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BOR - Papers in Press, published online ahead of print October 29, 2003.
Biol Reprod 2003, 10.1095/biolreprod.103.021162
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BIOLOGY OF REPRODUCTION 70, 640–648 (2004)
DOI: 10.1095/biolreprod.103.021162
© 2004 by the Society for the Study of Reproduction, Inc.

Ovary

Sequential Gonadotropin Treatment of Immature Mice Leads to Amplification of Transforming Growth Factor ß Action, Via Upregulation of Receptor-Type 1, Smad 2 and 4, and Downregulation of Smad 6

Xavier Guéripel, Mohamed Benahmed, and Alain Gougeon1

INSERM U-407, Faculté de Médecine Lyon-Sud, 69921 Oullins, France


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The present study was designed to establish the cellular localization and expression of transforming growth factor ß (TGFß) signaling pathway components, including TGFß1 and ß2; TGFß receptors type I (TßRI) and II (TßRII); and Smads 2, 3, 4, and 6 during gonadotropin-induced follicular maturation and ovulation in the mouse ovary. Immature 21-day-old mice were sequentially treated with recombinant human FSH, 5 IU daily for 3 days, and hCG once at Day 24 of life. Immunohistochemical experiments revealed a TGFß1 staining in granulosa cells (GC) and theca interna cells (TIC) as well as in oocytes, whereas that of TGFß2 was mainly localized in oocytes and GC. Strong immunostaining for both TßRI and -RII was observed in the TIC and, to a lesser extent, in GC. Whereas oocytes did not exhibit any staining for TßRII, their TßRI immunostaining was strong. Smads were detected in oocytes, GC, and luteal cells and in a lesser amount in TIC; the immunostaining for Smad 4 was the strongest. Western blotting and reverse transcription-polymerase chain reaction analyses indicated that, in response to gonadotropins, TGFß2, TßRI, Smad 2 and Smad 4 mRNA and protein levels increased, while those of Smad 6 decreased in ovarian homogenates. In conclusion, these results show that, in a model of immature mouse exposed to a sequential gonadotropin treatment, FSH and LH increased the expression of the TGFß signaling system through the increase of TGFß2, TßRI, stimulatory Smad 2, and common Smad 4 expression, which occurred concomitantly with a decrease of the inhibitory Smad 6 expression.

follicle, follicle-stimulating hormone, growth factors, luteinizing hormone, ovary


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Peptides of the large transforming growth factor ß (TGFß) superfamily regulate multiple biological processes in mammals, such as development, growth, differentiation, and apoptosis (reviewed in [1, 2]). In the ovary, many members of the TGFß superfamily have been detected, such as anti-Müllerian hormone [3], bone morphogenetic proteins (BMPs), growth differentiation factors [4], inhibin subunits {alpha} and ß [5], and TGFßs [6]. For TGFßs, the signaling pathway is initiated by the binding of the ligand to cell-surface serine/threonine kinase receptors, type I and type II (TßRI and TßRII) [7], that activate the Smad signaling pathway [8]. The Smads are a family of related intracellular proteins, ranging from about 400 to 500 amino acids in length, critical for transmitting to the nucleus signals from the receptor after binding of the ligand [9]. Smads are classified into three subfamilies: the receptor-regulated Smads 1, 2, 3, 5, 8, and 9; the common Smad 4; and the inhibitory Smads 6 and 7 (I-Smads).

In rodents, TGFß1 and TGFß2 have been imunolocalized in theca interna cells (TIC), granulosa cells (GC), and luteal cells as well as in oocytes of mice [6, 10], rats [11–14], and hamsters [15, 16]. However, intensity level of TGFß immunostaining appears to be highly variable among species, cycle day, and stage of follicular development. In the presence of TGFßs, TßRI and TßRII form a heterodimer that is essential in mediating the actions of TGFßs (reviewed in [17]). The TßRI has been detected in GC and luteal cells of the mouse ovary, but the main sites of immunoreactive TßRI are TIC and oocytes [6]. In mice, despite the fact that TßRII protein has been localized at a high level in oocytes and at a lower level in TIC, GC, and luteal cells [6], the TßRII mRNAs have been localized at a strong level in TIC and at only a weak level in GC and oocytes [18]. Little information is available on the expression of Smad proteins in the ovary. Smad 2 protein has been localized in hen GC [19]; in human oocytes [20]; and in oocytes, GC, and TIC of both immature [21, 22] and adult rats [22]. Smad 3 mRNA has been detected in mouse GC [23]. Similar to Smad 2, Smad 3 [22] and 4 [21] proteins have also been localized to oocytes, GC, and TIC of both immature and cycling rats.

In laboratory rodents, the effects of TGFßs on the ovary appear to be largely paracrine and generally enhance follicular growth [24–27]. TGFßs also alter ovarian steroidogenesis; they stimulate progesterone synthesis in rat ovaries [28] but inhibit androgen production by TIC [28].

The gonadotropins LH and FSH are major regulators of ovarian function, but there are no data related to the hormonal regulation of TGFßs, TßRs, and Smads mRNA and proteins in mice. In the hamster ovary, FSH and LH stimulate the production of TGFß2 and TGFß1, respectively [15]. FSH has been shown to inhibit TGFß2 expression [29] and production [30] by rat GC cultured in vitro. LH, which inhibits TGFß2 expression in rat GC, has either no effect on [12] or inhibits [30] TGFß1 production by TIC. Taken together, available data indicate that the relationships between the endocrine system and the expression of different components of the ovarian TGFß system remain to be clarified.

The present study was undertaken to test the hypothesis that the complete TGFß system, including the signaling proteins Smads, is expressed in the mouse ovary and is regulated by gonadotropins. Consequently, 1) the cellular localization of TGFß1, TGFß2, TßRI, and TßRII and Smads 2, 3, 4, and 6 was determined in the mouse ovary. Regulated Smads 2 and 3 were investigated because they are phosphorylated by activated activin and TGFß receptors whereas Smads 1, 5, and 8 are phosphorylated by activated BMP-receptors [21]. 2) Expression of TGFß1, TGFß2, TßRI, TßRII, and Smads 2, 3, 4, and 6 was analyzed in terms of mRNA and protein by using as a model immature mice sequentially treated with gonadotropins FSH and LH/hCG.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animals and Treatments

Twenty-one-day-old OF1 mice (Iffa Credo, l'Arbresle, France) were treated by a daily s.c. injection of 5 IU recombinant human FSH (r-hFSH; Gonal F, Serono, Boulogne, France) on 3 consecutive days (Days 21, 22, and 23). On Day 24, they were injected i.m. with 5 IU hCG (Pergonal, Serono). Treated mice were killed by cervical dislocation at Days 22 (24 h after r-hFSH injection: F24), 23 (F48), 24 (F72), 25 (24 h after hCG injection: H24), and 26 (H48) of age, and untreated control animals were killed at Days 21, 22, 23, 24, 25, and 26 of age. Ovaries were carefully dissected to separate ovarian tissue from surrounding fat tissue, ovarian bursa, and Fallopian tubes. The freshly dissected ovaries were either fixed for 24 h in 10% buffered formalin or Bouin solution, or snap frozen on dry ice then kept at -80°C for further biochemical studies. Investigations and animal care procedures were in accordance with the INSERM (French National Institute for Health and Medical Research) Animal Care Committee (decrees 2001-486 and 2001-464).

