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. 2008 Oct 9;150(2):906–914. doi: 10.1210/en.2008-0880

Presence, Actions, and Regulation of Myostatin in Rat Uterus and Myometrial Cells

Pasquapina Ciarmela 1, Ezra Wiater 1, Sean M Smith 1, Wylie Vale 1
PMCID: PMC2732292  PMID: 18845635

Abstract

Myostatin, a member of the TGF-β superfamily of proteins, is known to suppress skeletal muscle mass and myocyte proliferation. The muscular component of the uterus is the myometrium, a tissue that regulates its mass in response to different physiological conditions under the influence of sex steroids. Recently, our laboratory reported effects of activin-A, another TGF-β family member, on signalling and proliferation of rat uterine explants and human myometrial cell lines in culture. Here, we explore the expression, actions, and regulation of myostatin in uterine smooth muscle. Myostatin mRNA was demonstrated to be expressed in a myometrial cell line, pregnant human myometrial 1 cell line (PHM1). Functional assays showed that myostatin induced phosphorylation of Smad-2 and reduced proliferation of PHM1 number in a time and dose-dependent manner. Furthermore, myostatin activated smad-2 specific signalling pathways in rat uterine explants. To expand on our in vitro findings, we found that myostatin is expressed in rat uterus and determined that myostatin mRNA expression varies as a function of the phase of the estrous cycle. Uterine levels of myostatin peaked during late estrus and were the lowest at proestrus. Ovariectomy increased myostatin expression; estrogen treatment strongly decreased myostatin levels, whereas progesterone weakly decreased myostatin expression. In conclusion, myometrial cells are myostatin sensitive, myostatin mRNA levels are modulated in vivo in rats during the estrous cycle, and in response to steroid deprivation and replacement.

Myometrial cells are myostatin-sensitive; myostatin mRNA levels are modulated in rats during the estrous cycle and in response to steroid deprivation and replacement.

Myostatin is a highly conserved TGF-β family member that functions as an endogenous inhibitor of muscle growth in diverse species (1,2,3,4,5,6). Targeted disruption of the myostatin gene in mice leads to muscle hypertrophy and hyperplasia with an approximate doubling of muscle mass (3). Similarly, natural myostatin mutations in cattle have been associated with a “double-muscling” phenotype (1,2,4). Most recently, a new mutation in myostatin has been found in the whippet breed of dog that results in a double-muscled phenotype known as the “bully” whippet (6). The function of myostatin as a negative regulator of muscle growth has been conserved also in humans through the identification of a hyper-muscular child with a loss-of-function mutation in the myostatin gene (5).

The myostatin gene is predominantly expressed in cells of skeletal muscle lineage throughout embryonic development as well as in adult animals (1,2,3,4,5,7). Myostatin is also expressed in cardiac myocytes (8) and in placenta (9).

The muscular component of the uterus is the myometrium. The myometrium undergoes changes in size under the influence of steroid hormones, during each menstrual cycle (10), throughout pregnancy (11,12), and menopause (13). Myometrial mass is modified also in tumoral conditions such as leiomyoma and leiomyosarcoma (14,15). The leiomyoma is a benign form but highly diffuse and with high morbidity. The leiomyosarcoma is rare, but malignant (16). Changes in myometrium mass have been regulated by a wide array of endocrine and paracrine factors (17,18,19,20,21,22).

Activin-A, another TGF-β family member, exerts a negative regulatory role on myometrial cell proliferation (23). Myostatin and activin-A share some receptors and signaling transduction molecules. Signaling events, initiated in response to activin, require binding to two types of transmembrane serine/threonine receptor kinases classified as type II [activin receptor type (ActR) II or ActRIIB)] and I [activin receptor-like kinase (ALK) 4]. Activin type II receptors are the primary ligand binding proteins. ActRII binds activin with high affinity in the absence of type I receptors (24). In the receptor complex, the constitutively active type II receptor kinase phosphorylates ALK4, and this phosphorylation leads to activation of the ALK4 kinase (25). Once activated, ALK4 phosphorylates Smad-2, which forms part of the post-receptor signal transduction system (26). Myostatin, like activin, stimulates target cells by assembling a cell surface receptor complex containing type I and II receptors. Myostatin binds the type II Ser/Thr kinase receptor, ActRIIB, and then partners with a type I receptor, either ALK4 (or ActRIB) or ALK5 (TGFβRI). These complexes induce phosphorylation of Smad-2 and activate an activin/TGF-β-like signaling pathway (27).

