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Molecular Endocrinology 19(10):2610–2623
Copyright © 2005 by The Endocrine Society
doi: 10.1210/me.2005-0047
Activin Regulates Luteinizing Hormone ␤-Subunit
Gene Expression through Smad-Binding and
Homeobox Elements
Djurdjica Coss, Varykina G. Thackray, Chu-Xia Deng, and Pamela L. Mellon
Departments of Reproductive Medicine and Neuroscience (D.C., V.G.T., P.L.M.), Center for
Reproductive Science and Medicine, University of California, San Diego, La Jolla, California 920930674; and Genetics of Development and Disease Branch (C.-X.D.), National Institute of Diabetes and
Digestive and Kidney Diseases, National Institutes of Health, Bethesda, Maryland 20892
LH ␤-subunit (LH␤), which is essential for ovulation
and reproductive fitness, is synthesized specifically in pituitary gonadotropes. In this study, we
show that LH␤ gene expression is induced by activin in mouse primary pituitary cells if the cells are
treated within 24 h after dispersion in culture. Furthermore, male mice deficient in Smad3, and therefore in activin signaling, have lower expression of
both LH␤ and FSH␤ mRNAs compared with their
wild-type littermates. Using the L␤T2 immortalized
mouse gonadotrope cell line that endogenously
expresses LH, we identify specific elements in the
regulatory region of the rat LH␤ gene necessary for
its induction by activin. Activin responsiveness is
conferred by a promoter-proximal region located
ⴚ121/ⴚ86 from the transcriptional start site. Maximal LH␤ induction by activin requires a homeobox
element (HB) and a 5ⴕ-early growth response (Egr)
site found in this region of the promoter. Juxtaposed to the HB are three Smad-binding elements
(SBEs), which are essential for LH␤ induction. Interestingly, two of the SBEs are also critical for
basal expression of the LH␤ gene. We demonstrate
that Smad proteins are necessary and sufficient for
activin induction of the LH␤ gene. Furthermore,
Smad proteins can bind one of the identified SBEs.
In addition to binding this SBE, Smad proteins interact with pituitary homeobox 1 (Ptx-1) and orthodenticle homeobox 1 (Otx-1), which can bind the
HB located close to the Smad-binding site. Thus,
activin induction of LH␤ gene expression requires
a combination of several transcription factors,
both basal and activin induced, as well as cooperation between multiple DNA elements. (Molecular
Endocrinology 19: 2610–2623, 2005)
L
initially described as functioning in gonadal feedback
on gonadotropin synthesis. Activin increases release
of FSH (6) from the pituitary and induces FSH␤ expression in gonadotrope cells (7), whereas inhibin antagonizes activin action. Follistatin, a potent activinbinding protein (8), can inhibit the biosynthesis and
secretion of FSH (9). Interestingly, activin, follistatin,
and inhibin are expressed in the mature pituitary gonadotrope and can function in an autocrine manner
(10–12). Follistatin is also synthesized by folliculostellate cells in the pituitary and regulates activin availability in a paracrine manner (13, 14).
TGF␤ family members, including activin, activate
signaling molecules known as receptor-associated
Smads, which, in the case of activin, are Smad2
and/or Smad3 (15). Smad2 or Smad3 then associate
with a common Smad, Smad4 (DPC4). The activated
heteromeric Smad complex translocates into the nucleus, where it binds a Smad-binding element GTCTAGAC, or either half of this palindrome, within DNA to
regulate the expression of target genes (16).
As mentioned above, there is considerable evidence
that activin regulates FSH synthesis and secretion.
However, the initial reports regarding the role of activin
in gonadotropes failed to detect an effect of activin on
LH secretion (6, 17). In contrast, a number of subse-
H IS ESSENTIAL for steroidogenesis and reproductive function in both males and females, because a lack of this hormone leads to hypogonadism
and infertility in both sexes (1). LH synthesis is restricted exclusively to the anterior pituitary gonadotropes (2). It is a heterodimeric glycoprotein, composed of an ␣-subunit, in common with FSH and TSH,
and a unique ␤-subunit, which confers biological
specificity. Transcription of the ␤-subunit is the ratelimiting step for LH production (3, 4). LH ␤-subunit
(LH␤) gene transcription is induced by GnRH and repressed by gonadal steroids (5).
Activin and inhibin, members of the TGF␤ family of
growth factors, also play important roles in the modulation of gonadotropins. These glycoproteins were
First Published Online June 16, 2005
Abbreviations: BSA, Bovine serum albumin; Egr, early
growth response; GAPDH, glyceraldehyde-3-phosphate dehydrogenase; GFP, green fluorescent protein; GST, glutathione-S-transferase; HB, homeobox element; LH␤, LH ␤-subunit; NP-40, Nonidet P-40; Otx, orthodenticle homeobox;
Ptx, pituitary homeobox; SBE, Smad-binding element; SF-1,
steroidogenic factor 1.
Molecular Endocrinology is published monthly by The
Endocrine Society (http://www.endo-society.org), the
foremost professional society serving the endocrine
community.
2610
Coss et al. • Activin Regulation of LH␤
quent studies suggest that activin can influence LH
synthesis, both in vivo and in cell culture. Stouffer et al.
(18) have shown an acute and sustained increase in LH
secretion in response to activin in female rhesus monkeys. McLachlan et al. (19) reported that 2-d activin
infusion in adult male monkeys significantly increased
both LH and FSH release in response to GnRH injection. Attardi and Miklos (20) demonstrated that treatment of primary rat pituitary cell cultures with activin
results in significant increases in both LH secretion
and protein content in the cells, as well as LH␤ mRNA
levels. More recently, this observation was confirmed
in human fetal primary cell culture, where recombinant
human activin caused a significant increase in LH secretion into the medium. In addition, inhibin decreased
FSH and LH secretion, but the LH response to inhibin
was less prominent than that of FSH (21). Furthermore,
our laboratory has previously shown that an LH␤ reporter gene is induced by activin in transiently transfected L␤T2 cells (22), and, recently, this finding was
extended in a study in which the endogenous LH␤
gene was induced by activin in this mature gonadotrope cell model (23). Despite these reports, activin is
still generally regarded as a selective regulator of FSH.
