Estrogen Receptor-ß mRNA Expression in Rat Ovary: Down-Regulation by Gonadotropins

Michael Byers, George G. J. M. Kuiper, Jan-Åke Gustafsson and Ok-Kyong Park-Sarge

Department of Physiology (M.B., O-K.P-S.) University of Kentucky Lexington, Kentucky 40536
Center for Biotechnology (G.G.J.M.K.) and Department of Medical Nutrition (J-Å.G.) Karolinska Institute S-14186 Huddinge, Sweden


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
We have examined the expression and regulation of the two estrogen receptor (ER{alpha} and ERß) genes in the rat ovary, using Northern blotting, RT-PCR, and in situ hybridization histochemistry. Northern blotting results show that the ovary expresses both ER{alpha} and ERß genes as single (~6.5-kb) and multiple (ranging from ~1.0-kb to ~10.0-kb) transcripts, respectively. ER{alpha} mRNA is expressed at a level lower than ERß mRNA in immature rat ovaries. This relationship appears unchanged between sexually mature adult rats and immature rats. In sexually mature adult rats undergoing endogenous hormonal changes, whole ovarian content of ERß mRNA, as determined by RT-PCR, remained more or less constant with the exception of the evening of proestrus when ERß mRNA levels were decreased. Examination of ERß mRNA expression at the cellular level, by in situ hybridization, showed that ERß mRNA is expressed preferentially in granulosa cells of small, growing, and preovulatory follicles, although weak expression of ERß mRNA was observed in a subset of corpora lutea, and that the decrease in ERß mRNA during proestrous evening is attributable, at least in part, to down-regulation of ERß mRNA in the preovulatory follicles. This type of expression and regulation was not typical for ER{alpha} mRNA in the ovary. Although whole ovarian content of ER{alpha} mRNA was clearly detected by RT-PCR, no apparent modulation of ER{alpha} mRNA levels was observed during the estrous cycle. Examination of ER{alpha} mRNA expression at the cellular level, by in situ hybridization, showed that ER{alpha} mRNA is expressed at a low level throughout the ovary with no particular cellular localization.

To further examine the potential role of the preovulatory pituitary gonadotropins in regulating ERß mRNA expression in the ovary, we used immature rats treated with gonadotropins. In rats undergoing exogenous hormonal challenges, whole ovarian content of ERß mRNA, as determined by RT-PCR, remained more or less unchanged after an injection of PMSG. In contrast, a subsequent injection of human CG (hCG) resulted in a substantial decrease in whole ovarian content of ERß mRNA. In situ hybridization for ERß mRNA shows that small, growing, and preovulatory follicles express ERß mRNA in the granulosa cells. The preovulatory follicles contain ERß mRNA at a level lower than that observed for small and growing follicles. In addition, there is an abrupt decrease in ERß mRNA expression in the preovulatory follicles after hCG injection. The inhibitory effect of hCG on ERß mRNA expression was also observed in cultured granulosa cells. Moreover, agents stimulating LH/CG receptor-associated intracellular signaling pathways (forskolin and a phorbol ester) readily mimicked the effect of hCG in down-regulating ERß mRNA in cultured granulosa cells.

Taken together, our results demonstrate that 1) the ovary expresses both ER{alpha} and ERß genes, although ERß is the predominant form of estrogen receptor in the ovary, 2) ERß mRNA is localized predominantly to the granulosa cells of small, growing, and preovulatory follicles, and 3) the preovulatory LH surge down-regulates ERß mRNA. These results clearly implicate the physiological importance of ERß in female reproductive functions.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Estrogen critically affects the growth and development of ovarian follicles during the female reproductive cycle (reviewed in Refs. 1–6) by stimulating the proliferation of granulosa cells from small follicles (7), increasing granulosa cell gonadotropin receptor levels and thus responsiveness of granulosa cells to gonadotropins (8, 9), modulating progesterone production by granulosa cells (10) and androgen production by theca cells (11), enhancing gap junction formation among granulosa cells (12), and modulating luteal steroidogenic capacity (reviewed in Ref.13). These intraovarian actions of estrogen indicate the presence of specific receptor molecules interacting with this steroid in the ovary. Indeed, the ovaries of a number of species have been shown to express estrogen-binding molecule(s) as determined by specific retention of radiolabeled estrogen (14, 15), and specific binding capacity of estrogen in ovarian extracts (16, 17, 18, 19). These binding studies suggest that ovarian estrogen receptor (ER)-like molecules exert steroid specificity and binding affinity similar to those of the well characterized conventional ER (20), recently renamed ER{alpha} (21). ER{alpha} is believed to mediate many of estrogen’s actions in a variety of reproductive tissues, including the uterus (22, 23). If the same scenario is applied to the ovary, ER{alpha} protein should be present at a detectable level in the ovary and elimination of the ER{alpha} gene in vivo should disrupt all ovarian functions requiring estrogen action, such as folliculogenesis. Although ER immunoreactivity has been found in ovaries of a limited number of species, such as the baboon and human (24, 25, 26), disruption of the ER{alpha}-gene in vivo did not eliminate the ability of small follicles to grow as evident from the presence of secondary and antral follicles in ER{alpha}-knockout mice (27, 28), arguing for the possibility that intraovarian action of estrogen may be mediated by molecular mechanisms requiring estrogen-binding molecules other than ER{alpha}. This possibility is further supported by experimental results from our laboratory (29, 30) and others (31, 32) demonstrating that in the ovary, in contrast to the uterus, estrogen does not stimulate one of the best known genes targeted by ER, the progesterone receptor (PR) gene. In light of these observations, the identification of a second subtype of ER (ERß) that is expressed in the ovary (21) may provide a fundamental understanding of the longstanding critical role of estrogen in ovarian function. This receptor has been shown to bind to estrogen with high affinity and steroid specificity very similar to that of the ER{alpha} (Ref. 21 and G.G.J.M. Kuiper, unpublished data). Thus, we sought to examine the expression and hormonal regulation of these two ERs (ER{alpha} and ERß) in the rat ovary. We have used ovaries of immature rats treated with gonadotropins to stimulate folliculogenesis, ovulation, and luteinization, as well as of sexually mature rats during the estrous cycle. Our results demonstrate that 1) the primary ER subtype in the ovary is ERß, although the ovary expresses both ER{alpha} and ERß genes, 2) ovarian expression of ERß mRNA is decreased during follicular development and differentiation, and 3) activation of LH/CG receptors down-regulates ERß mRNA expression in cultured rat granulosa cells.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Differential Expression of ER{alpha} and ERß mRNA Species in the Rat Ovary
To determine whether the two ERs (ER{alpha} and ERß) are expressed at a comparable level in the rat ovary, we performed Northern analysis (Fig. 1AGo) and RT-PCR (Fig. 1Go, B and C) to specifically detect ER{alpha} and ERß transcripts in rat ovarian RNA. An equal amount of rat uterine RNA was used as a positive control since it expresses ER{alpha} mRNA at a high level. Forty micrograms of total RNA were fractionated on a 1% denaturing gel that was transferred onto a nylon membrane and probed for ER{alpha} using the hormone-binding domain of the rat ER{alpha} (20) or for ERß using the EcoRI/PstI fragment (~750 bp most 5'-end) of the rat ERß (21). The results of Northern analyses performed under stringent conditions show that the ovary contains an ER{alpha} transcript (~6.5 kb), as the uterus does, but at an extremely low level (Fig. 1AGo, left panel). When the same blot was reprobed for ERß mRNA, the ovarian RNA was found to contain multiple transcripts of the ERß gene (ranging from ~1.0 kb to ~10.0 kb) at a detectable level whereas little ERß mRNA signal was detected in uterine RNA (Fig. 1AGo, middle panel). These multiple ERß transcripts were better shown in RNA of isolated granulosa cells as compared with whole ovaries (Fig. 1AGo, right panel), indicating that ERß mRNA is enriched in granulosa cells. Similar quantification results were obtained using RT-PCR assays performed under conditions generating a linear range of specific amplification of ER{alpha} and ERß mRNAs. A typical example of the relationship between cycle numbers and RT-PCR amplification of ER{alpha} and ERß mRNAs from a fixed concentration of ovarian RNA (1/4 of cDNA generated from 5 µg RNA) is shown in Fig. 1BGo. This concentration of cDNA is within a linear range of amplification of ER{alpha} and ERß mRNAs (data not shown). Linear amplification of ER{alpha} and ERß mRNAs in ovarian samples was obtained using up to 30 cycles of PCR and thus, subsequent experiments were performed using 25 cycles of amplification. ER{alpha} and ERß RT-PCR products from the same ovarian or uterine cDNA samples show that the uterus predominantly expresses ER{alpha} mRNA while ERß is the predominant ER subtype expressed in the ovary (Fig. 1CGo). These reactions amplified specific ER{alpha} and ERß PCR products of the expected size based upon our results using ER{alpha} [the human ER{alpha} (33)] or ERß [the rat ERß (21)] cDNA plasmids as controls.



