-binding site.
Interactions established during cell cycle progression were analyzed
using protein extracts from growth-arrested and S phase cells (Fig.
6B). Site Sp1.3 formed an abundant complex with S phase cell
extracts, whereas a minor proportion of the probe interacted with
factor(s) from growth-arrested cells (Fig. 6B, lanes 1-2); these results did not reflect a lower content of transcription factors
in extracts from G0, compared with S phase, cells, because the TATA box-binding protein had a comparable abundance in extracts from both sources (Fig. 6B, lanes 3-4). When the E2F sites
were incubated with extracts from both starved and restimulated cells (Fig. 6B, lanes 5-12), site E2F-b was found to assemble
more abundant nucleoprotein complexes (lanes 5-8) than did
site E2F-c, the highest proportion of which migrated as free probe
(lanes 9-11), indicating a generally higher avidity for
DNA-binding proteins of site E2F-b compared with E2F-c. Regardless of
their different affinity, both E2F sites interacted with proteins in a
cell cycle-regulated manner: S phase complexes were more abundant than
seen with growth-arrested cell extracts. Qualitative differences were
also apparent between G0 and S phase in the assembly of
high molecular weight complexes, the migration of which was compatible
with that of multimeric complexes containing E2F/DP/pocket proteins
(Fig. 6B, compare lanes 5 and 7 and
lanes 9 and 11), indicating that
G0-specific interactions had been disrupted and replaced by
S phase-specific complexes.
To define these interactions in more detail, binding reactions with
both E2F sites were carried out in the presence of antibodies directed
against particular E2F or pocket members. Representative panels are
shown in Fig. 7. G0 complexes
assembled with both E2F probes essentially reacted with antibodies
directed against E2F-4, p130, and pRb. When S cell extracts were used,
however, the probes displayed a different antibody reactivity: E2F-b
effectively interacted with pRb as indicated by the abundance of
supershifted complexes, whereas the reactivity to anti-pRb antibody was
significantly lower with the E2F-c probe (compare Fig. 7A, lane
12, and 7B, lane 15). Because the absolute amount of
pRb protein in the binding reaction is identical, this observation
suggests that reactivity to the preferential partner of pRb in S phase
cells, i.e. E2F-1, differed for the two probes. Indeed,
E2F-c interacted with E2F-4 and, to a lesser extent, E2F-5 (as revealed
on a prolonged gel shift exposure, see Fig. 7B, lane 10)
among E2F family members. Complexes assembled with site E2F-b were also
supershifted by anti-E2F-4 and, in addition, were recognized by
anti-E2F-1, which interfered with the assembly of DNA-binding complexes
with E2F-b, but not E2F-c, probe (compare Fig. 7A, lane 9, to Fig. 7B, lane 12). This differential interference of the
anti-E2F1 antibody was confirmed in several independent experiments
using different preparations of S phase cell extracts and different
lots of antibody (data not shown). We reasoned that the absence of a
discrete supershift might have reflected the low relative abundance of
the E2F-1 species compared with other E2F family members. We therefore
decided to assess enriched extracts from asynchronously cycling cells
transfected with constructs expressing both E2F-1 and DP1: gel shift
experiments (Fig. 8) showed that the
E2F-b probe now assembled an abundant complex that was reactive to
anti-E2F-1 (lanes 1 and 2); binding experiments
were also carried out in conditions of partial competition with the
heterologous E2F-c probe, with the expectation that a low molar excess
of heterologous site would competitively inhibit those DNA-binding
complexes that have similar affinity for both sites, yet would not
interfere with the assembly of high affinity complexes. Indeed, a
10-fold excess of unlabeled E2F-c site competitively inhibited most
DNA-binding complexes assembled with probe E2F-b, yet left a discrete
E2F-1 supershift (Fig. 8, lanes 3 and 4), indicating that the interaction of factor E2F-1 with site E2F-b was
substantially unaffected. In the reverse experiment using E2F-c as the
probe, no DNA-binding component reactive to anti-E2F-1 antibody was
visualized (Fig. 8, lanes 5-8). These results indicate that
E2F-1 binds site E2F-b but not, or only very poorly, E2F-c. The
reactivity of the distal site to anti-E2F-1 antibody was further increased when a longer probe was generated that included both adjacent
E2F-b and Sp1.3 sites (data not shown), suggesting that the
simultaneous binding of both factors to the DNA increases the complex
stability.
