The Hypersensitive Glucocorticoid Response Specifically Regulates
Period 1 and Expression of Circadian Genes
Timothy E. Reddy,a,b,dJason Gertz,aGregory E. Crawford,bMichael J. Garabedian,cand Richard M. Myersa
HudsonAlpha Institute for Biotechnology, Huntsville, Alabama, USAa; Institute for Genome Sciences & Policy, Duke University, Durham, North Carolina, USAb; Departments
of Microbiology and Urology, NYU School of Medicine, New York, New York, USAc; and Department of Biostatistics and Bioinformatics and Institute for Genome Sciences
& Policy, Duke University, Durham, North Carolina, USAd
64). Glucocorticoids bind the receptor with high affinity, leading
to nuclear translocation, direct binding to DNA, and ultimately
regulation of gene expression. The affinity of glucocorticoid for
the GR is a primary determinant of activation. Many studies have
characterized GR binding and transcriptional regulatory activity
at saturating concentrations of hormone at which most of the GR
is bound (7, 38, 43, 48, 50, 56). While these studies have revealed
hundreds of genes that are differentially expressed in response to
tisol concentrations vary throughout the day as part of circadian
rhythms (70) and are often near or below the dissociation con-
stant (Kd) for the GR (46).
Low doses of glucocorticoids that are well below the Kdof the
GR have been shown to elicit GR-mediated gene regulation for a
small number of genes (47, 55). Such findings indicate that GR
activity can be tuned to different doses of glucocorticoids in a
tissue-specific manner, potentially in order to regulate expression
of specific genes at different points in the circadian cycle. For
example, subsaturating doses of the synthetic glucocorticoid
dexamethasone (DEX) can drive expression of the tyrosine ami-
notransferase (TAT) gene in FU5-5 cells but not in the HTC rat
hepatoma cell line (44). In addition, doses of DEX as low as 0.001
nM, well below the Kdof DEX for the GR (Kd, ?3 to 5 nM [31,
inhibit the activity of vasoactive intestinal peptide (VIP) in pri-
mary rat pituitary cells (52). Numerous studies have shown that
gene for Period 1 (PER1), a major component of the mammalian
circadian clock, in humans and rodents (3, 9, 14, 32, 57). Gluco-
corticoids also regulate PER1 in primary tissues in response to
acute physical stress (72) and as part of circadian rhythms (58).
We also recently showed that expression of PER1 is sensitive to
sponse to the steroid hormone cortisol (reviewed in reference
A prominent mechanism for tuning the glucocorticoid re-
sponse involves interaction between the GR and cofactors.
of the GR, the nuclear receptor coactivators 1 and 2 (NCOA1 and
NCOA2; also known as SRC-1 and TIF2, respectively), or of the
CREB binding protein (CREBBP), can dramatically sensitize the
GR-mediated response (47, 61, 63). More recently, a tubulin ty-
was found to cooperate with the NCOA proteins and further en-
hance sensitivity to glucocorticoids (22). Nearby DNA sequence
elements can also tune glucocorticoid responses. For example, a
DNA sequence known as the glucocorticoid modulatory element
sensitivity of the local glucocorticoid response (28, 49, 62). The
modulation of sensitivity typically applies equally well to gene
induction and repression, indicating that the mechanisms of tun-
ing the corticosteroid response may be shared (59). Dose-specific
responses are general to other nuclear receptor-mediated hor-
mone responses, and similar mechanisms can alter expression re-
sponses to estrogens and mineralocorticoids as well (69).
Based on these results, we hypothesized that the dose at which
GR binds DNA and regulates gene expression varies throughout
Received 12 January 2012 Returned for modification 7 February 2012
Accepted 9 July 2012
Published ahead of print 16 July 2012
Address correspondence to Richard M. Myers, firstname.lastname@example.org
Supplemental material for this article may be found at http://mcb.asm.org/.
Copyright © 2012, American Society for Microbiology. All Rights Reserved.
mcb.asm.orgMolecular and Cellular Biologyp. 3756–3767September 2012 Volume 32 Number 18
the genome. To investigate, we measured low-dose specific
changes in GR occupancy and related changes in gene expression
across the human genome. We found genome-wide sensitivity of
with cooccupancy of numerous cofactors. We mapped in detail
the minimal enhancer element responsible for driving the GR-
mediated regulation of the expression of PER1, the most sensitive
glucocorticoid-responsive gene in A549 cells. Doing so revealed
that a complex set of nucleotide signals across a 274-bp minimal
enhancer cooperate to tune the glucocorticoid response. Finally,
we show that targeted expression of PER1 regulates expression of
other circadian rhythm genes many hours later, suggesting a role
in circadian biology.
MATERIALS AND METHODS
Cell growth. A549 cells were grown at 37°C and 5% CO2in F-12K me-
cin (Gibco). AtT-20 cells were grown in F-12K medium with 2.5% fetal
bovine serum, 15% horse serum, and 1% penicillin-streptomycin
(Gibco). Dexamethasone and cortisol were dissolved in ethanol at a
with an equal volume (0.02% [vol/vol]) of ethanol to control for solvent
as described previously with minor modifications (50). Briefly, for
each immunoprecipitation mixture, we used 2 ? 107cells, 5 ?g anti-
body (see Table S4 in the supplemental material), and 200 ?l Dynal
sheep anti-rabbit or sheep anti-mouse antibody beads (Invitrogen).
DNA was prepared for sequencing on an Illumina Genome Analyzer 2
as described previously (50) but without PCR prior to agarose gel size
selection and with 15 rounds of PCR for final library amplification.
by using Bowtie (35) with the parameter “-m 1 –best –strata.” To
evaluate reproducibility between experiments and biological repli-
cates, we correlated the number of reads from each experiment that
aligned within 1,000 bp of a transcription start site. For GR ChIP with
sequencing (ChIP-seq) experiments, binding sites were called inde-
pendently for each biological replicate by using the QuEST Peak Caller
(65) with the “stringent” threshold setting. We then defined the final
set of binding sites for each experiment by requiring each peak summit
(i.e., the position of maximal ChIP-seq signal as determined by
QuEST) in one replicate to be within 50 bp of a peak summit in the
other replicate. We further classified GR binding sites as 0.5 nM, 5 nM,
or 50 nM, according to the lowest dose at which the binding site oc-
curred. For all other ChIP-seq experiments, reads from two biological
replicates were combined into a single data set, and binding sites were
called using QuEST as described above.
Transcription factor binding site motif detection. Motif detection
was performed by using BioProspector (39), followed by BioOptimizer
(26). First, we extracted the genomic sequence for each binding site and
masked low-complexity sequence by using the DUST program. To inves-
motif width of 18 on the 150 strongest binding sites in each class of GR
initial motif on the complete set of DUST-masked sequences for each
class. Motif prevalence was calculated by using FIMO (2), with a signifi-
cance threshold of P ? 1 ? 10?4.
To identify additional DNA binding motifs common to identified
binding sites, we masked GR binding sites identified with FIMO and
then used BioProspector to search for additional motifs. Motifs
matching AP-1 were commonly identified in this second round of
motif detection. For a third round of motif searching, AP-1 sites were
This third round yielded motifs matching that of the FOXA factors, at
motif widths of ?10 bp, and motifs matching that of CREB, at a motif
width of 7 bp.