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Figure 1. Expression of Estrogen Receptor (ER{alpha} and ERß) mRNAs in Rat Ovary and Uterus

Panel A shows Northern blotting. Approximately 40 µg RNA from ovaries (as well as isolated granulosa cells) and uteri of immature rats (23 days of age) were separated on a 1% denaturing agarose/formaldehyde gel, the RNA was transferred to a nylon membrane, and the membrane was probed with the hormone-binding domain of the rat ER{alpha} cDNA (500 bp) or with the most 5'-end (5'-UTR and N-terminal A/B region) of the rat ERß cDNA (~750 bp). Exposure was for 2 days. The same blots were subsequently reprobed to assess RNA equivalence using clone CHO-B. The ribosomal bands 28S and 18S are indicated. Panel B shows RT-PCR (66 C annealing temperature) of ER{alpha} and ERß transcripts in the ovary as a function of cycle numbers. An autoradiogram of a polyacrylamide gel on which the products of an RT-PCR assay have been separated and its quantification data are shown. The ER{alpha} or ERß PCR product is indicated as is the ribosomal protein S16 internal control. Panel C shows the specificity of our RT-PCR for ER{alpha} and ERß transcripts. ER{alpha} and ERß cDNAs were used as controls.

 
ER{alpha} and ERß mRNA Expression in Rat Ovaries in Vivo
To gain insights into whether ovarian expression of the ER{alpha} or ERß gene is temporally associated with particular stages of the female reproductive cycle, we examined ER{alpha} and ERß mRNA levels in ovaries of 1) sexually mature adult rats throughout the reproductive cycle and 2) immature rats treated with exogenous gonadotropin combination (PMSG plus human CG), by performing RT-PCR and in situ hybridization. After 25 cycles of amplification at 66 C annealing temperature, ERß as well as ER{alpha} RT-PCR products were detectable in ovaries at all stages of the reproductive cycle (Fig. 2Go). Again, ERß mRNA levels were higher than ER{alpha} mRNA levels in these samples. This is consistent with our observations showing a preferential amplification of ERß mRNA during coamplification of ER{alpha} and ERß mRNAs in the same samples (data not shown). ER{alpha} RT-PCR fragment was detected in all stages of the estrous cycle with less variation than that observed for ERß. Quantification of ERß mRNA levels shows little variation during the cycle with the exception of the evening of proestrus when ERß mRNA levels were decreased. This decrease in ERß mRNA levels occurred after the onset of the preovulatory LH surge (our LH RIA results show that the preovulatory LH surge was initiated at 1600 h and peaked at 1800 h of proestrus in these animals, data not shown). Our results employing in situ hybridization followed by liquid emulsion autoradiography show that ER{alpha} mRNA is expressed at a low level with no particular cellular localization within the ovary, and that ERß mRNA in small and growing follicles remained more or less similar throughout the estrous cycle (data not shown), whereas ERß and LH receptor (LH-R) transcripts are coexpressed in preovulatory follicles of cycling rats and are down-regulated during the transition from proestrus to estrus (Fig. 3AGo). Before the peak of the preovulatory LH surge (shown are 1400 h and 1800 h), preovulatory follicles express both ERß and LH-R mRNA in the granulosa cell layer. After the peak of the preovulatory LH surge, levels of both ERß and LH-R mRNA are substantially decreased (shown are 2200 h and 2400 h). An example of ER{alpha} and ERß mRNA expression in ovaries containing different follicular structures is shown in Fig. 3BGo. A subset of corpora lutea which express LH-R mRNA have been found to weakly express ERß mRNA (upper panel, shown are corpora lutea from metestrus 1800 h). Interestingly, small follicles expressing LH-R mRNA only in theca cells in the same field clearly expressed ERß mRNA in granulosa cells. Throughout folliculogenesis in adult rats during the estrous cycle as well as in immature rats during PMSG treatment, all growing follicles, regardless of their size (small, antral, or preovulatory), were found to express ERß mRNA [shown are ovarian sections of PMSG-treated (10 IU, 48 h) immature rats, lower panel]; as previously shown (34), preovulatory follicles express LH-R mRNA in granulosa cell layers (34). The colocalization of LH-R mRNA and ERß mRNA in follicles at different stages is shown at higher magnifications in Fig. 4AGo. Small follicles (indicated by an arrow, left panel) with LH-R mRNA in the theca, but not granulosa, cell layer (34) clearly express ERß mRNA in the granulosa, but not theca, cell layer. Preovulatory follicles (also indicated by an arrow, right panel) with LH-R mRNA in both granulosa and theca cell layers (34) also express ERß mRNA, although at a decreased level, in the granulosa cell layer. Figure 4BGo shows colocalization of LH-R or PR mRNA with ERß mRNA in preovulatory follicles in immature rats challenged with hCG. Three hours after hCG treatment, granulosa cell expression of LH-R mRNA is persistent in preovulatory follicles. In contrast, ERß mRNA expression in the same follicles is markedly decreased whereas nonpreovulatory follicles (absence of LH-R mRNA in granulosa cells) persistently express ERß mRNA. Six hours after hCG treatment, granulosa cell expression of PR mRNA is evident in preovulatory follicles (29), which express little ERß mRNA. Again, nonpreovulatory follicles (absence of PR mRNA expression) persistently expressed ERß mRNA.