Binding experiments thus far were carried out with whole cell
extracts. However, E2F-4 is functionally regulated by
compartmentalization (23-26). Therefore, it was important to assess
whether the binding of E2F-4, depicted as the most abundant DNA binding
activity in Fig. 7, was biologically significant. NIH/3T3 cells
were brought to growth arrest and restimulated for 15 h; nuclei
were isolated, and protein extracts were prepared from the nuclear and
cytoplasmic fractions (23). E2F-4 was similarly abundant in whole cell
extracts from both G0 and S phases in Western blot assays;
however, although it is nuclear in G0 cells, the bulk of
E2F-4 protein was found in the cytoplasm during S phase (Fig.
9). In order to ascertain that the
anti-E2F-4 reactive material depicted in Western assays was
biologically active in our cell cultures, as previously shown in
several cell lines (25, 26, 40), we assayed these extracts in gel shift
experiments. Most DNA binding activity reactive to anti-E2F-4 antibody
was found in extracts from G0 nuclei; however, the
distribution of anti-E2F-4 reactive protein was reversed in S phase
extracts, with the highest proportion of E2F-4 DNA binding activity
being detected in the cytoplasmic fraction (data not shown). These
findings together suggest that the highest proportion, if not all, of
the E2F-4 pool has exited the nucleus of S phase cells.
In summary, protein binding experiments indicate that all
RanBP1 promoter elements establish cell cycle-regulated
interactions with factors. The binding of Sp1 to site Sp1.3 is
quantitatively up-regulated in S phase. In addition, subtle sequence
differences affect the assembly of S phase complexes with the E2F
sites. Both sites interact similarly with complexes containing E2F-4
and pocket proteins in G0 cells; however, assay of nuclear
and cytoplasmic extracts indicate that E2F-4 is exported out of the
nucleus in S phase; at the same time, newly assembled complexes
containing E2F-1 preferentially interact with site E2F-b, which
actively contributes to S phase up-regulation of the RanBP1
promoter, but not with site E2F-c, which confers G0 repression.
The Activating Role of Site E2F-b Is Dependent on Its Position in
the Promoter Context--
We finally asked whether the DNA sequence
preference depicted in vitro for factor E2F-1 was the sole
determinant of the different genetic functions exerted by the two E2F
sites in control of RanBP1 transcription. To ask that
question, we moved site E2F-b, i.e. the preferred target of
the E2F-1 activator in vitro, to the proximal position that
is normally occupied by site E2F-c. Two novel reporter constructs were
generated (Fig. 10A): pBB
carries duplicated E2F-b, but no E2F-c, sites, whereas pmBB carries a
mutated E2F-b site in the distal region and a wild-type E2F-b sequence
replacing the proximal E2F-c site. CAT assays of these constructs in
growth-arrested and restimulated cells (Fig. 10B) showed
that pBB, despite carrying two copies of the E2F-b site, was not
up-regulated any more efficiently than the wild-type promoter in S
phase cells. In addition, in the absence of a functional distal region
(pmBB construct), the E2F-b site failed to reinstate S phase activity
from the proximal position normally occupied by site E2F-c, contrary to
what would have been expected if the activating function of the E2F-b
element was exerted in a position-independent manner. These results
suggest that the activating function of site E2F-b does not simply
require the element integrity but has to be exerted from its natural
position in the native promoter context and/or requires proximity with the flanking Sp1 site.
