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Department of Biochemistry and Physiology (G.Z., Y.L., S.M., H.A.W., A.E., J.M.S., R.G.S., D.E.M.) Department of Molecular Design & Diversity (R.C., J.D.H.) Merck Research Laboratories Rahway, New Jersey 07065
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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) as a
model, known agonists (thiazolidinediones and 
12, 14-PGJ2) induced a
specific interaction resulting in FRET between the fluorescently
labeled LBD and fluorescently labeled coactivators [CREB-binding
protein (CBP) or steroid receptor coactivator-1 (SRC-1)]. Specific
energy transfer was dose dependent; individual ligands displayed
distinct potency and maximal FRET profiles that were identical when
results obtained using CBP vs. SRC-1 were compared. In
addition, half-maximally effective agonist concentrations
(EC50s) correlated well with reported results
using cell-based assays. A site-directed AF2 mutant of PPAR (E471A)
that abrogated ligand-stimulated transcription in transfected cells
also failed to induce ligand-mediated FRET between PPAR LBD and CBP
or SRC-1. Using estrogen receptor (ER
) as an alternative system,
known agonists induced an interaction between ER LBD and SRC-1,
whereas ER antagonists disrupted agonist-induced interaction of ER
with SRC-1. In the presence of saturating agonist concentrations,
unlabeled CBP or SRC-1 was used to compete with fluorescently labeled
coactivators with saturation kinetics. Relative affinities for the
individual receptor-coactivator pairs were determined as follows:
PPAR-CBP = ER-SRC-1 > PPAR-SRC-1 >> ER-CBP.
Conclusions: 1) FRET-based coactivator association is a novel approach
for characterizing nuclear receptor agonists or antagonists; individual
ligands display potencies that are predictive of in vivo
effects and distinct profiles of maximal activity that are suggestive
of alternative receptor conformations. 2) PPAR interacts with both
CBP and SRC-1; transcriptional activation and coactivator association
are AF2 dependent. 3) Nuclear receptor LBDs have distinct affinities
for individual coactivators; thus, PPAR has a greater apparent
affinity for CBP than for SRC-1, whereas ER interacts preferentially
with SRC-1 but very weakly with CBP. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Ligand-induced transactivation is mediated through interactions with
members of a growing family of coactivator proteins including
CREB-binding protein (CBP/p300) (6, 9), steroid receptor coactivator-1
(SRC-1)/nuclear receptor coactivator-1 (NcoA-1) (6, 10, 11),
transcriptional intermediary factor 2 (TIF2)/glucocorticoid receptor
interacting protein 1 (GRIP-1)/NcoA-2 (12, 13), and p300/CBP
interacting protein (p/CIP) (14). Upon agonist binding, a
conformational change in the LBD creates a coactivator binding surface;
transcriptional activation occurs after recruitment of coactivator(s)
to the receptor. This interaction is viewed as necessary and sufficient
for receptor-mediated transcriptional activation. The binding of
antagonist ligands to nuclear receptors results in a conformational
state that does not allow coactivators to interact (and may also
promote interactions with corepressor molecules). The coactivator CBP
can form a bridge between nuclear receptors and basal transcriptional
machinery (6). In addition, CBP and SRC-1 also contain intrinsic
histone acetyltransferase activity that results in local chromatin
rearrangement that is crucial for transcriptional activation (14, 15, 16).
A ligand- and AF2-dependent interaction between selected nuclear
receptors — retinoid X receptor (RXR), retinoic acid receptor (RAR),
glucocorticoid receptor (GR), vitamin D receptor, thyroid
receptor (T3R), and estrogen receptor (ER) — and
CBP or SRC-1 has been demonstrated using in vitro
glutathione-S-transferase (GST) pull-down experiments and
with far-Western assessment of protein-protein interactions (6). Thus,
AF2 mutants of T3R, ER, or RAR that impair the
transcriptional function are also deficient with respect to coactivator
interactions (6, 17). The important biological role of nuclear receptor
coactivators was elegantly shown by Xu et al. (18) who
characterized defects in steroid hormone action in SRC-1 null mice.
