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Female Reproductive Tract |
Center for Animal Biotechnology and Genomics, Department of Animal Science, Texas A&M University, College Station, Texas 77843-2471
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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subunit, ßA subunit, and ßB subunit were expressed in antral follicles between PNDs 0 and 56. These results led to rejection of the hypothesis that the ovary does not influence endometrial adenogenesis. Rather, the ovary and, thus, an ovarian-derived factor regulates, in part, the coiling and branching morphogenetic stage of endometrial gland development after PND 14 and expression of specific components of the activin-follistatin system in the neonatal ovine uterus that appear to be important for that critical process.
activitin, follistatin, inhibin, ovary, uterus
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Jost [7] first established, based on results from studies of rabbits, the concept that prenatal urogenital tract development in female mammals is an ovary-independent event. Since then, studies of several species have revealed that uterine development and endometrial adenogenesis can proceed normally in the absence of the ovary and, by default, ovarian steroids for varying periods of time during early postnatal life. In the prepubertal ewe, the ovary contains growing and antral ovarian follicles at birth that decline to PND 14, increase and peak in number on PND 28, remain high from PNDs 42 to 56, and then decline thereafter [8, 9]. These changes in ovarian follicles correlate with the ontogeny of endometrial gland development in the ewe lamb [3]. However, ovariectomy of the ewe at birth does not affect uterine wet weight [9] or the initial stages of endometrial gland tubulogenesis [10] on PND 14, but it does affect uterine growth after PND 14 [11]. Postnatal uterine growth and endometrial adenogenesis are ovary- and steroid-independent in rodents [12–14] and pigs [15]. Similarly, recent results indicate that postnatal uterine growth and endometrial adenogenesis are estrogen-independent from birth to PND 56, although coiling and branching morphogenesis after PND 14 is, in part, dependent on activated estrogen receptor (ER) [16]. Although the ovary plays a role in postnatal uterine growth, to our knowledge the role of the ovary in endometrial gland development after PND 14 has not been investigated in the ewe.
Recent results from studies of the neonatal ovine uterus implicate follistatin, activins, and activin receptors as autocrine and paracrine regulators of endometrial gland morphogenesis [17]. Activins and inhibins are members of the transforming growth factor (TGF) ß superfamily and regulate growth and differentiation of many branched epitheliomesenchymal organs via autocrine, paracrine, and perhaps, endocrine mechanisms [18–24]. Activins and inhibins are dimeric proteins (for review, see [25, 26]). Activin consists of two ß subunits, ßA and ßB, that homodimerize or heterodimerize to form activin A (ßA:ßA), activin B (ßB:ßB), or activin AB (ßA:ßB). Inhibin consists of an subunit that heterodimerizes with a ß subunit to form either inhibin A (:ßA) or inhibin B (:ßB). The biological activity of activins is mediated by receptor complexes consisting of activin receptor (ActR) type IA or ActRIB and ActRII. One of the key features that distinguish the effects of activins from those of TGFß is that binding of activins to their receptors can be blocked if activin binds to follistatin or if inhibin subunit binds to ActRs [27–29]. Follistatin binds to activins with high affinity and neutralizes their activity [30–32]. Inhibin subunit, activins, and follistatin are synthesized and secreted by ovarian follicles in the neonate and the adult [25, 26, 33–36]. Therefore, these ovarian factors could act in an endocrine manner to regulate uterine growth and endometrial adenogenesis via the activin-follistatin system to complement endocrine and paracrine effects of the activin-follistatin system in the neonatal ovine uterus.