E2F-1 Up-regulates Endogenous RanBP1 mRNA
Transcription--
Cotransfection assays in which both the promoter
and the E2F expression constructs are overexpressed elicit the
responsiveness of particular cis-active sequences to E2F family
members, yet do not indicate which E2F factor actually regulates
transcription of the endogenous RanBP1 gene in
vivo. That question was examined in NIH/3T3 cell lines that were
infected with retroviral vectors directing the synthesis of individual
E2F members. Cell lines overexpressing E2F-1 to E2F-5 factors were
generated, and several independent clones were isolated for each E2F
member. We ascertained that expression levels of the exogenous E2Fs
were equal among cell lines; in addition, the gene encoding cyclin E, a
known target gene of E2F-1, -2, and -3 factors (41-43), showed a
5-10-fold induction;2 a more
detailed characterization of the cell lines will be reported elsewhere.
Northern blot experiments showed that RanBP1 mRNA levels are indeed up-regulated in cell lines that overexpress E2F-1 compared with parental NIH/3T3 cells, or to cells infected with viral vector alone (M2 cell line). Whereas the RanBP1 and
GAPDH mRNA transcripts are expressed with comparable
abundance in control NIH/3T3 cells, RanBP1, but not
GAPDH, mRNA transcription shows significantly increased
levels in three independently selected E2F-1 overexpressing clones
(Fig. 11A). Cell lines
expressing other members of the E2F family show no significant
difference in the expression level of RanBP1 mRNA
compared with control cultures (Fig. 11B). Thus, RanBP1 is indeed a regulatory target of activation by E2F-1
in vivo.
In the present work, we have addressed the mechanisms controlling
cell cycle-regulated transcription of the RanBP1 gene. We have concentrated on two E2F-binding elements in two previously identified promoter regions that control G1/S up-regulation
(6, 7) and basal transcription (9), respectively. We have found that
each E2F site controls a distinct aspect of RanBP1
transcription: the distal E2F-b site acts as an activating element and,
together with the flanking Sp1.3 element, confers high levels of
transcriptional activity during S phase. The proximal E2F-c site
instead acts as a negative control element during quiescence.
Conclusions drawn from the genetic dissection of the RanBP1
promoter are schematized in Fig.
12A.
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EXPERIMENTAL PROCEDURES
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150 to
90 relative to TS-1 (6), henceforth referred to as the
distal promoter region. That region includes target sites for E2F/DP
(E2F-b site) and Sp1 (Sp1.3 site) factors, both of which are
footprinted in vivo in cycling cells (10). Thus, the E2F
sites in the RanBP1 promoter display different protein
binding properties in vivo.
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-galactosidase reporter constructs were routinely used. In
cotransfection experiments, constructs expressing E2F-1, E2F-4, DP1,
pRb, p130, and p107 were used. Various amounts (i.e. 0.2-2
µg) of effector plasmid DNA were initially tested, and routinely,
0.5-1 µg were used. In coexpression experiments, we used constructs
expressing E2F-4 and DP1 (1 µg each), together with constructs
encoding either p107 (1.5 µg) or p130 (1.5 µg or 3 µg). Vector
DNA was added to equalize the total DNA amount in all experiments. The
medium was replaced 6 h after transfection with either low serum
(0.5% FCS) to induce growth-arrest or complete medium (10% FCS) to
maintain asynchronous proliferation. To obtain S phase-enriched
cultures, cells that had been starved for at least 48 h were
stimulated to reenter the cycle by adding 15% FCS-containing medium
and collected after 15 h of restimulation. Transfected cells were
harvested and lysed by repeated freeze-thawing cycles. Proteins were
extracted from each sample in 100 µl of Buffer A (100 mM
Hepes, pH 7.9, 1.5 mM MgCl2, 10 mM
KCl); 10 µl were used for determining the extract concentration using
the Bradford assay kit (Bio-Rad), 80 µl were used to measure
synthesized CAT enzyme using the CAT enzyme-linked immunosorbent assay
(Boehringer Mannheim), and 10 µl were used to measure the amount of
-galactosidase synthesized from the cotransfected construct, using
the
-galactosidase enzyme-linked immunosorbent assay (Boehringer
Mannheim). Promoter strengths were quantified by calculating the ratio
of CAT/
-galactosidase activities from each transfected sample. To
compare results from different experiments and calculate mean and S.D.
values, promoter values are expressed relative to that of the wild-type
RanBP1 promoter (pTS-A construct), which was taken as 100%
in most experiments unless otherwise indicated in the text.
tubulin (Amersham Pharmacia Biotech) and anti-histone H1 (Upstate
Biotechnology, Lake Placid). All primary antibodies were used at 0.5 µg/ml, except for anti-
tubulin, which was used 0.05 µg/ml.
