 |
INTRODUCTION |
The differential chromatin structure and nuclease
accessibility at a promoter region has long been recognized as a key
distinguishing feature between active and inactive genes (reviewed in
Ref. 1). Almost invariably, the promoter regions of active genes are
marked experimentally by hypersensitivity to restriction endonucleases (2, 3) or, more commonly, DNase I (4). Although this hypersensitivity
can be caused by torsional or topological stress in the DNA and
distortions in the chromatin structure resulting from transcription
factor binding, it is generally caused by the absence of nucleosomes or
their remodeling (1, 4). The paucity of nucleosomes, in turn, is a
direct consequence of the repressive effect that nucleosomes have on
the transcriptional machinery and the requirement to alleviate that
effect for productive transcription to occur (5-7). Thus, inactive
genes usually have promoters that are nucleosomal, are insensitive to
DNase I, and are regarded as being in a closed confirmation
whereas active genes usually have core promoters that are
nucleosome-free, hypersensitive to DNase I, and in an open
configuration. Consequently, one of the hallmarks of gene
induction mechanisms is the remodeling of the nucleosomal
architecture of a promoter as it is activated (5, 6, 8).
Enormous strides have been made in the last decade in understanding
transcriptional activation at the chromatin level. In particular, the
activation of expression of many, although not all (9-11), inducible
eukaryotic genes is consistent with what can collectively be called
recruitment models of gene activation (3, 7, 8) (reviewed in
Ref. 12). These models usually require, in response to some
extracellular cue, the induction of a specific transcription factor and
the subsequent interaction of that factor with its cognate response
element, which is invariably a cis-acting sequence located
within the minimal promoter region of the relevant target genes. These
transcription factors then recruit either chromatin remodeling factors
(13, 14) or histone transacetylases
(HATs1; reviewed in Ref. 15)
or both, which catalyze the opening of the chromatin at the promoter.
The subsequent (7) recruitment of additional coactivators, general
transcription factors, and RNA polymerase II then facilitates gene
expression. Implicit in all of these models is the assumption that
chromatin alterations at the promoter will accompany and/or are
required for gene induction (16, 17).
Although the signal transduction pathways responsible for the cellular
response(s) to genotoxic stress are complex (reviewed in Refs. 18 and
19), it has become apparent that the p53 tumor suppressor lies at the
heart of the matter. In particular, p53 suppresses tumorigenic growth
by transcriptionally inducing genes that facilitate either survival or
death of an injured cell (reviewed in Refs. 20-22). Active p53 binds,
albeit with varying affinities (23), to consensus response elements
(p53 REs) within genetic regulatory loci. p53 REs consist of tandem
palindromic decamers of 5'-PuPuPuC(A/T)(A/T)GPyPyPy-3' (where Pu
represents purine and Py represents pyrimidine) (24). Binding of p53 to
p53 REs alters the expression of a host of genes (reviewed in Ref. 25), which fall into five main categories and include genes 1) that act to
arrest the cell cycle at G1/S and G2, 2) that
are involved in the induction of the G2/M cell cycle
checkpoint, 3) that are involved in DNA repair, 4) that play roles in
the induction or suppression (26) of apoptosis, and 5) that are
involved in autoregulation (20-22).
p53 is normally expressed at very low steady-state levels, because it
is rapidly turned over via proteosome-mediated degradation. This
degradation requires the specific MDM2-mediated ubiquitination of p53
(27, 28). The disruption of the inhibitory interaction between MDM2 and
p53, which can occur by phosphorylation of either MDM2 (29, 30) or p53
(reviewed in Refs. 21 and 31), permits the opportunistic access of HAT
complexes to the N terminus of p53. Binding of HATs to p53 results in
acetylation of the C-terminal regulatory domain of p53, which strongly
activates the latent specific DNA binding activity of p53 in
vitro (32). The biological consequences of p53 acetylation
in vivo, however, are less clear, and it has been argued
that at least one important aspect of p53-HAT interactions may
rather be the ability of p53 to facilitate the delivery of the HATs to
the adjacent p53-inducible promoters (33, 34) (reviewed in Ref. 35).
Indeed, the association of p53 with HATs suggests that modulation of
the promoter chromatin structure may be essential for the activation of
p53-responsive genes (33, 34). Thus, implicit in many models of p53
activation is the assumption that alteration or remodeling of the
nucleosomal architecture of a promoter is likely to be a critical
feature of the mechanism of gene activation.
We have experimentally begun to address the question of how p53
influences the chromatin structure of promoters and p53 REs of p53
target genes. Here we demonstrate that the chromatin at three separate
target promoter elements was constitutively open and accessible to
DNase I, regardless of whether gene expression was induced and/or
whether p53 was present. In contrast, the chromatin domains of p53 REs
were closed and were not altered following genotoxic stress or by p53
binding. These experiments demonstrate that p53 activation of gene
expression does not require extensive chromatin alterations at either
the RE or the promoter of the target gene.
 |
EXPERIMENTAL PROCEDURES |
Cells--
A set of matched p53+/+ and
p53
/
HCT116 cell lines (36, 37) were generously
provided to us by Dr. Bert Vogelstein (Johns Hopkins University). All
cells were cultured in McCoy's 5A medium supplemented with 100 µg/ml
penicillin and streptomycin, and 2 mM
L-glutamine at 37 °C with 5% CO2.