Measuring the gene expression response to DEX. To evaluate the
gene expression response to DEX, we treated A549 cells for 1 h with 0.1
nM, 0.5 nM, 1 nM, or 5 nM DEX or with ethanol as a vehicle control and
measured gene expression with RNA sequencing as described previously
saline (PBS, pH 7.4), lysed them in buffer RLT (Qiagen) with 1%
?-mercaptoethanol, and extracted total RNA using Qiagen RNeasy
Mini columns according to the manufacturer’s protocol, including the
DNase-mediated DNA digestion step. Total RNA was then poly(A)
selected, fragmented, reverse transcribed to cDNA, and sequenced to a
depth of ?20 million 36-bp reads. We then aligned reads to RefSeq
transcripts by using Bowtie (35), with parameters “-n 2 -k 1 -m
10 –best,” and calculated expression in units of aligned reads per ki-
lobase of exon and per million aligned reads (RPKM).
To calculate differential expression for each gene at each dose
point, we filtered and normalized the expression measurements and
identified genes with expression changes greater than would be ex-
pected by chance according to biological replicates. To do so, we first
filtered transcripts shorter than 150 bp and also lowly expressed genes
(average RPKM ? 2 across all measurements), because they were un-
satisfactorily noisy in our results. Then we normalized expression val-
ues by using variance stabilization as implemented in the vsn package
for the R statistical language. After variance stabilization, we further
filtered out genes with a coefficient of variation between replicates that
was greater than 0.1, removing 83 of 11,225 genes. To model noise
between biological replicates, we used a normal distribution fit to the
difference in transformed RPKM values between biological replicate
experiments (? ? ?2.5 ? 10?3; ? ? 0.12). We then calculated the
statistical significance of differential expression as the probability that
background noise model by using a two-sided comparison. To correct for
multiple hypotheses, we calculated the false discovery rate (FDR) for each
Glucocorticoid reporter constructs. Reporter plasmids were con-
structed either by traditional cloning or by custom synthesis. For the
reporters containing the endogenous promoter as well as for many of
the enhancer constructs, we used PCR with Pfu Ultra II DNA polymer-
ase (Stratagene) to amplify the indicated regions from the A549 ge-
nome and to tail amplified regions with unique restriction enzyme
sites. We then used restriction enzyme digestion of PCR products and
plasmid, followed by ligation to insert PCR products into versions of
the pGL4-luc2p luciferase reporter plasmids (Promega). For the 160-
bp, 135-bp, 100-bp, and the PER1-intronic enhancer constructs that
were modified to match regions of the PER1 upstream enhancer re-
nology) and subcloned into the luciferase reporter plasmid. Reporters
containing the endogenous transcription start site were inserted into
the pGL4.24 vector, whereas enhancer regions were inserted in the
pGL4.14 vector, which contains a minimal promoter. Lastly, we con-
firmed the oligonucleotide sequence of all reported plasmids with
Sanger sequencing. The mouse mammary tumor virus (MMTV) re-
porter we used was pGL4.36 from Promega.
cleotide sequence of enhancer regions, we performed site-directed mu-
tagenesis with the Stratagene QuikChange kit, with the modification of
propagating plasmid in TOP10 chemically competent cells (Invitrogen).
We verified correct mutations by Sanger sequencing.
Transient transfection and reporter dose response. To measure ac-
tivity of each construct, 5,000 A549 cells were seeded into each well of a
Low-Dose Glucocorticoids Regulate Period 1
September 2012 Volume 32 Number 18mcb.asm.org 3757
96-well plate. After incubating cells overnight, we transiently transfected
cells with 100 ng luciferase reporter and 4 ng Renilla-simian virus 40
control plasmid (Promega) by using a 6:1 ratio of FuGene to DNA, ac-
and replaced with medium containing corticosteroids or ethanol as a ve-
hicle control. After 4 h of incubation at 37°C, cells were lysed using Pro-
mega Dual-GLO reagent, and luciferase expression was measured using a
tion efficiency, luciferase signal was normalized to the Renilla signal in
each well, and log2ratios over ethanol-treated control wells were deter-
mined. All experiments were repeated in 8 replicates and at 12 doses. To
1 nM, 2.5 nM, 5 nM, 10 nM, 25 nM, 50 nM, 100 nM, 250 nM, 500 nM, 1
?M, and 2.5 ?M.
Measuring gene expression in response to transient PER1 induc-
tion of PER1 expression, we seeded A549 cells into wells of a 6-well plate.
Twenty-four hours later, we began treating individual wells for 1 h with
0.5 nM DEX. After a 1-h treatment, cells were washed twice with 5 ml of
fresh medium and incubated in 2 ml of fresh medium for 0 to 48 h, as
indicated below. Treatments were scheduled such that all time points for
a biological replicate concluded at the same time, thus limiting effects
due to differing times in culture. Cells were then lysed with buffer RLT
Qiagen RNeasy Mini columns according to the manufacturer’s protocol
and including an on-column DNase digestion. RNA was reverse tran-
scribed using a SuperScript VILO cDNA synthesis kit (Invitrogen), and
gene expression assays (Invitrogen) in a 384-well plate and with an assay
and the average threshold cycle (CT) was used for downstream calcula-
tions. Expression measurements were then normalized to GAPDH to
control for differences in cDNA amount, and finally expression was cal-
culated relative to cells harvested at the same time but that were never
treated with DEX. For the 12-h time course (see Fig. 7E), the procedure
was repeated five times on different days, and we report the relative ex-
pression as the means and standard deviations of those five biological
replicates. For the 48-h time course (see Fig. S6 in the supplemental ma-
terial), six biological replicates were performed.
iments are available both on the UCSC genome browser (http://genome
.ucsc.edu/cgi-bin/hgGateway; human genome version hg19) as well as
at the HudsonAlpha website, http://hudsonalpha.org/sites/default/files
The GR binds the genome in a dose-dependent manner. To un-
derstand the activity of the GR at physiologically low doses of
glucocorticoids, we used ChIP-seq to measure glucocorticoid re-
ceptor binding in response to 0.5 nM (sub-Kd), 5 nM (near the
At the highest dose (50 nM), we found 5,898 GR binding sites,
similar to our previous studies of GR binding with 100 nM DEX
(50). Of the sites we found at the highest dose of DEX, the GR
bound 145 (2.5%) and 1,449 (24.5%) at 0.5 nM and 5 nM DEX,
sites varies throughout the human genome by at least 3 orders of
magnitude. We classified sites by the lowest dose of DEX at which
der of the manuscript as hypersensitive (bound at 0.5 nM DEX),
medium sensitive (bound at 5 nM DEX), or low sensitive (bound
at 50 nM DEX) GR binding sites (Fig. 1B; see also Table S2 in the
the sensitivity of GR binding sites to follow a continuum that
would become evident with the study of many more dose points.