In this report, we used primary mouse pituitary cells
enzymatically dispersed in culture to address this discrepancy in the literature by showing that the LH␤
gene can be induced by activin treatment. Moreover,
using Smad3-deficient mice, we confirmed a role for
activin in regulation of LH␤ expression in vivo, when
we found that these mice have lower expression of
LH␤ mRNA, in addition to lower levels of FSH␤, compared with their wild-type littermates. We then sought
to identify the molecular mechanism of activin induction of LH␤ gene expression using L␤T2 cells, the
mature gonadotrope cell model derived from a transgenic mouse pituitary tumor, which endogenously expresses LH␤. We found that in L␤T2 cells, LH␤ responds to activin in a time- and dose-dependent
manner, and the response maps between ⫺86 and
⫺121 bp from the transcriptional start site. This is an
active region of the promoter that contains a homeobox element (HB) at ⫺100 that is crucial for basal
and cell-specific expression of the LH␤ gene (24). In
close proximity, on either side of the HB, are tandem
steroidogenic factor 1 (SF-1) and early growth response (Egr) sites (25). The SF-1 sites are located at
⫺127 and ⫺59 and are outside the ⫺121/⫺86 region
where activin response maps. The Egr sites are located at ⫺112 and ⫺50 and convey GnRH responsiveness, upon induction of Egr proteins by GnRH (26,
27). GnRH regulation of the LH␤ gene occurs through
the combinatorial action of several transcription factors that bind the HB element, the SF-1 and Egr sites,
because they show synergistic interaction on the induction of the LH␤ gene (26–28). The 5⬘-Egr site is
found in the ⫺121/⫺86 region of the promoter where
activin response is located. We found that mutation of
either the HB or the 5⬘-Egr site dramatically decreases
LH␤ induction by activin, in addition to decreasing
Mol Endocrinol, October 2005, 19(10):2610–2623 2611
basal expression. In this region, we also identified
three Smad-binding elements (SBEs), and mutation of
any one of these sites completely abolished activin
response. Interestingly, two of these SBEs are also
important for basal expression of LH␤. Finally, we
demonstrate that overexpressed Smad proteins activate LH␤, bind a SBE in gel-shift assay, and interact
with orthodenticle homeobox (Otx) and pituitary homeobox (Ptx) proteins, which can bind the HB. Therefore, activin regulation of LH␤ gene expression, similar
to LH␤ regulation by GnRH, requires multiple, composite DNA elements and involves a combinatorial
action of several DNA binding proteins.
RESULTS
Activin Induces LH␤ in Primary Mouse Pituitary
Cell Culture
There is little doubt that activin is a major regulator of
FSH synthesis; however, examination of the effects of
activin on LH has resulted in conflicting reports. Some
studies have shown that activin regulates LH␤ induction
in animals and cell culture, both in rat primary pituitary
cells and a gonadotrope cell model derived from the
mouse; however, others did not detect this response. In
this study, we sought to provide a possible explanation
for these differing results regarding activin regulation of
LH synthesis as well as determine whether activin regulates LH␤ induction in mouse primary cells. Using mouse
cells instead of rat primary cells allowed us to directly
correlate our results with experiments in genetically
modified mice and more detailed mechanistic studies
using the mouse-derived gonadotrope cell line, L␤T2.
We enzymatically dispersed mouse pituitary cells and
treated them with activin or GnRH for 5 h, either 1 d or 3 d
after the dispersion. When cells were treated within 24 h
after dispersion, both GnRH and activin induced robust
expression of LH␤ compared with the vehicle-treated
cells (Fig. 1). Therefore, activin, as well as GnRH, induces
LH␤ gene expression. However, when cells were allowed to recover for 66 h before treatment with GnRH or
activin, GnRH treatment still caused a significant induction of LH␤, although to a lesser extent than at 24 h,
whereas activin induction failed to reach a significant
level. Because the level of LH␤ [normalized to glyceraldehyde-3-phosphate dehydrogenase (GAPDH)], did not
differ dramatically in the control cells between d 1 and d
3, the cells’ responsiveness to GnRH and activin seems
to have decreased from d 1 to d 3. Thus, it is possible
that the length of time in culture before treatment provides an explanation for the contradictory results obtained in other studies of LH␤ induction by activin.
Activin Induces LH␤ Gene Expression through
Proximal Regulatory Sequences
Activin is critical for gonadotrope cell function, and
LH␤ is crucial for both male and female reproduction
2612
Mol Endocrinol, October 2005, 19(10):2610–2623
Fig. 1. LH␤ Is Induced by Activin in Primary Mouse Pituitary
Cells
Primary pituitary cells were dispersed enzymatically from
8-wk-old male mice and plated in culture. They were treated
with vehicle (white bars), 30 nM GnRH (hatched bars), or 200
ng/ml activin (black bars) for 5 h, on the first day or on the
third day after dispersion. Total RNA was purified and reverse
transcribed, and the level of LH␤ expression was assayed by
real-time PCR. In each sample the amount of LH␤, calculated
from the standard curve, was compared with the amount of
GAPDH. Asterisk indicates significant difference from the
same day control as determined by one-way ANOVA and
Tukey’s post hoc test.
(1). Thus, it was important to determine the molecular
mechanisms by which activin induces LH␤ gene expression. The proximal 1800 bp of the rat LH␤ 5⬘regulatory region linked to a luciferase reporter gene
(LH␤-luc) was transiently transfected into L␤T2 cells.
This region of the LH␤ promoter is sufficient for gonadotrope-specific expression and for GnRH regulation
(24, 29). L␤T2 cells transiently transfected with LH␤luc were treated with various concentrations of activin
to determine whether this region of the LH␤ gene is
also sufficient for activin responsiveness. Treatment
with activin over a range of doses and at 3, 6, or 24 h
revealed that this regulatory region contains elements
that allow response to activin in a time- and dosedependent manner. Maximal induction is achieved after 6 h of treatment and the induction is maintained for
at least 24 h of activin treatment (Fig. 2A). There appears to be a decreasing trend at 24 h as compared
with the induction at 6 h of treatment; however, the
difference is not statistically significant.
Because L␤T2 cells secrete activin and express the
activin receptors, thus having a fully functional autocrine loop (22), we assessed the contribution of endogenous activin on LH␤ expression. We treated the
cells with follistatin for 24 h and measured the expression of LH␤-luc (Fig. 2B). Consistent with our previous
report, follistatin treatment does not diminish LH␤ levels in these cells (22). This is in contrast to FSH␤ in
which follistatin treatment decreases expression of
FSH␤ by about 50%. The concentration of follistatin
used in this experiment was sufficient to inhibit induction of either LH␤ (data not shown) or FSH␤ expres-
Coss et al. • Activin Regulation of LH␤
Fig. 2. The LH␤ Promoter Responds to Activin Treatment
A, The 1800-bp rat LH␤ promoter was linked to a luciferase
reporter gene (LH␤-luc) and transiently transfected into
gonadotrope-derived L␤T2 cells. The thymidine kinase ␤galactosidase reporter was cotransfected as an internal control. After overnight starvation in serum-free DMEM, the cells
were treated with 1 ng/ml activin (dashed line, solid triangles),
10 ng/ml activin (solid line, open squares), or 100 ng/ml
activin (dashed line, solid circles) for different lengths of time
(indicated on the abscissa). Luciferase activity in the lysates
was measured and normalized to ␤-galactosidase. Results
represent the mean ⫾ SEM of at least three independent
experiments, each performed in triplicate, and are presented
as fold induction from vehicle control. Statistical significance
was determined by one-way ANOVA followed by Tukey’s
post hoc test. B, L␤T2 cells were transiently transfected with
either ⫺1800 bp rat LH␤ or a ⫺398 bp mouse FSH␤ promoter
linked to a luciferase reporter and treated with vehicle (white
bars) or with 100 ng/ml of follistatin (gray bars) for 24 h, after
which the luciferase levels were measured and normalized to
the internal control. Results represent the mean ⫾ SEM of at
least three independent experiments, each performed in triplicate, and are presented as fold induction from vehicle control. *, Statistically significant difference from vehicle control,
as calculated by Student’s t test.
sion by exogenous activin (30). It is possible that the
response of LH␤ expression differs with endogenous
vs. exogenous activin. It is also possible that there is a
threshold level of activin needed to achieve induction
that is higher for LH␤ as compared with FSH␤ and that
this threshold is not achieved by endogenous activin
produced by L␤T2 cells under our conditions.