Bands were detected using horseradish peroxidase-conjugated secondary
antibodies (Santa Cruz Biotechnology) and revealed using the enhanced
chemiluminescence detection system (ECL-plus reagents, Amersham
Pharmacia Biotech).
-32P]ATP-labeled oligonucleotide and
7-10 µg of protein extract in a 20-µl reaction; 5 µg of protein
were used in binding reactions using extracts enriched in E2F-1 and
DP1. Gel shift conditions using the Sp1 and TATA oligonucleotides were
described previously (9). For supershift experiments, antibodies (0.1 µg/µl of reaction) were added to the binding mixture for 3 h
on ice. The following antibodies were used: pRb (C-15), p107 (SD9),
p130 (C-20), E2F-1 (KH95), E2F-4 (C-20 sc866), E2F-5 (C-20), and Sp1
(PEP2) (all from Santa Cruz Biotechnology).
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31 to
20 relative to the
TS-1 transcription start) flanks an element termed HFE, acting as the
target of retinoid receptors and required for basal transcription (9);
site E2F-b (position
115 to
106), near a putative Sp1-binding site
(Sp1.3, position
100 to
92), falls in a region previously
identified for conferring G1/S up-regulation of
transcription (6). To investigate the contribution of each element to
RanBP1 promoter activity, several promoter constructs
directing transcription of a CAT reporter sequence were synthesized by
site-directed mutagenesis: the distal elements were mutated both
individually in the E2F-b (pmE) and Sp1.3 (pmS) sites, or
simultaneously (pmES); the proximal E2F-c site was mutated in the pGi
construct, which maintained both wild-type distal sites; and finally,
the pQl construct carries mutated versions of both E2F sites. Promoter
reporter constructs were transfected in asynchronously cycling NIH/3T3
cells, and levels of synthesized CAT enzyme were measured (see under
"Experimental Procedures" for details). Results in Fig.
1B show that transcriptional activity of all clones mutated
in the distal sites was drastically reduced compared with the wild-type
promoter; destruction of the Sp1.3 site impaired promoter activity
somewhat more significantly than that of the E2F-b site; the
simultaneous inactivation of the adjacent Sp1.3 and E2F-b sites did not
amplify the effect of single mutations, indicating that both the Sp1.3
and E2F-b distal sites are important promoter elements in asynchronous
cell cultures. In contrast, transcription from the E2F-c mutated
construct (pGi) was comparable to, or slightly more efficient than,
that of the wild-type promoter (Fig. 1B). Thus, efficient
RanBP1 promoter activity in cycling cells requires the
integrity of both distal E2F-b and Sp1.3 sites.
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Fig. 1.
Activity of wild-type and mutated
RanBP1 promoter constructs in asynchronous NIH/3T3
cells. A, map of assayed promoters; crosses
indicate mutagenized sites, and arrows indicate the
transcription start (TS-1). B, bars show the activity of
reporter constructs relative to that of the wild-type promoter (pTS-A
construct), which was taken as 100%. In these and all following
experiments, absolute promoter strengths are calculated as
CAT/ -galactosidase activity in each transfected sample. Mean and
S.D. values were calculated from at least six independent assays for
each construct.
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Fig. 2.
