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Regular Article |
a Department of Tissue Physiology and
b Laboratory of Veterinary Physiology, Faculty of Agriculture, Tokyo University of Agriculture and Technology, Fuchu, Tokyo 183-8509, Japan
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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subunit mRNA remained low throughout estrus, metestrus, and diestrus; abruptly increased on the morning of proestrus; then rapidly decreased when the primary gonadotropin surge occurred. A similar changing pattern was observed in inhibin/activin ßA subunit mRNA. On the other hand, inhibin/activin ßB subunit mRNA showed a different changing pattern. Levels of ßB subunit mRNA remained constant during metestrus and diestrus, abruptly decreased on the afternoon of proestrus, then quickly recovered from the nadir by 1100 h on estrus. Throughout the rat estrous cycle, especially during the periovulatory period, subunit mRNA levels were considerably higher than ßA and ßB subunit mRNA levels. In addition, changes in plasma concentrations of inhibin A and inhibin B were very similar to that in ovarian ßA and ßB subunit mRNA levels, respectively, with several-hour delays. These results suggest that levels of ß subunit mRNAs restrict secretion of dimeric inhibins. Levels of follistatin mRNA remained low from the midnight of metestrus to the midnight of diestrus, then increased until initiation of the primary gonadotropin surge. Thereafter, follistatin mRNA decreased, reached the nadir at 0200 h on estrus, then increased abruptly at 1100 h on estrus. Afterward, follistatin mRNA levels remained high until the morning of metestrus. The changing pattern of ovarian follistatin mRNA was similar to, and preceded, the changes in plasma concentrations of progesterone, suggesting that ovarian follistatin may modulate progesterone secretion during the rat estrous cycle.
activin, follistatin, inhibin, ovary, ovulatory cycle
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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subunit and either a ßA subunit to give rise to inhibin A or a ßB subunit to form inhibin B [1–4]. Inhibin is predominantly secreted from granulosa cells of ovarian follicles in female mammals [1–4] and is a major FSH-suppressing factor in the female rat [5–8]. The ß subunits of inhibin also form homo- and heterodimers termed activin A, activin AB, and activin B, which stimulate pituitary FSH secretion [1–4]. Although activins have been identified as FSH-releasing proteins, it has not been demonstrated that ovarian activins directly regulate FSH secretion. However, ovarian activins are thought to be important autocrine or paracrine factors regulating follicular development, because activin affects functions of ovarian cells [3, 4]. Inhibins also likely act as important autocrine or paracrine factors in the ovary [3, 4]. Follistatin is a monomeric glycoprotein originally isolated from bovine and porcine follicular fluid as an FSH-suppressing protein [9, 10]. Thereafter, follistatin proved to be an activin-binding protein that antagonizes the function of activin via binding to its ß subunits [11]. Follistatin also affects functions of ovarian cells, probably by inhibiting actions of activins [3, 4].
In the ovary, inhibin subunit mRNA is localized in granulosa cells of follicles of all classes; theca cells of secondary, tertiary, and preovulatory follicles; luteal cells of young corpora lutea; and interstitial gland cells [12]. In contrast to the wide distribution of subunit mRNA in the ovary, ß subunit mRNAs show a distribution that is more restricted. Inhibin/activin ßA subunit mRNA is confined to granulosa cells of secondary, tertiary, and preovulatory follicles. Inhibin/activin ßB subunit mRNA is confined to granulosa cells of secondary and tertiary follicles and tends to be expressed in small follicles compared to ßA subunit mRNA [12]. With respect to follistatin, its mRNA and protein are localized in granulosa cells of secondary, tertiary, and preovulatory follicle and in luteal cells of young corpora lutea [13].
Previous studies have demonstrated that ovarian expression of mRNAs encoding inhibin/activin subunits and follistatin shows noticeable changes during the rat estrous cycle [12–15]. However, to our knowledge, quantitative analysis showing detailed changes in these mRNAs in the ovary has not been performed. Especially, quantitative changes in ßB subunit mRNA during the rat estrous cycle have not been demonstrated. In addition, only one point was examined on each day of the rat estrous cycle for quantification of ovarian follistatin mRNA in a previous study [15]. Furthermore, it is important to elucidate the changes in relative levels of ovarian inhibin/activin subunit mRNAs during the estrous cycle, because the proportion of and ß subunits is likely to determine which form (inhibins or activins) will be secreted [16].
