Department of Pathology Erasmus University 3000 DR Rotterdam, The Netherlands
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ABSTRACT |
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INTRODUCTION |
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Two functionally active AR-binding sites (androgen response elements or AREs) were identified in the proximal PSA promoter, at positions -170 (ARE-I) and -394 (ARE-II), respectively (11, 13). Although the proximal PSA promoter, including ARE-I and ARE-II, is more active in LNCaP prostate cells than in nonprostate cells, its activity is relatively low. This low level of activity suggested that the proximal PSA promoter is not sufficient to account completely for androgen regulation of the endogenous PSA gene, as observed in LNCaP cells (11). This indicated to us that additional cis-acting control elements residing outside the proximal promoter might contribute to androgen-regulated PSA gene expression. For several strong, tissue-specific promoters, like those of the ß-globin and tyrosine amino transferase (TAT) genes, it has been well established that important control elements are located in regions far upstream of the proximal promoter (14, 15, 16). These distal enhancers cooperate with the proximal promoter for high expression of the specific gene.
To identify putative regulatory elements upstream of the PSA gene, DNaseI- hypersensitive sites (DHSs) were mapped in chromatin from LNCaP prostate cells. Functional analysis of a DNaseI-hypersensitive region far upstream of the PSA gene showed the presence of a complex, androgen-regulated enhancer. In this study we present a detailed analysis of this strong enhancer, which contains a functionally active, high-affinity AR-binding site (ARE-III). Furthermore, we compare the AR-binding affinity and the functionality of this novel ARE with that of the previously identified ARE-I and ARE-II (11, 13). An abstract describing parts of this work has been published previously (17).
While this work was in progress, Schuur et al. (18) reported the identification of a 1.6-kb upstream enhancer fragment (-3.7 to -5.3). This fragment encompasses the 440-bp core enhancer region, which is the basis of the present study.
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RESULTS |
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To identify additional regulatory elements, we mapped DHSs in the 31-kb
region between the PSA and KLKI genes in chromatin from the
prostate-derived cell line LNCaP, grown in the presence and absence of
androgens, and in HeLa cell chromatin. DNA from DNaseI-treated nuclei
was digested with EcoRI and evaluated for location of DHSs
by Southern blot analysis with the appropriate hybridization probes.
With two different probes, DHSs could be found (Fig. 1).
No other DHSs were detected over the 31-kb region with any of the
probes tested (data not shown). Hybridization of
EcoRI-digested DNA from LNCaP cells with a 1.1-kb
HindIII-EcoRI fragment, spanning exon 1 and
intron 1 of the PSA gene, showed one DHS (DHSIV), which was most
prominent in the presence of R1881 (Fig. 1C
). This DHS mapped to the
proximal promoter region. Hybridization of genomic DNA from
R1881-treated LNCaP cells with a 0.5-kb
EcoRI-HindIII probe (-6 kb) revealed the
presence of a cluster of three DHSs, approximately 4 kb upstream of the
PSA gene (Fig. 1A
). The position of this cluster of DHSs could be
confirmed by hybridization with a more downstream located probe (data
not shown). Analysis of the same region in chromatin from LNCaP cells
grown in the absence of hormone showed that DHSII at -4.2 kb is
clearly androgen regulated. Intensity of DHSI, at approximately -4.8
kb, is also influenced by the presence of R1881 during LNCaP culturing.
The weak DHSIII (at -3.8 kb) could be found both in the absence and
presence of R1881. Although weak, DHSI and DHSIII might also be present
in chromatin from HeLa cells, which do not express PSA (Fig. 1B
). In
contrast, DHSII was clearly absent in HeLa cell chromatin, indicating
cell specificity.
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To determine whether core enhancer activity was directly androgen
regulated, the BstEII-PstI fragment was linked in
both orientations to the TK promoter (TK-85-s-LUC and TK-85-as-LUC,
respectively), and LNCaP cells were transfected with these constructs
(Fig. 3C). The result clearly showed that this 440-bp fragment
contained orientation-independent, intrinsic androgen- responsive
enhancer activity. This observation correlated with the strong androgen
regulation of DHSII, linking results of the transient transfection
studies with the activity of the promoter of the endogenous PSA gene.
