Prolactin receptor gene expression and immunolocalization of the ...

Molecular Human Reproduction Vol.7, No.10 pp. 1033–1038, 2001

Prolactin receptor gene expression and immunolocalization of the prolactin receptor in human luteinized granulosa cells Nikos P.Vlahos1,3, Elizabeth M.Bugg2, Michael J.Shamblott2, John Y.Phelps1, John D.Gearhart2 and Howard A.Zacur1 1Division

of Reproductive Endocrinology and 2Division of Developmental Genetics, Department of Gynecology and Obstetrics, The Johns Hopkins Medical Institutions, Baltimore, MD 21287, USA

3To

whom correspondence should be addressed at: Division of Reproductive Endocrinology, The Johns Hopkins Medical Institutions, 600 N Wolfe Street/Houck 246, Baltimore, MD 21287, USA. E-mail: [email protected]

Prolactin is mainly known for its role in breast development and lactation, but has been also implicated in other physiological functions such as immunoregulation and ovarian steroid production. Although prolactin and prolactin receptor (PRL-R) transcripts have been previously identified in the human ovary, the spatial localization of the receptor is unknown. To investigate the presence of PRL-R within the follicular apparatus, human luteinized granulosa cells were obtained at the time of follicular aspiration from women undergoing ovarian stimulation for IVF. RNA extracted from these cells was subjected to reverse transcriptase–polymerase chain reaction (RT–PCR) using specific primers for the PRL-R gene. In addition, paraffin sections of isolated granulosa cells and sections of premenopausal human ovaries were immunostained with a mouse anti-human PRL-R monoclonal antibody. PRLR were immunolocalized to the cell membrane of isolated luteinized granulosa cells and PRL-R transcripts were detected in the extracted RNA. No detectable staining was noted in secondary and early antral follicles in archived paraffin sections. These findings confirm the presence of PRL-R in human luteinized granulosa cells and suggest a localized role for PRL within the mature follicle. The absence of PRL-R in the early follicle suggests that the effects of prolactin are exerted around the time of ovulation. Key words: follicle/granulosa cells/ovulation/prolactin/prolactin receptor

Introduction Prolactin, which is primarily known for its role in the process of breast development and lactation (Glasier and McNeilly, 1990; Short, 1993; Clevenger and Plank, 1997; Cox et al., 1999; Horseman 1999), is also involved in a wide variety of physiological functions ranging from water transport by fetal membranes (Tyson et al., 1984), to ovarian steroid production (McNatty et al., 1974) and immunoregulation (Clevenger et al., 1998). Prolactin exerts its effects at the molecular level by binding and inducing homodimerization of the prolactin receptor (PRL-R). As a member of the cytokine receptor family, the PRL-R consists of three separate domains: an extracellular region with five cysteine residues containing the prolactin binding site, a transmembranic domain and a cytoplasmic region, the length of which appears to influence ligand binding and regulate cellular function. In the rat, two distinct PRL-R isoforms have been isolated; a long and a short form that differ in the length of the cytoplasmic domain (Boutin et al., 1988, 1989; Clevenger et al., 1998). A third © European Society of Human Reproduction and Embryology

intermediate isoform has also been detected in the Nb2 transformed rat T cell line. This is a truncated form of the long receptor missing the last 198 amino acids of the cytoplasmic domain due to a deletion of the last exon (Ali et al., 1991). A single gene located in chromosome 5 encodes the human PRL-R. A long isoform (599 amino acids) has been isolated from hepatoma (Hep G2) and breast cancer (T47D) cell lines, and this shares significant sequence identity with the growth hormone receptor, thus indicating a common ancestral origin (Boutin et al., 1989). In addition, an intermediate form (325 amino acids) of PRL-R has been recently isolated from the human breast carcinoma cell line T47D. This isoform results from the deletion of 573 bp from the long transcript due to an RNA splicing event. The splice induces a frame-shift causing the addition of 13 heterologous amino acids and a stop codon that leads to the deletion of a 274 codon segment of the intracellular domain (Kline et al., 1999). The role of PRL-R in the end-reproductive organs such as 1033

N.P.Vlahos et al.