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Figure 2. Expression of the ER{alpha} and ERß Genes in the Ovary of Adult Rats during the Estrous Cycle

Upper panel shows an autoradiogram of a polyacrylamide gel on which the products of an RT-PCR assay have been separated. The ER{alpha} and ERß PCR products are indicated as is the ribosomal protein S16 internal control. Times during the estrous cycle are indicated at the top. Bottom panel shows the quantification of the data in panel A. Band intensities were measured on a PhosphoImager, and the ER{alpha} or ERß signals were normalized to the S16 internal control for each time point. Relative mRNA levels were shown as the value at E1100 being 1.0. Sera from these animals were used to determine LH concentrations; the onset of the LH surge was observed at 1600 h of proestrus, and the peak was observed at 1800 h of proestrus.

 


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Figure 3. Colocalization of LH-R, ER{alpha}, and ERß mRNAs in Various Ovarian Structures as Determined by in Situ Hybridization Using an 35S-Labeled Antisense RNA Probe for the Rat ERß, the Rat ER{alpha}, or the Rat LH-Receptor

Panel A shows darkfield photographs (50x magnification) of preovulatory follicles of adult rats during the transition from proestrus to estrus. Time of death and the probes are indicated. Panel B shows brightfield and darkfield photographs (50x magnification) of corpora lutea of an adult rat killed at Metestrus 1800 h (upper panel) and of preovulatory follicles of an immature rat treated with PMSG (10 IU) for 48 h (lower panel). The probes are indicated.

 


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Figure 4. Colocalization of LH-R (or PR) and ERß mRNAs in Follicular Structures

Panel A shows small (left, an immature rat treated with PMSG for 48 h plus hCG for 3 h) and preovulatory (right, an immature rat treated with PMSG for 48 h) follicles expressing both LH-R mRNA and ERß mRNA. In each case, the follicle indicated by an arrow is shown at higher magnifications. Granulosa (G) and theca (T) cell layers are also indicated. Panel B shows colocalization of LH-receptor (or progesterone receptor) mRNA and ERß mRNA in serial ovarian sections of immature rats treated with PMSG (10 IU, 48 h) followed by hCG (10 IU) for 3 or 6 h. Some preovulatory follicles are indicated by arrows. The probes are indicated, and all photographs were taken at 50x magnification.

 
To further determine the potential role of the pituitary gonadotropins, FSH and LH, in regulating ERß mRNA expression in the ovary, we examined immature rats treated with PMSG followed by hCG. ERß mRNA expression in these rat ovaries was determined initially by RT-PCR (Fig. 5Go). PMSG treatment (up to 48 h) did not significantly alter total ovarian content of ERß mRNA levels. In contrast, a subsequent injection of hCG (up to 12 h) in rats similarly treated with PMSG (48 h) substantially decreased ERß mRNA levels. To examine the effects of PMSG and hCG on ERß mRNA expression at a cellular level, we performed in situ hybridization on ovaries of similarly treated rats. As shown in Fig. 6Go, the granulosa cells of preantral as well as antral follicles of untreated immature rats clearly express ERß mRNA at a detectable level. During folliculogenesis stimulated by PMSG treatment, ERß mRNA expression in growing follicles appears to decrease. Forty-eight hours after PMSG treatment, the granulosa cells of preovulatory follicles clearly express LH-R mRNA as well as ERß mRNA. During short-term treatment with hCG (3 h), down-regulation of ERß mRNA is evident in the preovulatory follicles still expressing LH-receptors, suggesting that ERß mRNA is rapidly down-regulated; six h after hCG injection, both LH-R and ERß mRNA levels were reduced.



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Figure 5. Modulation of Ovarian ERß mRNA Expression by Gonadotropins in Immature Rats

Upper panel shows an autoradiogram of a polyacrylamide gel on which the products of an RT-PCR assay have been separated while lower panel shows the quantification of the data. Band intensities were measured on a PhosphoImager, and the ERß signal was normalized to the S16 internal control for each experimental group. The ratio of ERß/S16 of control rats with no hormonal treatment was considered 1.0. Shown are the mean ± SE (n = 4).