RNA Extraction

Total RNA was extracted from whole ovaries with TRIzol reagent, a monophasic solution of phenol and guanidine isothiocyanate. This reagent is an improvement of the single-step RNA isolation developed by Chomczynski and Sacchi [31]. The amount of RNA was estimated by spectrophotometry at 260 nm.

Reverse Transcription

Single-stranded cDNAs were obtained from reverse transcription (RT) of 2 µg of total RNA using random hexanucleotides as primers (5 µM) in the presence of deoxynucleotide triphosphates (250 µM) (Invitrogen, Cergy-Pontoise, France), dithiothreitol (10 mM) (Life Technologies, Eragny, France), and Moloney murine leukemia virus reverse transcriptase (10 U/µl) (Life Technologies) for 1 h at 37°C.

Polymerase Chain Reaction Coamplification Analysis

Complementary DNAs (2 µl RT mixture) were amplified by polymerase chain reaction (PCR) with Taq polymerase (0.025 U/µl) (Promega, Charbonnière-les-Bains, France), deoxynucleotides (250 µM), 0.75 µCi [{alpha}-33P]deoxy-ATP (Amersham, Buckinghamshire, UK), specific primers (0.5 µM) and ß-actin primers (forward: TTGCTGATCCACATCTGCTG; reverse: GACAGGATGCAGAAGGAGAT). The mixture was first heated at 94°C for 5 min followed by different numbers of cycles (n) of 30 sec each at 94°C, 30 sec at melting temperature (Tm), 30 sec at 72°C, and 7 min at 72°C (see Table 1). The amplification was performed in a thermocycler (PCR express; Hybaid, Middlesex, UK). The n, the Tm, and the ß-actin concentrations were determined for each pair of primers after having tested temperatures between 50 and 70°C, cycle number between 20 and 40, and ß-actin concentration between 10 and 200 nM. PCR products were separated on an 8% polyacrylamide gel. Dried gels were exposed to phosphor screens (Packard, Meriden, CT). Intensity of bands was estimated by densitometric scanning using the Cyclone Storage Phosphor scanner (Packard). The data are expressed as interest molecule:ß-actin mRNA ratio.


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  		TABLE 1. Sequence of primers used for RT-PCR experiments

PCR analyses were carried out from the logarithmic phase of amplification. PCR-amplified products were checked by direct sequencing (ABI Prism; 310 genetic analyzer; Applied Biosystems, Courtaboeuf, France). RT-PCR primers were designed inside separate exons to avoid any bias caused by residual genomic contamination. Moreover, for all primers, no amplification was observed when PCR was performed on RNA preparations.

Western Blot Analysis

Protein was extracted by mechanical homogenization of ovaries in the presence of 200 µl ice-cold hypotonic buffer (25 mM Tris-HCl, pH 7.4, 1 mM EDTA, protease inhibitor cocktail 1% [Sigma, l'Isle d'Abeau, France]). Protein concentrations were obtained by using a colorimetric Bradford method.

Proteins (30 µg) were resolved on 12% SDS/polyacrylamide gels and electrophoretically transferred to nitrocellulose membranes (Hybond-C extra; Amersham, Les Ulis, France) using 25 mM Tris and 185 mM glycine (pH 8.3) containing 20% methanol. The transfer was performed at a constant voltage of 100 V for 2 h. After transfer, the membranes were incubated in blocking buffer (tris-buffered saline [TBS] buffer containing 10% fat-free dry milk) for 1 h at room temperature. The membranes were rinsed (3 x 10 min) with TBS/Tween 20 0.1% and incubated with primary antibodies (TGFß1, TGFß2, TßRII, Smads 2 and 3: 0.4 µg/ml, Smads 4 and 6: 0.2 µg/ml, TßRI: 0.1 µg/ml) overnight at 4°C. The membranes were rinsed with TBS/Tween 20 0.1% (3 x 10 min) and then incubated with horseradish peroxidase-labeled goat or rabbit antibodies (1:2000) as appropriate. Bound antibodies were detected by chemiluminescence using a Covalab kit (CovalAb; Oullins, France) and Biomax MR-1 film (Eastman Kodak Co., Rochester, NY). The protein loading was checked by reprobing the blot with an actin antibody (1:500). The molecular weight of each protein was determined using biotinylated protein markers (CovalAb). This allowed validation of all antibodies used.