Using homogeneous time-resolved fluorescence (HTRF) technology (19), we
developed a novel approach for characterizing the ligand-dependent
protein-protein interaction between nuclear receptors and coactivators.
This approach was employed to characterize differential effects of
specific ligands for peroxisome proliferator-activated receptor-
(PPAR) and ER on receptor function, to determine the requirement
of the PPAR AF2 domain for coactivator association, and to assess
the affinity and relative specificity of these receptors for CBP
vs. SRC-1.
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RESULTS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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maxex = 307 nm,
maxem = 620 nm); second, a long fluorescence
lifetime, as long as 1 msec, or about 5 to 6 orders of magnitude longer
than that of organic fluorophores, thereby allowing measurements to be
carried out in a time-resolved mode (20, 21). These two properties
result in extremely low backgrounds allowing assays of high sensitivity
where quantities as small as tens of attomoles have been detected (22, 23). HTRF technology also incorporates a second fluorophore, XL665
(maxex = 620 nm,
maxem = 665 nm), which mediates the
fluorescence resonance energy transfer (FRET). An additional benefit of
using lathanide in HTRF FRET measurements is that it has a large R0
value (R0, the distance at which 50% of the energy is transferred).
The R0 for the Eu(K)-XL665 pair is about 90 Å. As a consequence,
larger complexes can be detected, and the use of appropriately labeled
antibody partners becomes much more facile. For the HTRF-based
measurement of receptor-coactivator interaction, excitation of Eu(K) at
337 nm results in emission at 620 nm; XL665 can absorb the 620-nm
emission if both molecules are close to one another (near or within
R0), and subsequently emit at 665 nm. Therefore, FRET between the two
fluorophores is a measure of the proximity of the molecules. The extent
of the specific FRET is measured as a ratio of the emission intensity
at 665 nm vs. that at 620 nm. This ratiometric readout
provides additional advantages, in that 620 nm signal serves an
internal standard, to minimize interference from variations due sample
impurities (color, turbidity), probe concentrations, excitation
intensity, and emission sensitivity. As a result, variation between
multiple determinations is typically less than 2%.
To mimic the functional interaction between nuclear receptors and
coactivators in an HTRF-based format, we used PPAR-CBP,
PPAR-SRC-1, and ER-SRC-1 as model systems (Fig. 1
). The LBDs of either PPAR or ER,
each of which is also predicted to contain the ligand-dependent
coactivator-binding domain, were expressed in Escherichia
coli (PPAR) or yeast (ER) and purified as GST-fusion
proteins (GST-PPARLBD and GST-ERLBD) where the NH2
terminus of the LBDs was fused to the COOH terminus of GST. The nuclear
receptor-binding domains of human CBP, CBP1-453 (amino
acids 1–453), and human SRC-1, SRC568-780 (amino acids
568–780) were also expressed in Escherichia coli and
biotinylated after purification. As shown in Fig. 1, GST-PPARLBD or
GST-ERLBD was indirectly linked to Eu(K) through an anti-GST
antibody, which was covalently liked to Eu(K). CBP1-453 or
SRC568-780 was indirectly linked to XL665 through a
streptavidin (SA)-biotin adapter. Thus, agonist-induced interaction
between the nuclear receptor and its coactivator would bring the two
fluorogenic partners together and result in the nuclear receptor
ligand-dependent FRET. The stoichiometry of biotin labeling that
resulted in maximal ligand-induced FRET was determined for each
nuclear receptor-coactivator pair and was generally 2–2.5 biotins to
one coactivator molecule.