RT-PCR--
RT-PCR was performed essentially as described (38).
Total RNA was isolated from p53+/+ and p53
/
HCT116 cells at 0, 2, 4, and 6 h post-IR (10 Gy) treatment via the
TRIzol (Invitrogen) protocol. 5 µg of total RNA was further purified with the DNA-Free RNA kit (Zymo Research; Orange, CA), assayed
for the presence of contaminating genomic DNA, and then used in
downstream protocols. cDNA synthesis reactions were performed via
random hexamer priming with SuperScriptTM (Invitrogen) reverse transcriptase (for p21, 14-3-3
, and
-actin) or via gene-specific priming with ThermoScriptTM (Invitrogen) thermostable reverse
transcriptase at 55 °C (for KARP-1,
5'-CTTATTCCCCGACCGCACCATGTTGCCGGT-3'). The cDNA from each sample
was then subjected to one round of PCR amplification consisting of an
initial melting step (94 °C for 5 min) followed by a number,
optimized for each amplicon, of repetitive cycles consisting of
denaturation at 94 °C for 30 s, annealing at 55 °C for 1 min, and extension at 72 °C for 30 s using primers (see below)
specific to different exons within p21, 14-3-3
, KARP-1, and
-actin (the latter to control for quantification of cDNA pools)
messages. 26 PCR cycles were utilized for p21 and 14-3-3
, 30 for
KARP-1, and 24 for the
-actin control.
Chromatin Immunoprecipitations--
Modifications of the
published method (39) included harvesting in phosphate-buffered saline
solution, pH 7.6, and cross-linking with 2% formaldehyde.
Cross-linking was quenched by addition of 125 mM glycine
for 5 min followed by two ice-cold washes with phosphate-buffered
saline. Cells were permeabilized in lysis buffer (50 mM
HEPES-KOH, pH 7.5, 140 mM NaCl, 1 mM EDTA, 1%
Triton X-100, 0.1% sodium deoxycholate/protease inhibitors) and kept
on ice. Chromatin was sheared by sonication until the DNA was an
average length of 600-1000 bp as assessed by agarose gel
electrophoresis. Cellular debris was removed by centrifugation for 5 min at 13,000 rpm and then the supernatant was centrifuged again for 15 min. A 1/20 aliquot of the cleared extract was reserved as an
input control for PCR reactions, and the remaining extract was
incubated overnight with antibodies at 4 °C on a rotator. 50 µl of
a protein A-Sepharose slurry was added and incubated for 1.5 h at
4 °C while rotating and then the Sepharose beads were pelleted by
centrifugation. The beads were then washed at room temperature twice
with lysis buffer, once with lysis buffer containing 500 mM
NaCl, once with wash buffer (10 mM Tris-HCl, pH 8.0, 250 mM LiCl, 0.5% Nonidet P-40, 0.5% sodium deoxycholate, 1 mM EDTA), and once with 10 mM Tris-HCl, pH 8.0, 1 mM EDTA. Beads were resuspended in 200 µl of elution
buffer (50 mM Tris-HCl, pH 8.0, 10 mM EDTA),
and precipitated protein-DNA complexes were eluted from the
antibodies/beads by incubation at 65 °C for 30 min. The resulting
supernatants, along with input aliquots, were subjected to cross-link
reversal by heating to 65 °C overnight followed by treatment with
100 µg of proteinase K for 2 h at 37 °C. DNA was purified by
phenol/chloroform extraction followed by addition of 2 µg of glycogen
carrier for EtOH precipitation with a 1/10 volume of 3 M sodium acetate. Precipitated DNA was resuspended in 100 µl of water, and 1/50 of the immunoprecipitates or
1/10,000 of the input were used as templates in PCR reactions with locus-specific primers.
DNase I Hypersensitivity Assays--
The basic protocol has been
described (40), and all preparatory steps were carried out at 4 °C.
Briefly, 1.5 to 2 × 108 trypsinized cells were
centrifuged at 2000 rpm for 10 min. The cells were washed twice by
resuspension in 25 mM Mg2+/Ca2+-free
phosphate-buffered saline and centrifugation at 1500 rpm for 10 min.
The cells were then resuspended in freshly made douncing buffer (20 mM Tris-HCl, pH 7.4, 3 mM CaCl2, 2 mM MgCl2, 0.3% Nonidet P-40) and transferred
to a Dounce homogenizer. Following incubation on ice for 10 min and
removal of an aliquot to quantitate cellular breakage, cells were
broken apart with ~ 6 hard strokes and checked by visual
observation under a microscope. Following quantitative breakage, nuclei
were isolated by centrifugation at 900 rpm for 7 min. Nuclei were
washed twice in 25 ml of resuspension buffer (10 mM
Tris-HCl, pH 7.4, 10 mM NaCl, 3 mM
MgCl2) followed by centrifugation at 900 rpm for 7 min.