The hypersensitive GR binding sites had stronger binding sig-
nals overall (see Fig. S1 in the supplemental material). Therefore,
in some cases, we may have erroneously classified hypersensitive
sites, because they are also easier to detect, even when weakly
bound. However, many of the medium and low sensitive sites
ultimately had stronger ChIP-seq signals after treatment with 50
in the supplemental material.
The GR commonly binds to a consensus DNA binding motif
sequence of the bound GRE can drive conformational changes in
of corticosteroids, we used de novo DNA binding motif detection
to identify the consensus GR binding motif in each group (39).
While we found a clear match to the known GRE in each class of
sites, the motif did not substantially differ between the classes of
GR binding sites or from that of previous reports (33, 50). The
a specific version of the GRE, as might be expected in light of
recent studies (43) (Fig. 1C). An alternative hypothesis is that
ing site may explain the high sensitivity of some sites. That is
unlikely, however, because the fraction of GR binding sites with a
consensus GR binding sequence did not differ significantly be-
tween the different classes (59%, 62%, and 57% of sites in the 0.5
FIG1 Dose-specific GR binding across the human genome. (A) Intensities of
ChIP-seq signals across all GR binding sites. Briefly, A549 cells were treated in
biological duplicate for 1 h with doses of DEX as indicated in the columns.
of DEX at which a binding site was called, and that class is indicated on the
right as hypersensitive (white), medium sensitive (orange), and low sensitive
(blue). The color within the ChIP-seq data indicates the ChIP-seq signal in-
tensity, expressed in units of aligned reads per kilobase of the binding site and
per million aligned ChIP-seq reads (RPKM). (B) The total number of binding
sites in each class, using the same color scheme as in panel A. (C) The consen-
sus GR binding motifs identified in each sensitivity class of GR binding sites.
Reddy et al.
mcb.asm.orgMolecular and Cellular Biology
test), nor did the number of GR binding sequences per binding
sought to identify additional features responsible for modulating
the corticosteroid sensitivity of GR binding sites across the ge-
Chromatin accessibility is associated with sensitivity of GR
binding. Sequences outside the core GR binding motif may in-
able open chromatin state or that may act as cofactors to stabilize
GR-DNA interactions. We reasoned that the regions of open
chromatin prior to treatment with glucocorticoid would be more
available to the GR, thus enabling GR binding at lower doses of
DEX. To test this hypothesis, we measured genomic accessibility
to DNase I in A549 cells in the absence of steroid, and we defined
open chromatin regions as those with significantly increased
DNase I accessibility. Overall, we found that open chromatin ac-
tivity, similar to observations in mouse cells (27). However, when
we evaluated each class of GR binding site individually, we found
a strong association between chromatin availability and binding
site sensitivity, with nearly all (144 of 145, or 99%) of the hyper-
sensitive GR binding sites occurring in already open chromatin
(Fig. 2A). Chromatin accessibility appeared important for the
I hypersensitivity signal in each GR binding site tracked closely
showing that open chromatin largely predetermines GR occu-
pancy suggested that cobinding transcription factors (TFs) are
important contributors to that sensitivity (27). For example, it
pioneer factor activity of the GR is limited to sites that are only
active at high concentrations of glucocorticoids (13, 67).
Transcription factors synergistic with the GR are prebound
to regions of increased GR sensitivity. Numerous additional
DNA binding proteins may also contribute to modulation of the
sensitivity of GR binding by acting synergistically with GR to sta-
bilize DNA interactions. To identify such factors, we searched for
DNA binding motifs that were enriched in hypersensitive GR
binding sites (39). We found motifs matching the forkhead box
(FOX), the AP-1 binding motif, and cyclic AMP response ele-
the FOX family, the FOXA1 and FOXA2 TFs, both expressed in
A549 cells, have been associated previously with modulating GR
well-documented factor known to enhance GR binding (1, 6, 17,
the CRE binding protein (CREB) (19, 25).
To investigate the role of the FOXA, AP-1, and CREB tran-
scription factors in sensitizing the glucocorticoid response, we
used ChIP-seq to measure genomic occupancy of FOXA1,
sites for each factor are listed in Table S5 of the supplemental
was highly correlated between biological replicates as well as be-
tween the related FOXA1 and FOXA2 factors, but less correlated
To examine the association between the binding of synergistic
transcription factors and the DEX sensitivity of GR binding, we
evaluated the fraction of GR binding sites in each sensitivity class
negative control, we also performed ChIP-seq for USF1, a factor
stronger enrichment for GR binding sites for each factor studied,
compared to USF1 (Fig. 2C). Occupancy of additional TFs was
associated with increased DEX sensitivity of the GR binding site,
indicating that these factors may contribute to genome-wide GR
recruitment. While FOXA1 and FOXA2 had the greatest overlap
with both hypersensitive and medium sensitive GR binding sites,
FIG 2 Overlap of hypersensitive GR binding with open chromatin and tran-
scriptional cofactors (A) Fraction of GR binding sites in each sensitivity class
binding site. The inset shows the mean ? standard deviation of each class of
GR binding site. (C) Fraction of GR binding sites in each sensitivity class that
with the GR, and dashed lines indicate the fraction of overlap with USF1 for
each class. (D) Receiver operating characteristic (ROC) curve, showing the
sensitivity and specificity of predicting hypersensitivity of a GR binding site
based on the occupancy of each cofactor. The ROC was calculated for the
versus predicting a medium sensitive GR binding site.
Low-Dose Glucocorticoids Regulate Period 1
September 2012 Volume 32 Number 18mcb.asm.org 3759
CREB1 occupancy was specific for the hypersensitive sites. We
evaluated if the factors were able to distinguish hypersensitive
from medium sensitive GR binding and found that, of the factors
evaluated, occupancy levels of CREB1 and JUND were the best
predictors of hypersensitive GR binding (Fig. 2D). Therefore,
may more moderately sensitize GR binding. Our work confirmed
that GR binding sites in preopen chromatin are enriched for oc-
ing of chromatin (i.e., the low-sensitivity GR binding sites in our
study) are more likely to act through GR binding alone (27). Spe-
cifically, the FOXA proteins are known pioneer factors that play a
role to open chromatin in advance of nuclear receptor occupancy
(40, 54), and these factors may assist in the occupancy of the GR
hypersensitive sites (13, 67).
PER1 expression is uniquely sensitive to DEX in A549 cells.