Available evidence from the ewe and other domestic and laboratory animals supports the working hypothesis that endometrial adenogenesis is not regulated by the ovary and, thus, ovarian factors. To test this hypothesis, studies were conducted to determine the following: 1) the effects of removing the ovary at PND 7 on subsequent uterine growth and endometrial adenogenesis at PND 56, 2) the effects of ovariectomy on the activin-follistatin system in the uterus, and 3) the expression of the activin-follistatin system in the neonatal ovary.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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All experiments and surgical procedures were in accordance with the Guide for the Care and Use of Agriculture Animals and approved by the University Laboratory Animal Care Committee of Texas A&M University. Cross-bred Suffolk ewes were mated to Suffolk rams between the months of September and November 2001. Pregnant ewes were maintained according to normal husbandry practices. Ewes in the following experiments were born between the months of February and March 2002. In study 1, ewes were assigned randomly at birth (PND 0) to undergo either a sham surgery as a control (CX) or bilateral ovariectomy (OVX) on PND 7 (n = 6 per treatment). Beginning on PND 0, blood samples were collected by jugular venipuncture every 7 days into Vacutainer tubes (Becton-Dickinson, Franklin Lakes, NJ). On PND 56, all ewe lambs were weighed and necropsied. The uterus was obtained; trimmed free of the broad ligament, oviduct, and cervix; and weighed. Sections (thickness, 
1 cm) from the midportion of each uterine horn were fixed in fresh 4% paraformaldehyde at room temperature for 24 h and processed for histology. The remainder of the uterus was frozen in liquid nitrogen and stored at -80°C. In study 2, ovaries were obtained from ewes (n = 45) on PNDs 0 (n = 6), 7 (n = 4), 14 (n = 5), 21 (n = 5), 28 (n = 5), 35 (n = 5), 42 (n = 5), 49 (n = 5), or 56 (n = 5) as described previously [17].
Radioimmunoassay
Blood samples were allowed to clot for 1 h at room temperature. Serum was then separated by centrifugation (3000 x g for 30 min at 4°C), removed, and stored at -20°C.
Concentrations of estradiol-17ß (E2) in plasma were determined using methods described previously [3]. Assay sensitivity was 3 pg/ml, and the intra- and interassay coefficients of variation were 5% and 12%, respectively. Assay results were calculated using the AssayZap Version 3.1 program (Biosoft, Ferguson, CA).
Histology and Morphometry
After fixation, uteri or ovaries were changed to 70% ethanol for 24 h and then dehydrated and embedded in Paraplast Plus (Oxford Labware, St. Louis, MO). Uteri from study 1 were sectioned (thickness, 5 µm) and stained with hematoxylin and eosin as described previously [2]. Sections (n = 4) of the uterus from each ewe were photomicrographed, and images were analyzed using Scion Image software (Scion Corporation, Frederick, MD) as described previously [37]. Measurements were standardized using the image of a stage micrometer at the same magnification. The number of superficial ductal invaginations of GE from the LE into the stroma was determined. Endometrial gland number was determined by counting the total number of uterine glands in a complete cross-section of the uterine horn. The observation of a gland cross-section with a visible open lumen was counted as a single gland. Endometrial gland density was determined by counting the number of glands in a 200 µm2 area in the stratum compactum and stratum spongiosum areas of the intercaruncular endometrium. The number of ductal gland invaginations, endometrial gland number, and gland density estimates were generated for at least three areas within four nonsequential sections from each uterine horn. Intra- and intersection repeatability estimates for determination of ductal gland invaginations and endometrial gland number by a single observer were 0.85 and 0.8, respectively. The thickness or width of the endometrium and myometrium (inner circular and outer longitudinal layers) in the intercaruncular endometrial areas was measured using the Scion Image software from multiple points (n = 3–4) of at least 10 nonsequential uterine sections. Ovaries from study 2 were sectioned (thickness, 5 µm) and stained with Masson trichrome as described previously [38].