the ovary and endometrium has been the subject of intense investigation. Prolactin as well as PRL-R have been immunolocalized in the granulosa cells surrounding the oocyte rat mature follicles and in all luteal cells studied (Dunaif et al., 1982). Transcripts of the long form of PRL-R have also been found predominantly in the mural granulosa cells of rat Graafian follicles and in corpora lutea (Clarke et al., 1993). In the human, synchronized temporal expression of prolactin and PRL-R has been described in the endometrium, with an up-regulation during the midluteal phase (Jabbour et al., 1998; Jones et al., 1998). Prolactin is present in the follicular fluid of the pre-ovulatory follicle (Laufer et al., 1984), and seems to exhibit a direct effect on steroidogenesis of cultured luteal cells (McNatty et al., 1974; Alila et al., 1987). Recently, Schwarzler et al. were able to identify PRL-R transcripts in RNA extracted from homogenized human ovaries (Schwarzler et al., 1997). Since qualitative analysis of PRL-R mRNA reflects the total pool of PRL-R mRNA in the ovary, this work does not provide any information about the specific identity of the cells expressing the PRL-R within the ovary. Furthermore, since white blood cells express PRL-R (Pellegrini et al., 1992; Dardenne et al., 1994), contamination of their samples with white blood cell RNA during the process of homogenization cannot be excluded. In this study we confirm the presence of PRL-R within the human ovary and clarify the spatial localization of the receptor in a specific cell type, by reporting PRL-R mRNA and PRL-R protein expression in human luteinized granulosa cells.

Materials and methods Tissue collection Follicular fluid was collected from six women undergoing transvaginal oocyte retrieval for IVF following ovarian stimulation. The technique for ovarian stimulation and oocyte retrieval has been described elsewhere (Garcia et al., 1990). Briefly, starting on the second day of their menstrual cycle, these women received a daily dose of 0.75 mg of leuprolide acetate (TAP, Deerfield, IL, USA), followed by a daily dose of 300 IU of highly purified FSH (Fertinex; Serono Laboratories Inc., Randolph, MA, USA) starting on menstrual day 5. When at least three follicles reached 18 mm in diameter, 10 000 IU of human chorionic gonadotrophin (HCG, Profasi; Serono Laboratories, Inc., Rome, Italy) was administered. Transvaginal oocyte retrieval was performed under sonographic guidance 36 h later. Granulosa cells were obtained from the follicular aspirates after the removal of the oocytes. Individual follicles were not distinguished and all follicular fluid from the same individual was pooled and centrifuged at 400 g for 5 min. The thin layer of follicular cells overlaying the red blood cells was gently aspirated using a glass Pasteur pipette and washed twice in cell culture medium 199 (GibcoBRL, Gaithersburg, MD, USA) containing Earle’s salts, 2200 mg/l sodium bicarbonate and 25 mmol/l HEPES. Cells were suspended in 3 ml of medium 199, overlaid upon 5 ml of 50% Percoll solution (Amersham Pharmacia Biothech, New Jersey, USA) and centrifuged at 400 g for 30 min to pellet the blood components. Follicular cells visible in the interphase were collected using a Pasteur pipette and washed twice in medium to remove the Percoll. To further reduce contamination of adherent peripheral mononuclear cells, the collected cells were incubated on a serum-coated 60 mm diameter culture dish (Falcon; Becton Dickinson Co., Lincon Park, NJ, USA), pretreated