Having shown that the GR binds throughout the genome in re-
sponse to subsaturating DEX concentrations, we next sought to
determine the genome-wide effects of low doses of DEX on gene
expression. We treated A549 cells for 1 h with 0.1 nM, 0.5 nM, 1
sion response compared to a mock treatment. At the lowest dose
(0.1 nM DEX), we identified no genes with a significant gene ex-
see also Table S6 in the supplemental material). Of the genes re-
sponding to 5 nM DEX, expression levels of 23 (79%) were en-
hanced rather than repressed. For comparison, earlier genome-
wide screens showed that saturating (100 nM) DEX led to
increases in transcript levels of 59% of the responsive genes (50,
68). That low concentrations of DEX enhanced expression more
ing can also repress gene expression. Genes responding to low
doses of DEX play a role in diverse functions, including inflam-
mation (e.g., NFKBIA and TNFAIP3) and adipogenesis (e.g.,
CEBPB and CEBPD), but we observed no significant enrichment
for any gene ontology (GO) categories, likely due to the small
number of genes (FDR, ? 0.05 after Bonferroni correction [23,
Of all DEX-responsive genes in A549 cells, PER1 was the most
sensitive to low doses of DEX. We estimated 50% of the PER1
expression response (EC50) occurred at 0.47 nM DEX, a concen-
tration more than 6-fold lower than the mean EC50of 3.1 nM for
all other identified genes. PER1 also had the greatest change in
expression across the doses tested, with a 2.8-fold induction of
mRNA expression at 0.5 nM DEX and a 5.8-fold induction at 5
nM DEX, as well as a corresponding increase in PER1 protein
the next most sensitive gene, for angiopoietin-related protein 4
and 3.3-fold higher at 5 nM DEX. Notably, numerous genes that
sion to DEX is independent of the magnitude of expression re-
We found two GR binding sites near the PER1 transcription
start site (TSS), one located in the first intron and the second
sites contain a GR binding motif near the site of maximal ChIP-
seq signal. Both sites are conserved in the mouse genome, where
the syntenic binding sites are thought to regulate glucocorticoid-
mediated Per1 expression in peripheral mouse tissues (36) (see
Fig. S4 in the supplemental material). While both sites are bound
by the GR at 5 nM DEX, only the upstream site is bound with 0.5
for the hypersensitivity of PER1 expression to DEX.
A single GR binding site is sufficient for the hypersensitive
response is driven by DNA sequence alone, we cloned regions
of the nearby GR binding sites into a destabilized luciferase re-
porter vector. As a control, we also cloned the promoter and GR
binding site for the gene for stomatin (STOM), a gene that is
expressed only in response to high doses of DEX (Fig. 4A). After
transient transfection of each construct into A549 cells, the cells
nM. In all experiments, the reporter expression clearly reached
saturation by the 100 nM dose. Regions containing the hypersen-
site upstream of PER1 were substantially less responsive to DEX
moter (Fig. 4B). These results indicate that the DNA sequence of
the GR binding region upstream of PER1 is sufficient to drive
hypersensitive expression of PER1.
The specific GR binding sequence at the site upstream of the
by 3 nucleotides (1 nucleotide in the GR consensus sequence and
our earlier results, that the specific GR binding is not important
sequences while leaving the local DNA context intact and looked
for an effect on reporter sensitivity. Converting the upstream
binding sequence to the intronic sequence did not diminish the
hypersensitivity of the response (Fig. 4C). Therefore, consistent
FIG 3 Expression of PER1 is uniquely sensitive to low doses of glucocortico-
ids. (A) Dexamethasone dose-response curves for all 28 genes responsive to 5
nM DEX, as determined by RNA sequencing. Cells were treated for 1 h with
increasing doses of DEX (x axis), and gene expression relative to control-
treated cells was plotted (y axis). Of the responsive genes, PER1 (indicated in
blue) was particularly sensitive and showed the strongest response to ?5 nM
DEX. (B) Immunoblot of PER1 and actin (as a loading control) after 4-h
ChIP-seq, in the region surrounding the PER1 transcription start site. Absent
DEX, no occupancy was detected in the region (top line), whereas binding
upstream of PER1 occurred after addition of 0.5 nM DEX (second line), and
binding in the first intron of PER1 only occurred after addition of 5 nM DEX.
Reddy et al.
mcb.asm.orgMolecular and Cellular Biology
with the lack of a GR binding sequence specific to hypersensitive
GR binding sites, the particular version of the GR binding se-
quence is not responsible for the observed differences in the DEX
sensitivities between the two binding sites flanking the PER1 TSS.
Having confirmed that the specific sequence of the GR DNA
recognition motif did not alter sensitivity to DEX, it may instead
binding site relative to the TSS or from interaction with the en-
dogenous promoter. To test this hypothesis, we cloned DNA
flanking each of the PER1 GR binding sites in front of a minimal
promoter driving luciferase (Fig. 5A). We found that a minimal
274-bp region flanking the upstream GR binding site was suffi-
cient to drive expression, with an EC50of 0.25 nM DEX (Fig. 5B).
Cortisol produced a similar response, with an EC50of 21 nM for
first intron of PER1 (Fig. 5C). The result was important, as it
showed that the difference in sensitivity between the two sites is
not an artifact of DEX. Enhancer regions shorter than 274 bp
incrementally reduced the sensitivity of the reporter to DEX, and
the response of the shortest region tested (69 bp) was indistin-
MMTV promoter (Fig. 5D).
known as the Hill coefficient. A Hill coefficient of 1 indicates mo-
nomeric association of GR with DNA, while greater Hill coeffi-
cients imply the presence of an allosteric activator that increases
the affinity of corticosteroids to the GR (47). While the normally
responsive enhancers had a Hill coefficient close to 1, the hyper-
sensitive enhancers responded with a Hill coefficient near 2 (Fig.
5E), suggesting that proteins may interact with the GR away from
the DNA as part of effecting the more sensitive response. In this
case, it may be that the hypersensitive GR enhancer is both pre-
bound by cofactors that establish and maintain an open chroma-
tin state and also contains a composite recognition sequence spe-
cific to a particular GR-containing complex that has a greater
affinity to corticosteroids.
To probe for key nucleotides within the minimal necessary
hypersensitive enhancer and that are important for tuning the
sensitivity of GR binding, we used site-directed mutagenesis to
introduce 2-bp mutations proximal to the GR binding site (Fig.
6A). Mutation of one site located 13 nucleotides upstream of the
GR binding sequence had a mild effect on dose sensitivity. Inde-
pendently mutating a second site, located 4 nucleotides upstream
and the higher-order dose-response kinetics observed with the
wild-type enhancer (Fig. 6C and D), indicating the presence of a
nearby cofactor binding site essential for hypersensitivity. While
we observed binding of CREB and FOXA1 in the same enhancer
region, the introduced mutation appeared to disrupt a consensus
DNA binding motif for the NFI family of transcription factors.
Members of the NFI family have been shown to interact with the
GR, and it is not yet clear if that interaction may contribute to
sensitizing the PER1 response to low concentrations of glucocor-
ticoids (37). To test if the mutated sequences were sufficient to
sensitize a GR binding site, we introduced select sequences flank-
ing the GR binding sequence from the hypersensitive enhancer
FIG 4 The GR binding sites flanking the PER1 transcription start site are sufficient to drive gene expression outside the genome. (A) Diagram of the genomic
regions that were cloned into a promoterless luciferase reporter vector. Three regions were cloned from the PER1 locus. Two regions (“Full” and “Upstream,”
the intronic GR binding site. For both the full and intron reporter, luciferase was maintained in phase with the PER1 start codon; for the upstream reporter,
cloning included the endogenous promoter and stopped at the first nucleotide of the annotated transcription start site. As an additional control, a pair of GR
? SEM for 8 biological replicates, and lines indicates the fitted dose-response curve with variable Hill slopes.