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Agonists Induce Dose-Dependent FRET Resulting from
Interactions between PPAR LBD and CBP or SRC-1 (24, 25). TZDs bind PPAR with high
affinity in vitro and can mediate transactivation of PPAR
response element-luciferase reporter constructs in mammalian
cells. We first established that TZDs can induce the interaction of
PPARLBD with GST-CBP1-453 using a standard pull-down
approach (Fig. 2A); this demonstrated
that PPARLBD and nuclear receptor-binding domain of CBP were
sufficient to mediate the agonist-dependent interaction. Figure 2, B
and C, shows that, in an HTRF approach, TZDs induced a specific
interaction between PPAR-LBD and CBP1-453 or
SRC568-780 in a dose-dependent manner. The
EC50s obtained from this HTRF-based approach correlated
very well with published IC50s from receptor-ligand
binding-experiments and EC50s obtained via transactivation
assays. Thus, the rank order of potency for these different compounds,
TZDA > TZDB > TZDC = TZDD > TZDE, was preserved
relative to published results (24, 25, 26). In addition, these effects
correlate well with previously reported glucose-lowering potencies in
diabetic animals (25). The differential potencies of these PPAR
ligands were also nearly identical when data derived using CBP
vs. SRC-1 were compared. The naturally occurring, putative
endogenous ligand of PPAR, 15-deoxy12,14 PGJ2 (27, 28) also
induced the PPAR-CBP interaction, whereas a related compound, PGJ
A2, which is reportedly inactive as a PPAR agonist, failed to
mediate FRET between labeled PPAR LBD and labeled CBP (Fig. 2D).
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AF2 Domain Is Required for Transcriptional Activation
and Coactivator Association AF2 domain is similarly required for
transactivation and interactions with coactivators, a PPAR mutant
(E471A) was made, in which glutamic acid at position 471, located in a
conserved AF2 core sequence, was substituted by alanine via
site-directed mutagenesis. This mutant, as well as the wild-type
receptor, was expressed in COS1 cells as fusion protein constructs
containing the DNA-binding domain of the glucocorticoid receptor and
the ligand-binding domains of human PPAR: GR/PPARLBD and
GR/PPARLBDE471A. As observed with other tested nuclear receptors,
this mutant completely lost its ability to transactivate the reporter
gene (Fig. 3A). Western blot analysis
showed that the mutant chimeric receptor protein was expressed at a
level similar to the wild-type chimer (data not shown).
GST-PPARLBD-E471A protein was subsequently expressed in
Escherichia coli and purified. Using the HTRF assay, it was
shown to have lost its ability to interact with both CBP and SRC-1
(Fig. 3B), indicating that ligand-mediated PPAR interactions with
both CBP and SRC-1 depend on the AF2 domain of the receptor.
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LBD was used (Fig. 1
).
As shown in Fig. 4, the ERLBD-SRC-1
interaction was also ligand dose-dependent. The EC50s for
17ß-estradiol (E2) and diethylstilbestrol were 0.25
nM and 0.6 nM, respectively, which correlates
well with their reported potencies using ligand-receptor binding and
transactivation assays (29, 30). 4-OH-tamoxifen and clomiphene have
been shown to antagonize the activity of ER by binding to ER LBD and
preventing agonist-mediated recruitment of coactivators (31, 32).
Figure 4B shows that the E2-induced ER-SRC interaction
was effectively blocked by the addition of 4-OH-tamoxifen or
clomiphene. The IC50s obtained using this approach were
also similar to reported results obtained from binding determinations
or functional antagonism in transactivation experiments (31, 32).