Additionally, the results indicated synergistic cooperation between the
upstream enhancer and the proximal PSA promoter, because PSA-61-LUC and
PSA-85-LUC were considerably more active than TK-85-LUC (Figs. 2A
and 3
, A and C, and data not shown).
Identification of an ARE in the Core Enhancer Region
To identify candidate AR-binding sites, DNaseI footprints were
determined over four overlapping core enhancer segments, utilizing the
purified AR DNA-binding domain (AR-DBD). The only clear protection that
was observed was located in the middle part of the fragment, over the
sequence 5'-ACTCTGGAGGAACATATTGTATCGATT-3', directly upstream of the
ClaI site (Fig. 4A). The protected area
contained the sequence GGAACAtatTGTATC, which shows high homology
(overall 9 of 12 bp), with the consensus sequence GGT/AACAnnnTGTTCT for
high-affinity AR binding (19). Competition was found with a 100-fold
excess ARE consensus oligo, but not with an excess of an NF-1 consensus
oligo (Fig. 4A
, lanes 4 and 5), indicating specificity of the
interaction. Although both the BstEII-SalI and
EcoRV-PstI subfragments contributed to maximal
activity of the core enhancer (Fig. 3A
), AR binding was not observed in
one of these fragments (data not shown). Gel retardation analysis of a
double-stranded oligonucleotide encompassing the upstream AR-binding
site (ARE-III: ggaGGAACAtatTGTATCgat) with AR-DBD confirmed that this
fragment contains a specific, high-affinity AR-binding site (Fig. 4B
).
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Comparison of ARE-I, ARE-II, and ARE-III
The presence of (at least) three AREs [ARE-I(-170):
AGAACAgcaAGTGCT; ARE-II(-394): GGATCAgggAGTCTC, and ARE-III (-4200):
GGAACAtatTGTATC] in the PSA promoter raised the question of relative
AR-binding affinities of the individual AREs and their separate
contribution to overall androgen regulation of PSA promoter activity.
To compare AR binding to these AREs, gel retardation analyses were
performed with serial dilutions of purified AR-DBD (Fig. 5A). ARE-I and ARE-III turned out to be high-affinity
AR-DBD-binding sites, with comparable AR-binding affinity. The in
vitro interaction of AR-DBD with ARE-II was much weaker.
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Mutational Analysis of ARE-I, -II, and -III
To investigate the role of the individual AREs in overall
androgen- induced transcriptional responsiveness of the 6-kb PSA
promoter, for each individual ARE two different knock out mutations
were introduced in PSA-61-LUC (see Materials and Methods for
sequences of mutated AREs). Transient transfection of LNCaP cells with
the resulting mutated PSA promoter-LUC constructs showed that all three
AREs contributed to androgen regulation. ARE-I(-170) mutations
resulted in an 80% reduction of promoter activity (Fig. 6). Both mutations in ARE-II(-394) had a limited effect
(50% or less reduction). Mutations in ARE-III had by far the most
dramatic effect. As compared with wild type PSA-61-LUC, less than 1%
of activity was retained in the mutated promoter. This finding
indicated that ARE-III is not only a key element in the 440-bp upstream
core enhancer, as shown in Fig. 4C
, but also in the context of the 6-kb
PSA promoter.
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DISCUSSION |
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An important goal of the present study was the analysis of the chromatin structure in a 31-kb region upstream of the PSA gene by the identification of DHSs. Although other explanations are possible, DHSs in chromatin, which reflect structural alterations, are a strong indication for the interruption of the nucleosome structure due to binding of transcription factors to the DNA. In chromatin from LNCaP cells, three DHSs were found clustered in the area from 3.8 to 4.8 kb upstream of the PSA gene. DHSIII (at -3.8 kb) is weak and also present in chromatin from HeLa cells, which do not express PSA. DHSI (at -4.8 kb) is clearly androgen induced in LNCaP cells. DHSII (at -4.2 kb) is by far the most prominent: it is strongly androgen induced in LNCaP cells and absent in HeLa cells. The differences in structure between chromatin from LNCaP cells, grown in the presence and absence of R1881, and from HeLa cells indicated to us a functional role of the DHS cluster in androgen-regulated and cell-specific expression of the endogenous PSA gene.