Low-Dose Glucocorticoids Regulate Period 1
September 2012 Volume 32 Number 18mcb.asm.org 3761
However, we observed no increase in sensitivity in the hybrid en-
hancer (see Fig. S4 in the supplemental material). It is therefore
likely that within the minimal 274-bp enhancer identified, addi-
tional unidentified protein recognition sequences also contribute
to the enhancement of gene expression at low doses of corticoste-
To map key nucleotides throughout the hypersensitive en-
hancer region, we introduced mutations along the length of the
hypersensitive region (see the methods described in the supple-
mental material) and measured sensitivity to DEX (Fig. 6E). The
mutation scanning confirmed the effects of the 2-bp mutations
tested in Fig. 6A, as well as identifying numerous other regions
to hypersensitivity were found throughout the enhancer region,
additional recognition motif, such as the previously reported
GME (28). Instead, it is more likely that a combination of DNA
sequences contribute to the expression of PER1 at low concentra-
tions of glucocorticoids.
Hypersensitive PER1 expression is general to cell line and
conserved in the mouse. To determine if the sensitivity of PER1
were needed to drive reporter expression in ECC-1 cells, consis-
tent with expectations, given the ?20-fold-lower expression of
diminished glucocorticoid sensitivity in ECC-1 cells, the hyper-
sensitive enhancer element derived from the region upstream of
PER1 still responded to lower concentrations of DEX than the
in sensitivities of the upstream versus intronic enhancer elements
in ECC-1 cells was less than the 6-fold increase in sensitivity we
found in A549 cells and may indicate intermediate sensitivity of
FIG 5 Minimal enhancer region sufficient to drive a hypersensitive glucocorticoid response. (A) Regions surrounding the hypersensitive GR binding site
upstream of the PER1 transcription start site were cloned in front of luciferase with a minimal promoter. Also, regions surrounding the nonhypersensitive GR
to control-treated cells and then normalized to the maximal response observed. Error bars indicate standard errors of the means, and curves indicate the fitted
hypersensitive GR binding site upstream of PER1 shown in the same colors as in panel A. MMTV results show the dose-response curve of the MMTV promiter,
driving the same luciferase. (C) Dose-response curves of the PER1 enhancer regions in response to cortisol treatment. (D) EC50s, estimated by fitting dose-
Reddy et al.
mcb.asm.org Molecular and Cellular Biology
PER1 expression in ECC-1 cells. In addition, evidence of a coop-
erative response that we observed in A549 cells was not recapitu-
lated in ECC-1 cells (Fig. 7C). Therefore, we hypothesize that the
hypersensitive expression of PER1 results from the combination
of at least two mechanisms, one that increases sensitivity without
altering response kinetics and another that increases both sensi-
sion in A549 cells is conserved in the mouse genome, suggesting
that mouse Per1 expression may also be more sensitive to gluco-
mice, we used RNA sequencing to measure genome-wide gene
pituitary cell line AtT-20. Confirming our hypothesis, we found
that four genes were unusually sensitive to DEX in the mouse
pituitary cells, including Per1 (Fig. 7D). The other genes (Irs2,
homeostasis (20, 21), and clathrin-mediated endocytosis (51), al-
beit primarily in tissues other than mouse pituitary. Studies of
neuroendocrine function of the insulin response and Irs2 in par-
ticular have pointed to a possible role of insulin in regulating
gonadotropin levels that may be linked with infertility resulting
from diet-induced obesity (8, 10, 71). Many have noted that go-
nadotropin release varies diurnally during and after puberty, and
while primarily due to hypothalamic control of gonadotropin-
releasing hormone (42), corticosteroids may offer an additional
ing sites near PER1 are conserved in the mouse and likely explain
the observed sensitivity of Per1 expression (36) (see Fig. S4 in the
supplemental material). We did not find GR binding sites near
is either tissue or species specific. Together, these results show
both that that the regulation of PER1 expression by low concen-
trations of glucocorticoids is general to multiple cell types, con-
served in the mouse, and also part of a larger set of similar re-
sponses in other tissues.
Targeted expression of PER1 by low-dose glucocorticoids
at which we observed PER1 overexpression in A549 cells was sim-
suggests that triggering PER1 expression at low levels of cortisol
may have a specific impact on circadian rhythms in peripheral
tissues. To test if targeted induction of PER1 expression by corti-
costeroids affects circadian gene expression, we treated A549 cells
for 1 h with 0.5 nM DEX to induce expression of PER1. We then
washed the cells thoroughly with fresh medium and followed the
expression of a panel of additional circadian rhythm genes over
the following 12 h.
The transcriptional regulation of both the core and peripheral
circadian rhythms consists of a negative feedback loop involving
both regulatory and physical interactions between the Period
genes PER1 and PER2, the cryptochrome genes CRY1 and CRY2,
and the complex between CLOCK and ARNTL (also called
BMAL1). Over the 12 h following withdrawal of 0.5 nM DEX, we
observed significant changes in mRNA levels of all of these genes
except for CLOCK (Fig. 7E). The Period genes followed a similar
pattern of expression, with both elevated PER1 and PER2 expres-
sion over the 3 h immediately following DEX treatment, followed
DEX treatment. The cryptochrome genes exhibited significantly
increased expression for the first 6 h after DEX removal. For
ARNTL, expression was significantly decreased between 1 and 4 h
after treatment, reaching a minimum expression 3 h after treat-
ment. Expression of ARNTL then began to rise, and by 12 h, we
found significant overexpression compared to untreated cells. In-
terestingly, increased expression of CRY1 and CRY2 coincides
with repression of PER1, PER2, and ARNTL expression. A recent
FIG 6 Select nucleotides within the minimal hypersensitive enhancer are neces-
sary for the low-dose response. (A) Diagram of mutations introduced into the
hypersensitive GR response element via site-directed mutagenesis. The blue box
indicates the GR binding motif in the enhancer. WT indicates the wild-type se-
quence and, for both mutated sequences, the changed bases are colored and un-
derlined. (B) A549 cells were transiently transfected with mutated plasmids and
then treated for 4 h with increasing doses of DEX (x axis), and luciferase activity
response, whereas the mutated sequences (blue and orange) had less hypersensi-
tive and normally sensitive responses, respectively. Black indicates a 69-bp en-
bars indicating 95% confidence intervals. While the wild-type enhancer had sec-
ond-order kinetics, one mutant (blue) had intermediate kinetics, and the second
mutant (orange) had first-order kinetics that were indistinguishable from the
69-bp enhancer region. (E) Mutation scanning results for the enhancer region.