In the present study, to extend our knowledge of the inhibin/activin system in the rat, ovarian inhibin/activin subunit and follistatin mRNA levels during the rat estrous cycle were measured by ribonuclease (RNase) protection assay using digoxygenin-labeled RNA probes. Furthermore, plasma concentrations of inhibin A and inhibin B were measured, and the relationship between plasma levels of inhibins and ovarian levels of inhibin subunit mRNAs was examined.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Adult cycling female rats of the Wistar strain weighing 220–270 g were used. They were kept under controlled temperature (25 ± 2°C) and lighting (lights-on from 0500 to 1900 h). Food and water were available ad libitum. Vaginal smears were checked daily, and only rats that showed at least two consecutive 4-day estrous cycles were used.
Rats were killed by decapitation at 1100 and 2300 h on each day of the estrous cycle; at 0500, 1400, 1700, and 2000 h on proestrus; and at 0200 and 0500 h on estrus (n = 5–6 for each point). Trunk blood was collected and plasma separated immediately and stored at -20°C until assayed for FSH, LH, inhibin A, inhibin B, estradiol, and progesterone. Both ovaries were immediately immersed in liquid nitrogen and stored at -80°C until isolation of total RNA. Total RNA was isolated with Trizol reagent (Life Technologies, Tokyo, Japan) and quantified by spectrophotometric measurement at 260 nm.
The experimental protocol was approved in accordance with the Guide for the Care and Use of Laboratory Animals prepared by the Tokyo University of Agriculture and Technology.
Probes
Rat inhibin , ßA, and ßB subunit and follistatin cDNA clones were obtained from Dr. Kelly E. Mayo (Northwestern University, Evanston, IL). Rat cyclophilin cDNA clone was provided by Dr. James Douglass (Vollum Institute, Portland, OR). The 1572-base pair (bp) rat inhibin subunit cDNA (rINA-13) [17], the 1628-bp rat inhibin/activin ßA subunit cDNA (rINB-5) [17], the 2100 bp rat inhibin/activin ßB subunit cDNA (rINBB-2) [18], the 730-bp rat follistatin cDNA (rFST-4a), and the 680-bp rat cyclophilin cDNA (1B15) [19] clones were subcloned into a pBluescript KS (-) vector (Stratagene, La Jolla, CA). Template inhibin subunit, inhibin/activin ßA subunit, and ßB subunit cDNAs represent 320-bp subclones between restriction sites KpnI and PstI, 192-bp subclones between restriction sites HindIII and BamHI, and 281-bp subclones corresponding to the 5' region from the first restriction site RsaI, respectively. Template follistatin and cyclophilin cDNA clones represent 393-bp subclones between restriction sites RsaI and 304-bp subclones between restriction sites PstI and NcoI, respectively. Transcription of probe (antisense) and reference (sense) RNAs was done in vitro from linearized template DNA using bacteriophage T3 and T7 RNA polymerase (Roche Diagnostics, Tokyo, Japan). Digoxygenin (DIG)-labeled RNA probes were synthesized using DIG RNA labeling mix (Roche Diagnostics) according to the recommended procedures. The reaction was terminated by adding RNase-free deoxyribonuclease I (Roche Diagnostics), and the probe was purified by passing through a 5% acrylamide denaturating gel. Probe was eluted out of the gel with probe elution buffer (0.5 M ammonium acetate, 1 mM EDTA, and 0.2% SDS) for 2 h at 37°C. Reference RNAs were extracted with phenol/chloroform and precipitated twice in ethanol after adjusting the final concentration of ammonium acetate to 0.5 M. The purified reference RNAs were quantified by spectrophotometric measurement at 260 nm. Molecular weights of the reference RNAs were calculated as sodium salt from their deduced base composition.