Immunohistochemistry

Paraffin sections (5 µm) of either 10% neutral formalin- or Bouin-fixed ovaries were mounted on polysin-coated glass slides (Polysin; Menzel-Glaser, Braunscheig, Germany). Sections were deparaffinized in xylen, hydrated, treated 20 min at 98°C in pH 6 citric buffer, rinsed in osmosed water (2 x 5 min), washed (2 x 5 min) in PBS Tween 20 0.1%, then incubated 10 min at 37°C in the peroxidase blocking reagent (DAKO, Trappes, France), washed (2 x 5 min) in PBS-Tween 20 0.1%. After which the primary antibody, appropriately diluted 1:100–1:200 (1–2 µg/ml) was added and the sections incubated overnight at 4°C. Dose-response studies indicated that these dilutions of the antibodies gave optimal labeling results. When possible, several primary antibodies of either different origins or lots were used. Each of them was tested on both Bouin fluid and neutral formalin-fixed sections. When different staining between the two fixative types was observed, the primary antibody was tested on frozen sections postfixed in glacial acetone for 5 min. Immunohistochemistry was subsequently performed on the fixed sections that displayed the same immunostaining as the frozen sections. Each selected antibody was controlled by Western blotting to ensure that only one band at the expected molecular weight for the peptide against which it was directed was obtained. The selected antibodies were purchased from Santa Cruz Biotechnology Inc. (Santa Cruz, CA) and were the following: TGFß1 (sc-146, lot E170), TGFß2 (sc-090, lot BO603), TßRI (sc-398, lot D135), TßRII (sc-220, lot E222), Smad 2 (sc-6200, lot AO603), Smad 3 (sc-6202, lot L1702), Smad 4 (sc-1908, lot K238), and Smad 6 (sc-6034, lot L111). After incubation with the primary antibody, the sections were rinsed, washed (2 x 5 min) in PBS-Tween 20 0.1%, then incubated 30 min at 37°C in the presence of the secondary antibody. For rabbit polyclonal antibodies (TGFß2, TßRI, and TßRII), we used secondary rabbit immunoglobulins attached to a peroxidase-conjugated polymer backbone (Envision+ kit; DAKO). After incubation with the secondary antibody, the sections were rinsed; washed (2 x 5 min) in PBS-Tween 20 0.1%; incubated 10 min at room temperature with AEC (3-amino-9-ethylcarbazole, DAKO), which generated a red color at the site of peroxidase activity; rinsed; washed (2 x 5 min) in osmosed water; and then the nuclei were counterstained with Mayer hematoxylin (DAKO). For goat (TGFß1 and Smads) polyclonal antibodies, we slightly modified the procedure mentioned above for rabbit antibodies by incubating the sections 30 min at 37°C in the presence of a rabbit anti-goat serum (DAKO) diluted at 1:1000 just before incubation with the secondary antibody. Sections were coverslip mounted using Faramount (DAKO). Control sections received either buffer or serum (goat or rabbit, as appropriate) diluted appropriately, in place of the primary antibody. For each control or treatment day, one ovary from five mice was used, two to four sections per ovary were examined for each antibody. The intensity of the immunostaining was graded as weak, moderate, or strong.

Follicles were classified as follows: primordial follicles containing an oocyte surrounded by a single layer of flattened squamous GC; early growing follicles including primary, secondary, and preantral follicles; small antral follicles; and preovulatory follicles larger than 0.4 mm in diameter.

Data Analysis

All data are presented as the mean ± SD of at least three separate RT-PCR and Western blot experiments for each animal. Statistical analyses were carried out by ANOVA and post hoc Bonferroni/Dunn test (Statview software; SAS Institute Inc., Cary, NC).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
For all antibodies, the replacement of the primary antibody by either the antibody diluent (Fig. 1A) or rabbit or goat normal serum, as appropriate, led to absence of any immunostaining in all ovarian tissues.



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  		FIG. 1. Immunolocalization of TGFß1 and TGFß2 in the 21- to 26-day-old mouse ovary. Bar = 70 µm. O, Oocyte; GC, granulosa cells; TIC, theca-interna cells; P, primordial follicles. A) In control section where the primary antibody has been omitted and replaced by the antibody diluent, no immunostaining is observed (Bouin fluid fixation). B) The immunostaining for TGFß1 in a 26-day-old mouse is strong in primordial oocytes (inset, bar = 12 µm), moderate in GC and growing oocytes, and weak in TIC (neutral formalin fixation). C) The immunostaining for TGFß2 in a 21-day-old mouse is strong in primordial oocytes (inset, bar = 12 µm), moderate to strong in growing oocytes, moderate in GC from some but not all antral follicles (AF); it is weak in GC from early growing follicles (early GF) and in TIC (neutral formalin fixation). D) By 48 h after the first r-hFSH injection, the TGFß2 immunostaining is strong in GC of all growing follicles and in oocytes and weak in TIC (neutral formalin fixation)

TGFß1 and TGFß2

No significant changes were observed in ovarian TGFß1 mRNA and protein levels in gonadotropin-treated mice when compared with control animals (data not shown). In control mice, the TGFß1 immunostaining was similarly present at a moderate level in oocytes and GC, but was weak in TIC (Fig. 1B). After sequential treatment with gonadotropins, neither immunolocalization of TGFß1 nor its staining intensity were changed. Corpora lutea, which appeared after hCG treatment, were, similar to preovulatory follicles, stained at a moderate level.

In control mice, neither TGFß2 mRNA nor TGFß2 protein levels exhibited significant changes from Day 21 to Day 26 (Fig. 2). In gonadotropin-treated animals, the ovarian levels of TGFß2 mRNA were significantly increased as soon as 24 h after the first r-hFSH injection when compared with control values at Day 21 (P < 0.0006) and Day 23 (P < 0.0001); they reached a plateau by 72 h after the first r-hFSH injection (Fig. 2A). The TGFß2 protein levels increased significantly (P < 0.0001) only 72 h after the first r-hFSH injection and remained at the same level after hCG injection (Fig. 2B). In control mice, the TGFß2 immunostaining was mainly observed in oocytes of both nongrowing and growing follicles; it was weak in both TIC and GC, but sometimes, GC in some antral follicles displayed a moderate staining (Fig. 1C). By 48 h after the first injection of r-hFSH, the TGFß2 immunostaining increased in both early growing and antral follicles (Fig. 1D); after hCG injection, it was moderate to strong in corpora lutea.



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  		FIG. 2. Effect of gonadotropins on TGFß2 mRNA (A) and TGFß2 protein (B) levels in the mouse ovary. Ovarian TGFß2 mRNA and protein levels in gonadotropin-treated mice (F24, F48, F72, H24, H48: open bars) were compared with those in untreated 21- (C21) to 26-day-old (C26) control mice (black bars). The histogram A represents the mean ± SD values of the TGFß2:ß-actin mRNA ratio of six animals for each control and treatment day; a was different from b and c at P < 0.0001, and from C21 and C23 control values at P < 0.006 and P < 0.0001, respectively; b was different from c and control values at P < 0.0001. The histogram B represents the mean ± SD values of the TGFß2:ß-actin protein ratio of five animals for each control and treatment day; a was different from b at P < 0.0001, but not different from control values. Lower panels, a representative RT-PCR (A) or Western blot (B) is shown

TßRI and TßRII

There were no changes in ovarian TßRI mRNA and protein levels from Day 21 to Day 26 in control mice (Fig. 3). By 48 h after the first r-hFSH injection, the ovarian levels of TßRI mRNA and protein were significantly increased when compared with control values (P < 0.0001). A plateau was reached by 72 h after the first r-hFSH injection (Fig. 3). In control mice, a strong TßRI immunostaining was seen in both oocytes and TIC; it was weak in GC, but moderate in basal GC from some antral follicles (Fig. 4A). Whereas the TßRI immunostaining remained strong in both TIC and oocytes in response to both r-hFSH and hCG, by 48 h after the first injection of r-hFSH, it increased in GC (Fig. 4B). When they were present, after hCG injection, the corpora lutea exhibited a moderate to strong immunostaining for TßRI. Whereas the TßRI immunostaining remained strong in oocytes exhibiting a germinal vesicle, it disappeared from oocytes displaying a germinal vesicle breakdown in atretic follicles (Fig. 4C).