1034

with autologous serum, for 60 min at 37°C in 95% air/5% CO2 atmosphere. The cells were collected by mild washing and resuspended in a small volume of the same medium. Cell viability was determined by Trypan Blue exclusion test and was found to be consistently ⬎90%. These cells were used for subsequent immunohistochemistry and RNA extraction. The cells from four women were retained separately for RNA extraction and the cells of the remaining two women were prepared for immunohistochemical analysis. In addition, archived paraffin blocks from the ovaries of four premenopausal women were sectioned and prepared for immunohistochemical analysis. Fixation and immunohistochemical analysis of these sections was performed according to the protocol described below for the granulosa cell suspensions. The collected cells were a by-product of the IVF– embryo transfer procedure and normally would have been discarded. As they were used in this study as coded samples with the identifiers of the participants unavailable, this study was granted an exemption from the institutional review board of the Johns Hopkins Medical Institutions. Immunohistochemisty Granulosa cells were re-suspended in a small volume of phosphatebuffered saline (PBS; Gibco-BRL) and mixed in 1% low melting point agarose prepared in PBS and brought to 42°C. The mixture was pipetted into moulds and allowed to solidify at room temperature. The agarose blocks were fixed overnight in 10% phosphate buffered formalin at room temperature. The agarose moulds were processed, embedded in paraffin and 6 µm sections were placed on microscope slides (ProbeOn Plus; Fisher Scientific, Pittsburgh, PA, USA). Sections were dewaxed through descending grades of ethanol to distilled water, transferred into trays containing sodium citrate buffer (HIER buffer Ventana-Biotek Solutions Inc., Tucson, AZ, USA) and steamed at 80°C for 20 min. Immunohistochemical analysis was performed in a BioTek Mate 1000 automatic stainer (Ventana-Biotek Solutions Inc., Tucson, AZ, USA) using a mouse anti-human PRL-R monoclonal antibody (Clone U5; Affinity Bioreagents, Inc., Golden, CO, USA) at a concentration of 2 µg/ml (dilution 1:500). To evaluate white blood cell contamination, additional sections were stained with a monoclonal mouse anti-human leukocyte common antigen (DACOLCA; DACO, Carpinteria, CA, USA) and by a monoclonal mouse anti-human myeloid/histiocyte antigen (DACO-MAC387; DACO). For further identification sections were also stained with a mouse anti-human inhibin A antibody (Serotec Inc., Raleigh, NC, USA). Primary antibodies were detected by using a biotinylated rabbit antimouse secondary antibody (dilution 1:500), strepavidin-conjugated horseradish peroxidase, and diaminobenzidine chromagen (VentanaBioTek Solutions Inc.). Slides were counterstained with haematoxylin. RNA extraction Total RNA was extracted from granulosa cells by using a commercially available kit (Qiagen Inc., Santa Clarita, CA, USA). Additional RNA, extracted from a human breast carcinoma cell line T-47D (HTB-133; American Type Culture Collection, Manassas, VA, USA), known to expresses the gene for PRL-R, was used as positive control. The quantity of the extracted RNA was assessed by measuring the optical density at Å 260 nm. The quality of the RNA was evaluated by electrophoresis on a 6.5% formaldehyde agarose gel and the integrity of the 18S and 28S rRNA was confirmed. Reverse transcription–polymerase chain reaction (RT–PCR) RNA (2 µg) from granulosa cells from each woman as well as from the T-47D cell line were used for the reaction. To eliminate genomic DNA contamination, all samples were treated with RNase-free DNase