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or ER with Different
Affinities shows that
under the same experimental conditions, ligand binding induced the
interaction of ER only with SRC-1, whereas PPAR interacted with
both CBP and SRC-1. This finding may be the result of an intrinsic
difference in affinity between each of these nuclear receptors and the
two coactivators. The affinity between PPAR or ER and between CBP
or SRC-1 was analyzed using saturating concentrations of potent
agonists, TZDA for PPAR and E2 for ER. As shown in
Fig. 5, B–D, unbiotinylated CBP or SRC-1 effectively competed for the
TZDA-induced FRET with saturation kinetics. The PPAR-CBP interaction
showed a higher apparent affinity than PPAR-SRC-1 interaction in
both experimental paradigms, either using biotinylated-CBP or
biotinylated-SRC-1. In contrast, unbiotinylated CBP had little effect
on the ER biotinylated-SRC-1 interaction, although unbiotinylated
SRC-1 could compete efficiently with biotinylated-SRC-1. Further
evidence of ER specificity for SRC-1 vs. CBP was obtained
from data depicted in Fig. 5E. In this experiment, unlabeled
full-length ER was used to compete for interactions of PPAR with
biotinylated-SRC-1 or -CBP. In the presence of E2,
unlabeled ER effectively competed with PPAR for
biotinylated-SRC-1 by disrupting TZDA-induced FRET. In contrast,
increasing ER concentrations did not affect the PPAR-BioCBP
interaction. These data show that under our experimental conditions,
there was a distinct profile of preferred receptor-coactivator
interactions, which follows a rank order of
PPAR-CBP1-453 = ER-SRC-1568-780 >
PPAR-SRC-1568-780 >> ER-CBP1-453.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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The HTRF nuclear receptor strategy described here provides a novel
quantitative tool for studying functional interactions of nuclear
receptors with coactivator molecules (and possibly corepressors as
well). The components of this system, purified recombinant nuclear
receptors and nuclear receptor binding domains of coactivators, are
simple compared with transcription complexes that exist within the
nucleus of cells. We achieved further simplification by using only the
LBDs of the PPAR and ER. However, we have not observed
significant differences when comparing results obtained using
full-length PPAR2 rather than PPARLBD (data not shown). Compared
with other approaches for measuring nuclear receptor-coactivator
interactions noted above, the HTRF-based strategy provides the first
in vitro homogeneous method allowing for quantitation of
such events under equilibrium conditions.
Importantly, the HTRF-based assessment of nuclear receptor activation
was shown to be specific and predictive of in vivo receptor
function, as evidenced by the following three sets of experimental
results. First, EC50s of TZDs and ER agonists obtained
using the HTRF approach correlated very well with other data available
from ligand-binding and cell-based transactivation assays as well as
with in vivo potencies in the case of TZD PPAR agonists.
In addition, the putative natural ligand for PPAR, 15-deoxy12,14
PGJ2, was fully active with an EC50 that reflects its
reported activity using other approaches (27, 34), whereas PGJ A2,
which is reported to be functionally inactive, was unable to induce
FRET. Second, AF2 function, which is required for ligand-dependent
function of nuclear receptors in cells, was shown to be necessary for
transcriptional activation of a chimeric GR-PPARLBD protein in
transfected cells and for interaction of PPAR with both tested
coactivators in the cell-free HTRF context (Fig. 3). Third, two known
ER antagonists, 4-OH-tamoxifen and clomiphene, failed to induce the
association of ER with SRC-1 and could efficiently block the
E2-induced ER-SRC-1 interaction (Fig. 4B). These effects occurred
with half-maximal activities that could be predicted from their
reported potencies as ligands and antagonists in cell-based
assays (31, 32). It is interesting to note that different agonists of
PPAR resulted in different degrees of maximal FRET induction. As
shown in Fig. 2, B and C, testing of TZDE repeatedly yielded 50–60%
maximum FRET induced by TZDA, using either CBP or SRC-1; for other
PPAR ligands, distinct profiles of maximal FRET that differed from
TZDA or TZDF were also observed using SRC-1 (Fig. 2C). Since FRET is a
measure of the proximity of the receptor LBD and coactivator,
saturating concentrations of individual ligands may have resulted in
some variation in net proximity achieved. Alternatively, subtle changes
in the receptor-coactivator affinity may underlie these differences. In
either case, alternative conformational states adopted by PPAR upon
binding to structurally distinct ligands can be implicated. It is clear
that functionally distinct classes of ER ligands (agonists
vs. partial agonists vs. antagonists) exert
alternative receptor conformations as measured by protease protection
(35). Furthermore, we have observed that certain PPAR ligands that
appear to function as partial agonists in cell-based assays also
exhibit reduced maximal FRET in the HTRF assay (J. Berger and G. Zhou,
unpublished). Further understanding of potential variation in
ligand-induced PPAR conformation might provide a molecular basis for
variation in biological responses to in vivo treatment with
different agonists.