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  		FIG. 3. Effect of gonadotropins on TßRI mRNA (A) and TßRI protein (B) levels in the mouse ovary. Ovarian TßRI mRNA and protein levels in gonadotropin-treated mice (F24, F48, F72, H24, H48: open bars) were compared with those in untreated 21- (C21) to 26-day-old (C26) control mice (black bars). The histogram A represents the mean ± SD values of the TßRI:ß-actin mRNA ratio of six animals for each control and treatment day. The histogram B represents the mean ± SD values of the TßRI:ß-actin protein ratio of five animals for each control and treatment day. For both histograms (A and B), a was different from c at P < 0.0001 but not from b and control values; b, which was not different from a and c, was different from control values. Lower panels, a representative RT-PCR (A) or Western blot (B) is shown



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  		FIG. 4. Immunolocalization of TßRI and TßRII in the 21- to 26-day-old mouse ovary. Bar = 70 µm. O, Oocyte; GC, granulosa cells; bGC, basal granulosa cells; TIC, theca-interna cells; P, primordial follicles. A) The immunostaining for TßRI in a 23-day-old mouse is strong in oocytes from primordial and growing follicles and in TIC, moderate in basal GC of some antral follicles, but weak in GC of most follicles (Bouin fluid fixation). B) By 48 h after the first r-hFSH injection, TßRI immunostaining had increased to a moderate level in GC of all follicles and remains strong in both TIC and oocytes (Bouin fluid fixation). C) In an ovary sampled 48 h after hCG injection, the TßRI immunostaining is strong in oocytes with a germinal vesicle (GV) whereas it is absent from oocytes displaying germinal vesicle breakdown (GVBD) in atretic follicles (Bouin fluid fixation). D) The immunostaining for TßRII in a 23-day-old mouse is strong in TIC while the oocytes are not stained. In antral follicles, basal GC display a weak to moderate immunostaining while antral GC are weakly stained (Bouin fluid fixation)

No significant changes were observed for either ovarian TßRII mRNA or protein levels in gonadotropin-treated mice when compared with control animals (data not shown). TßRII immunostaining was strong in TIC but absent in oocytes; it was weak in GC of growing follicles but stronger in basal than in antral GC (Fig. 4D). The immunostaining for TßRII changed neither after r-hFSH treatment nor after hCG injection.

Smads

In control mice, from Day 21 to Day 26, there were no significant changes in ovarian protein and mRNA levels for Smad 2 (Fig. 5), Smad 3 (data not shown), Smad 4 (Fig. 6), and Smad 6 (Fig. 7). From 48 h after the first r-hFSH injection onward, the ovarian levels of Smad2 mRNA and proteins were significantly increased (P < 0.0001) and reached a maximum by 72 h after the first r-hFSH injection (Fig. 5). Whereas the treatment of mice with hCG did not induce any changes of Smad 2 mRNA levels, it induced a transitory decrease (P < 0.0001) of Smad 2 protein levels by 24 h after hCG. However, 48 h after hCG injection, these levels increased (P < 0.0001) back to those observed 72 h after the first r-hFSH injection (Fig. 5B). In response to either FSH or hCG, the Smad 3 mRNA and protein levels did not exhibit any changes when compared with control values (data not shown).



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  		FIG. 5. Effect of gonadotropins on Smad 2 mRNA (A) and Smad 2 protein (B) levels in the mouse ovary. Ovarian Smad 2 mRNA and protein levels in gonadotropin-treated mice (F24, F48, F72, H24, H48: open bars) were compared with those in untreated 21- (C21) to 26-day-old (C26) control mice (black bars). The histogram A represents the mean ± SD values of the Smad 2:ß-actin mRNA ratio of six animals for each control and treatment day; a was different from b and c at P < 0.0001, but not different from control values; b was different from c and control values at P < 0.0001. The histogram B represents the mean ± SD values of the Smad 2:ß-actin protein ratio of five animals for each control and treatment day; a was different from b, c, d at P < 0.0001, but not different from control values; b was different from d (P < 0.002) and from c and control values at P < 0.0001). c was different from d at P <= 0.0003. Lower panels, a representative RT-PCR (A) or Western blot (B) is shown



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  		FIG. 6. Effect of gonadotropins on Smad 4 mRNA (A) and Smad 4 protein (B) levels in the mouse ovary. Ovarian Smad 4 mRNA and protein levels in gonadotropin-treated mice (F24, F48, F72, H24, H48: open bars) were compared with those in untreated 21- (C21) to 26-day-old (C26) control mice (black bars). The histogram A represents the mean ± SD values of the Smad 4:ß-actin mRNA ratio of six animals for each control and treatment day; a was different from b at P < 0.0001 and from control values at P <= 0.0007. The histogram B represents the mean ± SD values of the Smad 4:ß-actin protein ratio of five animals for each control and treatment day; a was different from b, c, and control values at P < 0.0001; b was different from c and control values at P < 0.0008. Lower panels, a representative RT-PCR (A) or Western blot (B) is shown



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  		FIG. 7. Effect of gonadotropins on Smad 6 mRNA (A) and Smad 6 protein (B) levels in the mouse ovary. Ovarian Smad 6 mRNA and protein levels in gonadotropin-treated mice (F24, F48, F72, H24, H48: open bars) were compared with those in untreated 21- (C21) to 26-day-old (C26) control mice (black bars). The histogram A represents the mean ± SD values of the Smad 6:ß-actin mRNA ratio of six animals for each control and treatment day. The histogram B represents the mean ± SD values of the Smad 6:ß-actin protein ratio of five animals for each control and treatment day. For both histograms (A and B), a was different from b at P < 0.0001, but not from control values. Lower panels, a representative RT-PCR (A) or Western blot (B) is shown

The ovarian levels of Smad 4 mRNA significantly (P < 0.0001) increased 24 h after the first injection of r-hFSH and reached a plateau by 48 h after the first r-hFSH injection. The slight decrease of Smad 4 mRNA levels observed 24 h after hCG injection was not significant (Fig. 6A). The levels of Smad 4 protein significantly (P < 0.0001) increased 48 h after the first injection of r-hFSH and reached a plateau 72 h after the first r-hFSH injection (Fig. 6B). From 48 h after the first r-hFSH injection to 48 h after hCG injection, the ovarian Smad 6 mRNA levels continuously and significantly (P < 0.0001) decreased (Fig. 7A). The ovarian Smad 6 protein levels were significantly (P < 0.0001) decreased by 72 h after the first r-hFSH injection and by 48 h after hCG injection when compared with levels at 24 and 48 h after the first r-hFSH injection and with control values. However, by 24 h after hCG, the Smad 6 protein levels were similar to control values (Fig. 7B).