In transient transfection experiments, the -4.8 to -3.8 region showed
strong, androgen-regulated enhancer activity (PSA-64-s-LUC and
PSA-64-as-LUC in Fig. 3A). It is not clear whether sequences
corresponding to DHSI and DHSIII are present in the shorter active
enhancer construct PSA-73-LUC. Most likely, DHSI maps very close to the
PstI site, which is at the 5'-border of PSA-73-LUC; DHSIII
maps close to the BamHI site, which determines the 3'-border
of PSA-73-LUC. An even shorter fragment, PSA-85-LUC, lacks both DHSI
and DHSIII, but contains DHSII, which maps close to the ClaI
site. The finding that PSA-85-LUC still shows approximately 50% of the
enhancer activity suggests that, in transient transfections, DHSI and
DHSIII sequences play no or only a minor part in PSA promoter activity.
Whether they are required for proper expression of the endogenous PSA
gene remains to be determined. The finding that ARE-III at -4.2 kb
corresponds to DHSII in the chromatin suggests that ARE-III is not only
essential in the PSA promoter in transient transfections, but also in
androgen-regulated expression of the endogenous PSA gene.
The upstream core enhancer most probably has a complex structure. We
were unable to narrow down the size of the core enhancer to less than
440 bp without loosing substantial activity. Combined with the
essential role of ARE-III in the enhancer, at least three separate
active regions can be identified in the core enhancer: ARE-III, and the
5'- and 3'-end fragments (see Fig. 3). In each of the two end
fragments, one or more binding sites of ubiquitous or prostate-specific
transcription factors might be located. The possibility that these
fragments contain one or more weak, so far not identified, AR-binding
sites cannot be excluded. In cooperation with ARE-III, and additional
cis-acting sequences within the 130-bp
SalI-EcoRV fragment, a complex enhancer might be
formed with cooperative interactions between the different components.
Further experiments are obviously required to elucidate the detailed
composition of this core enhancer.
A functional ARE-III is a prerequisite for high activity of the upstream enhancer. Inactivation of ARE-III almost completely abolished core enhancer activity. In contrast, truncation of the 5'- and 3'-fragments resulted only in a partial reduction of enhancer activity. However, there is little doubt about a synergistic cooperation of ARE-III with other cis-acting elements in the core enhancer: when attached to the TK promoter, the core enhancer, containing one ARE-III, is superior to even three copies of ARE-III coupled to the TK promoter. A mechanism explaining the central role of ARE-III could be AR-induced DNA bending, enabling the direct or indirect interaction between other transcription factors in the core enhancer. Alternatively, ARE-III-bound AR might be a key element in the interaction of the upstream enhancer with the proximal promoter region or in the recruitment of the RNA polymerase II holoenzyme to the PSA promoter (20, 21). In this respect the PSA upstream enhancer might function as a classic complex, steroid hormone- regulated control region. Similar upstream glucocorticoid-regulated enhancers, composed of GR-binding sites, binding sites for the liver-enriched transcription factors HNF-3 and HNF-4, and binding sites for more ubiquitous transcription factors, have been identified for the TAT gene, which is highly expressed in liver parenchymal cells (15, 16). The enhancer motifs restrict the hormonal activation of the TAT gene to liver cells, not only in cultured cells, but also in transgenic mice (22). The PSA gene is the first example of an androgen-regulated, prostate-specific gene for which such a potent upstream enhancer is documented. Further study of this enhancer may be of great help in the identification of prostate-specific transcription factors and in the elucidation of the mechanism of cooperative interaction between the AR and other transcription factors.