An important question concerns the extent to which different receptors
(or different conformations of the same receptor in response to binding
of alternative ligands) can differentially interact with subsets of the
spectrum of coactivators. In other words, does signaling specificity
reside at the level of direct interactions between receptors and
coactivator? There appear to be clear differences in the ability of
individual receptors to interact with corepressors; thus, unlike TR or
RAR, PPAR does not readily associate with nuclear receptor
corepressor (N-CoR) (36). In addition, ligand-induced
association of coactivators (SRC-1 and p140) with RXR is differentially
regulated by the heterodimeric partner; this is permitted in the case
of PPAR/RXR but not in the case of RAR/RXR heterodimers (36).
However, RAR was shown to actually block binding of ligands to RAR/RXR
heterodimers. Therefore, the ability of any given receptor LBD to
exhibit differential interactions with multiple coactivators has not
been adequately explored.
Several investigators have reported that PPAR exhibits a
ligand-induced association with SRC-1 (11, 33, 36). Our data indicate
that PPAR can also be induced to strongly associate with CBP,
suggesting an important role for both SRC-1 and CBP in PPAR-mediated
regulation of gene transcription. In addition, the results we obtained
with either SRC-1 or CBP using several compounds from the two important
classes of PPAR ligands appeared to be nearly identical. CBP has
been shown to be a universal coactivator or integrator required for
many transcription factors, including nuclear receptors, AP-1 (6, 37),
signal transducer and activator of transcription-1 (STAT-1) (38),
cAMP-regulated enhancer binding protein (CREB) (39), and nuclear factor
B (NF-B). Since cellular CBP concentrations may be limiting,
recruitment of CBP to PPAR in response to agonist stimulation might
antagonize the activity of other CBP-requiring transcription
factors. This is a potential explanation for recently reported
observations concerning the ability of 15-deoxy12,14 PGJ2 and
synthetic PPAR ligands to inhibit macrophage activation and cytokine
production (40, 41).
In contrast to results obtained with PPAR, known ER agonists
promoted the interaction of ER with SRC-1 but not with CBP. Further
characterization of the relative affinities of PPAR and ER for
either coactivator revealed that PPAR interacted with CBP with
higher affinity than with SRC-1. Moreover, CBP1-453, which
did not interact efficiently with ER, was unable to compete for the
ER-SRC-1 interaction. The overall rank order of affinities was:
PPAR-CBP = ER-SRC-1 > PPAR-SRC-1 >> ER-CBP.
Poor interaction between a peptide motif derived from the nuclear
receptor-binding motif of CBP with ER has been documented using a
yeast two-hybrid system (17). However, the pattern of potential
coactivators that may interact directly with ER has not been
otherwise assessed. The fact that ER does not interact with CBP with
high affinity does not exclude a role for CBP in ER function. CBP,
via a discrete domain, constitutively interacts with the CBP-binding
domain of SRC-1 (6). Thus, the high-affinity interaction of ER with
SRC-1 could result in CBP recruitment to the transcriptional complex.
In this regard, it is interesting to note that RAR has been shown to
bind to the NH2-terminal domain of CBP; however, this
binding is not required for its function since a CBP
NH2-terminal deletion mutant retained the ability to
augment RAR activation (42).