Semiquantitative Reverse Transcription-Polymerase Chain Reaction
Expression of mRNAs for ßA subunit, ßB subunit, ActRIA, ActRIB, ActRII, and follistatin was assessed in uterine total RNA using semiquantitative reverse transcription-polymerase chain reaction (RT-PCR) with methods [4, 5] and primers [17] as described previously [4, 5]. Total cellular RNA was isolated from frozen uteri using Trizol (Gibco-BRL, Grand Island, NY) according to manufacturer's recommendations. Briefly, cDNA was synthesized from total cellular RNA (5 µg) isolated from neonatal uteri using random (Life Technologies, Gaithersburg, MD) and oligo-dT primers and SuperScript II Reverse Transcriptase (Life Technologies). Newly synthesized cDNA was acid-ethanol precipitated, resuspended in 20 µl of water, and stored at -20°C. The cDNAs were diluted (1:1 or 1:10) with water before use in PCR. Primers were designed to amplify partial cDNAs for ovine follistatin, ßA subunit, ßB subunit, ActRIA, and ActRII as described previously [17]. ß-Actin primers were ATGAAGATCCTCACGGAACG (forward) and GAAGGTGGTCTCGTGAATGC (reverse), which amplified a 270-base pair product. The PCR reactions were performed using AmpliTaq DNA polymerase (Applied Biosystems, Foster City, CA) and Optimized Buffer D (Invitrogen, Carlsbad, CA) for ß-actin; Optimized Buffer E (Invitrogen) for ActRIB; Optimized Buffer F (Invitrogen) for follistatin, ActRIA, and ActRII; and Optimized Buffer J (Invitrogen) for ßA subunit and ßB subunit according to manufacturer's recommendations. The amount of cDNA template, annealing temperature, and number of cycles used for PCR were initially optimized to ensure that final PCR conditions were within the linear range of amplification for each primer pair. Follistatin and ßA subunit PCR reactions contained 1.5 µl of cDNA (1:10), ßB subunit reactions 2.5 µl of cDNA (1:1), ActRIA reactions 2 µl of cDNA (1:10), ActRIB reactions 2 µl of cDNA (1:1), ActRII reactions 3 µl of cDNA (1:10), and ß-actin reactions 1 µl of cDNA (1:10). All PCR reactions were performed at 95°C for 30 sec, 55–59°C for 1 min [17], and 72°C for 1 min. Cycle number was 25 for ß-actin; 27 for ActRII; 30 for follistatin, ßA subunit, and ActRIA; and 35 for ActRIB and ßB subunit. In negative-control reactions, RT cDNA was substituted by inclusion of uterine total RNA or water. Following PCR, equal amounts of reaction product were analyzed using a 2% agarose gel, and PCR products were visualized by ethidium bromide staining. The amount of DNA present was quantified by measuring the intensity of light emitted from correctly sized bands under ultraviolet light using an AlphaImager (Alpha Innotech Corporation, San Leandro, CA), and data are expressed as relative light units. All RT-PCR products were cloned into pCRII (Invitrogen) and fully sequenced in both directions to confirm identity.
In Situ Hybridization Analysis
Expression of mRNAs in the uterus (study 1) and ovary (study 2) was determined by in situ hybridization as described previously [39]. Briefly, deparaffinized, rehydrated, and deproteinated cross-sections (thickness, 5 µm) of the uterus (study 1) or ovaries (study 2) from each ewe were hybridized with radiolabeled sense or antisense cRNA probes generated from linearized plasmid templates containing partial cDNAs using in vitro transcription with [35S-]UTP. Partial cDNAs for ovine inhibin subunit, ßA subunit, ßB subunit, ActRIA, ActRII, and follistatin were generated by RT-PCR as described in the companion paper [17]. After hybridization, washing, and ribonuclease A digestion, slides were dipped in NTB-2 liquid photographic emulsion (Eastman Kodak, Rochester, NY), stored at 4°C for 2–28 days, and developed in Kodak D-19 developer. Slides were then counterstained with Gills modified hematoxylin (Stat Lab, Lewisville, TX), dehydrated through a graded series of alcohol to xylene, and protected with a coverslip. Images of representative fields in sections hybridized with antisense or sense cRNAs were recorded under bright- or dark-field illumination with a Nikon Eclipse 1000 photomicroscope (Nikon Instruments, Inc., Lewisville, TX) fitted with a Nikon DXM1200 digital camera using constant image acquisition parameters to ensure accurate comparison.