In the control mouse ovary, immunostaining for Smads 2, 3, 4, and 6 was observed in the same cell types. In oocytes and GC, immunostaining for Smads 2 (Fig. 8A) and 4 (Fig. 8C) was stronger than that for Smads 3 (Fig. 8B) and 6 (Fig. 8F). For all Smads, the level of immunostaining was weak in TIC. There was a heterogeneous staining of GC between follicles; for Smads 2 and 4, the staining varied from weak to moderate, whereas for Smads 3 and 6, it varied from absent to weak. No relationships between follicle size and Smad immunostaining were observed. While the levels of immunostaining for Smads 3 and 6 did not consistently change in gonadotropin-treated mice when compared with control animals, those for Smads 2 and 4 (Fig. 8D) appeared stronger in gonadotropin-treated than in control mice. After hCG treatment, corpora lutea were positively stained for Smads, and the staining was the strongest for Smad 4 (Fig. 8E).



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  		FIG. 8. Immunolocalization of Smads 2, 3, 4, and 6 in the 21- to 26-day-old mouse ovary (neutral formalin fixation). Bar in each = 70 µm. O, Oocyte; GC, granulosa cells; TIC, theca-interna cells; P, primordial follicles; CL, corpus luteum. A) The immunostaining for Smad 2 in a 23-day-old mouse is strong in primordial oocytes, moderate in growing oocytes and GC, and weak in TIC. B) The immunostaining for Smad 3 in a 23-day-old mouse is weak to moderate in oocytes, GC, and TIC. C) The immunostaining for Smad 4 in a 23-day-old mouse is moderate to strong in both GC and oocytes of some, but not all follicles, and weak in TIC. D) By 48 h after the first FSH injection, the Smad 4 immunostaining appears to be increased in GC of all growing follicles, whereas it remains weak in TIC. E) By 48 h after hCG injection, the Smad 4 immunostaining is moderate to strong in both follicles and corpora lutea. F) The immunostaining for Smad 6 in a 23-day-old mouse is often strong in primordial oocytes but weak in growing oocytes, in TIC, and GC; however, in some follicles, GC show a moderate immunostaining


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
This is, to our knowledge, the first report on the expression of the ovarian TGFß system, including ligands, receptors, and Smads, in terms of cellular immunolocalization as well as mRNA and protein levels in immature mice either untreated or following a sequential hormonal treatment with FSH and LH/hCG. Indeed, gonadotropic stimulation of the 21-day-old mouse ovary induces strong changes in the follicular population, leading to preovulatory maturation within 48–72 h [32]. During this time interval, GC actively proliferate, then differentiate when follicles reach the large antral stage. Thus, changes in levels of certain proteins and mRNA need to be considered in this context of both active follicular growth and terminal differentiation.

The localization of TGFß1 protein in the immature ovary is consistent with previously published studies in the mouse [6, 10], showing that this peptide is present in oocytes, TIC, and GC. The present data showing that TGFß1 was regulated neither by FSH nor by hCG are also consistent with previous studies in immature rats [12] and with porcine ovarian cells [33]. However, they differ from a previous immunohistochemical study in the immature mouse where eCG was reported to decrease, whereas eCG plus hCG increased, the TGFß1 immunostaining in TIC [10].

TGFß2 immunostaining was mainly detected in oocytes and at a lesser extent in GC, but it was very low in TIC. These data are in contrast with previous studies for the localization of TGFß2 in either the TIC [6, 10] or the oocyte [6], but are consistent with a high level of TGFß2 mRNA in oocytes, but apparently none in TIC [18]. The TGFß2 mRNA levels significantly increased from control levels as soon as 24 h after the first FSH injection, while those of the protein increased 48 h later. The TGFß2 immunostaining, which was weak in GC from early growing follicles in control mice, became moderate by 48 h after the beginning of the FSH treatment. Taken together, our data are consistent with those of Roy et al. [15], who observed that TGFß2 protein is upregulated by FSH in hamster GC, but they appear to be contradictory with data from rats, where FSH has been reported to inhibit both total and active TGFß production by cultured GC from hypophysectomized 21-day-old rats [30]. Nevertheless, in the latter experiment, GC were cultured in vitro and all TGFßs were assayed, leading to the impossibility of distinguishing which TGFß isoform production was altered by FSH. Finally, our data and previous studies performed in rodents show that important differences may exist in the regulation of TGFß1 and TGFß2 expression by gonadotropins. The presence of a cAMP regulatory element (CRE) sequence in the mouse TGFß2 gene promoter [34, 35] suggests that TGFß2 can be directly stimulated by FSH at the transcriptional level, although it cannot be excluded that FSH increases TGFß2 mRNA stability, as previously demonstrated in cultured porcine Sertoli cells [36, 37]. No CRE sequence was noted in the promoter region of the mouse Tgfb1, but various data [38, 39] show that TGFß1 regulation by the cAMP pathway is complex and remains unclear. Whether TGFß1 stimulation requires different conditions than TGFß2 to be activated by FSH and LH in the mouse ovary requires further studies, but the present data suggest that TGFß2 may play a more critical role than TGFß1 during mouse ovarian follicle development.

The immunolocalization of TßRI in ovaries of 21- to 26-day-old mice is consistent with previously published data in the mouse [6]. For TßRII, contrary to Juneja et al. [6], who described strong immunostaining in oocytes while GC and TIC displayed moderate staining, the present data show that there is no immunostaining in oocytes, strong staining in TIC, and weak to moderate staining in GC. Consequently, our results are rather in agreement with the study showing that TßRII mRNA is mainly expressed in TIC, barely expressed in GC, and not expressed in oocytes [18].

In response to FSH, the TßRI mRNA and protein levels increased 48 h after the first injection of FSH, whereas those for TßRII did not change. Simultaneous with these changes, TßRI immunostaining increased in GC. In response to hCG, TßRI immunostaining as well as mRNA levels remained high. Taken together, the present findings indicate that, following a sequential FSH/LH-treatment, TßRI but not TßRII mRNA and protein levels increase. Because in the rat the promoter of the gene coding for TßRI contains several Sp1 binding sites [40], it appears that this gene may be directly regulated by gonadotropins. However, the possibility also exists that FSH increases TßRI mRNA via an increase in TGFß ligand levels because, in various cellular models, TGFß increases TßRI mRNA levels [41, 42].