To identify which promoter elements convey activin
responsiveness, we mapped the regions of the LH␤
gene promoter that confer activin response using truncation/deletion analysis. L␤T2 cells were transiently
transfected with a series of truncations of the LH␤
gene 5⬘-flanking region, ranging in length from 1800
bp to 86 bp upstream of the transcription start site,
and the ability of activin to induce LH␤ gene transcrip-
Coss et al. • Activin Regulation of LH␤
tion was tested (Fig. 3). Samples treated with activin
were normalized to the vehicle-treated samples for
each truncation, and results are represented as fold
induction because the basal levels of expression of the
truncations vary (24). The response of LH␤ to activin
mapped between ⫺86 and ⫺121. The 121 most proximal bp of the rat LH␤ promoter linked to luciferase
was induced about 2-fold, as was the longer 1800 bp
of the promoter, whereas truncation to ⫺86 reduced
the induction to the level of the vector control.
Homeobox and Smad-Binding Elements Are
Critical for LH␤ Induction by Activin
Because the activin response maps directly to this
region of 35 bp, we then determined what promoter
elements in this region are necessary for this response. The proximal region of the LH␤ promoter contains a homeobox element, GATTA, with tandem SF-1
and Egr sites on either side. The Egr-1 sites are important for GnRH induction of LH␤ because Egr-1 is
induced by GnRH. SF-1 is expressed specifically in
the gonadotropes of the pituitary and is important for
expression of genes in these cells. The truncations of
86 and 121 bp from the transcription start site of LH␤
gene were originally created to narrow this region and
limit the number of transcription factor-binding sites.
Only the 5⬘-Egr site and the HB are located in this
small region, as illustrated in Fig. 4A with uppercase
letters. We created mutations in the Egr and the HB
sites and named them mEgr and mHB, respectively
(Fig. 4B). As has been shown previously, mutations in
these sites decreased basal expression of the LH␤
gene. In addition, they decreased the induction by
activin (Fig. 4C). The HB site mutation significantly
decreased the fold induction by activin by 57%,
Mol Endocrinol, October 2005, 19(10):2610–2623 2613
whereas the mutation of the 5⬘-Egr site decreased fold
induction by activin by 33%. Thus, the 5⬘-Egr site and
especially the HB site are involved in the activin regulation of the LH␤ gene. The homeodomain protein,
which binds the HB in the gonadotrope cell, was determined to be either an Otx-1-like protein the identity
of which is not known, or potentially a posttranslationally modified Otx-1. This protein is expressed at a high
basal level in L␤T2 cells (24). The involvement of the
HB site in activin regulation of LH␤ expression may,
therefore, involve this Otx-1-like protein. The involvement of the 5⬘-Egr in activin induction is not clear,
because Egr proteins are not expressed in the gonadotrope cell until they are induced by GnRH. Therefore, it was important to assess whether activin can
also induce Egr-1 (Fig. 4D). Egr-1 is the major Egr
family member induced by GnRH to bind the Egr site
in the LH␤ promoter. We performed a Western blot to
detect Egr-1 protein following GnRH (G) or activin (A)
treatment of L␤T2 cells. Because Egr-1 is an immediate early gene, induced very rapidly after GnRH treatment (25), we used the same duration of activin treatment to assess the activin affect on Egr-1 protein
levels in L␤T2 cells. We determined that activin, contrary to GnRH, did not induce Egr-1 protein under
these conditions.
To identify promoter elements that may further contribute to LH␤ induction by activin, we examined the
DNA sequence in this region more closely. As mentioned, activin signals primarily through Smad2 and/or
Smad3, which, together with Smad4, bind SBE,
GTCTAGAC, or its half-sites. Sequence analysis revealed that the ⫺121/⫺86 region contains three SBEs,
which may play roles in activin induction of LH␤ gene
expression (Fig. 5A). Starting from the 5⬘-end, we
Fig. 3. Activin Response Localizes to a 35-bp Region in the LH␤ Proximal Promoter
Different lengths of the LH␤ regulatory region were transiently transfected into L␤T2 cells and, after overnight starvation, the
cells were treated with 10 ng/ml activin for 6 h. The results are represented as fold induction from vehicle-treated cells for each
truncation. *, Significantly different induction in the treated cells vs. the control cells for each truncation, as determined by
Student’s t test. Results represent the mean ⫾ SEM of three independent experiments each performed in triplicate.
2614 Mol Endocrinol, October 2005, 19(10):2610–2623
Coss et al. • Activin Regulation of LH␤
termed them S1, S2, and S3 and mutated each of
these SBEs to investigate their roles in activin responsiveness (Fig. 5B). Each mutation abolished activin
responsiveness of the LH␤ gene. Surprisingly, two of
these SBEs, S2 and S3, also had a strong influence on
LH␤ basal expression. The basal expression was reduced by 50% by either mutation as compared with
the wild-type sequence.
Smad Proteins Are Essential for LH␤ Induction
by Activin
Fig. 4. The Homeobox and 5⬘-Egr Sites Are Required for
Maximal Induction of LH␤ by Activin
A, Rat LH␤ promoter sequence from ⫺130 to ⫺86 bp from
the start site of transcription is shown. The truncations between which the activin response maps are marked with
uppercase letters; however, the larger sequence was presented to identify the SF-1 site. The 5⬘-Egr and the homeobox element (HB) are indicated with an oval and a rectangle, respectively. B, Mutations in the homeobox element
(mHB) and Egr (mEgr) were introduced into the LH␤-luc vector, and transfections were performed in L␤T2 cells. Cells
were treated with vehicle (white bars) or with 10 ng/ml activin
(black bars) for 6 h, after which the luciferase activity was
measured and normalized to ␤-galactosidase. The results are
normalized to the luciferase/␤-galactosidase ratio in the cells
transfected with the wild-type reporter and treated with vehicle control. Statistical significance was determined by oneway ANOVA followed by Tukey’s post hoc test. *, Significant
induction by activin in the treated cells vs. the control cells for
each reporter; #, a significant difference in expression from
the wild-type reporter in control-treated cells. Results represent the mean ⫾ SEM of four independent experiments, each
To determine whether activin signals to the LH␤ promoter through its classical pathway by phosphorylation of the Alk4 receptor and activation of the Smadsignaling pathway, we overexpressed Smad7 in L␤T2
cells and, after activin treatment, assessed the induction of LH␤. Smad7 serves as a dominant-negative
Smad because it prevents phosphorylation of receptor
Smads and formation of an active Smad heterodimer
of Smad2/3 and Smad4 (31, 32). Complete inhibition
of activin signaling by overexpression of Smad7 would
indicate that activin uses the classical signaling pathway to induce LH␤ gene expression. Indeed, overexpression of Smad7 completely abolishes activin induction of the LH␤ gene, indicating that an active Smad
pathway is necessary for induction (Fig. 6A). Again,
consistent with our results with follistatin treatment,
Smad7 does not affect basal LH␤ levels, indicating
that endogenous activin does not play a role in the
regulation of LH␤ under these culture conditions.