RNase Protection Assay
The RNase protection assay using DIG-labeled RNA probes was done according to the procedure of Wundrack and Dooley [20] with modifications. An aliquot of DIG-labeled RNA probe (1 ng each) was added to a known amount of ovarian total RNA (5 µg for inhibin subunit and cyclophilin; 20 µg for inhibin/activin ßA and ßB subunits and follistatin) and precipitated with ethanol. To obtain standard curves, a known amount of yeast total RNA (5 µg for inhibin subunit and cyclophilin; 20 µg for inhibin/activin ßA and ßB subunits and follistatin) containing 2.5–320 pg of appropriate sense strand RNA was used instead of ovarian total RNA. The resulting pellet was air-dried and dissolved in 10 µl of hybridization solution containing four parts deionized formamide and one part 5x stock solution (200 mM piperazine-N,N'-bis[2-ethane-sulfonic acid] [pH 6.4], 2 M NaCl, and 5 mM EDTA). Samples were denatured for 5 min at 90°C, then hybridized at 45°C overnight. After the incubation, 100 µl of RNase digestion buffer (10 mM Tris-HCl [pH 7.5], 5 mM EDTA, 300 mM sodium acetate, 0.33 kunitz units/ml of RNase A, and 100 U/ml of RNase T1) were added. Single-strand RNAs were then digested with RNase A and T for 30 min at 37°C. Thereafter, RNases were inactivated by adding 5 µl of 10% SDS and 1.25 µl of proteinase K solution (10 mg/ml in 10 mM Tris-HCl [pH 7.5]). After incubation at 37°C for 30 min, protected RNA probes were precipitated with ethanol in the presence of 10 µg of yeast RNA. The resulting precipitate was redissolved in 8 µl of gel loading buffer (95% deionized formamide, 0.5% SDS, 0.5 mM EDTA, 0.025% xylene cyanol, and 0.025% bromophenol blue). After denaturation at 90°C for 5 min, the sample was loaded onto a 6% denaturating polyacrylamide gel, then electrotransferred on a nylon membrane (Highbond N+; Amersham Pharmacia Biotech, Tokyo, Japan). The membrane was reacted with ovine anti-DIG antibody conjugated with alkaline phosphatase (Roche Diagnostics) according to the procedures recommended by Roche Diagnostics. Disodium 3-(4-methoxyspiro{1,2 dioxethane-3,2'-(5'-chlolo)triclo[3.3.1.13,7]decan}-4-yl)phenyl phosphate (CSPD; Roche Diagnostics) was used as a substrate. Membranes were exposed to x-ray film for 2 h for inhibin subunit and cyclophilin mRNAs and 6 h for inhibin/activin ßA and ßB subunit and follistatin mRNAs. The intensity of the bands was measured using a laser densitometer. Hybridization of inhibin/activin ßA and ßB subunit and follistatin mRNAs for ovarian samples was carried out in one tube. Signal variation between membranes was checked by loading an equal amount (20 pg) of DIG-labeled RNA on each gel. The coefficient of variation between membranes was 5.7%. All samples were measured in a single assay.
RIA of FSH and LH
Plasma concentrations of FSH and LH were measured using NIDDK RIA kits (Bethesda, MD) for rat FSH and LH. Iodinated preparations were rat FSH-I-5 and LH-I-5. The antisera used were anti-rat FSH-S-11 and anti-rat LH-S-10. Results were expressed as rat FSH RP-2 and rat LH RP-2. The intra- and interassay coefficients of variations were 4.8% and 11.4%, respectively, for FSH and 5.4% and 6.9%, respectively, for LH.
Concentrations of progesterone and estradiol in plasma were determined by double-antibody RIAs with 125I-labeled radioligands as described previously [21]. Antisera to progesterone (GDN 377) and estradiol (GDN 244) provided by Dr. G.D. Niswender (Colorado State University, Fort Collins, CO) were used. The intra- and interassay coefficients of variations were 3.1% and 12.1%, respectively, for progesterone and 5.3% and 18.1%, respectively, for estradiol.
ELISA of Inhibin A and Inhibin B
Inhibin A and inhibin B assay kits were purchased from Serotec (Oxford, Oxon, U.K.). These dimer-specific assays have been shown to be applicable to the rat [22]. Amounts of inhibin A and inhibin B were expressed in terms of recombinant human inhibin A and recombinant human inhibin B, respectively. All samples were measured in a single assay. The intraassay coefficients of variation were 4.4% for inhibin A and 2.2% for inhibin B.