We hypothesized that myostatin is a uterine factor able to influence myometrial mass and cell proliferation. In this report we investigated the presence of myostatin in myometrial cells and explored possible functions on myometrial cells, including the ability to initiate downstream signaling and regulate cell proliferation. We have also investigated the changes of myostatin mRNA levels in rat uterus during the estrous cycle, and in response to steroid deprivation and administration.

Materials and Methods

Materials

NuPAGE gels, molecular weight standards, CyQUANT Cell Proliferation Assay kit, pcDNA3.1 plus hygro expression vector were from Invitrogen Life Technologies, Inc. (Carlsbad, CA). Phospho-Smad-2 (Ser465/467) antibody (catalog no. 3101) was purchased from Cell Signaling, Inc. (Danvers, MA). The rabbit polyclonal Smad-2–3 antibodies were raised against a peptide conserved between smad-2 and smad-3 spanning amino acids 159-175 of human smad-3, and were produced by Joan Vaughan (Peptide Biology Laboratories, Salk Institute). β-Estradiol 3-benzoate, progesterone (P), sesame oil, high-molecular weight polyethyleneimine (PEI), and FLAG-tag antibody (M2; catalog no. A2220) were purchased from Sigma-Aldrich Corp. (St. Louis, MO). Pregnant human myometrial 1 cell line (PHM1) (28) was generously provided by Dr. Barbara Sanborn (Colorado State University, Fort Collins, CO).

Tissue culture

PHM1 cells were grown on tissue culture-treated plastic (Corning, Inc., Corning, NY) in 5% CO2 in a 37 C humidified incubator and were cultured in high-glucose DMEM (Salk Institute, La Jolla, CA) supplemented with 10% fetal bovine serum (FBS), 1% penicillin/streptomycin glutamine, and 0.1 mg/ml Geneticin (Invitrogen Life Technologies).

Myostatin production and purification

The full-length human myostatin cDNA was amplified, and the FLAG-tag was incorporated in the mature region with an overlapping PCR strategy. The gel-purified PCR product was cloned in pcDNA 3.1 plus hygro vector (Invitrogen Life Technologies). The DNA was fully sequenced before use for transfection. Human embryonic kidney (HEK) 293T cells were grown in 5% CO2 in a 37 C humidified incubator in complete DMEM containing 10% bovine calf serum. Transfection of FLAG-tagged myostatin construct was performed using a modified high-molecular weight PEI protocol, as previously described (29). Briefly, HEK293T were plated at in 10-cm plates coated with poly-d-lysine. After recovery overnight, cells were transfected with a total of 12 μg DNA. DNA was diluted in 600 μl serum-free DMEM. PEI (frozen as a 10 mg/ml stock) was diluted to 1 mg/ml in H2O and added to the DNA solution to a final concentration of 20 μg/ml. The DNA/PEI solution was vortexed and incubated at room temperature for 10 min. During this incubation, media were removed from the plates, and 5.4 ml serum-free DMEM was added to each well. After 10 min, the DNA-PEI complexes were added directly to the plates and incubated for 4 h at 37 C in 5% CO2. After 4 h, plates were aspirated, and 7 ml DMEM containing 10% FBS was added. Plates were incubated at 37 C in 5% CO2 to allow cells to recover and express protein for 48 h before assay. A stable transfected cell line was generated using 200 μg/ml hygromycin treatment. FLAG-myostatin stable cell line was cultured in freestyle serum free expressing media (Invitrogen Life Technologies). Crude media were filtered through a 5-μm nylon filter to separate cell debris. A total of 1 m 2-N-M-Morpholino-ethanesulfonic acid buffer (pH 6.2) was added to the filtrate up to a final concentration of 50 mm, along with 0.5 ml M2 anti-FLAG-agarose beads suspension (Sigma-Aldrich). After shaking overnight at 4 C, the proteins bounded were elute with glycine-HCl (pH 2.8) and neutralized with Tris-HCl. The elute was subjected to HPLC purification. A reverse-phase HPLC C4 column was used, and the elution was done with trifluoroacetic acid/acetonitrile (CH3CN) gradient. Different fractions were collected, and the protein identity was tested by silver staining and Western blotting with flag antibody. Myostatin protein activity was tested using both its 3TP and p15(ink4B) promoter reporter luciferase activity in HepG2 cell line.