Immunohistochemistry
Expression of immunoreactive follistatin, inhibin subunit, ßA subunit, ßB subunit, ActRIA, ActRIB, and/or ActRII were detected in cross-sections (thickness, 5 µm) of the uterus (study 1) or ovaries (study 2) from each ewe with specific antibodies and a Super ABC Mouse/Rat Immunoglobulin G (IgG) Kit (Biomeda, Foster City, CA) using methods and antibodies described previously [17]. Multiple tissue sections from each ewe were processed as sets within an experiment. Relative staining intensity for immunoreactive protein expression was assessed visually in sections of the uterus or ovary from each ewe by two independent observers and scored as follows: absent (-, i.e., no staining above that of the IgG control), weak (+), moderate (++), or strong (+++) [5]. The scores from the two observers were averaged. If histologically discernable in the uterus, intercaruncular endometrial tissues (including LE, stroma, and GE), caruncular endometrial tissues (including LE and stroma), and myometrium were scored. In the ovary, cumulus-oocyte complex (COC) cells, granulosa cells (GC), oocyte, and theca cells (TC) were scored. Images of representative fields of sections probed with primary antibodies or IgG were recorded using a Nikon Eclipse 1000 photomicroscope fitted with a Nikon DXM1200 digital camera using constant image acquisition parameters to ensure accurate comparisons.
Statistical Analyses
All quantitative data were subjected to least-squares ANOVA using general linear model procedures of the Statistical Analysis System [40]. Serum E2 levels were log transformed before least-squares regression analyses. Uterine wet-weight data were analyzed using body weight as a covariate. Histomorphometrical data were analyzed using an overall model that included main effects of treatment, day, treatment by day interaction, section, and area. In all analyses, error terms used in tests of significance were identified according to the expectation of the mean squares for error. The RT-PCR data were subjected to one-way ANOVA utilizing the ß-actin values as a covariate in the model to correct for differences in amounts of RT cDNA analyzed for each uterus. A probability (P) value of 0.10 or less was accepted as indicating significance. Data are presented as least-square means of untransformed values with overall SEMs.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Circulating concentrations of E2 in plasma were low and affected by day (P < 0.10) but not by treatment (P > 0.10) or their interaction (P > 0.10). Serum E2 levels were highest on PND 0 (10 ± 1.5 pg/ml), declined to PND 7 (3 ± 1.5 pg/ml), and remained low to PND 56 (data not shown).
Ovariectomy Retards Uterine Growth and Endometrial Gland Morphogenesis
As summarized in Table 1, uterine wet weight was 52% lower (P < 0.01) in OVX compared to CX ewes on PND 56. The intercaruncular endometrium of CX ewes contained numerous coiled and branched glands extending radially from the LE through the stroma to the inner circular layer of myometrium (Fig. 1). In contrast, the intercaruncular endometrium of OVX ewes contained substantially lower numbers of coiled and branched endometrial glands.
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These histological observations were confirmed by histomorphometrical analyses, and the results are summarized in Table 1. The total number of endometrial glands in the uterine wall was reduced (P < 0.0001) by 44% in ovariectomized ewe lambs. The number of gland invaginations from the LE and density of glands in the stratum compactum stroma adjacent to the LE were not affected (P > 0.10) by ovariectomy. However, density of glands in the stratum spongiosum stroma adjacent to the inner circular layer of myometrium was reduced (P < 0.001) by 22% in OVX ewes. In the intercaruncular endometrium, the thickness and width of the endometrium and the myometrium were reduced (P < 0.05) by 22% and 16%, respectively, in uteri of OVX ewes.