Activity of the wild-type and mutated
RanBP1 promoter constructs in growth-arrested and S
phase cells. A, FACS analysis of asynchronously cycling
(C), serum-starved (G0) and S phase
(S) NIH/3T3 cells harvested 15 h after cell cycle
reentry. Upper panels show the DNA content of the cell
populations as determined by Pr I incorporation; lower
panels show a biparametric analysis of the cell cycle, in which
bromodeoxyuridine incorporation (indicating the extent of DNA
replication) is plotted versus the DNA content.
B, relative activity of the wild-type and mutated promoter
constructs in growth-arrested (shaded histograms) and S
phase restimulated (open histograms) cell cultures. The mean
value obtained for the wild-type promoter (pTS-A construct) in S
phase-cells was taken as 100%; mean and S.D. (bars) values
were calculated from the following number of experiments: 10 for pTS-A
and pmE; 7 for pmS, pmES, and pGi; and 4 for pQl.
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Fig. 3.
S phase-dependent up-regulation
by the E2F-b element in a heterologous promoter context.
A, maps of the pA10 construct, carrying the SV40 minimal
promoter, composed of two copies of the 21-base pair repeat
(hatched boxes) and a TATA element (filled box)
upstream of the CAT gene, and of the pE1 chimeric promoter construct.
The transcription start is arrowed. B, promoter activities
in G0 (shaded histograms) and S phase
(open histograms) NIH/3T3 cells; activities are expressed
relative to that of the pA10 promoter in G0 cells (taken as
1). Mean and S.D. (bars) values were calculated from three
independent assays.
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Fig. 4.
Activity of the wild-type and mutated
RanBP1 promoter constructs in the presence of
exogenous E2F-1 or E2F-4 plus DP1, factors. Histograms
represent the relative activity of reporter promoter constructs in the
presence or absence of constructs expressing either E2F1 or the
E2F4/DP1 combination in G0 cells; for comparison, promoter
activities were also assessed in restimulated cells collected during S
phase. pTS-A activity during S phase was taken as 100%. Mean and S.D.
(bars) values were calculated from nine (for pTS-A) or four
(for pmE, pmS, and pGi) cotransfection experiments with each set of
constructs.
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Fig. 5.
Activity of the wild-type and mutagenized
RanBP1 promoter constructs in the presence of
exogenous pocket proteins in S phase and G0 cells.
A, histograms represent the relative activity of the
wild-type (pTS-A) and mutagenized constructs in S phase and
G0 cells, in the presence of constructs expressing pocket
proteins as indicated. The activity of pTS-A in S phase cells was taken
as 100%. Mean and S.D. (bars) values were calculated from
four cotransfection experiments for each construct. Activity of pmS and
pmES were not assayed in the presence of p130. B, histograms
represent the relative activity of the pTS-A promoter in G0
cells alone or in the presence of the E2F-4/DP1 combination (1 µg
each), with or without constructs expressing pocket proteins: 1.5 and 3 µg of p130 construct, or 1.5 µg of p107 construct were used. The
activity of pTS-A with E2F-4/DP1 was taken as 100%. Mean and
S.D. (bars) values were calculated from three
cotransfection experiments.
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Fig. 6.
Protein binding features of the
RanBP1 promoter elements. A, gel shift
assays of the Sp1.3 oligonucleotide with 5 µg of NIH/3T3 extract from
asynchronously growing cells (lanes 1 and 7) and
increasing amounts of site Sp1.2 from the proximal RanBP1
promoter (lanes 2-3) or of SV40-derived Sp1 site (SV40,
lanes 4-6), and in the presence of anti-Sp1 antibody
(lane 8). The asterisk marks the Sp1 supershift.
B, gel shift experiments using extracts from growth-arrested
(G0) and S phase (S) NIH/3T3 cells, with Sp1.3
(lanes 1 and 2), TATA (lanes 3 and
4), E2F-b (lanes 5-8), and E2F-c (lanes
9-12) oligonucleotides; 5 µg (lanes 1-4) or 10 µg
(lanes 5-12) of protein extracts were used. The association
of TAFs (TATA box-binding protein (TBP)-associated factors)
in lanes 3 and 4 marks the assembly of
transcriptionally competent complexes, whereas the formation of the
TATA box-binding protein-TATA complex is a relatively constant event.