To further assess the role of Smad proteins in induction of LH␤ gene expression, we cotransfected a
Smad3 expression vector with LH␤-luc and assessed
luciferase levels with and without activin treatment. A
recent report has shown that Smad2 cannot be overexpressed in L␤T2 cells and that Smad3 alone is sufficient for FSH␤ induction, presumably because
Smad4 levels are not the limiting factor for activin
effect in these cells (33). Similar to FSH␤ regulation,
LH␤ is induced by 3.2-fold with overexpression of
Smad3 (Fig. 6B). Mutation of any one of the three SBE
sites diminished LH␤-luciferase induction by overexpression of Smad3. Specifically, mutation of the SBE 1
or 2 sites completely abolished induction, not only by
performed in triplicate. C, The same results from panel B,
presented as percent change in fold induction of the mutated
reporters from the induction of the wild-type reporter. #,
Statistically significant change in fold induction by one-way
ANOVA and Tukey’s test. D, Western blot of L␤T2 whole-cell
extract after 2-h treatment with vehicle control (C), 10 nM
GnRH (G), or 10 ng/ml activin (A). Equal amounts of protein
were run on the gel and, after transfer, membranes were
probed with antibodies specific for Egr-1 protein. After the
secondary antibody, enhanced chemiluminescence was performed and the blots were exposed to film. The experiment
was repeated three times with the same result and a representative gel was shown. WT, wild type.
Coss et al. • Activin Regulation of LH␤
Fig. 5. The SBE Sites Are Crucial to the LH␤ Response to
Activin
A, Rat LH␤ promoter sequence from ⫺121 to ⫺86 bp from
the start site of transcription is shown. In addition to the
indicated homeobox and Egr sites, SBEs (S1, S2 and S3
numbering from 5⬘ to 3⬘) are circled for clear observation. B,
Point mutations (2 bp) in each of the SBEs were separately
introduced into the reporter (mS1, mS2, and mS3), and their
induction by activin treatment was compared with the wildtype reporter. *, Significantly different fold induction in the
treated cells vs. the control cells for each reporter; #, a
significant difference in the level of luciferase expression from
the control treated cells transfected with wild-type reporter.
activin, as we have shown in the previous figure, but
also by Smad3 overexpression. The SBE 3 site mutation considerably reduced the level of induction. Promoter induction was reduced from 3.2-fold for the wild
type to 1.65-fold. None of the mutant promoters
showed an additional increase with treatment with
activin, which is similar to what was observed in Fig.
5B. This result indicates that all three sites play roles in
induction by activin and by Smad3 protein. As in previous reports studying FSH␤ induction (33, 34), overexpression of Smad3 alone is sufficient to reproduce
the effect of activin on LH␤ induction.
Smad Proteins Bind an SBE in the LH␤ Promoter
and Can Interact with Ptx-1 and Otx-1 Proteins
To assess whether Smad proteins can bind the SBE
sequences in the ⫺121/⫺86 region of the promoter,
we used EMSA. The whole ⫺121/⫺86 region was
used as a probe (Fig. 7A) with nuclear extracts from
Cos 1 cells transfected with the expression vector for
Smad4 or empty vector control. Smad proteins have
relatively low affinity in binding DNA, and overexpression is one way to overcome this limitation. Comparing
Mol Endocrinol, October 2005, 19(10):2610–2623 2615
Fig. 6. Smad Proteins Are Necessary and Sufficient for LH␤
Induction by Activin
A, Cells transfected with wild-type LH␤-luc were treated
for 6 h with vehicle (white bars) or 10 ng/ml activin (black
bars). Cells were cotransfected with an expression vector for
Smad7 or its empty vector control, to assess the necessity for
Smad protein activation in the induction by activin. Results
represent the means of three independent experiments each
performed in triplicate. *, Statistically significant difference
from the control cells transfected with the wild-type vector. B,
Cells transfected with wild-type LH␤-luc, or each SBE mutation introduced into LH␤-luc, were cotransfected with
Smad3 or empty vector control (control). Activin was added
to the cells 24 hr after transfection (filled bars are activin
treated and white bars are control) and after 6 h of treatment,
luciferase activity was measured and normalized to ␤-galactosidase. Results represent the means of three independent
experiments each performed in triplicate. *, Statistically significant induction by activin; #, statistically significant induction by Smad3 overexpression as analyzed by ANOVA and
Tukey’s post hoc test. WT, Wild type.
complexes that bind this region from cells transfected
with empty vector control (first lane) with cells transfected with the Smad4 expression vector (second
lane), we detected two strong bands not present in the
control lane (marked with arrows) (Fig. 7B). The same
pattern was observed when nuclear extract from Cos
1 cells transfected with both Smad3 and Smad4 was
used (data not shown). These bands are supershifted
(labeled “ss”) with antibodies to Smad4 protein (third
and last lane), indicating that Smad proteins are indeed a part of this complex. Thus, Smad proteins can
bind this region in vitro. To determine which nucleotides are needed for Smad protein binding, competi-
2616 Mol Endocrinol, October 2005, 19(10):2610–2623
Fig. 7. Smad Proteins Bind a SBE at ⫺116 in the LH␤ Promoter
A, An alignment of wild-type sequence (WT) from the rat
LH␤ promoter ⫺121/⫺86, and the oligonucleotides used as
competitors, labeled A–K, is shown. Scanning mutations
were introduced into oligonucleotides A–K, and these
changes are underlined. These unlabeled oligonucleotides
were used in 200-fold excess in EMSA experiments in panel
B, whereas wild-type sequence (WT) was used as a probe.
The SBE sites are indicated with a circle in the wild-type
sequence, and the HB site is indicated with a square. B, Nuclear
extracts from transfected Cos 1 cells were subjected to EMSA
with radiolabeled wild-type probe. Extracts from cells transfected with the control plasmid were used in the lane labeled C,
whereas extracts from Cos 1 cells transfected with Smad4
expression vector were used in the remaining lanes. Complexes
that change in Smad4-containing lysates are indicated with
arrows. Mutated oligonucleotides were used as competitors in
200-fold excess in the corresponding lanes. In the lanes labeled
1 and 2, the antibody to Smad4 was included to induce the
supershift, marked with “ss.” The antibody in the lane labeled 1
was obtained from Upstate USA, whereas the antibody in the
lane labeled 2 was purchased from Santa Cruz Biotechnology.
NE, Nuclear extract; ab, antibody.
Coss et al. • Activin Regulation of LH␤
tions with 200-fold excess of unlabeled mutant oligonucleotides were used. Scanning mutations (3 bp)
were introduced into the ⫺121/⫺86 oligonucleotide
and were termed mutations A–K (Fig. 7A). Oligonucleotides with mutations in the second SBE (S2) element
(Fig. 7, B and C) could not bind Smads and did not
compete with the labeled wild-type probe. Therefore,
SBE 2 at ⫺116 from the start site of the transcription
is the site that can be bound by Smad proteins with
high enough affinity to be detected in EMSA.