Protein extraction and immunoblotting

For immunoblotting assays, PHM1 cells were treated with myostatin for 30 min or left untreated. The immunoblotting was performed as previously described (23). Briefly, the cellular extract was solubilized in ice-cold radioimmunoprecipitation assay buffer [50 mm Tris-HCl (pH 7.4), 150 mm NaCl, 1 mm EDTA, 1% Igepal CA-630, and 0.25% Na-deoxycholate] supplemented with protease inhibitors (set III; Calbiochem, San Diego, CA) and phosphatase inhibitors (set II; Calbiochem). Samples were rocked at 4 C for 30 min, and insoluble cellular materials were removed by centrifugation. Rat uteri (collected randomly in animals during the estrous cycle) were rapidly excised, cut into fragments, washed in Hepes dissociation buffer, and treated with or without myostatin for 45 min in 200 μl DMEM supplemented with 10% FBS, 1% penicillin/streptomycin glutamine. Uteri fragments were flash frozen on dry ice, stored at −20 C, until crushed while frozen, and suspended in 200 μl ice-cold radioimmunoprecipitation assay buffer supplemented with protease and phosphatase inhibitors. Samples were rocked at 4 C for 30 min, and insoluble cellular materials were removed by centrifugation.

Soluble protein was quantified using a Bradford protein assay method (Bio-Rad Laboratories, Inc., Hercules, CA) with absorbance measured at 595 nm. Equal amounts of proteins were loaded onto 10% NuPAGE sodium dodecyl sulfate gels (Invitrogen Life Technologies) and resolved by SDS-PAGE under reducing conditions. Proteins were transferred in 0.2-μm nitrocellulose membranes. The membranes were blocked with 5% (wt/vol) nonfat milk powder in 50 mm Tris-HCl (pH 7.4), 150 mm NaCl, and 0.05% Tween 20, incubated overnight with 1:1,000 dilutions of primary antibodies against phosphorylated-Smad-2, or Smad-2/3 as noted and incubated with 1:10,000 dilutions of horseradish peroxidase-conjugated antirabbit IgG (Pierce, Rockford, IL) for 2 h. Immunoreactive proteins were visualized using Super Signal West Pico chemiluminescent substrate (Pierce).

Cell proliferation assays

Cellular growth curves were measured using the CyQUANT Cell Proliferation Assay kit according to the manufacturer’s instructions and at the condition previously set up for PHM1 cells (23). Briefly, PHM1 cells were seeded in 96-well plates at initial densities of 103 cells per well in a total volume of 100 μl DMEM supplemented with 10% FBS and doses of myostatin as noted. Cells were allowed to divide for the number of days indicated, with cellular growth media being replaced every 3 d, maintaining original serum concentrations and myostatin doses. At the indicated times, media were discarded, and plates were frozen. The day of the assay, plates were thawed, cells were lysed, and total cellular nucleic acid was measured using florescence at 520 nm emission after excitation at 480 nm.

Animals

Adult female (200–220 g) Sprague Dawley rats (Harlan Sprague Dawley, Indianapolis, IN) were used in experiments described in this study. Animals were maintained on a 12-h light, 12-h dark cycle (lights on at 0600 h), and provided rat chow (Harlan-Teklad, Madison, WI) and water ad libitum. The Salk Institute animal use and care committee approved all procedures described in this study.

Estrous cycle studies

Vaginal smears were examined daily between 0800 and 1000 h; animals were chosen for further examination only if they had exhibited at least three consecutive 4-d estrous cycles. The stage of cycle was determined by the characteristic presence or absence of inflammatory leukocytes, and the histological appearance of epithelial cells. Rats were killed by rapid decapitation, and the blood was collected from the trunk and successively used to validate the cycle stage by estrogen and P measurement.

Rat ovariectomy (OVX) and steroid replacement

Sprague Dawley rats were bilaterally ovariectomized or exposed to sham surgical procedure.

The ovaries were removed from adult female rats under isoflurane anesthesia using a dorsal approach. After an incision had been made in the skin, the body wall was pierced superficially to the position of the organ to be removed. To prevent excessive bleeding, ovaries were removed after the uterine horn was tied off. Ovariectomized animals were allowed to recover for 10 d before receiving hormone injections. Animals were injected sc with hormones dissolved in 200 μl sesame oil (Sigma-Aldrich). Animals were treated with either E (50 μg/rat β-estradiol 3-benzoate; Sigma-Aldrich), P (4 mg /rat; Sigma-Aldrich), both E plus P, and only sesame oil (no treatment, NT) at 1000 h for 2 or 4 consecutive days. Subsets of animals were implanted at ovariectomization with 1 mg E pellet (Hormone Pellet Press, Catalina, KS). After removal of the ovaries, a small incision was made on the dorsal aspect of the animal, and hormone pellets were deposited in the sc space between the shoulder pads. After surgery, animals were allowed to recover for 2 wk. Two weeks after surgery, rats were killed by rapid decapitation, blood was collected from the trunk, and the uterus was saved for further analyses.