Statistical Analysis
Values are represented as means ± SEM. To compare the mean values, results were subjected to one-way analyses of variance followed by Student-Neuman-Keuls test. Logarithmic conversion was carried out before the analysis. A value of P < 0.05 was considered to be significant. The correlation between parameters was determined using simple regression analysis. Parallelism of reference RNA and sample RNA in the RNase protection assay was examined by analysis of covariance.
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RESULTS |
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, ßA, and ßB subunits as well as follistatin and cyclophilin mRNAs obtained from the RNase protection assay are shown in Figures 1 and 2, respectively. Increases in signals in response to increasing amounts of reference RNAs and ovarian total RNA were observed (Fig. 1), and reference RNAs gave good standard curves (Fig. 2). Signals for ovarian total RNA also gave similar dose-response curves (data not shown). Analysis of covariance revealed parallelism between dose-response curves for reference RNA and ovarian total RNA.
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In rats used for the present study, the primary gonadotropin surge was observed at 1700 h on proestrus, and the secondary surge of FSH occurred during the early morning of estrus (Fig. 3). Inhibin subunit mRNA was maintained at low levels throughout metestrus and diestrus, increased noticeably at 1100 h on proestrus, then decreased abruptly when the primary gonadotropin surge occurred (Fig. 4A). Afterward, inhibin subunit mRNA reached a minimum level at 0200 h on estrus, then slightly increased until 1100 h on estrus. The changing pattern of inhibin/activin ßA subunit mRNA was similar to that of inhibin subunit mRNA (Fig. 4B). Levels of ßA subunit mRNA remained constant from metestrus to diestrus, abruptly increased at 1100 h on proestrus, then rapidly decreased with the primary gonadotropin surge. Thereafter, ßA subunit mRNA levels reached the nadir at 2000 h on proestrus, then gradually increased from 0500 h on proestrus until the morning of metestrus. Although the changing pattern of and ßA subunit mRNAs were similar to each other, subunit mRNA levels were relatively constant during estrus, metestrus, and diestrus compared to ßA subunit mRNA levels. On the other hand, the changing pattern of inhibin/activin ßB subunit mRNA was quite different from those of and ßA subunit mRNAs. Inhibin/activin ßB subunit mRNA remained at constant levels from the morning of estrus to the morning of proestrus, quickly decreased with the primary gonadotropin surge, and reached the nadir at 1700 h on proestrus (Fig. 4C). Thereafter, ßB subunit mRNA levels quickly recovered to the predecline levels by 1100 h on estrus. Ovarian follistatin mRNA levels remained low from 2300 h on metestrus to 2300 h on diestrus, then increased until 1400 h on proestrus (Fig. 4D). Afterward, follistatin mRNA decreased until 0200 h on estrus, then abruptly increased until 2300 h on estrus. After the increase, follistatin mRNA was maintained at high levels until 1100 h on metestrus.
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Plasma concentrations of inhibin A, inhibin B, progesterone, and estradiol are shown in Figure 5. Changing patterns of ovarian inhibin/activin ßA and ßB subunit and follistatin mRNAs are overlapped. Changes in plasma concentrations of inhibin A and inhibin B were similar to those reported previously [22] and were very similar to changes in ovarian inhibin/activin ßA and ßB subunit mRNA levels, respectively, with several-hour delays (Fig. 5, A and B). Plasma levels of inhibin A and inhibin B were correlated with levels of ßA (r = 0.63) and ßB subunit (r = 0.51) mRNAs, respectively. However, during the periovulatory period, levels of ovarian ß subunit mRNAs showed higher correlation with plasma concentrations of inhibins that were measured for later time points. Mean levels of ßA subunit mRNA at 1100, 1400, 1700, 2000, and 2300 h on proestrus and at 0200 h on estrus were highly correlated with mean values of plasma inhibin A for 3 h later (r = 0.96). Similarly, mean levels of ßB subunit mRNA at 1100, 1400, 1700, 2000, and 2300 h on proestrus and at 0500 h on estrus were well correlated with mean values of plasma inhibin B for 6 h later (r = 0.91).