In lanes 5-12, and + indicate the absence and presence,
respectively, of homologous competitor DNA (50-fold excess).
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Fig. 7.
Immunological analysis of complexes
interacting with sites E2F-b and E2F-c with growth-arrested
(G0) and S phase (S) cell
extracts. Assayed antibodies are specified above each lane.
Supershifts can be seen in lanes marked by the asterisk,
whereas antibody interference is indicated by . A,
supershift assays of complexes binding to the E2F-b probe.
B, supershift assays of complexes binding to the E2F-c
probe; all panels correspond to a standard 15-h exposure, except for
lanes 8-11, which show a longer exposure of lanes
2-4 to visualize the anti-E2F-5 antibody reaction.
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Fig. 8.
Supershift assays using extracts from cells
transfected with E2F-1/DP constructs. 5 µg of protein extract
were used with probes E2F-b (lanes 1-4) and E2F-c
(lanes 5-8) in the presence of anti-E2F1 antibody
( E2F-1, lanes 2, 4, 6, and 8),
with (lanes 3, 4, 7, and 8) or without
(lanes 1, 2, 5, and 6) heterologous competitor
site. A 10-fold molar excess of heterologous DNA was used to achieve a
partial competition; the asterisk marks the E2F-1
supershift.
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Fig. 9.
Distribution of the E2F-4 protein during the
cell cycle. Western blot of whole cell extracts (WCE)
during growth arrest (G0) and S phase (S)
analyzed with anti-E2F-4 antibody. Anti- -tubulin and anti-p27
antibodies were used to control protein loading and cell cycle reentry,
respectively. Subcellular compartmentalization was analyzed using
cytoplasmic (C) and nuclear (N) extracts from
synchronized G0 and S phase cultures and analyzed with an
antibody against E2F-4. Antibodies to
-tubulin and histone H1 were
used to control the purity of the cytoplasmic and nuclear
fractions.
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Fig. 10.
Activity of RanBP1 promoter
constructs carrying displaced elements in growth-arrested and S phase
cells. A, map of assayed promoters; crosses
indicate sites inactivated by mutagenesis. In constructs pBB and pmBB,
site-directed mutagenesis was employed to replace the E2F-c sequence
with E2F-b (c b box); the transcription start (TS-1) is
arrowed. B, relative activity of promoter constructs in
cells cultured as for Fig. 2A to induce growth arrest
(shaded histograms) and synchronous S phase progression
(open histograms). The mean value obtained for the wild-type
promoter (pTS-A construct) in S phase-cells was taken as 100%; mean
and S.D. (bars) values were calculated from four independent
experiments.
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Fig. 11.
Expression of the endogenous
RanBP1 gene in cells overexpressing E2F factors.
A, Northern blot analysis of the RanBP1 and
glyceraldehyde-3-phosphate dehydrogenase (GAPDH) mRNAs
in parental NIH/3T3 cell cultures and in three independently selected
clones overexpressing E2F-1. C, asynchronously cycling
cultures; G0, growth-arrested cultures; S,
restimulated cells collected 15 h after cell cycle reentry.
B, the histograms represent the
RanBP1/GAPDH mRNA levels, as quantified by
phosphorimager reading of Northern blots from different
E2F-overexpressing cell lines in high serum, compared with control
(NIH/3T3) and vector-infected (M2) cells. Overexpressed E2F factors are
indicated below each lane.
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Fig. 12.
Growth control and cell cycle control
elements in the RanBP1 promoter. A,
genetic functions of RanBP1 promoter elements identified in
loss of function experiments; the arrow indicates the major
start site of transcription. B, hypothetical model
summarizing the interactions of positively and negatively acting
factors with promoter elements in G0 and in S phase cells.
ssBP, single-stranded DNA-binding protein. × indicates
transcriptional repression in growth-arrested cells.