The identification of three AREs in the PSA promoter in the present and in our previous studies (11, 13) readily raised the question of relative AR- binding affinities and functional activities. ARE-I and ARE-III were found to be of similar potency, whereas ARE-II was less active. Mutational analysis indicated a clear synergistic cooperativity between ARE-I and ARE-III and, to a lesser extent, ARE-II. However, inactivation of ARE-III had a far more impressive effect on the 6-kb PSA promoter than mutation of ARE-I. From these findings it can be concluded that the context in which the ARE is present has a pivotal effect on its functional activity. As indicated above, this might involve interactions with other transcription factors, including the spacing between specific cis-acting elements.
Although the PSA gene is the first example of a gene containing a very
potent far upstream ARE, clear synergistic interaction between multiple
ARE sequences has also been found in the proximal (600-bp) PSA promoter
(see Ref. 13, as discussed above) and in the proximal (426-bp)
prostate-specific rat probasin (PB) promoter (23). Both the proximal
PSA and the PB promoter contain one high-affinity and one low-affinity,
functionally active AR-binding site (13, 23). Although much more active
in their natural setting, multimers of the different, separate AREs
from the PSA promoter are functionally active when fused to a
heterologous promoter (Fig. 5B, and Ref.13). In contrast, cooperative
binding of the AR to both AREs in the PB promoter is required for
androgen induction (24).
To investigate cell specificity of PSA upstream core enhancer, we compared its activity in several prostate and nonprostate cell lines. Transient transfection experiments in (PSA negative) PC3, DU145, Hep3B, and COS cells did not reveal any activity, although two of the cell lines (PC3, DU145) originate from a prostate background. The MMTV promoter showed a very limited activity in AR-cotransfected PC3 and DU145 cells, but was clearly active in COS and HeLa cells. Together these findings indicate the absence of one or more transcription factor(s) or coactivator(s) essential for PSA promoter activity in these cells. Alternatively, a specific inhibitor of PSA promoter activity is present. The specificity of DHSII provided additional evidence for cell-specific activity of the PSA promoter.
TK-85-LUC and PSA-61-LUC were both found to be active in T47D cells. In
T47D cells, PSA promoter activity was not only mediated through the AR
but also via the PR (Table 2). These results indicate that activity of
the 6-kb PSA promoter is not entirely prostate- and androgen-specific.
However, PSA promoter activity in LNCaP cells is superior to T47D cell
activity. The issue of receptor specificity should be investigated in
more detail in cell lines containing comparable amounts of PR and AR,
either endogenously or after cotransfection with the respective steroid
receptor expression plasmid. In a similar type of experimental setup,
we generated LNCaP sublines containing a stable transfected GR
expression plasmid. This resulted in dexamethasone induction of the
endogenous PSA gene and GR-regulated activity of the 6-kb PSA promoter
in transient transfections (K. B. J. M. Cleutjens, in preparation). In
summary, steroid hormone-regulated expression of the PSA promoter
depends on the properties of the cell line: the presence of AR, PR, or
GR is an essential, but not the only, factor. Absence of specific
inhibitors or presence of additional transcription factors and/or
coactivators will also be essential.
Although PSA expression was originally thought to be strictly restricted to prostate epithelial cells, low PSA expression in mammary tumor cells has been recently published (25, 26). The activity of the PSA promoter in transfected T47D cells, which are negative on Northern blots for endogenous PSA expression, could be in accordance with these findings. It would be of interest to determine whether, in mammary tumor tissue, PSA expression is progesterone regulated and could be used as a reliable marker for PR-positive tumors. To obtain more definite information on tissue specificity of the 6-kb PSA promoter in both normal and tumor cells, animal studies with PSA promoter constructs are required. If the upstream enhancer is able to confer preferential expression of a target gene to prostate cells, in vivo applications in humans, including gene therapy for delivery of pharmaceutical reagents to the prostate, can be further explored.
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MATERIALS AND METHODS |
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HeLa, T47D, and COS cells were grown in DMEM; Hep3B cells were grown in
MEM, supplemented with 5% FCS and antibiotics. PC3 and DU145 cells
were grown in RPMI 1640, supplemented with 7.5% FCS and antibiotics.