Expression of the Follistatin-Activin System in the Uterus Is Altered by Ovariectomy
Semiquantitative RT-PCR analyses were conducted using total RNA isolated from uteri of CX and OVX ewes to determine steady-state levels of activin-follistatin system components. The number of PCR cycles was optimized for each primer pair to ensure amplification within the linear range of detection, as representatively illustrated by follistatin (Fig. 2A). As shown in Figure 2B, each of the primer pairs used for RT-PCR amplified a single product of the expected size [17]. A partial cDNA for ActRIB was detected abundantly in total RNA isolated from the neonatal ovary, but at extremely low amounts in the neonatal ovine uterus (data not shown). The amplified products were sequenced to confirm identity (data not shown). As compared to mRNA levels in uteri of CX ewes (Table 2), uteri of OVX ewes had substantially lower levels of activin ßA subunit, ActRIA, ActRII, and follistatin on PND 56. In contrast, the level of activin ßB subunit mRNA was greater in uteri of OVX compared to CX ewes on PND 56.
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In situ hybridization and immunohistochemical analyses were performed to localize expression of follistatin-activin system mRNAs and protein in the uteri from CX and OVX ewes (Figs. 3 and 4). Results of immunohistochemical analyses were quantified and are summarized in Table 3. Expression of inhibin subunit mRNA or protein was not detected in the neonatal ovine uterus (data not shown). As compared to CX ewes, uteri of OVX ewes had lower levels of ßA subunit expression, particularly in the endometrial LE, GE, and myometrium (Fig. 3A and Table 3). In contrast, the expression of ßB subunit was more abundant in the endometrial LE and GE of OVX ewes (Fig. 3B). Follistatin mRNA and protein levels were lower in uteri of OVX ewes, particularly in the endometrial stroma and myometrium (Fig. 3C).
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Overall, expression of ActRIA and ActRIB was lower in the uteri of OVX ewes (Fig. 4, A and B). Specifically, ActRIA and ActRIB protein levels were reduced in both the endometrial glands and myometrium of OVX compared to CX ewes (Table 3). The low abundance of ActRIB mRNA was below the detectable limits of the in situ hybridization procedure (data not shown). The mouse anti-human ActRII antibody detects both ActRIIA and ActRIIB (R&D Systems, Inc., Minneapolis, MN). Although ActRII mRNA expression was lower in uteri of OVX ewes, immunoreactive ActRIIA/B protein abundance was not different between uteri from CX and OVX ewes (Fig. 4C).
Expression of Activin-Follistatin System Components in the Neonatal Ovary
As expected [6], the number of growing and antral follicles in the neonatal ovine ovary declined from PND 0 to PND 7, increased after PND 7 to PND 14, peaked on PND 28, and remained high to PND 56 (Fig. 5). On PND 28 and thereafter, a number of the follicles exhibited signs of atresia, including rupture and disorganization of the TC and GC layers. As expected, no corpora lutea were observed in the ovaries.
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In the ovaries, inhibin subunit mRNA and protein were detected only in GC and cells of the COC (Fig. 6A). Overall, expression of inhibin subunit mRNA and protein increased in antral and graafian follicles between PND 0 and PND 14 and remained abundant thereafter (Table 4). In the neonatal ovary, ßA subunit mRNA and protein were detected in GC, COC, and the oocyte, with low levels of protein in the TC (Fig. 6B). The ßB subunit mRNA and protein were detected in GC and COC (Fig. 6C). The most abundant levels of immunoreactive ßB subunit protein were in the zona pellucida of the oocyte. Follistatin mRNA and protein expression were detected predominantly in the GC and COC (Fig. 6D), but immunoreactive follistatin protein was also present in TC and tunica muscularis of blood vessels.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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In the present study, ovariectomy on PND 7 did not affect the number of superficial ductal invaginations of the GE from the LE or the density of endometrial glands in the stratum compactum area of the stroma on PND 56. Similarly, Bartol et al. [10] found that initial genesis and development of endometrial glands between birth and PND 14 was not dependent on the ovary or an ovarian factor. However, results of the present study clearly demonstrate that the total number of endometrial glands in the uterine wall and the density of the endometrial glands in the stratum spongiosum area of the stroma are reduced in uteri from OVX ewes, indicating that an ovarian-derived factor regulates, in part, the coiling and branching morphogenetic stage of endometrial gland development between PND 14 and PND 56. Postnatal uterine growth and endometrial adenogenesis are ovary-, adrenal-, and steroid-independent in rodents [12–14]. In the neonatal gilt, ovariectomy at birth inhibits uterine growth after PND 56 but does not affect genesis of uterine glands or related endometrial morphogenetic events before PND 120 [15]. Collectively, available results indicate that the precise role of ovarian factor(s) in uterine growth and endometrial adenogenesis is species-specific.