The Smad proteins were mainly detected in oocytes and GC. Our data are consistent with those of Li et al. [19], Osterlund et al. [20], and Drummond et al. [21], who detected Smad 2 in hen GC, in human oocytes, and in oocytes, GC, and TIC of immature 12-day-old rats, respectively. Similar to our own data, Drummond et al. [21] detected Smads 2 and 4 at similar levels in the same cell types. Smad 2 mRNA and protein levels become significantly different from control values after 48 h of FSH treatment. The decrease in Smad 2 protein levels observed by 24 h after hCG injection may have no physiological meaning; however, it cannot be excluded that hCG transitorily inhibits Smad 2 translation. Surprisingly, the situation in the mouse ovary seems to be different from that recently observed in immature and cycling rats [22], where Smad 2 and 3 proteins are downregulated by equine CG, while hCG has a dual effect, being first inhibitory and then stimulatory. For Smad 4, the mRNA levels significantly and strongly increased 24 h after the beginning of FSH treatment whereas the levels of the protein became significantly different from control levels after 48 h of FSH treatment. For Smad 2 and Smad 4, the changes in protein levels can be correlated to the increasing number of large follicles possessing GC, exhibiting a moderate immunostaining for both Smads 2 and 4. Very little is known about the regulation of Smad proteins. Smad 2 is upregulated by TGFß in avian GC [19] and in mammary carcinoma cells treated with the anticancer agent perillyl alcohol [43]. There are therefore some lines of evidence showing that not only TßRs can be induced in response to TGFß action, but that Smad 2 can also be upregulated by TGFßs. Given these different findings, we suggest, on one hand, that FSH stimulates TGFß2 production and, on the other hand, that TGFß2 may act on TßRI and Smad 2 expression to enhance its own actions in the mouse ovary. Nevertheless, it remains possible that FSH directly stimulates Smad expression, especially that of Smad 4, mRNA levels of which were rapidly and strongly increased in response to FSH.

Changes in Smad 6 mRNA and protein levels occur later than do changes in Smads 2 and 4; the significant decreases in Smads 2 and 4 occur 48 h and 72 h after FSH treatment, respectively. The increase in Smad 6 protein back to control level observed by 24 h after hCG injection may have no physiological meaning; however, it cannot be excluded that hCG transitorily stimulates Smad 6 translation. In mice, Smad 6 forms stable associations with TßRI and interferes with the phosphorylation of Smad 2 and the subsequent heteromerization with Smad 4 [44]. Surprisingly, contrary to the situation observed in various cellular models where I-Smads are activated to terminate either the TGFß, the activin, or the BMP signaling [45–48], the present study shows that the I-Smad 6 is downregulated in response to a gonadotropin treatment, leading to amplification of the TGFß action on ovarian cells. This suggests that a transcriptional repressor for Smad 6 may be operating in response to gonadotropins in the mouse ovary. Emerging evidence suggests that TGFß-inducible early gene (TIEG) is a transcriptional repressor that can downregulate the I-Smad 7 gene expression, a process that is Smad 4 dependent [49] and may involve the activation of Smad 2 [50]. Because in the ovary of mice sequentially treated with gonadotropins, Smad 6 is downregulated, it remains also to be demonstrated if TIEG can act as a transcriptional repressor for Smad 6.

The present data show that TGFß2, TßRI, Smad 2, and Smad 4 mRNA and proteins from whole ovarian extracts increase in response to FSH, which induces terminal folliculogenesis in immature mice. In the mammalian ovary, FSH receptors are present only on GC. In response to FSH, TGFß2, TßRI, Smad 2, and Smad 4 showed increased immunostaining in GC of growing follicles, consequently, it can be assumed that mRNA and protein changes observed in whole ovarian extracts likely reflect changes at the GC rather than at the TIC or oocyte level.

Although the present study shows that TIC exhibit a strong immunostaining for TßRI and TßRII, these cells exhibit weak immunostaining for Smads. A similar observation has been previously made in both immature and cycling rats for Smad 2 and Smad 3 [22]. Consequently, despite many effects of TGFßs on TIC having been reported in the literature (reviewed in [51]), our data suggest that, in vivo, TICs have poor ability to respond to TGFß, raising concerns about previous in vitro experiments. In addition, immunohistochemical observations of the present study have shown that oocytes contain TßRI but not TßRII, suggesting that the mouse oocyte may not be a target for TGFß, contrary to the situation in rats [52, 53].

In conclusion, the present study, showing that GC contain both TGF-ß receptors and Smad signaling proteins, suggests that this tissue is the preferential ovarian target for TGFßs either produced by itself (autocrine action) or by neighboring tissues (paracrine action). This study is, to our knowledge, the first report showing that a sequential gonadotropin treatment in the immature mouse can induce an autocrine loop leading to amplification of TGFß action at the ovarian level, via upregulation of TßRI, Smads 2 and 4 expression and production, and downregulation of the I-Smad 6 expression and production. Because GC are the only cells to possess FSH receptors, then LH receptors when they become preovulatory, it appears likely that this autocrine loop takes place at the GC level.


   ACKNOWLEDGMENTS
 
We thank Sonia Chater for her kind advices, Nicole Riu and Pierre Contard for their technical assistance, and Anne Florin-McLeer for her critical reading of the manuscript.


   FOOTNOTES
 
1 Correspondence: A. Gougeon, INSERM U-407, Faculté de Médecine, Centre hospitalier Lyon-Sud, 165 Chemin du Grand Revoyet, 69310 Pierre Bénite, France. FAX: 33 4 78 86 31 16; gougeon{at}lsgrisn1.univ-lyon1.fr Back

Received: 15 July 2003.

First decision: 8 August 2003.