Prolactin receptor expression in human granulosa cells (Boehringer Mannheim, Indianapolis, IN, USA) for 30 min at 37°C followed by incubation at 75°C for 5 min to inactivate the enzyme. The reverse transcription reaction was performed using Moloney Murine Leukemia Virus Reverse Transcriptase (Life Technologies, Carlsbad, CA, USA) for 60 min at 42°C, in the presence of polythymidine oligonucleotide primers (Life Technologies). PCR was performed using oligonucleotide primers complementary to the sequence encoding the extracellular domain of the human PRL-R and Taq polymerase (Life Technologies) for 40 cycles with denaturing, annealing and extension temperatures of 94, 48 and 72°C respectively. The sense strand oligonucleotide primer was 5⬘-ACT TAC ATA GTT CAG CCA GAC-3⬘ and the antisense strand primer was 5⬘-TGA ATG AAG GTC GCT GGA CTC C-3⬘ for annealing at nucleotide positions 364–383 and 673–652 respectively. The primers were designed to produce a 310 bp product. Internal nested PCR was performed for 30 cycles on the original PCR product with denaturing, annealing and extension temperatures of 95, 54 and 72°C respectively. The sense strand oligonucleotide primer was 5⬘-TTT GGA GCT GGC TGT GGA AG-3⬘ and the antisense strand primer was 5⬘-TGT CCT GGA TGT AGG CTG AGA ATC-3⬘ for annealing at nucleotide positions 390–409 and 599–576 respectively. This primer set was designed to produce a 210 bp product. To ensure that viable RNA was isolated, expression of the housekeeping gene glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was also assayed by RT–PCR. The sense strand primer was 5⬘-TCT CCT CTG ACT TCA ACA GCG AC-3⬘ and the antisense strand primer was 5⬘-AGT GAG GGT CTC TCT CTT CCT CTT G-3⬘. These primers were designed to produce a 220 bp product. To detect possible genomic DNA contamination, a duplicate tube, in which the reverse transcriptase enzyme was omitted, was assayed for each sample. Southern blot hybridization RT–PCR products were electrophoretically separated in 1% ethidium bromide agarose gel with reference to a molecular weight marker (123 bp DNA ladder; Gibco BRL) and transferred to an uncharged nylon membrane. The membrane was prehybridized in 6⫻SSC and 5⫻Denhardt’s solution for 4 h at 42°C. By using T4 polynucleotide kinase and [γ-32P]ATP, a 20 mer oligonucleotide, with a sequence of 5⬘-CCA CCT ACC CTG ATT GAC TT-3⬘ designed to anneal at PRL-R mRNA nucleotide positions 458–477 was end-labelled for 1 h at 37°C. The hybridization solution contained 6⫻standard saline citrate (SSC), 5⫻Denhardt’s, 100 µg/ml sheared and denatured herring sperm DNA, 0.05% sodium pyrophosphate and 0.1% sodium dodecyl sulphate (SDS). Hybridization was performed overnight at 42°C. The blot was subsequently washed at 42°C using a solution of 6⫻SSC, 0.05% sodium pyrophosphate, and 0.1% SDS. The hybridized blot was analysed using a Molecular Dynamic Storm phosphorimager (Molecular Dynamics, Sunnyvale, CA, USA).

Results Strong specific immunoreactivity for the PRL-R was detected in the cell membranes in all granulosa cell sections by immunohistochemistry (Figure 1A). Omission of the primary antibody did not reveal any staining in all examined sections (Figure 1B). All examined sections of isolated cells also demonstrated a strong signal for inhibin A, a well-characterized granulosa cell marker (Figure 1C) Careful inspection of several sections stained by DACOLCA and DACO-MAC387 showed minimal contamination by

Figure 1. Immunostained sections of granulosa cell preparations. (A) Strong specific prolatin receptor staining was observed in most of the cells. (B) Omission of the primary antibody revealed no detectable staining (negative control). (C) Strong specific staining for inhibin A confirmed the identity of the isolated cells. Scale bars ⫽ 50 µm.

white blood cells (Figure 2A and B). DACO-MAC387 would react with a human cytoplasmic antigen expressed in granulocytes, blood monocytes and tissue histiocytes as well as macrophages, whereas DACO-LCA would react with a common antigen (LCA) that is present on the surface of the majority of human leucocytes, thus labelling lymphoid cells and to a lesser degree macrophages and histiocytes. Evaluation of the ovarian sections showed numerous follicles in the early stages of development. Several primary and secondary follicles and a few early antral follicles (⬍3 mm in their largest diameter) were identified, but no specific PRL-R 1035

N.P.Vlahos et al.

Figure 2. Immunostained sections of granulosa cell preparations. Evaluation of white blood cell contamination by DACO-LCA and DACOMAC387. In all examined sections, staining for DACO-LCA was observed in only one cell in a single section (A) and no staining was identified for DACO-MAC387, indicating an essentially pure granulosa cell population (B). Scale bars ⫽ 50 µm. Figure 3. Human ovarian sections, immunostaining for prolactin receptor. Several follicles at different stages of development were observed; however, no detectable staining for prolactin receptor was observed in secondary (A) and early antral follicles (B). Scale bars ⫽ 50 µm.

staining was evident in these sections (Figure 3). In the examined sections we were unable to identify any intact preovulatory follicles. Inhibin A has been routinely used to stain similarly fixed ovarian sections (data not shown). Integrity of the mRNA-derived cDNA samples was verified by amplification of the housekeeping gene GAPDH in all patient samples as well as in the positive control. Following nested PCR for PRL-R mRNA, the expected 210 bp product was detectable by agarose gel electrophoresis in all patient samples as well as the breast carcinoma line T-47D (Figure 4). Southern blot analysis with a PRL-R-specific radiolabelled probe confirmed the identity of the nested PCR product in all four patient RNA samples as well as in the T-47D cell line. A weak signal was also detected in two of the first round PCR lanes (lanes 12 and 16). No hybridization signal was evident in any of the negative control lanes. Nucleotide sequence analysis identified the PCR product as the human PRL-R sequence.