The process of uterine morphogenesis is governed by a variety of hormonal, cellular, and molecular mechanisms, many of which remain to be defined (for review, see [4, 43]). Although changes in uterine ER gene expression can be correlated with endometrial adenogenesis [3], uterine growth and endometrial gland development is an estrogen-independent process in the ewe between birth and PND 56 [16]. However, endometrial gland coiling and branching morphogenesis between PND 14 and PND 56 is dependent, in part, on uterine ER [16]. In the present study, effects of ovariectomy on uterine growth and development observed in the present study were not caused by alterations in circulating E2. Moreover, expression of ER and progesterone receptor protein was not different between uteri of CX and OVX ewes (unpublished observations). Therefore, the effects of ovariectomy on uterine growth and endometrial adenogenesis cannot be attributed to alterations in either circulating E2 or the uterine ER system.
The ovarian factor that regulates uterine growth and endometrial adenogenesis is not known, but it could be inhibin, follistatin, or activins. In the neonatal ewe, the increase in ovarian weight between PND 14 and PND 56 is caused, in part, by the increase in the number and size of antral follicles and the associated accumulation of follicular fluid [8]. The stimulus for initiation and maintenance of ovarian follicular growth after birth is not known. During this period, low levels of LH can be detected in a pulsatile fashion, whereas FSH secretion is tonic [8, 11, 44]. The unregulated growth of ovarian follicles in the ewe after birth may be caused by endocrine stimuli, such as LH and FSH, as well as by intraovarian mechanisms. In the present study, ewes were born during seasonal anestrus and ovaries collected before puberty, but inhibin subunit, ß subunits, and follistatin were expressed in nearly all growing and antral follicles. In the ewe, follistatin levels rise in the fetus during parturition and remain high in the neonate [36]. In contrast, immunoreactive inhibin levels are low at 2 wk and then decline to 15 wk of life in the ewe [35]. In the present study, abundant expression of ß subunits and ActRs as well as of follistatin and inhibin subunit was detected in the neonatal ovary. These findings are similar to previously reported results [33, 34]. Undoubtedly, these factors regulate follicular development in the prepubertal ovine ovary through their established roles in GC proliferation and control of pituitary hormones [25, 26].
Recent evidence from studies of the neonatal ovine uterus implicates follistatin, activins, and ActRs as regulators of endometrial gland morphogenesis in the neonatal ovine uterus [17]. Follistatin, activins, and inhibins regulate growth and differentiation of many branched epitheliomesenchymal organs via autocrine, paracrine, and perhaps, endocrine mechanisms [18–24]. In general, exogenous activin inhibits gland development, whereas follistatin counteracts these inhibitory effects of activins by binding to the individual ßA and ßB subunits and preventing ActR activation [23, 29]. In a variety of cell types, expression of activin-follistatin system components can be modulated by activins, inhibin, and follistatin in a cell type-specific manner [45–49]. Activin can increase follistatin expression and differentially regulate ActR expression in pituitary cells [46]. In testicular tumor cells, activin regulates expression of ActRII and ßA subunit and inhibits cell proliferation [47]. In rat Sertoli cells, activin increases expression of ActR and follistatin and increases cell proliferation [49]. In human GC, activin A increases ßB subunit expression with no effects on ßA mRNA or inhibin subunit mRNA [45]. Available results support the working hypothesis that follistatin and activins from the ovary act on the uterus to regulate, in part, coiling and branching morphogenetic development of endometrial glands as well as overall uterine growth. This hypothesis is supported by findings in the present study that the expression of specific components of the activin-follistatin system were affected by ovariectomy and could be correlated with reduced endometrial gland development. Indeed, marked reductions in expression of follistatin, ßA subunit, ActRIA, and ActRII genes and an increase in ßB subunit expression were observed in the uterus of ovariectomized ewes. Furthermore, these changes could be correlated with, but not completely accounted for, by the decreases in uterine size and endometrial gland development in ovariectomized ewes. One or more of the ovarian factors likely is either follistatin, activins, or inhibin.