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Plasma concentrations of progesterone abruptly increased with the primary gonadotropin surge, then rapidly decreased by 0200 h on estrus (Fig. 5C). Thereafter, plasma levels of progesterone increased from the morning of estrus, remained at high levels during metestrus, and decreased on the morning of diestrus. It is interesting to note that the changing pattern of plasma progesterone was similar to that in ovarian follistatin mRNA with several-hour delays. Mean levels of follistatin mRNA at 1100, 1400, 1700, 2000, and 2300 h on proestrus and at 0200 h on estrus were highly correlated with mean values of plasma progesterone for 3 h later (r = 0.82). In addition, mean levels of follistatin mRNA during estrus, metestrus, and diestrus were also correlated with mean values of plasma progesterone for 12 h later (r = 0.93).
Plasma concentrations of estradiol remained low during estrus and metestrus, then increased from 2300 h on metestrus until initiation of the primary gonadotropin surge (Fig. 5D). As observed previously [22], correlation between plasma estradiol and inhibin A (r = 0.75) was higher than that between plasma estradiol and inhibin B (r = 0.46). Plasma estradiol was also correlated with ßA subunit mRNA (r = 0.65) but showed no correlation with ßB subunit mRNA (r = 0.02).
Ratios of ovarian subunit mRNA to ß subunit mRNAs are shown in Figure 6. Inhibin subunit mRNA was abundantly expressed in the ovary throughout the rat estrous cycle compared to ß subunit mRNAs. Inhibin subunit mRNA levels were approximately 5-fold greater than ßA or ßB subunit mRNA during metestrus and diestrus. Noticeable increases in the ratios of subunit mRNA to ß subunit mRNAs were observed during the periovulatory period.
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DISCUSSION |
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and ßA subunit mRNAs, the present results are consistent with previous observations [12, 14]; however, a minor discrepancy is observed. The maximum expression of inhibin subunit mRNA coincided with, or occurred after, the primary gonadotropin surge in the previous studies, whereas it was noted before the LH rise in the present study. Probably, the time of sample collection may be very critical during the periovulatory period. In addition, the previous studies were accomplished mainly by the in situ hybridization technique. Although Woodruff et al. [14] examined the changes in and ßA subunit mRNAs by RNase protection assay, they did not examine an internal standard control (e.g., cyclophilin, GAPDH, ß-actin, etc.). The differences of the methods may also be a cause of the discrepancy.
The present study showed that inhibin subunit mRNA levels increased abruptly in the late follicular phase, then rapidly decreased with the primary gonadotropin surge. It has been demonstrated that follicular expression of inhibin subunit mRNA continuously increases as follicles grow [12]. Therefore, the abrupt increase in subunit mRNA that was observed in the present study likely is a result of the growth of large antral follicles. During the other stages of the estrous cycle, levels of subunit mRNA were relatively constant compared to that of ßA subunit mRNA. The wide distribution of subunit mRNA in the ovary [12] may be, at least in part, responsible for the constant expression during estrus, metestrus, and diestrus. As well as subunit mRNA, ßA subunit mRNA showed an abrupt increase on the morning of proestrus. The previous study showed that ßA subunit mRNA is mainly localized in large tertiary follicles during diestrus and proestrus [12]. Furthermore, in the present study, plasma inhibin A was correlated with plasma estradiol, as observed previously [22]. These findings suggest that the growth of large antral follicles selected for ovulation is responsible for the abrupt increase in ovarian ßA subunit mRNA. On the other hand, ßB subunit mRNA lacked the proestrous increase that was observed in and ßA subunit mRNAs. The previous study demonstrated that the distribution of ßB subunit mRNA was similar to that of ßA subunit mRNA, but that ßB subunit mRNA showed a more rapid decrease with maturation of follicles [12]. Probably, the decrease in ßB subunit mRNA with follicle maturation is responsible for the lack of the proestrous rise in ßB subunit mRNA. During the primary gonadotropin surge, ßB subunit mRNA decreased noticeably, as did and ßA subunit mRNAs. However, as observed in a previous study [12], ovarian contents of ßB subunit mRNA immediately declined compared to and ßA subunit mRNAs. The decreases in inhibin/activin subunit mRNAs in preovulatory follicles have been attributed to the primary gonadotropin surge [23, 24]. The hCG-induced ovulation also decreases ovarian inhibin/activin subunit mRNAs in eCG-primed immature rats [25]. Although the detailed mechanism of the decline of the mRNAs is not known, the present results suggest that transcription of ßB subunit mRNA is more quickly arrested by the primary gonadotropin surge than that of and ßA subunit mRNAs. It is also possible that ßB subunit mRNA is unstable compared to the other inhibin/activin subunit mRNAs.