Tissue dissection

Samples of uteri were quickly excised, weighed, cooled in dry ice, and stored at −80 C for subsequent analyses. Samples of red and white muscle were obtained from dissection of gastrocnemius muscles that were quickly excised from the hindlimb of animals and dissected following the indications given by muscle color (corresponding to myoglobin concentration) into a red portion (derived from the deep part of the lateral head) and a white portion (derived from the superficial part of both lateral and medial head) (30).

RIA

Blood was collected into chilled polystyrene tubes containing EDTA and centrifuged. The resulting plasma was aspirated and stored frozen in plastic tubes at −20 C until assay for estradiol or P. Plasma estradiol and P were determined by RIA using a kit based on antibody coated tube methodology (Diagnostic Products Corp., Los Angeles, CA).

RNA extraction, PCR, and quantitative real-time PCR

Total RNA was extracted from PHM1 cells using an RNeasy Mini Kit (QIAGEN, Inc., Valencia, CA) according to the manufacturer’s instructions. Deoxyribonuclease (DNase) treatment was performed on-column using a ribonuclease-free DNase Kit (QIAGEN) according to the manufacturer’s instructions. For rat uteri and skeletal muscle tissues, total RNA was extracted using TRIzol reagent (Invitrogen Life Technologies) according to the manufacturer’s instructions. Samples were digested with a ribonuclease-free DNase (Promega Corp., Madison, WI), and the RNA was cleaned up and concentrated using an RNeasy Micro kit (QIAGEN). For qualitative PCR-RT, reactions were performed on 1 μg total RNA using SuperScript II (Invitrogen Life Technologies) according to the manufacturer’s instructions. Sequences of primers used with PHM1 cells were TTCGTCTGGAAACAGCTCCT (forward) and CATTTGGGTTTTCCATCCAC (reverse) for myostatin and GGTCTTGCCCATCTTCACAT (forward) and TCAGGGGCCATGTACCTTTT (reverse) for Alk 5, giving respectively a PCR product of 220 and 212 bp. Sequences of primers used with rat uteri were previously described for myostatin (31), and were AGTCTTCGGACGCAAGAAAA (forward) and AGCCACCAGAGCTTTTGAGA (reverse) for ribosomal protein S16, giving respectively a PCR product fragment of 223 and 250 bp. There were 35 cycles performed for 1 min at 94 C, 1 min at 55 C, and 1 min at 72 C, and amplification products were visualized on 2% agarose gel and stained with ethidium bromide. For newly designed primer pairs, computer analysis was performed to compare the selected primers with all human or rat sequences in the gene database of the National Center for Biotechnology Information using Basic Alignment Search Tool BLAST (32) to exclude homology to other genes. For PCR to detect expression, control reactions were performed omitting either the reverse transcriptase (RT) enzyme or template RNA to test for contamination with genomic DNA or nonspecific amplification. For real-time PCR performed to measure myostatin hypoxanthine phosphoribosyltransferase (HPRT) and activin β-A/HPRT expression level changes during the estrous cycle and in response to steroids treatment, we performed the RT using 1 μg RNA obtained with the high-capacity cDNA RT kit (Applied Biosystems, Foster City, CA), and we performed the TaqMan real-time PCR for all the genes. We used TaqMan gene expression assay Rn00569683_m1, Rn01538592_m1, and Rn01527840_m1 (Applied Biosystems), respectively, for myostatin, activin β-A, and HPRT, performing a fast thermal cycle protocol (initial denaturation at 95 C for 20 sec, followed by 40 cycles of 95 C for 1 sec and 60 C for 20 sec) in the ABI PRISM 7900 instrument (Applied Biosystems) using 50 ng cDNA in a final reaction volume of 10 μl. The standard curve for myostatin was done using RNA extracted from white muscle to have a wider range of the serial dilution. The blank for each reaction, consisting of amplifications performed in the absences of RT enzyme, was performed.

Data analysis

Data are presented as the mean ± sem, and a two-tailed t test was used for data analysis. Differences were considered significant when P < 0.05. The real time PCRs were done in triplicate for the cell proliferation assay (n = 6) and in all the in vivo experiments (n = 4). Experiments were repeated either two or three times.