 


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Figure 6. Modulation of Ovarian ERß mRNA Expression, at the Cellular Level, by Gonadotropins in Immature Rats

Animals were treated with PMSG (10 IU, s.c.) followed by hCG (10 IU, s.c.), and the ovaries were processed for in situ hybridization using an 35S-labeled antisense RNA probe for the rat ERß. Capital letters show brightfield photographs while lower case letters show darkfield photographs (50x magnification). A (a): untreated control; B (b): 6-h treatment with PMSG; C (c): 12-h treatment with PMSG; D (d): 48-h treatment with PMSG, E (e): 3-h treatment with hCG following by 48-h treatment with PMSG; F (f): 6-h treatment with hCG following 48-h treatment with PMSG; G (g): 12-h treatment with hCG following 48-h treatment; H (h): 24-h treatment with hCG following 48-h treatment with PMSG.

 
The Effect of hCG on ERß mRNA Expression in Cultured Rat Granulosa Cells
To examine the ability of hCG to modulate ERß mRNA expression by directly acting on granulosa cells cultured in vitro, we isolated differentiated granulosa cells from immature rats primed with PMSG (10 IU, 44 h) and cultured in vitro. Cells were treated with hCG (I IU/ml) for 3–12 h, and RNA was examined for ERß mRNA levels by RT-PCR. As shown in Fig. 7AGo (3-h incubation), granulosa cells contain a fairly high level of ERß mRNA, and incubation of these cells with hCG decreased ERß mRNA to 50% of the original value. Activation of LH receptor-associated signaling pathways by using activators for protein kinases A (forskolin, 10-5 M) and C [a phorbol 12-myristate 13-acetate (TPA), 10-7 M] results in ERß mRNA down-regulation in cultured rat granulosa cells. Interestingly, forskolin and TPA act together to further decrease ERß mRNA levels in these cells. This inhibitory effect of forskolin or TPA on ERß mRNA levels in granulosa cells lasts at least up to 12 h as shown in Fig. 7BGo.



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Figure 7. Modulation of ERß mRNA by hCG in Rat Granulosa Cells Cultured in Vitro

Granulosa cells were isolated from immature rats primed with PMSG (10 IU, 44 h), cultured in vitro for 15 h, and treated with the indicated reagents for 3 h. RNA samples were subjected to RT-PCR for ERß mRNA along with S16 as an internal control. Panel A shows an autoradiogram of a polyacrylamide gel on which the products of a RT-PCR assay have been separated and the corresponding quantification data. Shown are the mean ± SE (n = 4). The ratio of ERß/S16 of the control condition with no hormonal treatment was considered 1.0. Hormonal treatments are shown at the top. Treatments are control cells (Cont), forskolin (FSK) at 10-5 M, hCG at 1 IU/ml, a phorbol ester (TPA) at 10-7 M. Panel B shows the effect of forkolin (10-5 M) for 3–12 h in down-regulating ERß mRNA.

 

    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Predominant Expression of ERß in Granulosa Cells
Estrogen-signaling events are believed to critically affect a spectrum of physiological processes by interacting with an intracellular receptor (ER), a ligand-inducible transcription factor (35). Thus, the expression level of ER in a particular tissue has been used as an index of the degree of estrogen responsiveness (22, 23). In many female reproductive tissues, ER expression levels correlate with the ability of estrogen to stimulate expression of an array of well characterized target genes such as the PR gene (22, 23, 36). This tight cause-effect relationship between ER and PR expression appears to be uncoupled in the ovary. Although estrogen is essential for the growth of ovarian follicles (1, 2, 3, 4, 5, 6, 7) that clearly display specific binding sites for estrogen (14, 15, 16, 17, 18, 19), this steroid fails to directly stimulate the PR gene in ovarian follicles (30, 31, 32). In addition, the lack of expression of a full- length and functional ER{alpha} in ER{alpha} null mutant mice (27, 28, 37) did not prevent the growth of follicles, which resulted in high circulating estrogen levels (28, 37), suggesting the possibility that the ovary expresses estrogen-binding molecule(s) other than ER{alpha}. Consistent with this possibility are our results demonstrating that ERß is the primary ER subtype expressed in the ovary, in particular granulosa cells of small, growing, and preovulatory follicles. Thus, the ovarian responsiveness to estrogen in the ER{alpha} knockout mice (27, 37) may result from the expression of ERß. Our results, however, do not exclude the potential importance of ER{alpha} in ovarian functions. Indeed, experimental results of ours (this report) and others (38, 39) show the evident, although low, expression of the ER{alpha} gene in the ovary. Coexpression of ER{alpha} and ERß in the same follicles suggests the interesting possibility that ER{alpha} and ERß proteins may interact with each other and discriminate between target sequences leading to differential responsiveness to estrogen. Because both ER{alpha} and ERß proteins bind estrogen with an affinity of ~10-9 M and display the same steroid specificity (20, 21), it is highly unlikely that differential responsiveness to estrogen is achieved at the level of interaction between ligand and receptor. Because ERß, like ER{alpha}, when expressed in transfected cells, has been shown to be capable of transactivating a simple artificial estrogen response element-containing promoter (21), it is possible that differential responsiveness to estrogen may be achieved at the level of interactions between ER{alpha} and ERß or between ERs and DNA. Interestingly, PRA, the N-terminally truncated naturally occurring isoform of PRB, functions as a transactivator in some cells as a homodimer whereas it serves as a repressor of PRB when both isoforms are present (40, 41, 42). Perhaps similar mechanisms for estrogen responsiveness could be operating in cells expressing both ER{alpha} and ERß. In this regard, it is interesting to note that the N-terminal domain of the ERß protein is not homologous to, and is much shorter than, the corresponding region of the ER{alpha} protein. In cells in vivo, the ratio of ER{alpha}/ERß mRNA varies significantly among estrogen target tissues (Ref. 21 and our results). Our results show that the ratio of ERß/ER{alpha} in the ovary is much higher than in the uterus, a well known estrogen target. In uterine cells expressing high levels of ER{alpha} but extremely low levels of ERß, estrogen clearly stimulates multiple physiological processes including the synthesis of PRs (43). In addition, ER{alpha} knockout mice fail to initiate these physiological processes in response to estrogen (37). In contrast, in the ovary, which expresses relatively high levels of ERß but low levels of ER{alpha}, total elimination of ER{alpha} failed to disrupt normal growth of ovarian follicles (37). In addition, estrogen failed to directly stimulate PR gene expression in normal preovulatory follicles (30, 31, 32). Although one possible explanation is that these two cell types (uterus and ovary) express different coactivators facilitating the interactions between ERs and general transcription factors; another possibility is that these two cell types respond to estrogen differently because of the different ratio of ER{alpha}/ERß expression. It will be important to determine whether ER{alpha} and ERß proteins are coexpressed within a single cell, whether ER{alpha} and ERß could potentially dimerize with each other, and whether ER{alpha} and ERß homodimers, as well as ER{alpha}/ERß heterodimers, would exhibit cell context-dependent and promoter-dependent transactivation function.