Our results showed that changes in plasma concentrations of inhibin A and inhibin B were similar to changes in ovarian inhibin/activin ßA and ßB subunit mRNA levels, respectively. In addition, inhibin subunit mRNA was abundantly expressed in the ovary compared to ß subunit mRNAs throughout the estrous cycle, which is consistent with previous observations [25, 26]. These results suggest that levels of ß subunit mRNAs restrict dimeric inhibin secretion. On the other hand, dynamic changes in proportion of subunit mRNA and ß subunit mRNAs were observed during the estrous cycle. Noticeable increases in the ratios of to ß subunit mRNAs after ovulation are also reported in eCG-primed immature rats [25]. Because inhibins and activins share common ß subunits, high ratios of subunit mRNA to ß subunit mRNAs (ßA + ßB) are probably inappropriate for activin secretion. Although the present results show ratios of subunit mRNA to ß subunit mRNAs in the whole ovary, these results may partly reflect the ratios in individual cells. Both the high ratios of subunit mRNA to ß subunit mRNAs and the decreases in ß subunit mRNAs during the periovulatory period may result in a decline of activin secretion. It has been indicated that activin from secondary follicles inhibits development of small preantral follicles [27]. The present and previous results suggest that a decrease in activin secretion as well as the periovulatory surge of FSH may stimulate development of new cohorts of follicles.
To our knowledge, the present study is the first to demonstrate that follistatin mRNA levels show two surges during the estrous cycle, although the preovulatory rise is not statistically significant compared to the lowest value. It has been reported that follistatin is localized in granulosa cells of ovarian follicles and in young corpora lutea [13]. Therefore, the rise in proestrus may be due to development of follicles to be selected for ovulation, and the rise after the primary gonadotropin surge may be due to an increase in follistatin mRNA in young corpora lutea. Relatively high levels of ovarian follistatin mRNA in proestrus and estrus are consistent with previous results [15]. However, levels of ovarian follistatin mRNA remained high until the morning of metestrus in the present study, whereas those on the morning of metestrus were relatively low in the previous study [15]. In the present study, large variations of follistatin mRNA levels were noted from the night of estrus to the morning of metestrus. Therefore, it is possible that the timing of luteolysis may vary between individual rats in the present study, and this may be responsible for the difference between the results in the present and previous studies.
Interestingly, the changing pattern of ovarian follistatin mRNA was similar to, and preceded, the changes in plasma concentrations of progesterone. Although the preovulatory rise of plasma progesterone is induced by the primary gonadotropin surge, the present results suggest that ovarian follistatin may support progesterone secretion during the rat estrous cycle. It has been reported that follistatin stimulates progesterone secretion from human granulosa cells [28]. In addition, activin stimulates estradiol secretion, but inhibits progesterone secretion, from rat preovulatory granulosa cells [29]. The similar antiluteinizing effect of activin has been demonstrated in both human and bovine luteinizing granulosa cells [30, 31], and follistatin antagonizes the actions of activin [29–31]. Furthermore, it has been demonstrated that bone morphogenetic protein (BMP)-4, BMP-7, and their receptors are expressed in the rat ovary, and that the BMPs inhibit luteinization of granulosa cells in vitro [32]. Because follistatin is known to bind to activins [11] and BMPs [33], the rise of ovarian follistatin mRNA during estrus may support luteinization by blocking the antiluteinizing effects of the growth factors. The present data are, again to our knowledge, the first in vivo evidence that suggests physiological roles of ovarian follistatin in the regulation of progesterone secretion.