For transfection, PC3 and DU145 cells were grown in DMEM.
Mapping of DHSs
Cultured cells (LNCaP cells, grown in the presence and absence
of 1 nM R1881, and HeLa cells) were washed with ice-cold
PBS. Cells were suspended in 3 ml ice-cold HS buffer (15 mM
Tris-HCl, pH 7.4, 60 mM KCl, 15 mM NaCl, 0.2
mM EDTA, 0.2 mM EGTA, and 5% glycerol,
supplemented with 1 mM dithiothreitol, 0.15 mM
spermine, and 0.5 mM spermidine, directly before use). The
cells were disrupted by passing five to 10 times through a 0.5 x
16 mm (25G) needle. Disruption was monitored by light microscopic
examination. Nuclei were collected by centrifugation for 5 min at 2500
rpm and resuspended in HS buffer to a final concentration of 5 x
107 nuclei/ml. Limited DNaseI digestion was carried out in
a final volume of 0.5 ml HS buffer containing 5 x 106
nuclei, 5 mM MgCl2, and DNaseI (0800 U;
Boehringer Mannheim, Mannheim, Germany). The mixture was incubated for
30 min on ice, and the reaction was stopped by addition of 10 µl 0.5
M EDTA, 12.5 µl 20% SDS, and 50 µl Proteinase K (10
mg/ml). Next, the sample was incubated overnight at 37 C. Subsequent to
phenol/chloroform extraction, the DNA was collected by isopropanol
precipitation. The DNA was dissolved in 100 µl Tris-EDTA buffer and
digested with EcoRI. Restriction fragments were separated by
electrophoresis in a 1% agarose gel and transferred to a nylon
membrane (Hybond N+, Amersham, Cardiff, UK). Filters were
hybridized at high stringency with random primed
32P-labeled probes (as indicated in Fig. 1) using standard
procedures (28).
Construction of Plasmids
All plasmid constructs were prepared using standard methods
(28). The promoterless basic plasmid pLUC, PSA-4-LUC, TKLUC, the human
AR expression plasmid pSVARo, the AR(DBD) expression plasmid pRIT2TAR,
and pMMTV-LUC were described previously (13, 29, 30, 31). PSA-61-LUC was
generated by insertion of the HindIII/HindIII
(-6 kb/+12) fragment of the PSA promoter in the multiple cloning site
(MCS) of pLUC. PSA-1-LUC was generated by ligation of the
BamHI/HindIII (-2.2 kb/+12) fragment in the MCS
of pLUC. PSA-64-s-LUC and PSA-64-as-LUC
(XbaI-StuI, -5.4/-3.2 kb), PSA-73-LUC
(PstI-PstI, -4.7/-3.9 plus
PstI-BamHI -3.9/-3.8 kb), PSA-74-LUC
(PstI-PstI, -4.7/-3.9 kb), PSA-78-LUC
(PstI-EcoRV, -4.8/-4.1 kb), PSA-83-LUC
(SalI-BamHI, -4.25/-3.8 kb), and PSA-85-LUC
(BstEII-PstI, -4.35/-3.9 kb) were generated by
insertion of the appropriate fragments in front of the proximal PSA
promoter (-632/+12) in construct PSA-4-LUC. The artificial
SalI site (-4.25 kb) was derived from the 5'-end of a human
genomic DNA phage insert (4P1, see Ref.7).