It is clear that the differential capacity of certain nuclear receptor
classes (such as TR and RAR) to interact with corepressors has
important physiological consequences for in vivo repression
of gene transcription. Recently reported data suggest that the degree
to which receptors can associate with specific coactivators in response
to ligand binding can serve to modulate the subsequent biological
response. Thus, in HeLa cells in which 4-OH-tamoxifen functions as an
ER antagonist, increasing the SRC-1 expression level was able to confer
agonist activity upon this compound (31). The present studies provide a
new and precisely quantitative approach for characterizing interactions
between selected nuclear receptors and coregulators. Using this
approach, we have characterized the molecular pharmacology of
ligand-induced interactions between PPAR or ER and CBP or SRC-1.
Clearly, interaction of these receptors with coactivators that occurs
within a complex nuclear environment in vivo may differ from
the observations reported here. However, our results suggest that
differential affinity profiles may exist for any given combination of
receptor and coactivator. Such differences may contribute to cell
type-specific variation in biological responses that arise because of
relative changes in the pattern of coactivator expression.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Plasmids
pGEXKG-PPARLBD expressing GST fused with LBD (from amino
acids 176 of PPAR1 to 477) of human PPAR was constructed by
subcloning an XhoI-XbaI (XbaI was
blunt-ended with T4 DNA polymerase) fragment of pSG5- hPPAR/GR
into pGEXKG (43) digested with XhoI and HindIII
(HindIII site was blunt-ended with T4 DNA polymerase).
pGEXhCBP1–453, which expresses human CBP NH2-terminal 1–453 amino acids (hCBP1–453), was cloned by PCR. The primers for hCBP1–453 are: 5'-ACTCGGATCCAAGCCATGGCTGAGAACTTGCTGGACGG-3' and 5'-CTCAGTCGACTTATTGAATTCCACTAGCTGGAGATCC-3', and the expected DNA fragment is 1.5 kb. The template used for the PCR was human fetal brain cDNA libary (Stratagene, La Jolla, CA). The PCR-amplified 1.5 kb DNA fragment was digested with NcoI and HindIII and ligated with pGEXKG expression vector digested with NcoI and HindIII to generate pGEXhCBP1-453.
pGEXhSRC568-780 expressing GST fused with a human SRC-1 fragment containing amino acids 568–780 was prepared by subcloning an SRC-1 PCR fragment derived from the human fetal brain cDNA libary. The primers are: 5'-ACTCGGATCCAATTCACCTAGCAGATTAAATATACAACC-3' and 5'-CACATCT-AGATTACTGTTCTTTCTTTTCGACTTTCACC-3'. The 0.6-kb fragment was then digested with BamHI and XbaI and subcloned into the pGEX-4T-1 vector (Pharmacia).
pSG5-hPPAR/GR, which contains human PPARLBD fused to murine
GRDBD, was provided by Dr. Azriel Schmidt (Merck Research
Laboratories). pMMTV/luc, which contains the murine mammary tumor virus
(MMTV) promoter adjacent to the luciferase (luc) gene, was also
provided by Dr. Azriel Schmidt.
pESP1ERLBD expressing GST fused with the LBD (amino acids 302–595)
of human ER was constructed by performing PCR on a human ER clone
using primers HW234
(5'-CGCGGATCCAAGAACAGCCTGGCCTTGTCCCTG-3')
and HW209 (5'-TCAGACTGTGGCAGGGAAACCCTCTGCCTCCCCCGTGATG-3'). The product
was subcloned into pTGEM (Promega, Madison, WI) and sequenced by
PCR (Pharmacia) and subsequent gel electrophoresis (Stratagene Castaway
System). The clone was digested with BamHI (site introduced
by HW234) and SpeI (blunt ended using Klenow DNA polymerase)
and cloned into pESP-1(Stratagene) cut with BamHI and
SmaI.
Preparation of GST Fusion Proteins from E. coli
E. coli strain DH5 (GIBCO BRL, Gaithersburg, MD)
served as a host for either pGEXhCBP1–453 or
pGEXhSRC568–780 and BL21 (Stratagene) for pGEXhPPARLBD.