Each point represents the effect on sensitivity to DEX of replacing 10 bp of wild-
Low-Dose Glucocorticoids Regulate Period 1
September 2012 Volume 32 Number 18mcb.asm.org 3763
consistent with circadian rhythms (3). The regulation of other
circadian rhythm genes did not appear to result from long-term
retention of DEX in the cell culture medium, as PER1 remained
overexpressed after 4 h of continuous exposure to 500 pM DEX
(Fig. 7F). We also observed repression of CRY1, CRY2, and
ARNTL after continuous treatment with 500 pM DEX. The re-
pression is not likely to arise from direct regulation by GR, as our
previous has showed that high doses of DEX increased expression
of those genes (50), and it may instead result from continuous
To determine if transient induction of PER1 expression was
sufficient to establish oscillating patterns of gene expression, we
repeated the experiment over a 48-hour period. The results con-
firmed our initial findings for the first 12 h, but we found no
evidence for circadian oscillations in gene expression (see Fig. S6
in the supplemental material). These results showed that low
doses of glucocorticoids are sufficient to regulate expression of
circadian genes in a manner consistent with circadian rhythms,
lished circadian rhythms. A recent study in adrenalectomized ro-
dents supported our hypothesis, showing both that endogenous
daily cortisol administration restored rPer1 expression and en-
trained circadian rhythms in lung and kidney (58). Combined
with our study, these results suggest that GR-mediated responses
to low doses of corticosteroids are important for mammalian pe-
ripheral circadian physiology.
The glucocorticoid receptor is pivotal to the physiological re-
sponse to stress and peripheral circadian rhythms. The GR is
calculated as the average over four technical replicates, and expression was normalized to that of GAPDH and plotted relative to A549 cells prior to any
DEX treatment. Points indicate means, and error bars indicate standard deviations over the five biological replicates. (F) Expression of circadian rhythm
genes after 4 h of continuous exposure to 500 pM DEX. Columns indicate mean relative expression levels, and error bars indicate ranges observed across
3 biological replicates.
Reddy et al.
mcb.asm.org Molecular and Cellular Biology
provided by the bloodstream. Therefore, how the GR separates
stress and circadian effects remains an enigma. Here we have
shown that, in A549 cells, the GR can drive expression of many
glucocorticoid-responsive genes in a dose-dependent manner. Of
the genes responding to ?5 nM DEX, PER1 is uniquely sensitive
in A549 cells, exhibiting 2-fold overexpression at 0.5 nM DEX, a
dose at which no other genes have a significant response. To the
best of our knowledge, our study is the first to show that in some
tissue or cell types, low doses of glucocorticoids can directly reg-
upstream of PER1 is sufficient for a hypersensitive response to
additional GR binding sites nearby does not affect the response.
The region does not contain an instance of a previously reported
molecules through noncanonical binding motifs (60), that tune
the glucocorticoid response. Furthermore, we have found that
ever, the identity of those factors, and their specific roles in re-
modeling chromatin or otherwise modulating the glucocorticoid
responses, remains unclear.
eling is an integral component of GR activity (e.g., references 27
and 67). Pioneer factors, such as the FOXA family of TFs, actively
13, 40). The GR itself can also act to remodel chromatin at some
sites, and in doing so it appears to assist in the loading of addi-
tional factors into the same region (67). Our data revealed that
significantly more so than for medium sensitive GR binding sites
directs chromatin remodeling in response to DEX are uniformly
of sensitivity, suggesting that open chromatin only partially con-
tributes to tuning glucocorticoid responses and that genetic
mechanisms are also likely to be important. For example, muta-
number of DNA sequences outside the core DNA binding motif
contribute to the sensitivity of the corticosteroid response. These
sites may contribute both to the occupancy of pioneer factors as
of DEX would result in a low concentration of active GR in the
sensitive sites. Fully confirming the role of open chromatin will
require additional experiments to determine if disrupting the
chromatin state near hypersensitive GR binding sites affects the
sensitivity of those sites.
Given the potential importance of open chromatin for hyper-
sensitive GR binding, it is interesting that we see similar expres-
sion responses originating from plasmid reporters. Transiently
transfected plasmids are known to have incomplete nucleosome
structure and often to not faithfully model chromosomal DNA
structure (49, 62). It may be that the minimal enhancer element
on the plasmid, or that incomplete nucleosome structure on
transfected plasmids may be permissive to GR binding. Our ob-
served associations with open chromatin in the genome may
therefore reflect prebound TFs that help to recruit the GR rather
than a strict prerequisite for gene expression responses to low
glucocorticoid concentrations. Alternatively, while our motif
sibility that some GR binding motifs may limit remodeling and
influence sensitivity. Resolution of whether indeed plasmid chro-
matin structure matches that seen on the genome at the PER1
response elements may therefore provide further insights into if
sites and the extent to which that chromatin may tune glucocor-
ticoid responses. One possible approach is to determine how sta-
directly modify genomic GR binding regions, allowing study of
the genetics and epigenetics of glucocorticoid responses in the
native chromatin context (11, 37).
As described recently, monomeric GR interacting with DNA
response (47, 63). In that model, increased affinity of the GR for
DNA at hypersensitive sites would shift the dose-response curve
toward lower concentrations of DEX but would not cause an in-
crease in the steepness of the dose-response curve (i.e., the Hill
coefficient). The model further predicts, as we see in many of our
nonhypersensitive response curves, that a Hill coefficient of 1 is
characteristic of diffusion-mediated interactions of the GR with
DNA. In our results, however, the PER1 enhancer elements that
respond to a low concentration of DEX follow a steeper dose-
response curve in A549 cells. That change in the shape of the
dose-response curve suggests that cooperative binding of the GR
with other molecules also contributes to the sensitivity of the re-
sponse (47). It may be that a fraction of nuclear GR binds effector
to DNA and cooperative interactions with other molecules con-
tribute to the hypersensitive gene expression response, deletion
constructs that respond to higher DEX concentrations in A549
cells also lack the higher-order response. Meanwhile, in ECC-1
cells, the higher-order dose response is not evident despite in-
creased sensitivity, indicating that the two mechanisms may be
distinct and tissue specific. Together, the results point to a model
where a combination of chromatin state, occupancy of additional
transcription factors and cofactors, and cooperative interactions
genomic loci, including a locus responsible for the regulation of
ing site upstream of PER1 was 21 nM, below the normal range of
and is unavailable to activate the GR. Estimates of free cortisol
levels are as low as 5 nM in blood (3, 14, 57) and in tissues (57),
similar to the range that we expect would be required to dynami-
cally regulate PER1 expression through the day.
Low-Dose Glucocorticoids Regulate Period 1
September 2012 Volume 32 Number 18mcb.asm.org 3765
Cortisol released from the adrenal gland plays an important
role in the regulation of PER1 in some but not all mouse periph-
eral tissues and is an important messenger in peripheral circadian
rhythms (36, 57). Some peripheral clocks can be entrained inde-
of gene expression in tissue culture cells (3, 4) and that glucocor-
ticoids can also influence circadian rhythms in peripheral tissues
(3). Our work suggests that hypersensitive PER1 expression may
contribute to peripheral circadian timing by controlling the time
of day when PER1 is first expressed. Supporting our hypothesis, a
are essential for oscillation in rPer1 expression and that daily in-
jection of cortisol into the adrenalectomized rodents entrained
circadian rhythms (58). These results suggest that cortisol-medi-
ated regulation of PER1 is an important component of circadian
gene expression in peripheral tissues. Our time course study also
showed that triggering of PER1 by low-dose glucocorticoids is
sufficient to regulate expression of other circadian rhythm genes
many hours later. That regulation reveals coordinated expression
of PER1 and PER1, as well as of CRY1 and CRY2. In our study,
PER1/2 expression gave way to CRY1/2 gene expression, which
was followed by expression of ARNTL. While we did not observe
sion, the order of expression matched that of in vivo studies of
circadian gene expression (3, 58). Together, these studies suggest
of PER1 may help better dissect the regulatory network control-
ling circadian gene expression.