Constructs TK-85-s-LUC and TK-85-as-LUC were generated by insertion of the 440-bp BstEII-PstI fragment into the MCS of TKLUC. Constructs ARE-I-TKLUC and ARE-III-TKLUC were generated by cloning three copies of ARE-I and ARE-III oligonucleotides in TKLUC, respectively (sequences of oligonucleotides are shown below). ARE-II-TKLUC was generated by ligation of the double-stranded 3ARE-II oligonucleotide in the SalI site of TKLUC. ARE-I: 5' GATCCTTGCAGAACAGCAAGTGCTAGCTG3' 3' GAACGTCTTGTCGTTCACGATCGACCTAG 5' 3ARE-II: 5' TCGACAGGGATCAGGGAGTCTCACCAGGGATCA- 3' GTCCCTAGTCCCTCAGACTGGTCCCTAGT- GGGAGTCTCACCAGGGATCAGGGAGTCTCACG 3' CCCTCAGAGTGGTCCCTAGTCCCTCAGAGTGCAGCT 5' ARE-III: 5' TCGACGAGGAACATATTGTATCGAG 3' 3' GCTCCTTGTATAACATAGCTCAGCT 5'
pHS1, pHS2, pHS3, and pHS4, which were the starting material for footprint experiments, were obtained by insertion of the blunt-ended BstEII/ClaI, SalI/EcoRV, EcoRV/PstI, and ClaI/NcoI fragments, respectively, into the SmaI site of pTZ19 (Pharmacia, Uppsala, Sweden).
Generation of ARE Mutations
Mutations were introduced in ARE-I (-170), ARE-II (-394), and
ARE-III (-4200) essentially according to the PCR method of Higuchi
et al. (32). Standard amplification conditions were 30
cycles of denaturation for 1 min at 95 C, annealing for 1 min at 55 C,
and extension for 2 min at 72 C. The oligonucleotides that were used
for the generation of the different mutations are listed below. Two
different sets of outer primers were used, one set for ARE-I and -II
mutations and a separate set for ARE-III mutations. Substitutions in
complementary sets of inner primers ARE-I-1, -2; ARE-II-1 and -2;
ARE-III-1 and -2 are underlined (see below). PSA-61-LUC was
used as the template for the first PCR step. In the second PCR step,
appropriate samples of the purified products of the first amplification
reactions were mixed at a 1:1 ratio. The resulting PCR fragments were
cloned and, after sequencing, exchanged with the corresponding fragment
of PSA-61-LUC. ARE-I and -II outer primers:
forward primer: 5' CCACAAGATCTTTTTATGATGACAG 3'
reverse primer: 5' GCTCTCCAGCGGTTCCATCCTCTAG 3' ARE-III outer primers:
forward primer: 5' CTTCTAGGGTGACCAGAGCAG 3'
reverse primer: 5' GCAGGCATCCTTGCAAGATG 3' Inner primers: ARE-I-1: 5' GTAATTGCACATTAGCAATGGGTAACTCTCCC 3' 3' CATTAACGTGTAATCGTTACCCATTGAGAGGG 5' ARE-I-2: 5' GTAATTGCATAGTAGCAAAAGGTAACTCTCCC 3' 3' CATTAACGTATCATCGTTTTCCATTGAGAGGG 5' ARE-II-1: 5' GGTGCAGGCATAAGGGATGCTCACAATCT 3' 3' CCACGTCCGTATTCCCTACGAGTGTTAGA 5' ARE-II-2: 5' GGTGCAGGCATTAGGCAACCTGACAATCT 3' 3' CCACGTCCGTAATCCGTTGGACTGTTAGA 5' ARE-III-1: 5' CTCTGGAGCATAATATTTCAACGATTGTC 3' 3' GAGACCTCGTATTATAAAGTTGCTAACAG 5' ARE-III-2: 5' CTCTGGAGTAGTATATTACAGCGATTGTC 3' 3' GAGACCTCATCATATAATGTCGCTAACAG 5'
Transfections
Cells were transfected according to the calcium phosphate
precipitation method essentially as described (33), using 1 x
106 cells per 25-cm2 flask and 5 µg of the
appropriate PSA-LUC construct. After 4 h incubation with the
precipitate, the culture medium was replaced by PBS containing 15%
glycerol (incubation for 90 sec at room temperature). Subsequently,
transfected cells were incubated in culture medium in the absence or
presence of the appropriate hormone for 24 h. Transfections were
performed in duplicate. Experiments were repeated at least three times
using two independent plasmid isolates.