Hormonal Regulation of ERß mRNA Expression by Gonadotropins
In many estrogen target tissues such as the uterus and pituitary, estrogen and progesterone modulate levels of ER{alpha} protein as well as mRNA (22, 23, 36, 43). In the ovary, estrogen-binding sites have been shown to be modulated by the pituitary gonadotropins (17). The discrepancy between our RT-PCR results showing that ERß mRNA in whole ovary remained more or less similar during folliculogenesis (proliferation of granulosa cells) and our in situ hybridization results showing an apparent decrease in ERß mRNA in granulosa cells of large and preovulatory follicles may be explained by the fact that whole ovary contains an increased number of granulosa cells during folliculogenesis. This possibility is consistent with a published observation that estrogen-binding sites per granulosa cells decrease after PMSG treatment (48 h) (17). During PMSG treatment the follicular synthesis of estrogen significantly increases and thus the PMSG-induced decrease in granulosa cell expression of ERß mRNA may result from PMSG-induced estrogen production and/or PMSG-induced intracellular signaling pathways. Curiously, ERß mRNA levels in whole ovary are substantially decreased after the onset of the gonadotropin surge in cycling rats and after hCG injection in PMSG-primed immature rats. Our in situ hybridization data showing colocalization of ERß, LH-R, and PR mRNAs in immature rats treated with gonadotropins argue that LH/CG-induced down-regulation of ERß mRNA in whole ovaries is attributable, at least in part, to the loss of ERß mRNA in preovulatory follicles responding to LH/CG. In these experiments, only preovulatory follicles expressing LH-R mRNA lose ERß mRNA in response to hCG challenge. Similarly, all preovulatory follicles expressing PR mRNA in response to LH/CG (29, 30) express little ERß mRNA while neighboring follicles without PR mRNA (indicative of the lack of LH-R) express a normal level of ERß mRNA. This observation is consistent with a previous report demonstrating that hCG induces down-regulation of estrogen receptor-binding sites (17). Our results demonstrating the ability of protein kinase A (forskolin) and protein kinase C (TPA) activators to induce this type of down-regulation of ERß mRNA levels in cultured granulosa cells indicate that the intracellular signals generated by the preovulatory LH surge in differentiated granulosa cells initiate the molecular cascade of events leading to either rapid degradation of ERß mRNA and/or transcriptional turn-off of the ERß gene. In MCF-7 breast cancer cells in which the ER{alpha} gene has been shown to be down-regulated by estrogen (44), a phorbol ester TPA has also been shown to down-regulate ER{alpha} mRNA expression by facilitating rapid degradation of ER{alpha} messages (45). It remains to be determined whether similar mechanisms perhaps mediate ER{alpha} degradation in nontumor cells. Although the importance of the LH-induced down-regulation of ERß mRNA in preovulatory follicles is currently unknown, there must be another hormonal cue that increases ERß mRNA expression by increasing either mRNA stability or transcriptional rate, as evident from a subset of corpora lutea of cycling rat ovaries expressing levels of ERß mRNA that are slightly above background. Functional corpora lutea of pregnant rats also express ERß mRNA (M. Byers and O-K. Park-Sarge, unpublished observation). Thus, future studies will be required to determine the molecular mechanisms by which ERß mRNA expression is regulated in granulosa/luteal cells.

In summary, our results demonstrating that ERß is the primary ER subtype expressed in the ovary, that ERß is expressed in granulosa cells of virtually all healthy follicles, and that ERß is down-regulated by gonadotropins in granulosa cells suggests the possibility that in the ovary, the functional significance of estrogen action may be mediated primarily by ERß.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Animals and Hormone Treatments
Two sets of animals were used for these studies: immature female rats treated with gonadotropins and sexually mature adult female rats exhibiting regular 4-day estrous cycles. All animals are treated according to the NIH guidelines for the care and use of animals.

Immature Animals.
Sprague-Dawley female pups (21 days old) with a nursing mother were purchased from Harlan Breeding Company (Indianapolis, IN) and housed in a photoperiod of 14-h light, 10-h darkness, with lights on at 0500 h. Food and water were freely available. At 23 days of age, rats were injected s.c. with 10 IU PMSG (Sigma, St Louis MO) in 0.1 ml PBS. Forty-eight hours later, rats were injected s.c. with 10 IU hCG (Sigma) in 0.1 ml PBS. Rats were killed by decapitation at various time points throughout hormone treatments, and their ovaries were rapidly removed from surrounding fat and oviduct, frozen on dry ice and stored until use at -80 C.

Sexually Mature Adult Rats.
Adult female Sprague-Dawley rats (150–180 g body weight) were purchased from Charles-Rivers Breeding Company (Wilmington, MA) and were housed as above. Estrous cyclic stages were determined by daily examination of vaginal cytology, and only those animals demonstrating at least two consecutive 4-day cycles were used for the experiment. Rats were euthanized at specific time intervals throughout the estrous cycle. Trunk blood was collected for serum LH measurements, and ovaries were rapidly removed from surrounding fat and oviduct, frozen on dry ice, and stored until use at -80 C. Serum LH concentrations in these cycling rats were determined by an RIA using the NIH NIDDK kit with the exception of LH antibody CSU 120, which was generously provided by Dr. Terry Nett.