Accepted: 27 October 2003.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Dunker N, Krieglstein K. Targeted mutations of transforming growth factor-beta genes reveal important roles in mouse development and adult homeostasis. Eur J Biochem 2000 267:6982-6988[Medline]
  2. Chang H, Lau AL, Matzuk MM. Studying TGF-beta superfamily signaling by knockouts and knockins. Mol Cell Endocrinol 2001 180:39-46[CrossRef][Medline]
  3. Ueno S, Kuroda T, Maclaughlin DT, Ragin RC, Manganaro TF, Donahoe PK. Mullerian inhibiting substance in the adult rat ovary during various stages of the estrous cycle. Endocrinology 1989 125:1060-1066[Abstract]
  4. Elvin JA, Yan C, Matzuk MM. Oocyte-expressed TGF-beta superfamily members in female fertility. Mol Cell Endocrinol 2000 159:1-5[CrossRef][Medline]
  5. Findlay JK, Drummond AE, Dyson M, Baillie AJ, Robertson DM, Ethier JF. Production and actions of inhibin and activin during folliculogenesis in the rat. Mol Cell Endocrinol 2001 180:139-144[CrossRef][Medline]
  6. Juneja SC, Chegini N, Williams RS, Ksander GA. Ovarian intrabursal administration of transforming growth factor beta 1 inhibits follicle rupture in gonadotropin-primed mice. Biol Reprod 1996 55:1444-1451[Abstract]
  7. Wrana JL, Attisano L, Wieser R, Ventura F, Massagué J. Mechanism of activation of the TGF-ß receptor. Nature 1994 370:341-347[CrossRef][Medline]
  8. Heldin CH, Miyazono K, ten Dijke P. TGF-beta signalling from cell membrane to nucleus through SMAD proteins. Nature 1997 390:465-471[CrossRef][Medline]
  9. Attisano L, Lee-Hoeflich ST. The Smads. Genome Biol 2001 2:1-8
  10. Ghiglieri C, Khatchadourian C, Tabone E, Hendrick JC, Benahmed M, Ménézo Y. Immunolocalization of TGF-ß1 and TGF-ß2 in the mouse ovary during gonadotropin-induced follicular maturation. Hum Reprod 1995 10:2115-2119[Abstract/Free Full Text]
  11. Hernandez ER, Hurwitz A, Payne DW, Dharmarajan AM, Purchio AF, Adashi EY. Transforming growth factor-ß1 inhibits ovarian androgen production: gene expression, cellular localization, mechanism(s), and site(s) of action. Endocrinology 1990 127:2804-2811[Abstract]
  12. Mulheron GW, Danielpour D, Schomberg DW. Rat thecal/interstitial cell express transforming growth factor-ß type 1 and 2, but only type 2 is regulated by gonadotropin in vitro. Endocrinology 1991 129:368-374[Abstract]
  13. Teerds KJ, Dorrington JH. Immunohistochemical localization of transforming growth factor-ß1 and ß2 during follicular development in the adult rat ovary. Mol Cell Endocrinol 1992 84:R7-R13[CrossRef][Medline]
  14. Levacher C, Gautier C, Saez JM, Habert R. Immunohistochemical localization of transforming growth factor beta 1 and beta 2 in the fetal and neonatal rat ovary. Differentiation 1996 61:45-51[CrossRef][Medline]
  15. Roy SK, Ogren C, Roy C, Lu B. Cell-type localization of transforming growth factor-ß2 and transforming growth factor-ß1 in the hamster ovary: differential regulation by follicle-stimulating hormone and luteinizing hormone. Biol Reprod 1992 46:595-606[Abstract]
  16. Roy SK, Hughes J. Ontogeny of granulosa cells in the ovary: lineage-specific expression of transforming growth factor-ß2 and transforming growth factor-ß1. Biol Reprod 1994 51:821-830[Abstract]
  17. Massagué J, Blain SW, Lo RS. TGF-ß signaling in growth control, cancer and heritable disorders. Cell 2000 103:295-309[CrossRef][Medline]
  18. Schmid P, Cox D, Van Der Putten H, McMaster GK, Bilbe G. Expression of TGF-ßs and TGF-ß type ll receptor mRNAs in mouse folliculogenesis: stored maternal TGF-ß2 message in oocytes. Biochem Biophys Res Commun 1994 201:649-656[CrossRef][Medline]
  19. Li M, Li J, Hoodless PA, Tzukazaki T, Wrana JL, Attisano L, Tsang BK. Mothers against decapentaplegic-related protein 2 expression in avian granulosa cells is up-regulated by transforming growth factor beta during ovarian follicular development. Endocrinology 1997 138:3659-3665[Abstract/Free Full Text]
  20. Osterlund C, Fried G. TGFbeta receptor types I and II and the substrate proteins Smad 2 and 3 are present in human oocytes. Mol Hum Reprod 2000 6:498-503[Abstract/Free Full Text]
  21. Drummond AE, Le MT, Ethier JF, Dyson M, Findlay JK. Expression and localization of activin receptors, Smads, and betaglycan to the postnatal rat ovary. Endocrinology 2002 143:1423-1433[Abstract/Free Full Text]
  22. Xu J, Oakley J, McGee EA. Stage-specific expression of Smad2 and Smad3 during folliculogenesis. Biol Reprod 2002 66:1571-1578[Abstract/Free Full Text]
  23. Kano K, Notani A, Nam SY, Fujisawa M, Kurohmaru M, Hayashi Y. Cloning and studies of the mouse cDNA encoding Smad3. J Vet Med Sci 1999 61:213-219[CrossRef][Medline]
  24. Liu X, Andoh K, Abe Y, Kobayashi J, Yamada K, Mizunuma H, Ibuki Y. A comparative study on transforming growth factor-beta and activin A for preantral follicles from adult, immature, and diethylstilbestrol-primed immature mice. Endocrinology 1999 140:2480-2485[Abstract/Free Full Text]
  25. Dorrington JH, Chuma AV, Bendell JJ. Transforming growth factor ß and follicle-stimulating hormone promote rat granulosa cell proliferation. Endocrinology 1988 123:353-359[Abstract]
  26. Roy SK. Transforming growth factor-ß potentiation of follicle-stimulating hormone-induced deoxyribonucleic acid synthesis in hamster preantral follicles is mediated by a latent induction of epidermal growth factor. Biol Reprod 1993 48:558-563[Abstract]
  27. Saragueta PE, Lanuza GM, Baranao JL. Autocrine role of transforming growth factor beta 1 on rat granulosa cell proliferation. Biol Reprod 2002 66:1862-1868[Abstract/Free Full Text]
  28. Magoffin DA, Gancedo B, Erickson GF. Transforming growth factor-ß promotes differentiation of ovarian theca-interstitial cells but inhibits androgen production. Endocrinology 1989 125:1951-1958[Abstract]