Nuclei were then resuspended in 1 ml of resuspension buffer, and a
25-µl aliquot was diluted into 475 µl of 1% SDS for an
A260 measurement. The nuclear stock was diluted with resuspension buffer until the A260 of the
1:20 dilution equaled 0.5. A quenched nuclei sample was removed
and added directly to cell lysis solution (Gentra Systems, Minneapolis,
MN). The purified, diluted nuclei were divided into untreated and
treated aliquots and warmed to 37 °C. For the untreated aliquot, an
endogenous DNase I activity sample was collected at 5 min and
added to cell lysis solution. For the treated aliquot, RQDNase I (~1
µg/5 µl; Clontech) was added to a final
concentration of 1 µg/ml of nuclei. Aliquots of the treated nuclei
were removed at 30 s, 2 min, and 5 min and added to cell lysis
solution and mixed well. Genomic DNA was isolated from each sample as
described in the DNA isolation kit (Gentra Systems, Minneapolis, MN)
protocol (scaled up based on cell number) and then resuspended in DNA
hydration solution (10 mM Tris-HCl, pH 8.0) at 0.5 µg/µl. Genomic DNA was then analyzed by Southern blotting to detect
hypersensitive sites at particular loci.
Southern Blots--
All blots were performed with 10 µg of
genomic DNA/lane in 0.6 to 2.0% agarose gels following treatment with
the appropriate restriction enzyme(s) as shown in each figure legend.
Double digestions were performed simultaneously. Following
electrophoresis, agarose gels were transferred to nitrocellulose
membranes as described (41). Hybridizations were performed
according to the PerfectHyb Plus protocol (Sigma) with a 5-min
prehybridization at 64 °C with sheared salmon sperm DNA followed by
the addition of 32P radiolabeled probe prepared via the
random hexamer Klenow synthesis Prime-It II protocol (Stratagene) and
subsequently purified over a G-50 Sephadex column. Following overnight
hybridizations, the blots were washed twice at room temperature with
2× SSC for 5 min, twice (once at room temperature, once at 64 °C)
with 1× SSC for 10 to 20 min, and two to four times at 64 °C with
0.1× SSC. The exposure of the blots to Eastman Kodak Co.
autoradiographic film at
80 °C in the presence of an intensifying
screen varied from 1 h to 3 days.
Antibodies, Primer Sequences, and Southern and Northern Blot
Probes--
Antibodies used in chromatin immunoprecipitation analysis
included phosphoserine 15-p53 (Cell Signaling Technology, Beverley, MA)
and Ab421 and p21/WAF1 (Calbiochem) antibodies. The PCR primers used for the chromatin immunoprecipitation assays were as follows. For
the KARP-1-responsive element: KP-ChIP-9, 5'-AAGATGAGGAAGAGATGGGG-3' and KP-ChIP-10, 5'-TGAGTCAGAAGTGTGAGAGTG-3'; for the
p21/WAF1-responsive element: WAF-ChIP-1, 5'-TCCACCTTTCACCATTCCC-3' and
WAF-ChIP-2, 5'-ATAACTTCTA GCTCACCACCAC-3'; and for the
14-3-3
-responsive element: 1433-ChIP-1,
5'-AAAATCACTCACTCTCACTACCTC-3' and 1433-ChIP-2, 5'-TTCCTGCTTATCTGCCCCAC-3'. The KARP-1 primers generate a 200-bp fragment located 380 bp distal from the RE. The p21 and the 14-3-3 primers generate 586- and 368-bp fragments that span their respective REs.
DNase I hypersensitivity assay Southern blot probes were generated as
follows. For p21/WAF1: the primers WAF1-1, 5'-TCCTCACATCCTCCTTCTTC-3' and WAF1-2, 5'-CGCTCCTATACATCCAAACC-3' were used in a PCR
reaction to make an 882-bp product; for 14-3-3
, the primers 1433-1,
5'-TGTGTAGTGCCAGGTGAAG-3' and 1433-5, 5'-AAGGTGGAGGGGACAAAGAG-3' were
used to make a 1216-bp product. The probe used to detect the KARP-1
promoter locus was a XmaI-BglII 1064-bp
restriction derived from an 1812-bp ScaI GenomeWalker
(Clontech) product generated with the primers
K63-3, 5'-TCTTGACACCCGAACTAAAAACTTGAC-3' and K63-4,
5'-CTCCCTGCTCTGCCTCTCATTATTC-3'.
The PCR primers used for the RT-PCR reactions were as follows:
for p21, 5'-ACTGTGATGCGCTAATGGC-3' and 5'-ATGGTCTTCCTCTGCTGTCC-3'; for
14-3-3
, 5'-GTCTGATCCAGAAGGCCAAG-3' and 5'-CTCCTCGTTGCTTTTCTGCT-3'; for KARP-1, 5'-CGTACAAGAAGGGAGACAAGGACCACTGAC-3' and
5'-CTTATTCCCCGACCGCACCATGTTGCCGGT-3'; and for
-actin,
5'-ATCTGGCACCACACCTTCTACAATGAGCTGCG-3' and
5'-CGTCATACTCCTGCTTGCTGATCCACATCTGC-3'.
 |
RESULTS |
IR-induced Up-regulation of p21, 14-3-3
, and KARP-1 Gene
Expression in Wild-type, but Not p53-Null, HCT116 Cells--
HCT116 is
an immortalized human colon cancer cell line that is diploid, contains
wild-type p53 and p21 genes, and responds normally to DNA damaging
agents with respect to the induction of p53 and cell cycle arrest (36,
37, 42). RT-PCR analysis was used to characterize the induction of
mRNA for three p53-responsive genes in HCT116 cells exposed to 10 Gy of IR. The genes investigated were p21 and 14-3-3
, which
are involved in the DNA damage-inducible G1 and
G2 checkpoints, respectively (reviewed in Ref. 25), and KARP-1, a DNA damage-inducible isoform of the Ku86 DNA repair gene (41,
43).