Luciferase Assay
Cells were washed in PBS and lysed in 300 µl lysis buffer (25
mM Tris-phosphate, pH 7.8, 8 mM
MgCl2, 1 mM dithiothreitol, 1% Triton X-100,
15% glycerol). Next, 0.1 ml luciferin (0.25 µM) (Sigma,
St. Louis, MO)/0.25 µM ATP was added to 0.1 ml extract,
and luciferase activity was measured in a LUMAC 2500 M Biocounter
(LUMAC, Landgraaf, The Netherlands). After a delay of 2 sec (according
to supplier), the light emission was recorded for 5 sec. Luciferase
activities were corrected for variations in protein concentrations of
the cell extracts. Luciferase activities and relative induction factors
are expressed as mean and SEM of at least three independent
experiments.
DNAseI Footprint Analysis
Production and purification of AR(DBD) was done as described
previously (13, 30). Fragments for footprinting were generated by
digestion of pHS1, pHS2, pHS3, and pHS4 with XbaI and
SacI, or with SphI and EcoRI, in order
to identify protected windows on both the upper and lower strand.
Subsequently, fragments were filled in with MMLV-reverse transcriptase
(Boehringer) in the presence of [-32P]dATP and
isolated from nondenaturing polyacrylamide gel. The DNaseI footprinting
experiments were performed essentially according to Lemaigre et
al. (34). Labeled probe (50,000 cpm) was incubated with 1020
pmol AR(DBD) fusion protein for 30 min at 0 C, in the presence of 10
µM ZnCl2. In indicated cases, a 100-fold
excess competitor oligos (consensus ARE or consensus NF-1;
5'-GATCCAGGGAACAGGGTGTTCTACG-3', and 5'-ATTTTGGCTTGAAGCCAATATG-3',
respectively)) was added. Digestion with 0.04 U of DNaseI (Boehringer)
was for 60 sec at 20 C in a final volume of 50 µl. In the absence of
AR(DBD), 0.025 U of DNAseI was used. After phenol/chloroform extraction
and ethanol precipitation, DNA was dissolved in 5 µl formamide-dye
mix (98% formamide, 10 mM EDTA, 0.2% bromophenol blue,
and 0.2% xylene cyanol). After heating to 95 C for 2 min and rapid
cooling on ice, the DNA was separated on a denaturing (7 M
urea) 6% polyacrylamide gel. G and (G+A) sequence reactions according
to Maxam and Gilbert (35) of the same fragment were run as markers
alongside each footprint. After electrophoresis, gels were fixed,
dried, and exposed to x-ray film.
Gel Retardation Analysis
The gel retardation experiments were performed as described
previously (13). Double- stranded oligonucleotides used in gel
retardations: ARE-I: 5'
GATCCTTGCAGAACAGCAAGTGCTAGCTG 3' 3'
GAACGTCTTGTCGTTCACGATCGACCTAG 5' ARE-II: 5'
GATCCAGGGATCAGGGAGTCTCAGG 3' 3'
GTCCCTAGTCCCTCAGAGTCCTAG 5' ARE-III: 5'
TCGACGAGGAACATATTGTATCGAG 3' 3'
GCTCCTTGTATAACATAGCTCAGCT 5'
Shortly, probes were filled in with MMLV-reverse transcriptase in the
presence of [-32P]dATP and subsequently isolated from
nondenaturing polyacrylamide gel. Labeled probe, 50,000 cpm, was
incubated with AR(DBD) (30 fmol to 2 pmol). In indicated cases 100-fold
excess ARE or NF-1 competitor oligonucleotides were added. After
incubation for 20 min, samples were run on a 4% nondenaturing
polyacrylamide gel. Subsequently, gels were fixed, dried, and exposed
to x-ray film.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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This study was supported in part by a grant of the Dutch Cancer Society.
1 Current address: Department of Neurogenetics, Massachusetts General
Hospital, Charlestown, Massachusetts.
Received for publication July 12, 1996. Revision received October 28, 1996. Accepted for publication November 1, 1996.
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REFERENCES |
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