Granulosa Cell Isolation and Culture
Immature rats at 22–23 days of age were primed with a single s.c. injection of PMSG (10 IU) and 40–44 h later, granulosa cells were isolated by the method of follicular puncture (30, 46, 47). Ovaries were collected in cold serum-free medium (4F) consisting of 15 mM HEPES (pH 7.4), 50% Dulbecco’s MEM and 50% Ham’s F12 with bovine transferrin (5 µg/ml), human insulin (2 µg/ml), hydrocortisone (40 ng/ml), and antibiotics. After incubation in warm (37 C) 4F medium containing 0.5 M sucrose and 10 mM EGTA for 20–30 min to loosen cell junctions, ovaries were washed in fresh 4F medium, and individual follicles were punctured using 23-gauge needles. Extruded granulosa cells were collected, washed twice, and plated in 4F medium supplemented with 5% FBS (GIBCO, Grand Island, NY) at a density of approximately 2 x 106 cells per 100-mm dish, incubated overnight in the humidified atmosphere of 5% CO2 at 37C, and treated with various hormones.

RNA Blot Analysis
Total RNA was prepared from ovaries at the indicated time points by homogenization in guanidine isothiocynate and centrifugation through cesium chloride (48, 49). Approximately 40 µg of each was separated by electrophoresis on denaturing 1% agarose/formaldehyde gels. RNA was transferred to a nylon membrane (Schleicher & Schuell, Keene, NH), baked in a vacuum oven at 80 C for 2 h, and hybridized at 42 C with 32P-dCTP-labeled ER{alpha} [the ~500-bp hormone-binding domain (20)] or ERß [the ~200-bp 5'-untranslated region (UTR) and A/B region (21)]-specific probe in 50% formamide, 5 x SSPE (750 mM NaCl, 50 mM NaH2PO4, pH 7.4, 1 mM EDTA), 2 x Denhardt’s reagent, 10% dextran sulfate, 0.1% SDS, and 100 µg/ml salmon sperm DNA. The membranes were subsequently washed in 0.1 x NaCl-sodium citrate (SSC) at 65 C and exposed to Kodak XAR-5 film (Eastman Kodak, Rochester, NY). After removal of probe in 50% formamide at 65 C, the membranes were rehybridized to cDNA clone CHO-B (50), which detects the LLRep3 gene family (51), to assess the amount of RNA present in each lane.

In Situ Hybridization
Ovaries were removed from storage at -80 C and brought to -20 C, and 20-µm sections were cut using a Zeiss cryostat. Sections were mounted onto positively charged glass slides, fixed in 5% paraformaldehyde (pH 7.5) for 5 min, washed in 2 x SSC for 5 min, rinsed in distilled deionized water, washed in 0.1 M triethanolamine (pH 8.0), and incubated in 0.25% acetic anhydride in 0.1 M triethanolamine (pH 8.0) for 10 min. Sections were dehydrated through an ethanol series and vacuum dried until hybridization. Antisense [33P]UTP- or [35S]UTP-labeled RNA probes were synthesized using SP6 or T7 RNA polymerase (48, 49). Templates were an EcoRI/PstI subclone of the rat ERß cDNA encoding the 5'-UTR and N-terminal A/B region (21), a PCR clone encoding the hormone-binding domain of the rat ER{alpha} cDNA (20), the rat LH-R PCR clone (34), and the rat PR PCR clone (29). The RNA probe (2 x 107 cpm/ml in hybridization buffer: 50% formamide, 5 x SSPE (750 mM NaCl, 50 mM NaH2PO4, pH 7.4, 1 mM EDTA), 2 x Denhardt’s reagent, 10% dextran sulfate, 0.1% SDS, and 100 mg/ml yeast tRNA) was applied to the tissue sections, and the sections were overlaid with a coverslip. Slides were hybridized in a humidity chamber at 47 C for 16–18 h. After hybridization, the coverslips were removed and sections were treated with RNase A (20 µg/ml) at 37 C for 30 min, washed in increasingly lower concentrations of SSC down to 0.1 x SSC at 55 C, and dehydrated through an ethanol series. The slides were exposed to Kodak XAR-5 film for 2–3 days at room temperature and were then processed for liquid emulsion autoradiography using NTB-2 emulsion (Kodak). Slides were developed using Kodak D-19 developer and fixer and stained with hematoxylin.

RT-PCR Analysis
Oligonucleotide primer pairs of 20–22 nucleotides (40–60% GC content) were designed based on the sequences of the rat ERß (21) [amplifying nucleotides from 1018 to 1221 (21)], ER{alpha} [amplifying nucleotides from 1439 to 1847 or 1547 (20)], and rat ribosomal protein S16 [amplifying nucleotides from 59 to 109 (52)]. The predicted sizes of the amplified products are 203 bp (ERß), 115 or 400 bp depending on the primer sets (ER{alpha}), and 100 bp (S16). The conditions were such that amplification of the product was linear with respect to the amount of input RNA. Five micrograms of total RNA were reverse-transcribed at 37 C using random hexamer primers and MMLV reverse transcriptase (New England Biolabs, Boston, MA) in a 20-µl reaction. Five microliters of the cDNA samples were used for the subsequent PCR amplification of ER{alpha}, ERß, and S16 cDNAs. A 20 µl mix including the oligonucleotide primers (50 ng each), {alpha}-32P-dCTP (2 µCi at 3000 Ci/mmol), and Taq DNA polymerase (2.5 U) in 1 x PCR buffer (10 mM Tris, pH 8.3, 50 mM KCl, 1.5 mM MgCl2, 0.01% gelatin) was added to each cDNA sample and overlaid with light mineral oil. Amplification was carried out for 25 cycles using an annealing temperature of 65 C on a Perkin Elmer Cetus thermalcycler (Perkin Elmer Cetus, Norwalk, CT). The samples were then electrophoresed on a 8% polyacrylamide gel. After autoradiography, the gel was analyzed using a Molecular Dynamics PhosphoImager and ImageQuant version 3 software (Molecular Dynamics, Sunnyvale, CA). The intensity of the ER{alpha} and ERß signal was normalized to that of the ribosomal protein S16 internal control.


    ACKNOWLEDGMENTS
 
The authors wish to thank Drs. Nancy Krett and Kevin Sarge for insightful comments on this work, and Dr. Sandra Legan for performing LH RIA.


    FOOTNOTES
 
Address requests for reprints to: Dr. Ok-Kyong Park-Sarge, Department of Physiology, University of Kentucky, Lexington, Kentucky 40536-0084.