  29. Mulheron GW, Schomberg DW. Rat granulosa cells express transforming growth factor-ß type 2 messenger ribonucleic acid which is regulatable by follicle-stimulating hormone in vitro. Endocrinology 1990 126:1777-1779[Abstract]
  30. Magoffin DA, Hubert-Leslie D, Zachow R. Estradiol-17ß, insulin-like growth factor-I, and luteinizing hormone inhibit secretion of transforming growth factor ß by rat theca-interstitial cells. Biol Reprod 1995 53:627-635[Abstract]
  31. Chomczynski P, Sacchi N. Single-step method of RNA isolation by acid guanidium thiocyanate-phenol-chloroform extraction. Anal Biochem 1987 162:156-159[Medline]
  32. Greenwald GS, Roy SK. Follicular development and its control. In: Knobil E, Neill JD (eds.), Physiology of Reproduction, vol. 1, 2nd ed. New York: Raven Press; 1994:629–724
  33. May JV, Turzcynski CJ, Ramos L, Mau YH. Differential involvement of protein kinase C in the regulation of transforming growth factor-beta (TGF-beta) secretion by porcine theca and granulosa cells in vitro. Endocrinology 1995 136:1319-1322[Abstract]
  34. Kingsley-Kallesen ML, Kelly D, Rizzino A. Transcriptional regulation of the transforming growth factor-beta2 promoter by cAMP-responsive element-binding protein (CREB) and activating transcription factor-1 (ATF-1) is modulated by protein kinases and the coactivators p300 and CREB-binding protein. J Biol Chem 1999 274:34020-34028[Abstract/Free Full Text]
  35. Wilder PJ, Desler MM, Nowling TK, Rizzino A. Isolation and characterization of the murine transforming growth factor-beta2 promoter. Gene 2001 270:201-209[CrossRef][Medline]
  36. Boussouar F, Grataroli R, Ji J, Benahmed M. Tumor necrosis factor-a stimulates lactate dehydrogenase A expression in porcine cultured Sertoli cells: mechanisms of action. Endocrinology 1999 140:3054-3062[Abstract/Free Full Text]
  37. Benbrahim-Tallaa L, Tabone E, Tosser-Klopp G, Hatey F, Benahmed M. Glutathione S-transferase alpha expressed in porcine Sertoli cells is under the control of follicle-stimulating hormone and testosterone. Biol Reprod 2002 66:1734-1742[Abstract/Free Full Text]
  38. Kim SJ, Angel P, Lafyatis R, Hattori K, Kim KY, Sporn MB, Karin M, Roberts AB. Autoinduction of transforming growth factor beta 1 is mediated by the AP-1 complex. Mol Cell Biol 1990 10:1492-1497[Abstract/Free Full Text]
  39. Moreland RB, Gupta S, Goldstein I, Traish A. Cyclic AMP modulates TGF-beta 1-induced fibrillar collagen synthesis in cultured human corpus cavernosum smooth muscle cells. Int J Impot Res 1998 10:159-163[CrossRef][Medline]
  40. Ji C, Casinghino S, Chang D, Chen Y, Javed A, Ito Y, Hiebert SW, Lian JB, McCarthy TL, Centrella M. CBFa (AML/PEBP2)-related elements in the TGF-ß type I receptor promoter and expression with osteoblast differentiation. J Cell Biol 1998 69:353-363
  41. Bloom BB, Humphries DE, Kuang PP, Fine A, Goldstein RH. Structure and expression of the promoter for the R4/ALK5 human type I transforming growth factor-beta receptor: regulation by TGF-beta. Biochim Biophys Acta 1996 1312:243-248[Medline]
  42. Kleeff J, Korc M. Up-regulation of transforming growth factor (TGF)-ß receptors by TGF-ß1 in COLO-357 cells. J Biol Chem 1998 273:7495-7500[Abstract/Free Full Text]
  43. Ariazi EA, Satomi Y, Ellis MJ, Haag JD, Shi W, Sattler CA, Gould MN. Activation of the transforming growth factor beta signaling pathway and induction of cytostasis and apoptosis in mammary carcinomas treated with the anticancer agent perillyl alcohol. Cancer Res 1999 59:1917-1928[Abstract/Free Full Text]
  44. Imamura T, Takase M, Nishihara A, Oeda E, Hanai J, Kawabata M, Miyazono K. Smad 6 inhibits signalling by the TGF-beta superfamily. Nature 1997 389:622-626[CrossRef][Medline]
  45. Afrakhte M, Moren A, Jossan S, Itoh S, Sampath K, Westermark B, Heldin CH, Heldin NE, ten Dijke P. Induction of inhibitory Smad6 and Smad7 mRNA by TGF-beta family members. Biochem Biophys Res Commun 1998 249:505-511[CrossRef][Medline]
  46. Topper JN, Cai J, Qiu Y, Anderson KR, Xu YY, Deeds JD, Feeley R, Gimeno CJ, Woolf EA, Tayber O, Mays GG, Sampson BA, Schoen FJ, Gimbrone MA Jr, Falb D. Vascular MADs: two novel MAD-related genes selectively inducible by flow in human vascular endothelium. Proc Natl Acad Sci U S A 1997 94:9314-9319[Abstract/Free Full Text]
  47. Stopa M, Benes V, Ansorge W, Gressner AM, Dooley S. Genomic locus and promoter region of rat Smad7, an important antagonist of TGFbeta signaling. Mamm Genome 2000 11:169-176[CrossRef][Medline]
  48. Ishida W, Hamamoto T, Kusanagi K, Yagi K, Kawabata M, Takehara K, Sampath TK, Kato M, Miyazono K. Smad6 is a Smad1/5-induced Smad inhibitor. Characterization of bone morphogenetic protein-responsive element in the mouse Smad6 promoter. J Biol Chem 2000 275:6075-6079[Abstract/Free Full Text]
  49. Johnsen SA, Subramaniam M, Janknecht R, Spelsberg TC. TGFbeta inducible early gene enhances TGFbeta/Smad-dependent transcriptional responses. Oncogene 2002 21:5783-5790[CrossRef][Medline]
  50. Johnsen SA, Subramaniam M, Katagiri T, Janknecht R, Spelsberg TC. Transcriptional regulation of Smad2 is required for enhancement of TGFbeta/Smad signaling by TGFbeta inducible early gene. J Cell Biochem 2002 87:233-241[CrossRef][Medline]
  51. Shull MM, Doetschman T. Transforming growth factor-ß1 in reproduction and development. Mol Reprod Dev 1994 39:239-246[CrossRef][Medline]
  52. Feng P, Catt KJ, Knecht M. Transforming growth factor-beta stimulated meiotic maturation of the rat oocyte. Endocrinology 1988 122:181-186[Abstract]
  53. Tsafriri A, Vale W, Hsueh AJW. Effects of transforming growth factors and inhibin-related proteins on rat preovulatory graafian follicles in vitro. Endocrinology 1989 125:1857-1862[Abstract]



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