Discussion This study confirms the presence of PRL-R in human granulosa cells. PRL-R expression was shown in all granulosa cell preparations by immunohistochemistry and PRL-R mRNA 1036

Figure 4. Nested reverse transcription–polymerase chain reaction (RT–PCR) for PRL-R generated the expected 210 bp amplified product as identified by comparison with a molecular weight marker. Southern blotting followed by radionucleotide probe hybridization confirmed the identity of the amplicon in all patient samples. P1 to P4, patient samples; T47-D, human breast carcinoma breast line. Lanes 1, 5, 8, 12, 16: original PCR reaction. Lanes 2, 6, 9, 13, 17: nested PCR reaction. Lanes 3, 7, 10, 14, 18: negative controls (with RT omitted). Lanes 4, 11, 15: empty.

transcripts were confirmed in total mRNA extracted from these cells. In the human, the role of prolactin within the follicular machinery has been supported from studies that demonstrated

Prolactin receptor expression in human granulosa cells

a direct effect of prolactin in steroidogenesis. In granulosa cell cultures, neutralization of prolactin by addition of rabbit antihuman prolactin serum in the medium suppresses progesterone production, whereas addition of human prolactin in serially increasing concentrations from 25 to 100 ng/ml results in a progressive increase in progesterone concentrations (McNatty et al., 1974). A biphasic response to prolactin, as shown in cultures of midcycle corpora lutea cells (Alila et al., 1987), suggested the presence of a PRL-R that requires a ligandinduced dimerization in order to promote signal transduction. At physiological concentrations, a single prolactin molecule binds to two receptor molecules and facilitates dimerization; however, in the presence of an excess of prolactin, dimerization could be inhibited and the signalling process would be interrupted. PRL-R have been readily characterized in the rodent model. Early studies using radiolabelled ovine prolactin (Roy et al., 1987) were able to demonstrate specific binding to granulosa cells of isolated hamster follicles, indirectly indicating the presence of PRL-R. In these experiments, prolactin binding was maximal during the early stages of follicular development (one to four layers of granulosa cells) and dropped considerably in more mature follicles. By using a biotin-labelled riboprobe, Shirota et al. were able to identify transcripts of the long form of PRL-R in newly formed rat corpora lutea immediately after ovulation (Shirota et al., 1995). Long PRL-R transcripts were also present in granulosa and theca cells of follicles at different stages of development; however, the intensity of the signal was highest in pre-ovulatory follicles. By in-situ hybridization and quantitative RT–PCR, Clarke et al. have described the relative distribution of the two PRL-R isoforms in the rat ovary during the oestrous cycle (Clarke et al., 1993). They were able to demonstrate that the amount of PCR product from the long receptor mRNA was 6–10-fold higher than that from the short receptor mRNA. In their experiments, long PRL-R transcripts were detected only in a subset of corpora lutea as well as in many large pre-ovulatory follicles and preantral follicles, whereas short PRL-R transcripts were present in all cell types as well as in the cumulus of large pre-ovulatory follicles. Telleria et al. have been able to show that both PRL-R isoforms (predominantly the long one) are consistently expressed throughout pregnancy in isolated rat corpora lutea (Telleria et al., 1997). In animals that had undergone hypophysectomy on day 3 of pregnancy, 4 days of treatment with prolactin resulted in a significant increase in the expression of the long form of PRL-R only. Since there is a common primary transcript for both receptors it seems that prolactin exerts this effect at a post-transcriptional level. Another interesting observation in this study was a markedly decreased expression of both PRL-R isoforms after neutralization of the circulating serum LH by treating these animals with LH antiserum. This suggests a possible stimulatory role of LH in the regulation of PRL-R. In studies designed to evaluate PRL-R expression and postreceptor signalling during follicular development in the rat model (Russell and Richards, 1999), ovaries from hypophysectomized untreated animals contained near undetectable levels