-Actin mRNA expression was used to control for the quality
of the cDNA pools. In contrast to
-actin, the p21, 14-3-3
,
and KARP-1 mRNAs were expressed at low levels (Fig. 1A, lane 0) (data
not shown) in normally cycling cells. Upon exposure to X-irradiation,
the message levels for all three genes rose sharply in contrast to
-actin expression, which remained essentially unchanged (Fig.
1A). An isogenic HCT116 cell line in which the p53 gene had
been inactivated via somatic gene targeting (p53
/
cells) (36, 37) had very low or non-detectable resting state levels of
p21, 14-3-3
, and KARP-1 mRNAs, and no induction of any of the
genes was observed following IR exposure (Fig. 1B). This
analysis revealed the expected profiles of induction, and the
previously demonstrated dependence upon p53, for these well characterized p53-inducible genes.

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Fig. 1.
p53-responsive mRNAs are up-regulated in
HCT116 cells following IR exposure. A, wild-type HCT116
p53+/+ cells were exposed to 10 Gy of -radiation. At the
indicated time (in h), mRNA was isolated from the cells and used
for RT-PCR using primers specific for the indicated genes. RT-PCR using
-actin-specific primers was performed to ensure the quality of the
mRNA preparations. B, the same experiment as described
for A was carried out except that mRNA isolated from
HCT116 p53 / cells was utilized.
|
|
The Promoters, but Not the p53 REs, for p21, 14-3-3
, and KARP-1
Are Hypersensitive to DNase I--
The putative promoter regions for
p21 (reviewed in Ref. 44), 14-3-3
(45-48), and KARP-1 (41) have
been identified. These promoters correspond to CpG-rich domains
to which the likely start site of transcription (+1) has
been mapped (Fig. 2). Multiple putative
p53 REs reside within the vicinity of these promoters, and those REs
that have been defined functionally as responsive are indicated (Fig.
2). To investigate the chromatin structure of these p53-inducible
genes, we utilized DNase I hypersensitivity assays (reviewed in Refs. 4
and 49). In this technique, Southern blot analysis reveals the presence
or absence of a region of chromatin that is relatively open, and
therefore accessible, to the endonuclease DNase I. Nuclei,
prepared from wild-type HCT116 cells that had been mock treated or
X-irradiated (10 Gy), were harvested and mixed with DNase I for 0.5, 2, or 5 min. Genomic DNA was then purified, subjected to the indicated
restriction enzyme cleavage(s), and analyzed by Southern blotting.
Probes specific to each locus (Fig. 2) were used to detect changes in
the restriction enzyme cleavage patterns of DNA isolated from
irradiated versus non-irradiated cells at the varying
durations of endonuclease treatment. In the absence of DNA damage, the
promoter regions of p21, 14-3-3
, and KARP-1 were already
hypersensitive to DNase I indicating a relatively open (i.e.
non-nucleosomal) chromatin structure (see Figs. 2 and 3, Non-IR). In every case, the
major DNase I hypersensitive site (DHS) detected resided immediately
adjacent to the transcriptional start site (+1; see Fig. 2).
For p21, a second strong DHS was also detected 2 kb downstream of the
transcriptional start site within intron 1 (Fig. 3, EcoRI + EcoRV
digest; see also Fig. 5), but this site was not characterized
further. A distinct lack of DHSs was observed at all of the p53 REs
save for a few weak DHSs flanking the p53 RE in the 14-3-3
locus
(Figs. 2 and 3, Non-IR) (data not shown). To see whether the
chromatin structure of these genes was altered upon DNA damage, cells
were exposed to IR (10 Gy), nuclei were isolated, and DNase I
hypersensitivity assays were performed again. Although the intensities
of the DHSs varied somewhat from experiment to experiment, the only
consistent salient difference detected between the irradiated and
non-irradiated samples was that the irradiated samples were
unexpectedly more resistant to nuclease digestion (Fig. 3, compare
2' of DNase I Treatment lanes for IR
with Non-IR). This trait, which is poorly understood (see
"Discussion") was not characterized further. Most importantly, no
extra IR-inducible DHSs were ever observed. From these experiments we
concluded that the promoter regions for p21, 14-3-3
, and KARP-1 were
constitutively open to DNase I. In addition, we concluded that none of
the p53 REs were DNase I hypersensitive and that this attribute
did not change immediately following X-irradiation.

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Fig. 2.
Cartoons of the genomic loci of the three
p53-responsive genes studied. The genomic DNA is depicted as a
dark line. The starts of transcription are indicated as
+1. The thick vertical arrows correspond to the
location of the major DHS at each locus (see e.g. Fig. 3).