DH5 or BL21 was cultured in LB medium (GIBCO BRL) to a density of
OD600 0.7–1.0 and induced for overexpression by addition
of IPTG (isopropylthio-ß-galactoside) to a final concentration of 0.2
mM. The IPTG-induced cultures were grown at room
temperature for an additional 2–5 h. The cells were harvested
by centrifugation for 10 min at 5000 x g. The cell
pellet was used for GST-fusion protein purification according to the
recommended procedure from Phamacia Biotech using glutathione Sepharose
beads. hCBP1–453 and hSRC568–780 proteins
were generated by cleaving the corresponding GST fusion proteins with
thrombin.
Purification of GST-ER LBD from Yeast
The LBD of ER was expressed in yeast as a GST fusion protein.
Yeast strains were grown in fermentors at 28 C. The cell pellet (0.8
g/ml) was resuspended in TEGM [10 mM Tris, pH 7.2, 1
mM EDTA, 10% glycerol, 0.7% ß-mercaptoethanol, 1
mM dithiothreitol (DTT), 10 mM sodium
molybdate] containing one pellet/2 ml (40 µl/ml) Protease inhibitor
cocktail PI (Sigma, St. Louis, MO; 50 µg/ml antipain, 0.7 µg/ml
pepstatin, 2 µg/ml benzamidine), and lysed by shaking with glass
beads in a bead beater. Unbroken cells were removed by low-speed
centrifugation at 3000 x g for 10 min. The supernatant
was centrifuged at 100,000 x g for 60 min yielding
about 1 ml of cell extract per gram of cells. One milliliter of extract
was allowed to bind 1 h at 4 C to 2 ml of glutathione-Sepharose 4B
(Pharmacia) in TEGM containing 5 mM DTT, 0.1% Triton X-100
(Sigma), and PI cocktail. The beads were washed extensively with PBS
containing PI cocktail and finally eluted with the following buffer: 50
mM Tris, pH 8.0, 5 mM DTT, 0.1% Triton, 0.1% BSA, 0.1
M sodium chloride, PI, and 20 mM
glutathione.
Preparation of Eu(K)--GST
The anti-GST antibody (500 µg, 3.33 nmol) at 5 mg/ml in 150
mM NaCl-0.02% azide was buffer exchanged into PBS, pH 6.8,
with a BioSpin-30 (Bio-Rad, Richmond, CA) desalting column. To
this was added 0.9 µl of a 9.4 mg/ml solution of SPDP in absolute
ethanol (8.3 µg, 26.7 nmol, 8.0 eq), and the reaction was allowed to
stand at room temperature for 30 min. The thiopyridine moiety was then
cleaved by addition of 5 µl of 400 mM DTT in
H2O (final concentration, 20 mM). After 15 min
at room temperature, removal of excess DTT and buffer exchange into 50
mM Pi, 5 mM EDTA, pH 6.5, was
achieved by increasing the total sample volume to 200 µl with PBS and
then passaging through BioSpin 30 columns. To this was added activated
Eu(K) that had been prepared by reacting 7.8 µl of a 2.5 mg/ml
solution of Eu(K) in 10% DMF/PBS (19.6 µg, 13.4 nmol, 4 eq) with 1.3
µl of a 10 mg/ml solution of Sulfo-SMCC in H20 [13.0
µg, 29.8 nmol, 2.9 eq with respect to Eu(K)] at 4 C for 30 min. The
reaction was allowed to stand at 4 C overnight.
Purification was achieved by a two-step process. First, the material taken up to a total volume of 500 µl with PBS and buffer exchanged into PBS, pH 6.8, with a Nap-5 column. Subsequently, the protein was further purified by size exclusion chromatography using a BioSep-S2000 column coupled with a Waters 625 HPLC system using the same buffer. The materials were then frozen at -70 C until needed.