Results

Expression and purification of FLAG-tagged myostatin

To explore the effects of myostatin on myometrial cells, we cloned the full-length human myostatin cDNA with a FLAG tag into a pcDNA 3.1 plus hygro vector. Myostatin cDNA constructs were transfected into HEK293T cells to produce the protein. Myostatin expression was detected, under reducing conditions, as a 50-kDa band in cell extracts and as a 14-kDa band in the media of HEK293T cells, using a specific FLAG antibody (Fig. 1A). Immunoprecipitation of cell lysates with anti-FLAG-agarose beads identified bands of 50 and 14 kDa (Fig. 1B and lane 1 of Fig. 1D), whereas silver staining revealed other bands that did not interact with FLAG antiserum with an approximate size of 62 and 38 kDa. The bands at 38 and at 14 kDa correspond to the myostatin pro-peptide and mature form, respectively (33). We further purified myostatin proteins using HPLC separation, and examined the purity of fractions with silver staining (Fig. 1C) and Western blot analysis (Fig. 1D). Only the fractions containing the mature form (14 kDa) were used in the assays described in this study.

Figure 1.

Figure 1

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Analysis of human recombinant FLAG-tagged myostatin protein produced in HEK293T cells. Western blotting with FLAG antibody of cellular extract and culture media (A) and after immunoprecipitation (IP) (B) with anti-FLAG-agarose beads. Silver stained gel (C) and Western blotting with FLAG antibody (D) after HPLC separation. Lane 1, Sample before HPLC. Lanes 2–8, Different HPLC fractions. M: SeeBlue Plus2 Pre-Stained Standard for protein bands (Invitrogen Life Technologies).

Expression and functional activity of myostatin pathway components in myometrial cells

To examine if myostatin has a potential biological role in the myometrium, we first established the expression of this protein and the type I receptor, Alk 5, in PHM1 cells by RT-PCR. We have previously shown the presence of other components, including ActRIIA, ActRIIB, ALK 4, and Smad-2/3 in this cell line (23). As shown in Fig. 2, primer pairs spanning intron/exon junctions gave amplicons of predicted size corresponding to myostatin (Fig. 2B) and Alk 5 (Fig. 2A). Control experiments omitting RT did not result in amplification of products, confirming that we detected expression of processed mRNA (Fig. 2). To test the functional roles of these transcripts, we determined whether myostatin induced detectable Smad signaling in PHM1 cells. Myostatin treatment induced Smad-2 phosphorylation in PHM1 cells (Fig. 3A, upper panel), whereas the total level of Smad-2 and Smad-3 (Fig. 3A, lower panel) were not altered. Myostatin-induced phosphorylation of Smad-2 was also observed in uterine explants (Fig. 3B), suggesting that myostatin is able to initiate a signaling cascade in this tissue.

Figure 2.

Figure 2

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Expression of myostatin and its receptor in myometrial cells. Electrophoretic gel of the RT-PCR in PHM1 cells for the receptor ALK 5 (A) and myostatin (B) mRNAs. −RT consists of amplifications performed in absences of RT enzyme. The molecular DNA mass marker (M) is Hi-Lo ladder (Minnesota Molecular, Minneapolis, MN).

Figure 3.

Figure 3

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Myometrial responsiveness to myostatin (M). Myostatin induces phosphorylation (p) of Smad-2 in PHM1 (A) and in rat uterus explants (B). The levels of Smad-2 and -3, showed in the lower panels, instead resulted unchanged.

Myostatin inhibition of myometrial cell growth

Myostatin is a negative regulator of skeletal muscle cell proliferation and was hypothesized to produce similar effects on myometrial cells. To determine the effects of myostatin on myometrial cell proliferation, we examined PHM1 culture with a florescence-based assay (CyQUANT) that correlates with cell number. PHM1 cell growth media were left untreated or supplemented with 1 or 10 nm myostatin, and media were changed every 3 d. Cells were allowed to proliferate for 12 or 20 d, and then plates were assayed for total cell number. Myostatin treatment decreased cell number compared with untreated controls at both time points analyzed (Fig. 4A). Cell number between untreated cells and cells treated with 1 or 10 nm myostatin was statistically significant after both 12 and 20 d treatment (P < 0.05). To determine the potency of myostatin as myometrial growth inhibitor, we performed similar experiments using a range of myostatin doses. Florescence measurements after 20 d revealed that the inhibition of growth of PHM1 cells by myostatin was dose dependent, with an EC50 of approximately 1.1 nm (Fig. 4B), comparable to other reported myostatin responses (34).

Figure 4.