Thinner vertical arrows represent the weaker DHSs, with the
thickness of the arrow approximating the
intensity of the DHS. The promoter regions are embedded within CpG
islands, which are designated by rightward-hatched rectangles
below the dark line. The areas utilized as probes for
the DHS assays are shown as leftward-hatched rectangles
under the dark line. Relevant restriction enzyme sites
are shown as dashed vertical lines, and the distances
between these sites is shown with the thin lines. Exons are
represented as shaded rectangles. The p53 REs are shown as
small ovals. Shaded ovals (responsive) correspond
to REs that are required for transactivation and/or have been shown to
bind p53. In the case of p21, two REs have been identified, and these
are designated as distal and proximal based upon
their proximity to the start of transcription. Blackened
ovals (non-responsive) correspond to REs that are a good match for
the p53 consensus sequence but that do not appear to be involved in
either transactivation or p53 binding. The p53 REs are also
characterized by their homology to the 10-bp consensus sequence;
e.g. 10/10(0)8/10 indicates
an RE that consists of 10 nucleotides that are a perfect match
separated by 0 nucleotides from 10 nucleotides in which only 8 matches
exist.
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Fig. 3.
An open chromatin structure which is
independent of DNA damage exists over the promoters, but not the p53
REs, of p21, 14-3-3 , and KARP-1
genes. HCT116 cells were either left untreated
(Non-IR) or X-irradiated (IR; 10 Gy) and then
nuclei were isolated from the cells as described under "Experimental
Procedures." These nuclei were exposed to DNase I for 30 s
(30"), 2 min (2'), or 5 min (5') and
then genomic DNA was isolated, restricted with the appropriate
restriction enzyme(s) (shown on the left), and subjected to
Southern blot analysis using the probes indicated in Fig. 2. Sites
hypersensitive to DNase I are indicated with arrows. The
thickness of the arrow corresponds approximately
to the intensity of the DHS. An identical analysis for genomic DNA
isolated from cells not treated with DNase I is shown on the far
left of the top blot (Ctrl gDNA). In
addition, the positions (in kb) of the molecular markers used for these
gels are indicated on left side of each autoradiogram. The
approximate locations of the relevant promoter elements for each of the
loci are shown on the far right. p, proximal;
d, distal.
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|
The DNase I Hypersensitivity Profiles of p53 Target Gene Promoters
Is Not Altered Following X-irradiation--
To investigate whether
alteration of the chromatin structure of the p53 target gene promoters
required a lag or maturation phase, HCT116 cells were X-irradiated (10 Gy) and then DHS assays were carried out on cells isolated at varying
times post-irradiation. The DNase I hypersensitivity profiles at 15 min, 30 min, 1 h, or 1.5 h post-irradiation were
indistinguishable from cells that were analyzed immediately following
X-irradiation (Fig. 4) with the exception
that once again IR-treated samples were invariably somewhat more
resistant to DNase I digestion than non-IR samples (compare 0'
lanes with 15'; see Fig. 4). Importantly, however, even
as transcription for these genes rose over 5-fold (Fig. 1), there was
no corresponding detectable alteration of the chromatin structure of
the promoters or the p53 REs.

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Fig. 4.
The DNase I sensitivity profiles of the p21,
14-3-3 , and KARP-1 promoters and p53 REs do
not change over time. A DHS assay as described for Fig. 3 was
carried out except that the cells were harvested, and nuclei were
prepared at the times indicated post-exposure to IR (10 Gy). All
prepared nuclei were subjected to a 5-min DNase I treatment.
|
|
The Absence of p53 Does Not Alter the DNase I Hypersensitivity of
the p21, 14-3-3
, or KARP-1 Promoters--
In the absence of p53,
non-detectable levels of p53 target genes were expressed in cells
exposed to IR (Fig. 1). Thus, to extend these studies, the chromatin
structure of p53-inducible promoters was investigated in HCT116
p53-null cells, where the p53 alleles have been inactivated by two
rounds of gene targeting (36, 37). The DNase I hypersensitivity
patterns of p21, 14-3-3
, and KARP-1 loci in p53
/
cells were indistinguishable from those observed in wild-type cells
with the exception of the two weak DHSs at the 14-3-3
locus, which
could not reproducibly be detected in p53-null cells (compare Fig.
5 with Fig. 3). Moreover, like in
wild-type cells, the hypersensitivity pattern was not altered upon
X-irradiation (Fig. 5, compare Non-IR with IR).
Thus, despite a complete lack of detectable basal transcription of
these genes in p53-null cells, the discrete hypersensitive regions of
the respective promoters were present and indistinguishable from that
of wild-type cells.

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Fig. 5.
The absence of p53 does not affect the gross
chromatin structure of any of the p53-responsive genes studied.
HCT116 p53 / cells were analyzed for DHSs exactly
as described for wild-type cells in Fig. 3. All symbols are as in Fig.
3.
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|
p53 Binding to Its REs Following X-irradiation Is Detectable by
ChIP--
The absence of chromatin alterations at p53 REs as
detectable by DNase I accessibility compelled us to assess p53
interactions with its cognate binding sites by other techniques. Thus,
chromatin immunoprecipitation (ChIP; reviewed in Ref. 50) was utilized to analyze the in vivo occupancy of p53 REs. Preparations of
nuclei from either mock or IR-treated wild-type or p53-null HCT116
cells were cross-linked with 2% formaldehyde and subjected to
sonication to disrupt chromatin into ~600- to 1,000-bp pieces.
Subsequently, p53-DNA complexes were immunoprecipitated, the
cross-links were reversed, and the DNA was purified and then subjected
to PCR analysis for the enrichment of a particular genomic locus. To
ascertain whether a particular subset of modified p53 was associated
with p53 REs we carried out the ChIP experiments using monoclonal
antibodies specific for phosphoserine 15 p53 or by using monoclonal
Ab421, which is known to stabilize the binding of p53 to DNA (51). The
amount of phosphoserine 15 p53 associated with these p53 REs was
initially low in untreated cells, and it increased ~4- to 10-fold in
every case following IR exposure (Fig.