HTRF Assay Protocol
SA/XL665 was used as supplied by the manufacturer. Reaction
conditions were as follows: 198 µl of reaction mixture [100
mM HEPES, 125 mM KF· 0.125% (wt/vol)
3-[(3-cholamidopropyl) dimethyl-ammonio]-1-propanesulfonate
(CHAPS), 0.05% dry milk, 1 nM GST-PPARLBD or
GST-ERßLBD, 2 nM anti-GST-(Eu)K, 10 nM
biotin-CBP1–453 or biotin-SRC568–780, 20
nM SA/XL665] were added to each well, followed by addition
of 2 µl test sample in appropriate wells. Plates were mixed by hand
and covered with TopSeal. The reaction was incubated overnight at 4 C,
followed by measurement of fluorescence reading on a Discovery
instrument (Packard). One 96-well plate requires about 3 min. The
fluorescence signal is stable for at least 3 days at 4 C. Data were
expressed as the ratio, multiplied by a factor of 104, of
the emission intensity at 665 nm to that at 620 nm. The ratios were
directly used for plotting to obtain EC50 values.
Alternatively, the ratio obtained from each tested well was normalized
by subtracting results obtained from control (dimethylsulfoxide) wells,
followed by calculation of percent maximum activation (using maximal
ligand-induced FRET as 100%).
Transient Transfections
COS-1 cells were transfected with plasmid DNA constructs using a
standard lipofectamine method (GIBCO/BRL). COS-1 cells were seeded in
96-well plates at 1.2 x 104 cells per well in DMEM
containing 10% charcoal stripped FCS for 16–20 h before transfection.
Cells were cotransfected with pSG5-hPPAR/GR wild-type or
pSG5-hPPARE471A/GR mutant, pMMTV/luc reporter construct, and
pSV-ß-galactosidase as internal standard for variation in
transfection efficiency and sample toxicity. The DNA-lipid complex was
removed after 5 h. The cells were exposed to test compounds in
DMEM containing 5% charcoal-stripped FCS. Cell lysates were prepared
with Reporter Lysis Buffer (Promega, Madison, WI). Luciferase activity
in cell extracts was determined using Luciferase Assay Buffer (Promega)
in a ML3000 Luminometer (Dynatech Laboratories, Chantilly, VA).
GST Pull-Down Assay
GST-hCBP1–453 protein (1–2 µg) bound to
glutathione Sepharose (10 µl) was incubated with 0.2 µg of purified
hPPARLBD in 100 µl Buffer A [8 mM Tris, pH 7.4, 120
mM KCl, 8% glycerol, 0.5% (wt/vol) CHAPS, 1 mg/ml BSA]
for 12–16 h at 4 C. Samples were pelleted by centrifugation at
11,000 x g for 20 sec and washed four times with 300
µl cold Buffer A. The samples were then suspended in 20 µl Laemmli
sample buffer, heated for 5 min at 100 C, and electrophoresed in
4–20% SDS-polyacrylamide gels. Electrophoretically separated proteins
were electroblotted onto polyvinylidene difluoride (PVDF)
membrane. The membrane was blocked in TBST buffer (10 mM
Tris, pH 8.0, 150 mM NaCl, 0.2% Tween 20) containing 5%
dry milk for 1 h at room temperature. The primary rabbit antihuman
PPARLBD antibody was raised against purified recombinant
hPPARLBD. Primary antibody incubation was performed in TBST buffer
containing 5% dry milk at 4 C for 12–16 h. Secondary antibody
incubation and chemiluminescent detection with enhanced
chemiluminescence (ECL) (Amersham, Arlington Heights, IL) were
performed as described in instructions provided by the
manufacturer.
|
ACKNOWLEDGMENTS |
|---|
/GR and pMMTV/luc. We also
thank Dr. Barbara Leiting for providing a protein purification
procedure. We gratefully acknowledge the excellent technical assistance
of Sheng-Jian Cai and Gretel Salzmann. |
FOOTNOTES |
|---|
Received for publication April 17, 1998. Revision received June 5, 1998. Accepted for publication June 16, 1998.
|
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, TZDA; 
,
TZDB; 
, TZDC; •, TZDD; 
, TZDE. The experiment was repeated
three times with similar results.