In the next EMSA, we used nuclear extracts from
L␤T2 cells treated with vehicle control or activin, to
assess whether activin treatment causes changes in
complexes that bind this promoter region. Activin
treatment for 30 min, 2 h, or 6 h increased the intensity
of a complex indicated with an asterisk in Fig. 8A. The
same complex is supershifted with antibodies to
Smad4 (indicated with ”ss“ in the figure), confirming
the identity of the band as a Smad4-containing complex. Next, we introduced mutations into the wild-type
sequence to assess the ability of each of the SBEs to
bind Smad proteins. Subsequently, these mutants
were used as probes in comparison with the wild-type
sequence. Smad4 antibody was included with nuclear
extracts to allow clearer observation of the Smad
complex due to the supershift caused by Smad4 antibody (Fig. 8B). A supershift was observed when the
wild-type sequence was used as a probe, as demonstrated in Fig. 8A. When the mutated sequence that
contains only SBE 2 was used as a probe (2), a supershift with Smad4 antibody was retained, although
SBE 1 and SBE 3 were not present. On the other hand,
the probes containing only SBE 1 (1) or SBE 3 (3) sites,
but lacking SBE 2, did not bind Smad4 in this assay.
Thus, again, as with Cos-1 extract, the SBE 2 site is
critical for Smad protein binding of LH␤ promoter. The
SBE 1 and 3 may contribute to the affinity of binding
because the intensity of the complex binding probe 2
was lower than that with the wild-type probe. As expected, when all three SBE sites were mutated (Fig.
8A), Smad4 antibody did not cause a supershift, indicating that Smad proteins cannot be tethered to the
promoter solely through interaction with other promoter-binding proteins. Furthermore, when we used a mutant probe that contains only the three SBE sites, with
the Egr and HB sites mutated (Fig. 8B), Smad4 antibody caused a supershift, indicating that Smad proteins do not need the Egr or HB sites to bind the LH␤
promoter.
Herein, we have determined in transfection assays
that all of the three SBEs found in the ⫺121/⫺86
region of the LH␤ promoter are required for activin
responsiveness and that the HB element at ⫺100 is
necessary for maximal induction of LH␤ by activin.
We also determined by EMSA that Smad proteins
can bind one of these SBEs found in this region,
whereas previously, it has been shown that Ptx-1 or
Otx-1 proteins can bind the HB element (24, 35).
Further, it has also been shown that proteins that
bind this region of the promoter can interact syner-
Coss et al. • Activin Regulation of LH␤
Mol Endocrinol, October 2005, 19(10):2610–2623 2617
Fig. 8. Smad Proteins from Activin-Treated L␤T2 Cells Bind the LH␤ Promoter
A, EMSA was performed with nuclear extracts from activin-treated L␤T2 cells for the times indicated above each lane, using
the ⫺121/⫺86 region of LH␤ promoter as a probe. In the IgG lane, nonspecific antibody was included, whereas in the S4 lane,
antibody to Smad4 from Santa Cruz Biotechnology was included. “ss” indicates a supershift, whereas * marks a complex that
changed after activin treatment. B, Wild-type and mutant probes used in EMSA are illustrated. In the bottom panel, each lane
represents a corresponding probe used to assess binding in EMSA with activin-treated nuclear extract and Smad4 antibodies.
Ab, Antibody; WT, wild type.
gistically (27). Consequently, it was important to
assess whether Smad proteins can interact with Ptx
or Otx proteins. We used a glutathione-S-transferase (GST) pull-down assay with in vitro transcribed and translated Ptx-1 and Otx-1 proteins to
test whether they can interact with Smad2-, 3-, or
4-GST fusion proteins. GST pull-down assays indicate that both Ptx-1 and Otx-1 interact with the
Smad2, 3, and 4 proteins (Fig. 9). 35S-labeled Ptx-1
and Otx-1 precipitate with glutathione beads
through an interaction with GST-Smad2, 3, or 4. No
interactions were observed using GST alone, nor did
labeled green fluorescent protein (GFP), which
serves as a control, interact with any of the GSTfusion proteins. Thus, Ptx-1 and Otx-1 can form
heteromeric complexes with Smad proteins in vitro,
through protein-protein interaction.
Activin Is Critical for LH␤ Expression in Vivo
To validate our results obtained using the L␤T2 cells
and assess the role of activin in LH␤ gene expression
in vivo, we determined the level of LH␤ and FSH␤ gene
expression in Smad3-deficient mice. These mice lack
exon 8 of the Smad3 gene, which prevents synthesis
of the functional Smad3 protein (36). Females are infertile due to an ovarian phenotype (37) and, therefore,
pituitary effects are impossible to assess. For that
reason, we used only male animals in our studies at 8
wk of age. To determine the levels of LH␤ and FSH␤
mRNA in the pituitary, quantitative PCR was used, and
values were normalized to the levels of GAPDH mRNA
expression for each animal (Fig. 10). We determined
that both LH␤ and FSH␤ are expressed at a lower level
in Smad3-deficient animals than in their wild-type lit-
2618 Mol Endocrinol, October 2005, 19(10):2610–2623
Fig. 9. Smads2, 3, and 4 Can Interact Directly with Ptx-1
and Otx-1
35
S-labeled proteins, indicated on the left of the panels,
were used in the binding assay with GST-fusion proteins,
labeled above each lane. GST-fusion proteins were induced
with isopropyl-␤-D-thiogalactosidase overnight, and the bacterial pellets were sonicated. These proteins were bound to
Glutathione Sepharose beads and incubated with in vitro
transcribed and translated labeled proteins. After extensive
washing, the precipitates were run on a gel and subjected to
autoradiography. The experiment was repeated three times
with the same results and a representative experiment is
shown.
termates. LH␤ was decreased by 38%, whereas FSH␤
was reduced by 32%, in comparison with gonadotropin mRNA levels in wild-type mice. Thus, activin plays
a role in both LH␤ and FSH␤ expression in vivo.
Coss et al. • Activin Regulation of LH␤
Fig. 10. Smad3-Deficient Male Mice Have Lower Levels of
LH␤ and FSH␤ mRNA
Total RNA from individual pituitary was reverse transcribed
and LH␤, FSH␤, and GAPDH cDNA were quantified using
real-time PCR. For each animal, the amount of LH␤ or FSH␤
mRNA was calculated from a standard curve performed simultaneously. The amount was compared with the amount of
GAPDH also calculated from its standard curve. n ⫽ 9 animals per group; the level of LH␤/GAPDH ratio in ⫹/⫹ animals
was set to 1, to allow comparison of the change from wild
type. #, Significant difference from the wild-type littermates
by Student’s t test: P ⫽ 0.0007 for LH␤ and P ⫽ 0.008 for
FSH␤.
DISCUSSION
Recently, a number of studies have shown that activin
regulates both LH synthesis and secretion in vivo and
in primary pituitary cell culture, both in primates and in
rodents. However, due to initial reports that did not
detect increased LH synthesis after activin treatment,
activin is still generally regarded as a selective regulator of FSH. In gonadotrope-derived L␤T2 cells, both
endogenous LH␤ gene and a reporter gene under LH␤
promoter regulation were significantly induced by activin. Herein, we show that activin induces LH␤ mRNA,
if the primary pituitary culture is treated within 24 h
after enzymatic cell dispersion. Several possibilities
can be proposed to explain the differences in these
various studies. First, it is possible that higher activin
doses are necessary for stimulation of LH synthesis as
opposed to FSH, and the initial studies did not detect
the smaller, but significant, changes in LH due to the
lower activin concentrations used. In our studies, activin consistently induced LH␤ gene in primary cells
with the higher concentration (200 ng/ml), as presented in Fig. 1, whereas lower concentrations of activin (20 ng/ml) significantly increased LH␤ gene expression in about 50% of experiments. Second, the
difference may be due to the physiological state of the
animals from which the pituitaries were extracted or to
the stress level the animals experienced during transport or procedure. For example, the animals used
could have been from a stage of the estrous cycle
when ovarian follistatin is at its highest. On the other
Coss et al. • Activin Regulation of LH␤
hand, circulating activin may have been at a different
level. As demonstrated in Fig. 2B, autocrine, endogenous activin does not seem to play a role in LH␤
expression, whereas it contributes significantly to
FSH␤ expression. Circulating activin, however, may
have a different effect. It is also credible that different
levels of steroid hormones due to cycling of the female
animals may have contributed to the differential regulation of FSH and LH by activin. FSH synthesis has
been shown to be stimulated by progesterone or testosterone, which also synergize with activin to induce
FSH, whereas LH can be suppressed by progesterone
or testosterone in L␤T2 cells (Thackray, V. G., S. M.