6). Similarly, the monoclonal Ab421
detected an increase in p53 occupancy at these sites following
X-irradiation (Fig. 6), consistent with what has been reported
(23, 51). In all cases, no or very low levels of p53 were detected in
p53-null cells or when an irrelevant antibody (p21) was utilized (Fig.
6). From these experiments we concluded that p53 was indeed binding to
its REs in a DNA damage-inducible manner even though this event was not
accompanied by DNase I hypersensitivity.

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Fig. 6.
p53 interacts with p53 REs in DNA-damaged
HCT116 cells. Wild-type (+ p53 status) and
p53 / ( p53 status) HCT116 cells were
either X-irradiated (10 Gy; +IR) or mock treated
( IR). 30 min later the cells were harvested, and ChIP
analysis was carried out using either monoclonal Ab421 or an
epitope-specific monoclonal Ab to activated p53 (phosphoserine 15 p53). After reversal of the protein cross-links and DNA
purification, a PCR analysis for each of the indicated loci was carried
out with locus-specific primers. For p21, only the distal RE was
analyzed. Enrichment of the p53 RE in the immunoprecipitated DNA pool
indicates the interaction of p53 with the RE. PCR amplification with
primers for 14-3-3 was carried out for more cycles than the other
loci, indicating less enrichment for this locus in the
immunoprecipitation pool. Control input PCR reactions are shown
on the far left. These samples correspond to 1/300 of
the reaction volume utilized for the ChIPs.
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|
 |
DISCUSSION |
In this study we have established that the chromatin structures of
the promoters for three p53-inducible genes were accessible to DNase I
in human HCT116 cells growing under normal conditions and regardless of
whether the cells were damaged or p53 was present. We have demonstrated
that the binding of p53 to its cognate REs in response to DNA damage
occurs in the absence of detectable chromatin alterations at the RE.
These data confirm and significantly extend the previous observations
that the Mdm2 (52) and GADD45 (10) promoters, which are both dependent
upon p53 for DNA damage-inducible gene expression, are constitutively
nucleosome-free and DNase I hypersensitive. This study has important
implications for the mechanism of p53 transactivation.
Significance of DHSs at p53-regulated Promoters under Non-inducing
Conditions--
Empirically, it has long been recognized that the
promoters of genes that are being actively transcribed are
hypersensitive to DNase I, usually because of the absence of
nucleosomes (reviewed in Refs. 1 and 4). For example, the hypoxanthine
phosphoribosyltransferase gene is X-linked. On the active X chromosome,
where the gene is transcribed, the promoter is nucleosome-free (6) and
is hypersensitive to DNase I (6, 53). In contrast, the hypoxanthine
phosphoribosyltransferase gene on the inactive X chromosome is not
transcribed, and its promoter is occupied by nucleosomes, and it is not
DNase I hypersensitive. Reactivation of the inactive allele by
azadeoxycytidine is accompanied by the appearance of hypoxanthine
phosphoribosyltransferase mRNA and the DHS (54). Similarly,
-globin transcription is dependent upon the erythroid Krueppel-like
factor (EKLF) transcription factor. Cells lacking EKLF do not produce
globin transcripts and do not exhibit a DHS at the proximal
-globin
promoter (55, 56). Restoration of EKLF to EKLF-null cells results in
the reappearance of the DHS and of globin transcription (8). These and
similar studies, coupled with the observation that the degree of DNase I hypersensitivity often correlates with nucleosome remodeling (57),
demonstrate that the appearance of a DHS at a promoter usually implies
that the core promoter is nucleosome-free and that the gene is being
transcribed. Because the promoters for p21, 14-3-3
, and KARP-1
contain constitutive DHSs even before the genes are induced (Fig. 3) a
logical interpretation of this observation is that these promoters are
nucleosome-free and that the genes are being expressed in the absence
of DNA damage (Fig. 7B).
Whether the p21, 14-3-3
, and KARP-1 gene promoters are truly nucleosome-free and where the nucleosome-free boundaries reside will
require additional MNase footprinting studies, although these data are
completely consistent with the prior demonstration that the Mdm2 (52)
and GADD45 (10) promoters are constitutively hypersensitive to DNase I
and that positioned nucleosomes flank the GADD45 promoter, which itself
is nucleosome-free (10). Taken together, these studies suggest that the
vast majority of, if not all, p53-dependent DNA
damage-inducible promoters reside in open chromatin. Moreover, the
openness of these promoters is consistent with the genes being
transcribed at low, basal levels in non-damaged wild-type cells (Fig.
1) (data not shown). It should be noted, however, that the basal
expression of p21, 14-3-3
, and KARP-1 in p53-null cells was greatly
reduced over that observed in wild-type cells such that the transcripts
could no longer be detected by Northern blotting (data not shown). It
is unlikely that expression is reduced to zero, however, as the
expression of p21, albeit at significantly reduced rates, has been
reported in a p53-null cell line (33). Together, these data imply that
although p53 is not essential for the basal transcription of these
genes, its presence enhances constitutive expression. This conclusion
is consistent with a study in which a temperature-sensitive p53 protein was shown to induce transcription of the Mdm2 gene in the absence of
DNA damage only under permissive conditions (52). Whether p53 enhances
basal transcription via the same mechanism that it uses to augment
transcription following DNA damage (see below) is a subject that
deserves more study.