In summary, the present study is the first, to our knowledge, to demonstrate that levels of ovarian inhibin/activin ßB subunit mRNA remain constant during the rat estrous cycle, except during the periovulatory period, and that levels of follistatin mRNA show two surges during the rat estrous cycle. Furthermore, the present results confirm that ovarian inhibin/activin subunit mRNAs are differently regulated during the rat estrous cycle, and they suggest that ß subunit mRNA levels restrict dimeric inhibin secretion. A relationship between ovarian follistatin expression and progesterone secretion is also suggested.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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1 Correspondence and current address: Koji Y. Arai, Center for Reproductive Sciences, Dept. of Molecular and Integrative Physiology, University of Kansas Medical Center, 3901 Rainbow Blvd., Kansas City, KS 66160. FAX: 913 588 7180; karai{at}kumc.edu 
Accepted: November 9, 2001.
Received: August 15, 2001.
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REFERENCES |
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, ßA, and ßB subunits in ovarian cell types during the rat estrous cycle. Mol Endocrinol 1988; 2:1352-1363[CrossRef][Medline] and ß subunit complementary deoxyribonucleic acids and expression in the ovary. Mol Endocrinol 1987; 1:561-568[CrossRef][Medline], ßA, and ßB subunits in hormonally induced immature female rats. Mol Endocrinol 1989; 3:2062-2069[CrossRef][Medline], ßA, and ßB subunits in various tissues predicts diverse functions. Proc Natl Acad Sci U S A 1988; 85:247-251This article has been cited by other articles:
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  K. Y Arai and T. Nishiyama Developmental Changes in Extracellular Matrix Messenger RNAs in the Mouse Placenta During the Second Half of Pregnancy: Possible Factors Involved in the Regulation of Placental Extracellular Matrix Expression Biol Reprod, December 1, 2007; 77(6): 923 - 933. [Abstract] [Full Text] [PDF]
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 J. Yeh and B. Kim Increasing Blunting of Inhibin Responses to Dynamic Ovarian Challenge Is Associated With Reproductive Aging in the Rat Reproductive Sciences, January 1, 2007; 14(1): 10 - 19. [Abstract] [PDF]
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 H. A. Kenny and T. K. Woodruff Follicle Size Class Contributes to Distinct Secretion Patterns of Inhibin Isoforms during the Rat Estrous Cycle Endocrinology, January 1, 2006; 147(1): 51 - 60. [Abstract] [Full Text] [PDF]
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 K. Y Arai, H. Kishi, S. Onodera, W. Jin, G. Watanabe, A. K Suzuki, S. Takahashi, T. Kamada, T. Nishiyama, and K. Taya Cyclic changes in messenger RNAs encoding inhibin/activin subunits in the ovary of the golden hamster (Mesocricetus auratus) J. Endocrinol., June 1, 2005; 185(3): 561 - 575. [Abstract] [Full Text] [PDF]
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Y. Wang and W. Ge Developmental Profiles of Activin {beta}A, {beta}B, and Follistatin Expression in the Zebrafish Ovary: Evidence for Their Differential Roles During Sexual Maturation and Ovulatory Cycle Biol Reprod, December 1, 2004; 71(6): 2056 - 2064. [Abstract] [Full Text] [PDF]
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 R. C Hirst, M. H Abel, V. Wilkins, C. Simpson, P. G Knight, F.-P. Zhang, I. Huhtaniemi, T R. Kumar, and H. M Charlton Influence of mutations affecting gonadotropin production or responsiveness on expression of inhibin subunit mRNA and protein in the mouse ovary Reproduction, July 1, 2004; 128(1): 43 - 52. [Abstract] [Full Text] [PDF]
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B. Sisco, L. J. Hagemann, A. N. Shelling, and P. L. Pfeffer Isolation of Genes Differentially Expressed in Dominant and Subordinate Bovine Follicles Endocrinology, September 1, 2003; 144(9): 3904 - 3913. [Abstract] [Full Text] [PDF]
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K. Y. Arai, K. Tsuchida, K. Uehara, K. Taya, and H. Sugino Characterization of Rat Follistatin-Related Gene: Effects of Estrous Cycle Stage and Pregnancy on Its Messenger RNA Expression in Rat Reproductive Tissues Biol Reprod, January 1, 2003; 68(1): 199 - 206. [Abstract] [Full Text] [PDF]
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