Although the dual role of E2F elements in positive and negative control of transcription is well documented (reviews in Refs. 12-14; also see Ref. 44), few promoters contain E2F sites in which the functions are clearly separated in genetic terms. Different functional properties were previously attributed to two E2F sites in the p107 gene promoter (45): mutagenesis of the proximal site affected promoter activity more dramatically than that of the distal site, indicating a functional hierarchy between the sites, and the amplified effect of simultaneous compared with single site mutations suggested cooperation. In the RanBP1 promoter, the E2F sites actually exert opposite functions; hence, we have asked which molecular mechanisms underlie these distinct roles.
Both sites showed cell cycle-regulated interactions with factors, yet their protein binding specificity was different. G0 complexes formed with both sites showed a similar reactivity to antibodies, particularly against E2F-4 and p130. Most E2F-4 DNA binding activity exited the nucleus of NIH/3T3 cells during S phase, as revealed by Western blot assays (Fig. 9), in agreement with studies in different cell lines (23-26, 28, 40). At that stage, a differential reactivity to E2F-1 was depicted for each site, because antibody interference in crude extracts and supershift in overexpressing extracts indicated that only E2F-b, but not E2F-c, interacted with E2F-1 (Figs. 7 and 8). Thus, E2F-1 shows some DNA sequence preference, in agreement with results of recent CASTing experiments that pinpointed a differential affinity of specific E2F/DP heterodimers for particular E2F sequences (46). Preferences in E2F/DNA interaction are reflected by the composition of pocket-containing complexes: both sites showed a marked reactivity to the anti-p130 antibody with G0 extracts; in S phase extracts, most p130-containing complexes had been disrupted as indicated by the weak antibody reactivity. Complexes assembled with site E2F-b, but not E2F-c, were now supershifted by anti-pRb, further supporting the idea that E2F-b is a target of E2F-1, which interacts with high affinity with pRb.
The different protein binding abilities in vitro are paralleled by differences both in site occupancy in vivo and in the activity of mutagenized promoters. E2F-b falls in an extended genomic footprint also covering site Sp1.3 (10). Transcriptional activation of RanBP1 at the G1/S transition (7) is concomitant with the synthesis of new E2F-1 (47, 48). The E2F-b site is a target of E2F-1 in vitro, and mutational inactivation of that site impairs transcriptional activation in S phase (Fig. 2B). Finally, overexpression of E2F-1, but not of other E2F factors, activates endogenous RanBP1 transcription (Fig. 11). These data indicate that RanBP1 G1/S up-regulation is mainly controlled by E2F-1 via the E2F-b site.
The DNA sequence preference is not sufficient, however, to determine per se the functional role of E2F elements. The position in the promoter context is also relevant. Insertion of the E2F-b site away from the Sp1.3 site, near the HFE, did not support G1/S up-regulation in the absence of a functional distal site (Fig. 10). In contrast, insertion of site E2F-b in the SV40, TATA-dependent promoter near an Sp1-binding site contributed to S phase activation (Fig. 3). Previous reports (29, 30) indicate that neighboring E2F and Sp1 elements cooperate in cell cycle-regulated transcription. Karlseder et al. (29) showed physical interaction between Sp1 and E2F factors and demonstrated cooperation in DNA binding by genomic footprinting analysis of target promoter elements. In our experiments, mutation of the Sp1.3 site affected the RanBP1 promoter responsiveness to exogenous E2F factors, despite the integrity of the E2F-b site (Fig. 4), and inactivation of E2F-b (pmE), Sp1.3 (pmS), or both (pmES), similarly impaired transcriptional activity (Fig. 2B). These findings suggest that full S phase activity requires the simultaneous interaction of Sp1 and E2F-1 with their target sites in the distal region.