The ability to regulate circadian rhythms via a highly targeted
PER1 response may ultimately be pharmacologically useful. Ab-
nia (15), and metabolic syndrome (41). Correcting the circadian
expression of PER1 and to renormalize circadian rhythms and
related metabolic oscillations may provide a novel treatment op-
tion that is free of typical and serious side effects of high-dose
exogenous glucocorticoids (16). It may be that different cortisol
levels regulate different physiological functions. Testing that hy-
pothesis will require studies in diverse primary tissues to under-
nately regulate specific pathways or functions.
We thank Greg Barsh, Chris Gunter, and members of the Myers lab for
useful discussions and advice.
This work was funded by NHGRI ENCODE grant 5U54HG004576.
Support for T.E.R. was from NIH/NIAMS fellowship 5T32AR007450.
1. Almawi WY, Melemedjian OK. 2002. Negative regulation of nuclear
factor-?B activation and function by glucocorticoids. J. Mol. Endocrinol.
2. Bailey TL, et al. 2009. MEME Suite: tools for motif discovery and search-
ing. Nucleic Acids Res. 37:W202–W208.
3. Balsalobre A, et al. 2000. Resetting of circadian time in peripheral tissues
by glucocorticoid signaling. Science 289:2344–2347.
4. Balsalobre A, Damiola F, Schibler U. 1998. A serum shock induces
5. Belikov S, Astrand C, Wrange O. 2009. FoxA1 binding directs chromatin
structure and the functional response of a glucocorticoid receptor-
regulated promoter. Mol. Cell. Biol. 29:5413–5425.
6. Biddie SC, et al. 2011. Transcription factor AP1 potentiates chromatin
accessibility and glucocorticoid receptor binding. Mol. Cell 43:145–155.
7. Blind RD, Garabedian MJ. 2008. Differential recruitment of glucocorti-
coid receptor phospho-isoforms to glucocorticoid-induced genes. J. Ste-
roid Biochem. Mol. Biol. 109:150–157.
8. Brothers KJ, et al. 2010. Rescue of obesity-induced infertility in female
mice due to a pituitary-specific knockout of the insulin receptor. Cell.
9. Burioka N, et al. 2005. Dexamethasone influences human clock gene
expression in bronchial epithelium and peripheral blood mononuclear
cells in vitro. Chronobiol. Int. 22:585–590.
10. Burks DJ, et al. 2000. IRS-2 pathways integrate female reproduction and
energy homeostasis. Nature 407:377–382.
11. Cermak T, et al. 2011. Efficient design and assembly of custom TALEN
Res. 39:e82. doi:10.1093/nar/gkr218.
12. Christoffels VM, et al. 1998. Glucocorticoid receptor, C/EBP, HNF3, and
protein kinase A coordinately activate the glucocorticoid response unit of
13. Cirillo LA, et al. 2002. Opening of compacted chromatin by early devel-
opmental transcription factors HNF3 (FoxA) and GATA-4. Mol. Cell
14. Conway-Campbell BL, et al. 2010. Glucocorticoid ultradian rhythmicity
J. Neuroendocrinol. 22:1093–1100.
16. Debono M, et al. 2009. Modified-release hydrocortisone to provide cir-
cadian cortisol profiles. J. Clin. Endocrinol. Metab. 94:1548–1554.
17. De Bosscher K, Vanden Berghe W, Haegeman G. 2003. The interplay
between the glucocorticoid receptor and nuclear factor-?B or activator
protein-1: molecular mechanisms for gene repression. Endocr. Rev. 24:
manage depression. J. Psychopharmacol. 24:15–19.
19. Govindan MV. 2010. Recruitment of cAMP-response element-binding
protein and histone deacetylase has opposite effects on glucocorticoid
receptor gene transcription. J. Biol. Chem. 285:4489–4510.
20. Gray S, et al. 2007. Regulation of gluconeogenesis by Kruppel-like factor
15. Cell. Metab. 5:305–312.
21. Gum RJ, et al. 2003. Antisense protein tyrosine phosphatase 1B reverses
activation of p38 mitogen-activated protein kinase in liver of ob/ob mice.
Mol. Endocrinol. 17:1131–1143.
actions in glucocorticoid receptor-mediated induction and repression.
Mol. Cell. Biol. 27:1467–1485.
23. Huang da, W, Sherman BT, Lempicki RA. 2009. Bioinformatics enrich-
ment tools: paths toward the comprehensive functional analysis of large
gene lists. Nucleic Acids Res. 37:1–13.
24. Huang da, W, Sherman BT, Lempicki RA. 2009. Systematic and inte-
grative analysis of large gene lists using DAVID bioinformatics resources.
Nat. Protoc. 4:44–57.
25. Imai E, Miner JN, Mitchell JA, Yamamoto KR, Granner DK. 1993.
Glucocorticoid receptor-cAMP response element-binding protein inter-
action and the response of the phosphoenolpyruvate carboxykinase gene
to glucocorticoids. J. Biol. Chem. 268:5353–5356.
26. Jensen ST, Liu JS. 2004. BioOptimizer: a Bayesian scoring function ap-
proach to motif discovery. Bioinformatics 20:1557–1564.
27. John S, et al. 2011. Chromatin accessibility pre-determines glucocorti-
coid receptor binding patterns. Nat. Genet. 43:264–268.
28. Kaul S, Blackford JA, Jr, Cho S, Simons SS, Jr. 2002. Ubc9 is a novel
29. Kennaway DJ. 2010. Clock genes at the heart of depression. J. Psychop-
30. Kido Y, et al. 2000. Tissue-specific insulin resistance in mice with muta-
Reddy et al.
mcb.asm.orgMolecular and Cellular Biology
tions in the insulin receptor, IRS-1, and IRS-2. J. Clin. Invest. 105:199–
31. Koubovec D, Ronacher K, Stubsrud E, Louw A, Hapgood JP. 2005.
Synthetic progestins used in HRT have different glucocorticoid agonist
properties. Mol. Cell. Endocrinol. 242:23–32.
32. Koyanagi S, et al. 2006. Chronic treatment with prednisolone represses
the circadian oscillation of clock gene expression in mouse peripheral
tissues. Mol. Endocrinol. 20:573–583.
33. La Baer J, Yamamoto KR. 1994. Analysis of the DNA-binding affinity,
sequence specificity and context dependence of the glucocorticoid recep-
tor zinc finger region. J. Mol. Biol. 239:664–688.
34. Lamia KA, et al. 2011. Cryptochromes mediate rhythmic repression of
the glucocorticoid receptor. Nature 480:552–556.
35. Langmead B, Trapnell C, Pop M, Salzberg SL. 2009. Ultrafast and
memory-efficient alignment of short DNA sequences to the human ge-
nome. Genome Biol. 10:R25. doi:10.1186/gb-2009-10-3-r25.