McGillivray, and P. L. Mellon, manuscript submitted).
To eliminate fluctuation of hormones due to the estrous cycle, we used male animals for our studies.
Third, the studies that failed to detect activin induction
of LH␤ may have used less sensitive techniques to
analyze changes in the hormone or gene expression
levels. Lastly, the studies that did not observe activin
regulation of LH may have had a higher level of folliculostellate cells in their primary pituitary cultures due
to having been in culture longer before the experiment.
Traditionally, primary cells were left to recover for 48 h
after the pituitary dispersion and then were treated
with the stimulus of interest, because that was the
optimal time for studying secretory response to the
releasing hormones. Some studies in the literature still
follow this protocol, despite more recent studies with
primary pituitary cultures that have reported that secretory cells are rapidly overgrown by folliculostellate
cells, which secrete follistatin, a potent activin inhibitor
(14). Our results appear to correlate with the rationale
of overgrowth of folliculostellate cells, because the
LH␤ induction by activin was more robust when the
treatment was conducted 1 d after dispersion. When
the cells were treated on the third day after dispersion
in culture, induction of LH␤ was far less prominent,
perhaps due to high levels of follistatin secreted by the
predominant folliculostellate cells.
Smad3-deficient mice have lower levels of both LH␤
and FSH␤ mRNA when compared with their wild-type
littermates, confirming a role for activin in gonadotropin regulation in vivo. Female animals have a severe
ovarian phenotype, and any effect on the gonadotropes may be due to the lack of gonadal feedback. To
circumvent these problems, we have used only male
animals. Although fertile, Smad3-deficient males are
considerably smaller and we determined that they also
have lower gonadotropin levels than their littermates.
Because of the importance of activin in appropriate
gonadotropin gene expression and the necessity for
LH␤ in both male and female reproductive function,
we proceeded to identify the molecular mechanism of
activin regulation of LH␤ gene expression.
Activin response maps to a very active region of the
promoter that contains elements for basal, cell-specific, and GnRH regulation of the LH␤ gene. Mutation
of the HB site, GATTA, in this region, dramatically
reduces activin responsiveness. The HB is critical for
Mol Endocrinol, October 2005, 19(10):2610–2623 2619
basal regulation, and it is found to bind the Ptx proteins overexpressed in CV-1 cells (27) or an Otx-1related protein, expressed endogenously in the L␤T2
cells (24). Therefore, this site is also involved in cell
specificity of LH␤ expression, because L␤T2 cells,
which represent the most mature gonadotrope cells
and endogenously express LH␤, are the only gonadotrope-derived cell type that expresses the protein that
binds this site. A less mature gonadotrope cell model,
such as the ␣T3–1 cell, which does not express LH␤,
does not express this protein. An SF-1 site is found in
close proximity to the HB, and it may also contribute to
cell-specific expression, because SF-1 is limited to the
gonadotrope population within the pituitary. Ptx proteins were found to synergize with SF-1 to induce LH␤
gene expression in CV-1 cells, which do not express
endogenous SF-1 (28). L␤T2 cells express high levels
of endogenous SF-1, and thus it is difficult to observe
this regulation in these cells. Regardless, the SF-1 site
was not present in the ⫺121/⫺86 region of the promoter that remains activin responsive, and therefore
we concentrated on the HB and Egr sites. Here we find
that both HB and the 5⬘-Egr site have roles in activin
regulation of LH␤ gene expression, because mutation
of these reduces the magnitude of activin induction.
GnRH regulation of LH␤ occurs through the induction of Egr proteins, which bind their two sites also
found in this region of the promoter. Interestingly,
there is a functional difference between the 5⬘- and
3⬘-Egr sites. Mutation of the 3⬘-Egr site diminishes
GnRH induction, whereas the 5⬘-site does not seem to
play a role in GnRH regulation of LH␤ gene (38). Also,
in vitro binding to these sites by EMSA showed that
Egr-1 binds with high affinity to the 3⬘-Egr site,
whereas the intensity of binding to the 5⬘-site is orders
of magnitude lower (25, 39). This raises the question
whether the 5⬘-Egr site is bound in vivo by a different
transcription factor. Because the 5⬘-Egr site does not
seem to bind Egr proteins with high affinity nor is it
involved in GnRH induction of LH␤, this site may bind
a different member of the Egr/SP-1 superfamily. It is
interesting to speculate that this protein may be induced by activin and may play a role in activin regulation of LH␤. On the other hand, it has been demonstrated that Ptx can also synergize with Egr to induce
LH␤ (27). Interestingly, mutation of the Ptx element
has a more profound effect, and it is possible that the
Egr site contributes to the activin induction only because of its synergistic interaction with Ptx or Otx.
Mutation of any of the three SBE sites completely
abolishes induction by activin and dramatically diminishes induction by Smad3 cotransfection. This requirement for multiple sites has been found in other TGF␤/
activin-responsive promoters (40). Multiple SBEs in
close proximity to each other allow for cooperative
binding of Smad proteins and allow them to overcome
their low binding affinity. Furthermore, the requirements for other promoter elements for full activin response have also been reported (41). It is likely that a
similar situation occurs with the LH␤ promoter, in
2620
Mol Endocrinol, October 2005, 19(10):2610–2623
which the Ptx/Otx-binding homeobox site is required
for full activin response. Interaction of Smad proteins
with other transcription factors has been reported (42).
Most notably, Smad2 is known to interact with the
homeobox proteins Milk and Mixer (43). Herein we
detect that Smads 2, 3, and 4 can interact with homeobox proteins Ptx-1 and Otx-1. It is likely that,
because Smads can interact with Ptx-1 in addition to
Otx-1, they can also interact with a related family
member expressed endogenously in L␤T2 cell, the
Otx-1-like protein, the identity of which is not known.
Possibly, that interaction with other proteins stabilizes
Smad interaction with DNA and, in the case of LH␤,
may play a role in activin induction. Additionally, SBE
site 2, which has a dramatic effect both on basal and
activin-induced gene expression, and is bound by
Smad4 in EMSA, is juxtaposed to the 5⬘-Egr site. Thus,
it is possible that Smad proteins are interacting with
this putative Egr family member to regulate both basal
and activin-regulated LH␤ gene expression.