Figure 4

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Effect of myostatin on myometrial cell proliferation. A, Changes in PHM1 cell numbers after 12 and 20 d with (1 and 10 nm) and without (NT) myostatin treatment. B, Dose response of myostatin effect on PHM1 cell proliferation after 20 d treatment. arb, Arbitrary; flu, fluorescence.

Expression of myostatin mRNA in rat uterus: changes along the estrous cycle and correlation with activin-A mRNA levels

To address the question if myostatin is expressed in the uterine tissues, we examined myostatin mRNA expression using real-time PCR in female rats at multiple phases of the estrous cycle. To determine the stage of the estrous cycle, uterine weight, cell type, and serum steroid levels were determined for each animal. The proestrous-early estrous (PE) stage is characterized by increased uterine weight and high estrogen blood levels. In the postestrus, when the estrogen levels decrease, the uterus shrinks. The uterus remains small in diestrus (D), the quiescent period with uterine slow growth characterized by low steroid levels (Table 1) (35). We determined that uterine expression of myostatin changes during the estrous cycle with higher expression in postestrus, low expression in D, and almost no expression in PE (Fig. 5A). In a parallel experiment, in which the endometrial portion was excised from the tissue specimens, and the myometrial content was concentrated, there was a similar pattern during the estrous cycle, but overall expression of myostatin was higher. Because this expression pattern seemed to be opposite the one described for activin-A (36), a member of the same family that shares some receptors and signaling pathway, we checked the activin-A expression in the same samples. Interestingly, activin-A and myostatin exhibit an almost reciprocal expression pattern. Activin-A expression increased greater than 2-fold from the D to proestrous-estrous phases, thereafter levels decreased in the postestrous phase (Fig. 5B).

Table 1.

Value of uterine weight, uterine weight normalized with body weight, E, and P blood levels in cycling rats

Uterus weight (mg) Uterus/body weight (mg/g) E blood level (pg/ml) P blood level (ng/ml)
D 190.75 ± 20.3 0.83 ± 0.08 10.9 ± 3.2 10.14 ± 2.2
PE 285.2 ± 14 1.1 ± 0.2 29.2 ± 4 6.4 ± 2
Postestrus 154 ± 20 0.65 ± 0.1 9.8 ± 0.2 27.6 ± 7.1
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Values are presented as the mean ± sem. 

Figure 5.

Figure 5

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Expression of myostatin and activin-A in rat uterus along the estrous cycle. Results of real-time PCR for myostatin (A) and activin-A (B) in total rat uterus (light gray bars), and in myometrial-enriched specimens (dark gray bars) during the estrous cycle. D, PE, and postestrous (P) phases are considered. All mRNAs are normalized with the housekeeping gene HPRT.

Regulation of myostatin mRNA expression by steroids

To investigate the role of steroids in the regulation of myostatin expression, we performed OVX and steroid replacement experiments in female rats. The effects of OVX and steroid injection on uterine weight, uterine weight normalized with body weight, and resulting blood levels of gonadal steroids are summarized in Table 2. Because we obtained the expected effects on gross uterine morphology and steroid blood levels, we proceeded with the evaluation of myostatin expression. Electrophoretic analysis of RT-PCR products revealed a band corresponding to myostatin in ovariectomized (NT) animals and in P treated animals alone (Fig. 6A). Samples derived from animals treated with estrogen alone and in cotreatment with P (estrogen plus P) did not show any amplification products (Fig. 6A, upper panel). As a control, we verified that the housekeeping gene s16 was expressed in all the samples (Fig. 6A, lower panel). To perform a more precise analysis, quantitative real-time PCR with TaqMan chemistry was done. OVX increased myostatin expression. Real-time PCR data showed a relatively high expression in NT and P, and a very low expression (at the limit of detection) in estrogen and estrogen plus P treated rats. P treated animals showed less myostatin mRNA expression than NT, but this trend was not statically significant (Fig. 6B). Because estrogen injections increased circulating concentrations of the hormone above physiological levels (Table 1), we implanted a group of animals with 1 mg estrogen pellets. These animals had the estrogen levels around 40 pg/ml (47 ± 7), a concentration that decreases within the high physiological range of circulating hormone. In the uteri of these animals, myostatin mRNA was down-regulated, and the results are similar to the ones of the animals with much higher blood estrogen levels (data not shown). To test if the estrogen modulation of myostatin affected different types of muscles, we checked the myostatin expression in skeletal muscle of rats injected with estrogen. We examined myostatin expression in both red and white fibers of the gastrocnemius muscle. As previously reported (30), white fibers showed a much higher myostatin expression than red fibers. Interestingly, estrogen treatment did not affect myostatin expression in these tissues (Fig. 6C).