36. Le Minh N, Damiola F, Tronche F, Schutz G, Schibler U. 2001. Glu-
cocorticoid hormones inhibit food-induced phase-shifting of peripheral
circadian oscillators. EMBO J. 20:7128–7136.
targeted gene knockout and gene replacement in eukaryotes. Nucleic Ac-
ids Res. 39:6315–6325.
38. Li X, Wong J, Tsai SY, Tsai MJ, O’Malley BW. 2003. Progesterone and
glucocorticoid receptors recruit distinct coactivator complexes and pro-
mote distinct patterns of local chromatin modification. Mol. Cell. Biol.
39. Liu X, Brutlag DL, Liu JS. 2001. BioProspector: discovering conserved
DNA motifs in upstream regulatory regions of co-expressed genes. Pac.
Symp. Biocomput. 2001:127–138.
40. Lupien M, et al. 2008. FoxA1 translates epigenetic signatures into enhancer-
41. Maury E, Ramsey KM, Bass J. 2010. Circadian rhythms and metabolic
syndrome: from experimental genetics to human disease. Circ. Res. 106:
hormone secretion across puberty in girls: potential mechanisms and rel-
evance to the pathogenesis of polycystic ovary syndrome. J. Neuroendo-
43. Meijsing SH, et al. 2009. DNA binding site sequence directs glucocorti-
coid receptor structure and activity. Science 324:407–410.
44. Mercier L, Thompson EB, Simons SS, Jr. 1983. Dissociation of steroid
binding to receptors and steroid induction of biological activity in a glu-
cocorticoid-responsive cell. Endocrinology 112:601–609.
45. Mortazavi A, Williams BA, McCue K, Schaeffer L, Wold B. 2008.
Mapping and quantifying mammalian transcriptomes by RNA-Seq. Nat.
46. Naray-Fejes-Toth A, Fejes-Toth G. 1997. 11-?-Hydroxysteroid dehydro-
genase-2 is a high affinity corticosterone-binding protein. Mol. Cell. En-
47. Ong KM, Blackford JA, Jr, Kagan BL, Simons SS, Jr, Chow CC. 2010. A
Proc. Natl. Acad. Sci. U. S. A. 107:7107–7112.
48. Onica T, et al. 2008. Dexamethasone-mediated up-regulation of human
CYP2A6 involves the glucocorticoid receptor and increased binding of
49. Oshima H, Simons SS, Jr. 1992. Modulation of transcription factor
50. Reddy TE, et al. 2009. Genomic determination of the glucocorticoid
response reveals unexpected mechanisms of gene regulation. Genome
51. Ritter B, Blondeau F, Denisov AY, Gehring K, McPherson PS. 2004.
through subcellular proteomics. Biochem. Soc. Trans. 32:769–773.
52. Rotsztejn WH, Dussaillant M, Nobou F, Rosselin G. 1981. Rapid glu-
cocorticoid inhibition of vasoactive intestinal peptide-induced cyclic
AMP accumulation and prolactin release in rat pituitary cells in culture.
Proc. Natl. Acad. Sci. U. S. A. 78:7584–7588.
53. Roux J, Pictet R, Grange T. 1995. Hepatocyte nuclear factor 3 determines
the amplitude of the glucocorticoid response of the rat tyrosine amino-
transferase gene. DNA Cell Biol. 14:385–396.
54. Sekiya T, Muthurajan UM, Luger K, Tulin AV, Zaret KS. 2009. Nucleo-
the pioneer transcription factor FoxA. Genes Dev. 23:804–809.
55. Simons SS, Jr, Oshima H, Szapary D. 1992. Higher levels of control:
56. So AY, Chaivorapol C, Bolton EC, Li H, Yamamoto KR. 2007. Deter-
57. Stavreva DA, et al. 2009. Ultradian hormone stimulation induces gluco-
corticoid receptor-mediated pulses of gene transcription. Nat. Cell Biol.
58. Sujino M, et al. 2012. Differential entrainment of peripheral clocks in the
rat by glucocorticoid and feeding. Endocrinology. 153:2277–2286.
59. Sun Y, Tao YG, Kagan BL, He Y, Simons SS, Jr. 2008. Modulation of
Mol. Cell Endocrinol. 295:59–69.
60. Surjit M, et al. 2011. Widespread negative response elements mediate
direct repression by agonist-liganded glucocorticoid receptor. Cell 145:
61. Szapary D, Huang Y, Simons SS, Jr. 1999. Opposing effects of corepres-
sor and coactivators in determining the dose-response curve of agonists,
and residual agonist activity of antagonists, for glucocorticoid receptor-
regulated gene expression. Mol. Endocrinol. 13:2108–2121.
62. Szapary D, Oshima H, Simons SS, Jr. 1992. Modulation of glucocorti-
coid induction of stably transfected tyrosine aminotransferase gene con-
structs involves elements up-stream of the glucocorticoid-responsive ele-
ment. Endocrinology 130:3492–3502.
63. Szapary D, Xu M, Simons SS, Jr. 1996. Induction properties of a tran-
siently transfected glucocorticoid-responsive gene vary with glucocorti-
coid receptor concentration. J. Biol. Chem. 271:30576–30582.
thyroid receptor superfamily members. Annu. Rev. Biochem. 63:451–
65. Valouev A, et al. 2008. Genome-wide analysis of transcription factor
binding sites based on ChIP-Seq data. Nat. Methods 5:829–834.
66. Vedder H, Weiss I, Holsboer F, Reul JM. 1993. Glucocorticoid and
mineralocorticoid receptors in rat neocortical and hippocampal brain
cells in culture: characterization and regulatory studies. Brain Res. 605:
67. Voss TC, et al. 2011. Dynamic exchange at regulatory elements during
chromatin remodeling underlies assisted loading mechanism. Cell 146:
68. Wang JC, et al. 2004. Chromatin immunoprecipitation (ChIP) scanning
identifies primary glucocorticoid receptor target genes. Proc. Natl. Acad.
Sci. U. S. A. 101:15603–15608.
69. Wang Q, Anzick S, Richter WF, Meltzer P, Simons SS, Jr. 2004.
Modulation of transcriptional sensitivity of mineralocorticoid and estro-
gen receptors. J. Steroid Biochem. Mol. Biol. 91:197–210.
70. Weitzman ED. 1976. Circadian rhythms and episodic hormone secretion
in man. Annu. Rev. Med. 27:225–243.
71. Withers DJ. 2001. Insulin receptor substrate proteins and neuroendo-
crine function. Biochem. Soc. Trans. 29:525–529.
72. Yamamoto T, et al. 2005. Acute physical stress elevates mouse Period1
mRNA expression in mouse peripheral tissues via a glucocorticoid-
responsive element. J. Biol. Chem. 280:42036–42043.
73. Zhang L, Rubins NE, Ahima RS, Greenbaum LE, Kaestner KH. 2005.
Foxa2 integrates the transcriptional response of the hepatocyte to fasting.
Cell. Metab. 2:141–148.
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