Site E2F-c acts instead as the target of repressing factors during growth arrest, because E2F-c mutation derepressed the RanBP1 promoter and alleviated the requirements for exogenous activators in G0 cells. Other E2F sites acting as repressing elements include those of the genes encoding B-myb (49), cyclin A (50), and E2F-1 itself (51), and mutagenesis at those sites renders these promoters constitutively active. In these instances, E2F factors essentially act as vectors of repressor molecules to the promoter. In our experiments, E2F-4/DP complexes can activate or inhibit RanBP1 transcription depending on their ratio to pocket proteins: E2F-4/DP1 act as activating complexes when overexpression levels are such that endogenous pocket proteins are titrated (Fig. 4), yet exert an inhibitory role in the presence of co-expressed pocket proteins (Fig. 5B), implying that they can indeed bridge repressing factors to the RanBP1 promoter.
Together, the results support the model in Fig. 12B: in the proposed model, E2F-4 would essentially act as a vector of repressors in G0 cells. Although both E2F sites can interact with E2F-4, site E2F-c is essential for effective repression. E2F-c is flanked by the HFE, encompassing the TS-1 site. Previous characterization of that element showed binding by retinoid X receptor members in quiescent cells, associated with promoter inactivity (9). During transcriptional commitment, retinoid X receptors are displaced by single-stranded binding proteins that expose the TS-1 site, as depicted by the appearance of sensitivity to nuclease S1 in vivo (9). It is possible that pocket-bound E2F factor(s) interacting with site E2F-c and retinoid X receptors binding to the HFE stabilize each other and/or act in concert to determine promoter inactivity in G0 cells. During G1 progression, pocket protein phosphorylation and nuclear export of E2F-4 would free the promoter from repressing factors; at the same time, coordinated up-regulation of E2F-1 and Sp1 factors would yield a stable complex over the distal promoter elements and hence up-regulate RanBP1 transcription.
These results contribute to identify two levels of control for the
genes of the Ran signaling network. Examined members thus far are
regulated by growth-dependent mechanisms: Ran is
induced as an immediate-early responsive gene in the presence of serum (52), and RCC1 is induced by the Myc protein through an
E-box promoter element (53). In RanBP1, the alternative
binding either of retinoid receptors in quiescent cells or of
single-stranded DNA-binding proteins in cycling cells to the HFE act as
mutually exclusive signals from the proliferation apparatus to the
transcription machinery. These mechanisms may converge to coordinately
down-regulate the genes of the Ran network in quiescent and terminally
differentiated cells. In cycling cells, the Ran network is active in
cell cycle coordination (reviewed in Refs. 4 and 5). It has been shown that the molecular balance between RanBP1 and RCC1 is crucial in this
control (8, 54). We have shown here that this is essentially achieved
by the binding of E2F-1 and Sp1 to upstream promoter sites, yielding
high levels of RanBP1 transcription during S phase. These
findings further support the view that RanBP1 may act as a
pivotal gene linking cell cycle-regulated transcription by E2F factors
and the coordination of cellular processes controlled by the Ran network.
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ACKNOWLEDGEMENTS |
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We are grateful to Enrico Cundari and Rosamaria Mangiacasale for flow cytometry analyses, to Carmine Pittoggi for contributing to the initial phases of this work, to Filippo D'Ottavio and Graziano Bonelli for excellent technical assistance, and to Pidder Jansen-Duerr, Armando Felsani, and Willy Krek for the gift of expression constructs.
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FOOTNOTES |
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* This work was supported by grants from Associazione Italiana per la Ricerca sul Cancro (to P. L.), the Dutch Cancer Society (to R. B.), and the European Union (Contract BMH4-C796-1529 to P. L. and R. B.).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 Fondazione Buzzati-Traverso.
¶ Supported by a fellowship from the MURST (Ministry of University and Scientific and Technological Research).
** To whom correspondence should be addressed. Tel.: 39-06-445-7528; Fax: 39-06-445-7529; E-mail: lavia{at}axrma.uniroma1.it.
2 R. M. Kerkhoven and R. Bernards, unpublished data.
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ABBREVIATIONS |
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The abbreviations used are: RanBP1, Ran-binding protein 1; TS, transcription start; HFE, Htf9 footprinted element; FCS, fetal calf serum; FACS, fluorescence-activated cell sorting; CAT, chloramphenicol acetyltransferase.
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