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Fig. 7.
Models for transactivation of p53-regulated
genes. A, canonical recruitment model of
transactivation. A target gene is diagrammed in a closed
conformation with nucleosomes (flesh-colored ovals)
occluding the p53 RE (yellow square) and the transcription
initiation site (blue TATA rectangle and arrow).
Following DNA damage, p53 is activated and binds to its RE. It then
recruits HATs (green hexagon) or chromatin remodeling
complexes (CRCs; orange circle), which open the
chromatin occluding the promoter. p53 may then facilitate the
additional recruitment of RNA polymerase II and general transcription
factors (GTFs; pink oval). Transcriptional
activation then occurs, which may be facilitated by additional
coactivators, resulting in high levels of transcription (wavy
green lines). B, simple activation model. All symbols
are as in A. p53 target gene promoters are open, and low
levels of constitutive transcription occur. Following DNA damage,
activated p53 binds to its RE in the context of nucleosomal DNA. p53
then induces high levels of transcription potentially with the help of
additional coactivators. C, paused polymerase model. All
symbols are as in A. p53 target gene promoters are open and
occupied by a paused RNA polymerase II. Following DNA damage, activated
p53 binds to its RE in the context of nucleosomal DNA. p53 then induces
high levels of transcription by inducing re-initiation of
transcription.
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Importance of DHSs at p53-regulated Promoters under Inducing
Conditions--
In higher eukaryotes, chromatin is the physiological
template for transcription. The functional effect of nucleosomes on
transcription initiation is generally repressive, and the ability to
overcome these inhibitory effects is invariably required for
transcription to occur (reviewed in Refs. 1, 4, and 12). More often than not, transcription factors accomplish this task not through intrinsic enzymatic activities but through association with
coactivators, HATs, and chromatin remodeling complexes (1). This
mechanism of transcriptional activation is often referred to as
"recruitment," and a vast amount of data has demonstrated
that recruitment is actively utilized at a number of well characterized
yeast promoters and eukaryotic viral promoters (16, 17) (reviewed in
Refs. 12, 58, and 59). The documented association of p53 with HATs has
led to the hypothesis that p53 similarly activates transcription by
recruitment of factors that will facilitate chromatin opening at
promoters (33, 34) (Fig. 7A).
In this study, however, we have demonstrated that although p53 is
clearly essential for the induction of these genes following IR
exposure (Fig. 1) (reviewed in Refs. 20-22), it does so without grossly altering the chromatin structure of the promoters that it
transactivates (Fig. 3). These observations are virtually identical to
those reported for the GADD45 promoter following IR exposure (10).
Although our data cannot rule out subtle chromatin alterations at p21
and related p53-regulated promoters, they clearly do not support the
contention that significant chromatin alteration is needed for
p53-mediated gene activation. Indeed, the only reproducible alteration
observed following X-irradiation was an increased resistance to DNase I
immediately after irradiation (see Figs. 3 and 4). The mechanistic
basis for this increased resistance is unknown. One possibility is that
the exposure of cells to IR and general radioresistance have been
correlated with the phosphorylation of the histones H2AX (60) and H3
(61), respectively. Histone phosphorylation, in turn, has long been
associated with chromatin condensation, which would be consistent with
the observed reduction in nuclease accessibility. Importantly, however,
no other significant alterations in the DHS profile at the promoter
were observed following X-irradiation (see Figs. 3 and 4) even over a
time interval when p21, 14-3-3
, and KARP-1 transcription was known
to be induced manyfold (Fig. 1).
If the association of HATs with p53 is not required for promoter
opening, then HATs are presumably assisting transactivation through
another mechanism. One possibility would be through their ability to
act as factor acetyltransferases (reviewed in Ref. 15) and activate
either p53 (62-64) or associated cofactors. Thus, our data are most
consistent with a basic activation model in which, in their uninduced
state, p53 target gene promoters are open, and the genes are being
transcribed at a low level (Fig. 7B). Following DNA damage,
p53 is activated and induces transcription either through the
recruitment or stimulation of coactivators (33), including HATs,
general transcription factors, or RNA polymerase II without, however,
concomitant chromatin alterations (Fig. 7B).
Importance of DHSs at p53-regulated Promoters in p53-null
Cells--
The open chromatin structure of p53 target genes was
remarkably independent of p53 expression (Fig. 5). Although unexpected, there is precedent for the lack of a requirement for a transactivator in determining the chromatin structure of the target promoters. For
example, major histocompatibility complex class II genes can be induced
in a retinoblastoma tumor suppressor protein-dependent fashion (65, 66). In retinoblastoma
/
cells these genes
are not induced although the promoter region is constitutively open as
defined by DNase I hypersensitivity (67). Similarly, heat shock genes
are highly induced following exposure to elevated temperatures. The
promoters for these genes, however, are constitutively open and
accessible to DNase I even in the absence of the transactivator, heat
shock factor (reviewed in Ref. 68). In both of these cases the open
architecture of the relevant promoters is established by a second
transcription factor, whose expression precedes the expression of the
transactivator. Thus, regulatory factor X is responsible for
establishing the DHS at the HLA-DRA promoter (69) whereas GAGA
factor performs the same role at heat shock gene promoters (70, 71). A
logical extension of these observations is that there may be a
transcription factor that binds to p53-regulated promoters and
establishes the open chromatin confirmation prior to p53 activation.