Collectively, available results support the idea that a factor from the ovary regulates overall development, adenogenesis, and expression of the follistatin-activin system in the neonatal ovine uterus. It is tempting to speculate that the coordinate activities of the activin-inhibin-follistatin system in the ovary and uterus may be important in prolific breeds of ewes that possess an intrinsically high ovulation rate as well as an enhanced uterine capacity to maintain large litters [50]. Future experiments will be directed toward understanding the mechanistic aspects of the novel finding that an ovarian endocrine factor includes a member of the activin-inhibin-follistatin system that may act in concert with autocrine-paracrine effects of the uterine activin-follistatin system to regulate coiling and branching morphogenesis of the endometrial glands during postnatal development of the ovine uterus.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Correspondence: Thomas E. Spencer, Center for Animal Biotechnology and Genomics, 442 Kleberg Center, 2471 TAMU, Texas A&M University, College Station, Texas 77843-2471. Fax: 979 862 2662; tspencer{at}tamu.edu
3 These authors made equal contributions to the manuscript
Received: 12 February 2003.
First decision: 21 March 2003.
Accepted: 8 May 2003.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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and ßA subunits in the ovaries of postnatal and prepubertal lambs. Reprod Fertil Dev 1997 9:825-832[CrossRef][Medline]This article has been cited by other articles:
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  K. Hayashi, A. R O'Connell, J. L Juengel, K. P McNatty, G. H Davis, F. W Bazer, and T. E Spencer Postnatal uterine development in Inverdale ewe lambs Reproduction, March 1, 2008; 135(3): 357 - 365. [Abstract] [Full Text] [PDF]
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 S.-Y. Lin, R. G. Craythorn, A. E. O'Connor, M. M. Matzuk, J. E. Girling, J. R. Morrison, and D. M. de Kretser Female Infertility and Disrupted Angiogenesis Are Actions of Specific Follistatin Isoforms Mol. Endocrinol., February 1, 2008; 22(2): 415 - 429. [Abstract] [Full Text] [PDF]
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 K. Hayashi and T. E. Spencer Estrogen Disruption of Neonatal Ovine Uterine Development: Effects on Gene Expression Assessed by Suppression Subtraction Hybridization Biol Reprod, October 1, 2005; 73(4): 752 - 760. [Abstract] [Full Text] [PDF]
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K. Hayashi, K. D Carpenter, T. H Welsh Jr, R. C Burghardt, L. J Spicer, and T. E Spencer The IGF system in the neonatal ovine uterus Reproduction, March 1, 2005; 129(3): 337 - 347. [Abstract] [Full Text] [PDF]
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 K. Hayashi, K. D. Carpenter, and T. E. Spencer Neonatal Estrogen Exposure Disrupts Uterine Development in the Postnatal Sheep Endocrinology, July 1, 2004; 145(7): 3247 - 3257. [Abstract] [Full Text] [PDF]
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 T. E. Spencer and F. W. Bazer Uterine and placental factors regulating conceptus growth in domestic animals J Anim Sci, January 1, 2004; 82(13_suppl): E4 - 13. [Abstract] [Full Text] [PDF]
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K. Hayashi, K. D. Carpenter, C. A. Gray, and T. E. Spencer The Activin-Follistatin System in the Neonatal Ovine Uterus Biol Reprod, September 1, 2003; 69(3): 843 - 850. [Abstract] [Full Text] [PDF]
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