Table 2.

Effect of steroid injection on uterine weight, uterine weight normalized with body weight, and on E and P blood levels

Uterus weight (mg) Uterus/body weight (mg/g) E blood level (pg/ml) P blood level (ng/ml)
Oil 167.25 ± 11.7 0.7 ± 0.06 13.3 ± 7.2 1.9 ± 1.2
E 544.25 ± 83 2.44 ± 0.4 517.23 ± 135.2 4.2 ± 0.8
P 229.7 ± 84.8 1 ± 0.4 16.8 ± 8.3 24.36 ± 7.1
E + P 388.25 ± 17.6 1.7 ± 0.17 585.6 ± 117.4 53 ± 17.7
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Values are presented as the mean ± sem. 

Figure 6.

Figure 6

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Steroid regulation of gene expression. Representative electrophoretic gel of qualitative PCR (A, upper panel) and results of real-time PCR (B) for myostatin mRNA in ovariectomized rats after NT, or E and/or P injection. C, Real-time PCR results for myostatin mRNA in white and red fiber of skeletal muscle (gastrocnemius) in ovariectomized rat with (E) and without (NT) E injection. D, Results of real-time PCR for activin-A in ovariectomized and steroid-replaced rats. In all real-time PCRs, the mRNAs were normalized with the housekeeping gene HPRT. In A, the lower panel shows the electrophoretic gel of the housekeeping s16. The molecular DNA mass marker (M) is Hi-Lo ladder (Minnesota Molecular, Minneapolis, MN).

Data from estrous cycle experiments suggested that activin-A expression in uterus should be regulated differently from that of myostatin. Therefore, we also determined activin-A expression in uteri from ovariectomized and steroid-replaced rats. We observed an up-regulation of activin-βA subunit mRNA with estrogen treatment (Fig. 6D), and the estrogen effect was completely abrogated with P cotreatment. Interestingly, P treatment alone resulted in a slight, but not statistically significant, activin-β-A subunit mRNA down-regulation compared with the vehicle-treated animals. Similar results both for myostatin and activin-βA expression were obtained in the animals injected with steroids for 2 and 4 d.

Normal-cycling control rats that were sham operated resulted to be in D phase of the cycle. Myostatin and activin expression levels of these controls resulted similar to the cycling animals at the same phase of the cycle (data not shown).

Discussion

Targeted disruption of the myostatin gene in mice leads to muscle hypertrophy and hyperplasia with an approximate doubling of muscle mass (3). This increase seems to be the result of an increase in both the number (hyperplasia) and thickness (hypertrophy) of muscle fibers. These findings can be explained in part by the observation that myostatin can inhibit proliferation of myoblasts in culture (34,37).

Myostatin knockout mice have not been reported to have reproductive deficits, but a detailed analysis of uterine morphology and growth has not been performed in these animals (3). The myometrium is a dynamic tissue that undergoes changes in mass in response to different physiological and pathological conditions. Myostatin could represent a candidate protein that could play a role in regulating myometrial tissue. However, the expression and physiological effects of myostatin in uterus had not been investigated.

Here, we report that uterine tissues respond to myostatin, as measured by induction of phosphorylation of Smad-2. To ensure that myostatin induces intracellular transduction signaling in myometrial cells, we tested the induction of phosphorylation of Smad-2 also in PHM1. These cells, generated through expression of the E6 and E7 proteins of the human papilloma 16 virus in myometrial cells derived from a pregnant woman, retain a considerable array of morphological and biochemical properties of the parent cells (28). We also showed that myostatin reduces total PHM1 cell number in a time and dose-dependant manner, suggesting that myostatin may expert some functions in myometrial smooth muscle cells.

Because the uterine expression of myostatin has not been established, we tested the expression of myostatin mRNA in rat uterus. We demonstrated that uterus expresses myostatin and the levels of myostatin mRNA correspond to higher myometrial cell contents. Furthermore, we report that myostatin expression changes in a dynamic range during the estrous cycle, with higher expression in postestrus, low expression in D, and almost no expression in PE. These findings demonstrate that myostatin is expressed at higher levels only in some phases of the reproductive cycle. The pattern of myostatin expression that we observed during the estrous cycle suggested a regulation by steroid hormones, therefore, we examined myostatin expression in ovariectomized and steroid-replaced animals. We found that ovariectomized rats have higher myostatin expression than normal cycling rats and that estrogen treatment abrogates myostatin expression. Therefore, myostatin is not expressed in uterus in case of high estrogen levels, either pharmacologically induced or during physiological circumstances, such as the PE stage. Instead, myostatin expression is elevated in the presence of low steroid levels such as during the D, and this expression is even higher in the absence of steroids after OVX. In addition, myostatin expression correlates negatively with the growing phases of the uterus: its levels are low during the swelling, whereas its expression is high when the uterus shrinks.