We determined that Smad proteins bind the DNA in
the ⫺121/⫺86 region of the promoter using EMSA
both from Cos-1 cells with overexpressed Smad proteins and from activin-treated L␤T2 cells. Specifically,
we established that the middle SBE is needed for this
binding. This SBE is located 5⬘ to the HB and to the
Egr site. It is possible that the binding affinity for the
other SBEs is too low to detect in a gel shift, because
our transient transfection experiments indicate that all
of the SBEs are required for activin response. SBE 1
and 3 may contribute to the stability of Smad protein
binding, because, when only SBE 2 is present, the
intensity of binding is lower than when all three sites
are present.
Surprisingly, two of the three SBEs identified are
critical for LH␤ basal expression as well. These two
SBEs are located around the HB, which has been
shown to be crucial for the basal expression of LH␤
gene. However, these SBE sites are sufficiently separate such that the sequences do not overlap the HB,
and neither site is needed for binding of the homeodomain protein (24). Because neither follistatin nor overexpression of Smad7 can reduce the basal expression
of LH␤, it appears that endogenous activin does not
play a role in supporting basal expression under these
culture conditions, and therefore what we are measuring are changes in “true” basal expression due to
these cis mutations. It would be interesting to determine whether Smad proteins have a role in basal expression and the manner in which they accomplish it.
Smad4 has been shown to shuttle in and out of the
nucleus independent of activin treatment (44). Given
that Smad4 can bind the SBE 2 that is critical for basal
regulation, whereas Ptx or Otx bind the homeobox
element also involved in basal expression, Smad4 interaction with Ptx and Otx may contribute to basal
expression of LH␤ gene though interaction with these
homeodomain proteins. Alternatively, whether some
other proteins, which are involved in both basal and
Coss et al. • Activin Regulation of LH␤
activin induction, can bind these sites remains to be
determined.
In this report, we determined that activin plays a role
in LH␤ expression in vivo and induces LH␤ mRNA in
primary mouse pituitary cells before overgrowth of
secretory cells by folliculostellate cells, which may
diminish bioavailability of activin by secreting follistatin. Activin acts through three SBE sites, the mutation
of which abolishes activin induction, and, of which,
two are critical for basal expression of the LH␤ gene as
well. Interestingly, the region of the LH␤ promoter
where these SBE sites are found is also critical for
induction by GnRH and for basal expression. Transcription factors that bind elements in this region of
the promoter have been shown to physically interact,
forming a complex. Because Smads bind and induce
LH␤ and they can interact with homeodomain proteins, this suggests that Smad proteins are a part of a
large, transcriptionally active complex that is critical
for the regulation of the LH␤ gene transcription.
MATERIALS AND METHODS
Primary Mouse Pituitary Cell Culture and Treatment
Male C57 Black 6 mice (6 wk of age) were purchased from
Harlan Sprague Dawley (Indianapolis, IN), and housed for 2
wk in UCSD animal facility under standard conditions. At 8 wk
of age, mice were killed by decapitation and pituitaries were
collected in ice-cold Dulbecco’s A PBS (PBS). After a PBS
rinse, pituitaries were placed in Hank’s balanced salt solution
with 25 mM HEPES, pH 7.2, containing 0.25% collagenase
and 0.25% trypsin. After cell dispersion for 30 min at 37 C in
a shaking waterbath, the same volume of DMEM with 10%
fetal bovine serum was added to stop the reaction and then
deoxyribonuclease was added to a final concentration of 20
␮g/ml and incubated for another 15 min at 37 C. Subsequent
to removal of tissue debris, cells were pelleted by centrifugation and plated at 106 cells/2-cm2 well density. The medium was changed to serum-free DMEM with 0.1% bovine
serum albumin (BSA) 16 h before treatment. Cells were
treated with GnRH (Sigma Chemical Co., St. Louis, MO) or
activin (Calbiochem, La Jolla, CA) for 5 h, after which the cells
were lysed to obtain total RNA.
Smad 3-deficient mice and their littermates were housed in
our facility in accordance with approved animal protocols. At
8 wk of age, mice were killed and pituitaries were removed for
total RNA extraction.
Quantitative Real-Time PCR
RNA was obtained with Trizol reagent (Invitrogen, Carlsbad,
CA) according to the manufacturer’s instructions. Contaminating DNA was removed with DNA-free reagent (Ambion,
Inc., Austin, TX) and 2 ␮g RNA was reverse transcribed using
Superscript III First-strand Synthesis System (Invitrogen,
Carlsbad, CA). Quantitative real-time PCR was performed in
iCycler from Bio-Rad Laboratories, Inc. (Hercules, CA), using
QuantiTect SYBR Green PCR Kit (QIAGEN, Valencia, CA) and
the following primers:
LH forward: CTGTCAACGCAACTCTGG
LH reverse: ACAGGAGGCAAAGCAGC
GAPDH forward: TGCACCACCAACTGCTTAG
GAPDH reverse: GGATGCAGGGATGATGTTC
FSH forward: GCCGTTTCTGCATAAGC
Coss et al. • Activin Regulation of LH␤
FSH reverse: CAATCTTACGGTCTCGTATACC
under the following conditions: 95 C for 15 min, followed by
40 cycles at 95 C for 15 sec, 54 C for 30 sec, and 72 C for 30
sec. For LH␤ and FSH␤ measurements, equivalent to 50 ng
of starting RNA (as quantified before reverse transcription)
was used in each reaction, whereas for GAPDH 10 ng was
sufficient. Each sample was assayed in triplicate, and the
experiment was repeated four times. A standard curve with
dilutions of 10 pg/well, 1 pg/well, 100 fg/well, and 10 fg/well
of a plasmid containing LH␤ cDNA, FSH␤ cDNA, or GAPDH
cDNA was generated in each run with the samples. In each
experiment, the amount of LH␤ was calculated by comparing
threshold cycle obtained for each sample with the standard
curve generated in the same run. Replicates were averaged
and divided by the mean value of GAPDH in the same sample. After each run, a melting curve analysis was performed to
confirm that a single amplicon was generated in each
reaction.
Cell Culture and Transient Transfection
L␤T2 cells were plated in 12-well plates 1 d before transfection. Transfection was performed in DMEM with 10% FBS
using Fugene 6 reagent (Roche Molecular Biochemicals, Indianapolis, IN) following the manufacturer’s instructions.
Each well was transfected with 0.5 ␮g rat LH␤-luciferase or
mouse FSH␤-luciferase plasmid (45). Plasmid construction
and preparation were described previously (24). The sequences of all of the promoter fragments were confirmed by
dideoxynucleotide sequencing. An expression plasmid containing ␤-galactosidase driven by the Herpes virus thymidine
kinase promoter was co-transfected with LH␤-luc and used
as an internal control. Sixteen hours after transfection, the
cells were switched to serum-free DMEM supplemented with
0.1% BSA, 5 mg/liter transferrin, and 50 nM sodium selenite.
The following day, the cells were treated with 10 ng/ml activin
for 6 h, unless otherwise indicated, or follistatin (R&D Systems, Minneapolis, MN) for 24 h. The cells were then lysed
with 0.1 M K-phosphate buffer (pH 7.8) with 0.2% Triton
X-100. Equal volumes of each lysate were placed in 96-well
plates, and luciferase activity was measured on a luminometer (EG&G Microplate, Berthold Technologies, Oak Ridge,
TN) by injecting 100 ␮l of a buffer containing 100 mM Tris-HCl
(pH 7.8), 15 mM MgSO4, 10 mM ATP, and 65 ␮M luciferin per
well. Galactosidase activity was measured using the Galactolight assay (Tropix, Bedford, MA) following the manufacturer’s instructions. All transfection experiments were performed in triplicate and repeated at least three times.