The identity of this putative transcription factor is unknown. The p21
promoter has been analyzed exhaustively, and a plethora of
transcription factors have been implicated in its expression (reviewed
in Ref. 44). Unfortunately, much less is known about the promoters for 14-3-3
and KARP-1 (41), and so a direct comparison of potential common cis-acting binding sequences is not possible at this
time. It is interesting to note, however, that all three of these
promoters coincide within CpG islands (Fig. 2). CpG islands are
frequently rich in Sp1 or Sp1-like binding sequences (e.g.
GGGCGG). Moreover, Sp1 family members are known to interact with the
HATs, p300, and cAMP-response element-binding protein-binding protein
(44), and it is tempting to speculate that these HATs cooperate with Sp1 to establish the constitutive open chromatin structures observed at
p53-regulated promoters.
The constitutive open chromatin structure of p53-regulated genes also
suggests an alternative mechanism of transactivation. In the case of
heat shock genes (9, 72) (reviewed in Ref. 68), c-fos (73),
and c-myc (74, 75) activation is regulated by promoter
proximal pausing. For these genes, the chromatin structure of the
promoters is constitutively open and contains an engaged RNA polymerase
II, which is paused during elongation just downstream of the initiation
site. The subsequent binding of the relevant transactivator then
promotes rapid reinitiation of the stalled polymerase (Fig.
7C) (11). Clearly, the chromatin structure of the p21,
14-3-3
, and KARP-1 genes is strikingly similar to the chromatin
structure of the heat shock, c-fos, and c-myc
genes. Moreover, the genes regulated by promoter proximal pausing and p53 are genes that need to be induced in a rapid fashion in response to
a variety of cellular stresses and conditions. Given that, at least
in vitro, the assembly of the core transcriptional machinery takes several min, whereas the transition from initiation to elongation occurs within seconds (76), it is logical that these stress-related genes might be regulated similarly. There are, however, some subtle differences between p53-regulated genes and the genes regulated by
promoter proximal pausing. In the case of the promoter pausing, even
though a large nucleosome-free region exists over the proximal promoter
region, the subsequent initiation of transcription results in
downstream nucleosomal remodeling, which can be detected as additional
nuclease hypersensitivity (11) (reviewed in Ref. 68). In the case of
p53-regulated genes, no additional DHSs were detected (Fig. 3) even
when transcription had increased manyfold (Fig. 1). Moreover, a strong
argument has been made that the open chromatin region of genes
regulated by promoter proximal pausing has evolved specifically to
allow for a very rapid entry of the transactivator, whose
cis-acting RE invariably resides within the promoter region.
For p53-regulated genes, this is clearly not the case as many p53 REs
can and do reside kilobases away from the promoter and the nuclease
hypersensitive region (Fig. 2) (reviewed in Ref. 25). These differences
notwithstanding, it will be important to determine whether the p21,
14-3-3
, and KARP-1 promoters are regulated by promoter proximal
pausing (Fig. 7C).
Importance of the Lack of DHSs at p53 REs--
A second important
finding of these studies was the lack of chromatin alteration (see
Figs. 2-4) associated with p53 binding to its cognate REs (Fig. 6). In
all four cases examined, the REs appeared to contain a chromatin
structure that was inaccessible to DNase I, both before and after p53
binding. Most transcription factors are deleteriously affected, often
by factors of 1000 or more, in their binding to nucleosomal templates
as compared with naked DNA (77, 78). Thus, p53 appears to belong to a
more restricted set of transactivators that are capable of binding to
their cognate recognition sites within the context of nucleosomal DNA
(reviewed in Refs. 79 and 80). Interestingly, however, in many of these
latter cases the binding of the transcription factor to the
DNA-nucleosome complex resulted in weakened DNA-nucleosome interactions
that could be detected as increased nuclease sensitivity (81, 82). In
the case of p53, no additional DHSs were detected after X-irradiation
suggesting that either DNase I accessibility was not sensitive enough
to detect these changes, for which there is precedent (83), or that p53
binding was occurring in the absence of significant nucleosomal
alterations (10, 52). Our conclusion that p53 is capable of binding in
the context of nucleosomal DNA is consistent with the recent
observation that in vitro p53 binds preferentially to
nucleosomal DNA (34). The ability of a transcription factor to bind to
nucleosomes can be facilitated by the presence of multiple factor
binding sites (84) (reviewed in Ref. 79). Thus, it is probably not
fortuitous that p53 REs are composed of four half-sites and that p53
binds as a tetramer (85). The clustering of these sites and the
cooperativity required between p53 monomers may have evolved to
overcome the inhibitory nature of nucleosomes. This feature may also
explain why p53 REs elements can be, and often are, located outside of
the proximal promoter region where most cis-acting
regulatory sequences are generally found.
In conclusion, we have characterized the chromatin structure of p53 REs
and the core promoter regions for three p53-regulated genes. We report
that the promoter regions of these genes contain an open chromatin
configuration that is constitutive and unaffected by the presence or
absence of p53. These findings establish a paradigm for p53 transactivation.