Cellular proliferation and differentiation of the uterine tissues are considered to be regulated by ovarian steroids. The response of the uterine tissue to steroids is complex, involving many biochemical as well as morphological events resulting in uterine growth and differentiation (38,39,40). Although the most profound mitogenic effect of estrogen appears to be in the endometrial compartment, myometrium is also sensitive to the hypertrophic actions of steroids (38,39,41,42). The mitogenic action of steroids in their target tissues is considered to be mediated, with paracrine and/or autocrine mechanisms, through local production of growth factors (14,43). The modulation of growth factor expression by steroids suggests that proteins such as myostatin may be effectors of steroid hormone action. Myostatin could represent a growth factor that experts its action in myometrium in coordination with the other factors known to act on these cells, such as epidermal growth factor, platelet-derived growth factor (18), IGF (19), TGF-β (22), and activin-A (23). We demonstrate that rat uteri are able to respond to exogenous myostatin as assessed by increased Smad-2 phosphorylation, whereas detectable basal levels of phosphorylated Smad-2 are consistent with endogenous myostatin and/or activin affecting autocrine or paracrine signaling in this tissue. Clarifying the relative contribution of activin and myostatin to this Smad-2 phosphorylation must await improved assays to measure endogenous levels of activin and myostatin free of the myostatin pro-domain.

The roles of myostatin and activin could be distinguishable from one another because of differences in protein distribution and regulation. Although activin (23) and myostatin are both cytostatic on myometrial cells in vitro, our data also suggest that they are expressed and regulated differently. From observations of cycling rats, increased activin expression seems to correlate positively with the myometrial growth, whereas myostatin seems to correlate negatively with myometrial growth. In fact, it has been reported that the proliferative rate of rat myometrial cells display a single peak during proestrus, and a rapid decrease in postestrus (10), suggestive that the low myostatin levels may be causal, whereas the consequences of higher activin levels are either overridden by a lack of myostatin or have limited distribution.

To verify the involvement of myostatin and activin in the regulation of myometrial proliferation, further analysis of the expression patterns in different physiological and pathological conditions is needed. In particular, it is necessary to know how myostatin expression changes in the uterus when critical changes in mass occur, such as during pregnancy and in menopause. In pregnancy there is remarkable uterine growth, necessary to accommodate the growing fetus, whereas in menopause there is a uterine regression. It also would be interesting to test the involvement of myostatin in both benign (leiomyoma) and malignant (leiomyosarcoma) uterine smooth muscle tumors.

In conclusion, we present in vitro results demonstrating myostatin-induced myometrial cell responsiveness and reduction of cell proliferation rates. We also show that the expression of myostatin in uterus varies throughout the estrous cycle and in response to steroid hormones.

Acknowledgments

We gratefully acknowledge Drs. Barbara Sanborn and Chun-Ying Ku of Colorado State University for generously providing the pregnant human myometrial 1 cell line.

Footnotes

This project was supported by Award no. 5P01HD013527-28 from the National Institute of Child Health and Human Development. This research was supported in part by the Clayton Medical Research Foundation, Inc., and a grant from the Adler Foundation. W.V. is a Clayton Medical Research Foundation, Inc., Senior Investigator and is the Helen McLoraine Professor of Molecular Neurobiology.

The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institute of Child Health and Human Development or the National Institutes of Health.

Present address for P.C.: Institute of Normal Human Morphology, Faculty of Medicine, Polytechnic University of Marche, via Tronto 10/a, 60020 Ancona, Italy.

Disclosure Summary: W.V. is a cofounder, consultant, equity holder, and member of the Board of Directors of Neurocrine Biosciences, Inc. and Acceleron Pharma, Inc. P.C., E.W., and S.M.S. have nothing to declare.

First Published Online October 9, 2008

Abbreviations: ActR, Activin receptor type; ALK, activin receptor-like kinase; D, diestrus; DNase, deoxyribonuclease; E, estrogen; FBS, fetal bovine serum; HEK, human embryonic kidney; HPRT, human hypoxanthine phosphoribosyltransferase; NT, no treatment; OVX, ovariectomy; P, progesterone; PE, proestrous-early estrous; PEI, polyethyleneimine; PHM1, pregnant human myometrial 1 cell line; RT, reverse transcriptase.

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