Luciferase values from reporter gene-transfected cells were
consistently at least 100 times higher than values from mock
transfected cells. Results represent the mean ⫾ SEM of all
samples analyzed. Asterisk marks a statistically significant
difference from the control-treated cells, determined by oneway ANOVA analysis followed by Tukey’s post hoc multiple
range test for individual comparison with P ⱕ 0.05 or Student’s t test as indicated.
Western Blot
After overnight starvation and 2-h treatment with vehicle control, 10 nM GnRH, or 10 ng/ml activin, L␤T2 cells were rinsed
with PBS and lysed with lysis buffer [20 mM Tris-HCl (pH 7.4),
140 mM NaCl, 0.5% Nonidet P-40 (NP-40), 0.5 mM EDTA],
with protease inhibitors (aprotinin, pepstatin, leupeptin) at 10
␮g/ml each, and 1 mM phenyl-methyl-sulfonyl-fluoride
(PMSF). Protein concentration was determined with Bradford
reagent (Bio-Rad Laboratories), and an equal amount of protein per sample was loaded on an SDS-PAGE gel. After
proteins had been resolved by electrophoresis and transferred to a polyvinylidene fluoride membrane, they were
probed with specific antibodies for Egr-1 (Santa Cruz Biotechnology, Inc., Santa Cruz, CA). The bands were detected
Mol Endocrinol, October 2005, 19(10):2610–2623 2621
with secondary antibodies linked to horseradish peroxidase
and enhanced chemiluminescence reagent (Amersham Pharmacia Biotech, Piscataway, NJ).
EMSA
L␤T2 cells, after vehicle control or 10 ng/ml activin treatment
for the times indicated, or Cos 1 cells, transfected for 24 h
with expression vectors containing Smad 3 and/or Smad 4
cDNAs (kindly provided by Rik Derynck), or vector control,
were scraped in hypotonic buffer (20 mM Tris-HCl, pH 7.4; 10
mM NaCl, 1 mM MgCl2) with protease inhibitors (aprotinin,
pepstatin, leupeptin) at 10 ␮g/ml each, and 1 mM PMSF and
allowed to swell on ice. Cells were broken by passing through
a 25-gauge needle, and the nuclei were pelleted by centrifugation. Nuclear proteins were extracted in hypertonic buffer
(20 mM HEPES, pH 7.8; 420 mM KCl; 1.5 mM MgCl2 with
protease inhibitors; and 20% glycerol). Nuclear protein (2 ␮g
per sample) was used in the binding reaction (10 mM HEPES,
pH 7.8; 50 mM KCl; 5 mM MgCl2; 5 mM dithiothreitol; 0.1%
BSA; 0.1% NP-40 with 0.5 ␮g/ml polydeoxyinosinic deoxycytidylic acid; and 2 fmol per reaction of end-labeled probe).
Oligonucleotides were labeled with T4 kinase using ␥-32Plabeled ATP. In the competition experiments, competitor oligonucleotide was added 10 min before addition of the probe,
as were the antibodies in the supershift assays. The Smad4
antibody and nonspecific IgG were obtained from Santa Cruz
Biotechnology and from Upstate USA, Inc. (Charlottesville,
VA). The reaction was loaded on a 5% acrylamide gel in
0.25⫻ TBE and run on 0.5 V/ cm2 constant voltage. After
drying, gels were exposed to autoradiography.
Mutagenesis
Mutagenesis of the LH␤-luc plasmid was performed using
the QuikChange Site-Directed Mutagenesis kit (Stratagene,
La Jolla, CA) according to the manufacturer’s protocol. The
HB mutation and Egr site mutation were described in a previous report (24). From the wild-type sequence 5⬘-TGTCTGTCTCGCCCCCAAAGAGATTAGTGTCTAGG-3⬘, three SBEs
(S1, S2, and S3; GTCT, underlined) were individually mutated
to GTAG and named mutant SBE 1 (mS1, most 5⬘), mutant
SBE 2 (mS2, middle SBE), and mutant SBE 3 (mS3, most 3⬘).
Mutations were confirmed by dideoxyribonucleotide sequencing performed by the DNA Sequencing Shared Resource, UCSD Cancer Center.
GST Interaction Assay
The GST-Smad2, 3, and 4 in the pGEX vector were kindly
provided by Dr. Rik Derynck. The Otx-1 and Ptx-1 expression
vectors were obtained from Dr. Antonio Simeone and Dr.
Jacques Drouin, respectively. The GFP expression vector
was provided by Dr. Douglass Forbes. 35S-labeled proteins
were produced using the TnT T7 Coupled Reticulocyte Lysate System (Promega Corp., Madison, WI). Bacteria transformed with the pGEX vectors were grown to an OD of 0.6,
upon which protein expression was induced by addition of
0.25 mM isopropyl-␤-D-thiogalactosidase. Bacterial pellets
were sonicated in PBS with 5 mM EDTA and 0.1% Triton
X-100 and centrifuged, and the supernatant was bound to
glutathione sepharose beads (Amersham Pharmacia Biotech). Beads were washed four times with sonication buffer
followed by equilibration in the binding buffer (below) and
split equally between different samples and the control. 35Slabeled proteins were added to the beads and bound for 1 h
at 4 C in 20 mM HEPES (pH 7.8), with 50 mM NaCl, 10 mg/ml
BSA, 0.1% NP-40, and 5 mM dithiothreitol. After extensive
washing, samples were eluted from the beads by boiling in
Laemmli sample buffer and subjected to SDS-PAGE. Afterward, the gels were dried and autoradiographed.
2622
Mol Endocrinol, October 2005, 19(10):2610–2623
Acknowledgments
We thank Rik Derynck who kindly provided the GSTSmad2, 3, and 4 in the pGEX vector and Smad3 and 4
expression vectors, and Suzanne Rosenberg Jacobs for
many LH␤ reporter plasmids. We are grateful to Antonio
Simeone and Jacques Drouin from whom we obtained the
Otx-1 and Ptx-1 expression vectors, respectively, and to
Douglass Forbes for the GFP expression vector. We appreciate the time and insight of the Mellon laboratory members
and Janice Sue Bailey for helpful discussions.
Received January 18, 2005. Accepted June 7, 2005.
Address all correspondence and requests for reprints to:
Pamela L. Mellon, Department of Reproductive Medicine,
University of California San Diego, 9500 Gilman Drive, La
Jolla, California 92093-0674. E-mail: pmellon@ucsd.edu.
This work was supported by National Institute of Child
Health and Human Development/National Institutes of Health
(NIH) through a cooperative agreement (U54 HD12303) as
part of the Specialized Cooperative Centers Program in Reproduction Research (to P.L.M.). This work was also supported by NIH Grant R37 HD20377 (to P.L.M.). D.C. was
supported by NIH National Research Service Award F32
HD41301 and the Lalor Foundation. V.G.T. was supported by
NIH National Research Service Award F32 DK065437. University of California San Diego Cancer Center was funded in
part by National Cancer Institute Cancer Center Support
Grant P30 CA23100.
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