This work was supported, in part, by NIH Grant HD-30719 and NIH Research Career Development Award HD-01135 (to O-K.P-S.). M.B. was supported, in part, by the NIH Training Grant in Reproductive Sciences to the University of Kentucky (T32-HD07436). G.G.J.M.K. was supported, in part, by grants from the Netherlands Organization for Scientific Research (NWO) and by a visiting scientist fellowship from the Karolinska Institute. A.J.G. was supported in part, by Swedish Cancer Fund.

Received for publication September 10, 1996. Revision received November 4, 1996. Accepted for publication November 11, 1996.


    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 

  1. Richards JS 1980 Maturation of ovarian follicles: actions and interactions of pituitary and ovarian hormones on follicular cell differentiation. Physiol Rev 60:51–89[Free Full Text]
  2. Hsueh AJW, Adashi EY, Jones PBC, Welch Jr TH 1984 Hormonal regulation of the differentiation of cultured granulosa cells. Endocr Rev 5:76–127[Medline]
  3. Erickson GF, Magoffin DA, Dyer CA, Hofeditz C 1995 The ovarian androgen producing cells: a review of structure/function relationships. Endocr Rev 6:371–399[Medline]
  4. Gore-Langton RE, Armstrong DT 1994 Follicular steroidogenesis and its control. In: Knobil E, Neill JD (eds) The Physiology of Reproduction, ed 2. Raven Press, New York, pp 571–627
  5. Greenwald GS, Roy SK 1994 Follicular development and its control. In: Knobil E, Neill JD (eds) The Physiology of Reproduction, ed 2. Raven Press, New York, pp 629–724
  6. Zeleznik AJ 1993 Dynamics of primate follicular growth. In: Adashi EY, Leung PCK (eds) The Ovary. Raven Press, New York, pp 261–317
  7. Goldenberg RL, Vaitukaitis JL, Ross GT 1972 Estrogen and folllicle stimulating hormone interactions on follicle growth in rats. Endocrinology 90:1492–1498[Medline]
  8. Richards JS, Jonassen JR, Rolfes AI, Kersey KA, Reichert Jr LE 1979 Adenosine 3',5' monophosphate, LH receptor, and progesterone production during granulosa cell differentiation: effects of estradiol and FSH. Endocrinology 104:765–773[Medline]
  9. Richards JS, Ireland JJ, Rao MC, Bernath GA, Midgley Jr AR, Reichert Jr LE 1976 Ovarian follicular development in the rat: hormone receptor regulation by estradiol, follicle-stimulating hormone, and luteinizing hormone. Endocrinology 99:1562–1570[Abstract]
  10. Walsh Jr TH, Zhang L-Z, Hsueh AJW 1983 Estrogen augmentation of gonadotropin-stimulated progestin biosynthesis in cultured rat granulosa cells. Endocrinology 112:1916–1924[Abstract]
  11. Leung PCK, Armstrong DT 1980 Further evidence in support of a short-loop feedback action of estrogen on ovarian androgen production. Life Sci 27:415–420[CrossRef][Medline]
  12. Merk FB, Botticelli CR, Albright JT 1972 An intercellular response to estrogen by granulosa cells in the rat ovary;an electron microscopic study. Endocrinology 90:992–1007[Medline]
  13. Gibori G 1993 The corpus luteum of pregnancy. In: Adashi EY, Leung PCK (eds) The Ovary. Raven Press, New York, pp 261–317
  14. Stumpf WE 1969 Nuclear concentration of 3H-estradiol in target tissues. Dry-mount autoradiography of vagina, oviduct, ovary, testis, mammary gland and adrenal. Endocrinology 85:31–37[Medline]
  15. Saiduddui S, Milor Jr GE 1971 3H-Estradiol uptake by the rat ovary. Proc Soc Exp Biol Med 138:651–660
  16. Richards JS 1974 Estradiol binding to rat corpora lutea during pregnancy. Endocrinology 95:1046–1053[Medline]
  17. Richards JS 1975 Estrogen receptor content in rat granulosa cells during follicular development: modification by estradiol and gonadotropins. Endocrinology 97:1174–1184[Abstract]
  18. Kawashima M, Greenwald GS 1993 Comparison of follicular estrogen receptors in rat, hamster and pig. Biol Reprod 48:172–179[Abstract]
  19. Saidudduin S, Zassenhaus HP 1977 Estradiol-17ß receptors in the immature rat ovary. Steroids 29:197–213[CrossRef][Medline]
  20. Koike S, Sakai M, Muramatsu M Molecular cloning, characterization of rat estrogen receptor c DNA 1987 Nucleic Acids Res 15:2499–2513[Abstract]
  21. Kuiper GGJM, Enmark E, Pelto-Huikko M, Nilsson S, Gustafsson J-Å 1996 Cloning of a novel estrogen receptor expressed in rat prostate and ovary. Proc Natl Acad Sci USA 93:5925–5930[Abstract/Free Full Text]
  22. Katzenellenbogen B 1980 Dynamics of steroid hormone receptor action. Annu Rev Physiol 42:17–35[CrossRef][Medline]
  23. Walters MR 1985 Steroid hormone receptors and the nucleus. Endocr Rev 6:512–543[Medline]
  24. Iwai T, Nanbu Y, Iwai M, Taii S, Fujii S, Mori T 1990 Immunoreceptors in the human ovary throughout the menstrual cycle. Virchows Arch [A] 417:369–375
  25. Billiar RB, Loukides JA, Miller MM 1992 Evidence for the presence of the estrogen receptor in the ovary of the baboon Papio anubis. J Clin Endocrinol Metab 75:1159–1165[Abstract]
  26. Suzuki T, Sasano H, Kimura N, Tamura M, Fukaya T, Yajima A, Nagura H 1994 Immunohistochemical distribution of progesterone, androgen, oestrogen receptors in the human ovary during the menstrual cycle: relationship to expression of steroidogenic enzymes. Hum Reprod 9:1589–1595[Abstract]
  27. Lubahn DB, Moyer JS, Golding TS, Couse JF, Korach KS, Smithies O 1993 Alteration of reproductive function but not prenatal sexual development after insertional disruption of the mouse estrogen receptor gene. Proc Natl Acad Sci USA 90:11162–11166[Abstract]
  28. Korach KS 1994 Insights from the study of animals lacking functional estrogen receptor. Science 266:1524–1527[Medline]
  29. Park OK, Mayo KE 1991 Transient expression of progesterone receptor messenger RNA in ovarian granulosa cells after the preovulatory luteinizing hormone surge. Mol Endocrinol 5:967–978[Abstract]
  30. Park-Sarge O-K, Mayo KE 1994 Regulation of the progesterone receptor gene by gonadotropins and cyclic adenosine 3',5'-monophosphate in rat granulosa cells. Endocrinology 134:709–718[Abstract]
  31. Iwai M, Yasuda K, Fukuoka M, Iwai T, Takakura K, Taii S, Nakanishi S, Mori T 1991 Luteinizing hormone induces progesterone receptor gene expression in cultured porcine granulosa cells. Endocrinology 129:1621–1627[Abstract]
  32. Chandrasekher YA, Melner MH, Nagalla SR, Stouffer RL 1994 Progesterone receptor, but not estradiol receptor, messenger ribonucleic acid is expressed in luteinizing granulosa cells and the corpus luteum in rhesus monkeys. Endocrinology 135:307–314[Abstract]
  33. Tora L, Mullick A, Metzger D, Ponglikitmongol M, Park I, Chambon P 1989 The cloned human estrogen receptor contains a mutation which alters its hormone binding properties. EMBO J 8:1981–1986[Abstract]
  34. Camp TA, Rahal JO, Mayo KE 1991 Cellular localization and hormonal regulation of follicle stimulating hormone and luteinizing hormone receptor messenger RNAs in the rat ovary. Mol Endocrinol 5:1405–1417[Abstract]
  35. Beato M; Herrlich P; Schutz G 1995 Steroid hormone receptors: many actors in search for a plot. Cell 83:851–7[Medline]
  36. Katzenellenbogen BS, Norman MJ 1990 Multihormonal regulation of the progesterone receptor in MCF-7 human breast cancer cells: interrelationships among insulin/insulin-like growth factor-I, serum, and estrogen. Endocrinology 126:891–898[Abstract]
  37. Couse JF, Curtis SW, Washburn TF, Lindzey J, Golding TS, Lubahn DB, Smithies O, Korach KS 1995 Analysis of transcription and estrogen insensitivity in the female mouse after targeted disruption of the estrogen receptor gene. Mol Endocrinol 9:1441–1454[Abstract]
  38. Wu T-CJ, Wang L, Wan Y-JY 1993 Detection of estrogen receptor messenger ribonucleic acid in human oocytes and cumulus-oocyte complexes using reverse transcriptase-polymerase chain reacion. Fertil Steril 59:54–59[Medline]
  39. Hou Q, Gorski J 1993 Estrogen receptor and progesterone receptor genes are expressed differentially in mouse embryos during preimplantation development. Proc Natl Acad Sci USA 90:9460–9464[Abstract]
  40. Vegeto E, Shahbaz MM, Wen DX, Goldman ME, O’Malley BW, McDonnell DP 1993 Human progesterone receptor A form is a cell- and promoter-specific repressor of human progesterone receptor B function. Mol Endocrinol 7:1244–1255[Abstract]
  41. Tung L, Mohamed MK, Hoeffler JP, Takimoto GS, Horwitz KB 1993 Antagonist-occupied human progesterone B receptors activate transcription without binding to progesterone response elements and are dominantly inhibited by A-receptors. Mol Endocrinol 7:1256–1265[Abstract]
  42. Wen DX, Xu YF, Mais DE, Goldman ME, McDonnell DP 1994 The A and B isoforms of the human progesterone receptor operate through distinct signaling pathways within target cells. Mol Cell Biol 14:8356–8364[Abstract]
  43. Kraus WL, Katzenellenbogen BS 1993 Regulation of progesterone receptor gene expression and growth in the rat uterus: modulation of estrogen actions by progesterone and sex steroid hormone antagonists. Endocrinology 126:891–898[Abstract]
  44. Read LD, Greene GL, Katzenellenbogen BS 1989 Regulation of estrogen receptor messenger ribonucleic acid and protein levels in human breast cancer cell lines by sex steroid hormones, their antagonists, and growth factors. Mol Endocrinol 3:295–304[Abstract]
  45. Ree AH, Knutsen HK, Landmark BF, Eskild W, Hansson V 1992 Down-regulation of messenger ribonucleic acid (mRNA) for the estrogen receptor (ER) by phorbol ester requires ongoing RNA synthesis but not protein synthesis. Is hormonal control of ER mRNA degradation mediated by an RNA molecule? Endocrinology 131:1810–1814[Abstract]
  46. Erickson GF 1983 Primary cultures of ovarian cells in serum-free medium as models of hormone- dependent differentiation. Mol Cell Endocrinol 29:21–49[CrossRef][Medline]
  47. Park-Sarge OK, Sarge KD 1995 Cis regulatory elements conferring cAMP-induced transcription of the rat progesterone receptor gene in transfected rat granulosa cells. Endocrinology 136:5430–5437[Abstract]
  48. Sambrook J, Fritsch EF, Maniatis T 1989 Molecular Cloning: A Laboratory Manual, ed 2. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
  49. Ausubel FM, Brent R, Kingston RE, Moore DD, Seidman JG, Smith JA, Struhl K 1989 Current Protocols in Molecular Biology. John Wiley & Sons, New York
  50. Harpold MM, Evans RM, Salditt-Georgieff M, Darnell JE 1979 Production of mRNA in Chinese hamster cells: relationship of the rate of synthesis to the cytoplasmic concentration of nine specific mRNA sequences. Cell 17:1025–1035[Medline]
  51. Heller DL, Gianola KM, Leinwand LA 1988 A highly conserved mouse gene with a propensity to form pseudogenes in mammals. Mol Cell Biol 8:2797–2803[Medline]
  52. Batra SK, Metzgar RS, Hollingsworth MA 1991 Molecular cloning and sequence analysis of the human ribosomal protein S16. J Biol Chem 266:6830–6833[Abstract/Free Full Text]