of PRL-R mRNA. However, animals treated with oestradiol and FSH showed an impressive up-regulation of PRL-R in pre-ovulatory follicles (27-fold for the long form and 2-fold for the short). A further increase in PRL-R expression (4-fold for the long form and 10-fold for the short form) was noted when these animals were treated with an ovulatory dose of HCG. In contrast to previous findings during pregnancy (Telleria et al., 1997), treatment of these animals with prolactin resulted in a significant suppression of PRL-R. A significant amount of data has been collected about the temporal and spatial expression of PRL-R in the human endometrium where they seem to play an important role in implantation and subsequent placentation (Jabbour et al., 1998; Jones et al., 1998). Our knowledge about the distribution of PRL-R within the human ovary is still limited. In this study, we provide evidence for the presence of PRL-R on isolated luteinized human granulosa cells by immunohistochemical methods as well as by detection of PRL-R mRNA transcripts. In contrast with rodent data (Roy et al., 1987; Clarke et al., 1993; Shirota et al., 1995), we were unable to show staining in any of the ovarian sections containing primary, secondary or early antral follicles. Thus it is possible that PRL-R expression in the human may follow a different pattern with expression occurring during the late stages of follicular development. A possible role of HCG may also be suggested. In our study, all granulosa cells were obtained from mature follicles that had been exposed to HCG 36 h prior to retrieval. HCG has been used in ovarian stimulation protocols to mimic the action of the pre-ovulatory LH surge. In a natural cycle, the LH surge signals the process of ovulation and initiates a cascade of events that result in follicular and oocyte maturation. In contrast, all the follicles observed in the ovarian sections were in the early stages of development, therefore it is unlikely that they had been exposed to endogenous LH. It is possible that exposure of the follicular apparatus to LH or HCG stimulates PRL-R expression and subsequent PRL-R mediated mechanisms which, in combination with the high prolactin levels present in the pre-ovulatory follicle (Laufer et al., 1984), may have a role in luteinization and progesterone production. Rodent data (Telleria et al., 1997; Russell and Richards, 1999) support a stimulatory role for LH and HCG in the regulation of PRL-R. Whereas it is difficult to extrapolate from the rodent studies, it is possible that, in the human, either LH in a natural cycle or HCG in a stimulated cycle may up-regulate PRL-R in the pre-ovulatory follicle to facilitate luteal transition and to support the luteotrophic effect of prolactin. Another potential role for the expression of PRL-R during the late follicular phase may involve interleukin-1β (IL-1β). As an important mediator in the mechanism of ovulation, IL1β contributes to the follicular wall disruption by stimulating prostaglandin E2 (PGE2) and collagenase production (Takehara et al., 1994; Wallach and Vlahos, 1998). Follicular macrophages as well as granulosa cells are responsible for IL-1β production. Furthermore, there is evidence that granulosa cells may interact with macrophages to enhance IL-1β production (Machelon et al., 1995). Since PRL-R are present on tissue macrophages (Pellegrini et al., 1992; Dardenne et al., 1994), the appearance of PRL-R in luteinized granulosa cells prior to 1037

N.P.Vlahos et al.

ovulation may suggest a possible role for PRL in the cooperation of these cell populations. A major concern in our experiment is possible contamination of the RNA samples from cells of the haematopoietic line that express PRL-R. The issue of sample contamination represents a real threat to any project that involves PCR. To evaluate the extent of contamination, we used two different types of immunostaining to identify the presence of any tissue macrophages as well as any other cells that potentially could express PRL-R. Staining of the granulosa cell sections with DACOCLA and DACO-MAC387 demonstrated that our samples were relatively pure with minimal contamination. A strong positive inhibin A signal further confirmed the identity of those cells, and finally the strong specific staining for the PRL-R confirmed that the amplified transcripts were originating from the collected granulosa cells. In summary, our data confirm the presence of the PRL-R in human luteinized granulosa cells. The absence of staining for PRL-R in the preantral and early antral follicles and the presence of PRL-R in the luteinized cells suggest that the appearance of PRL-R is a late event in the process of folliculogenesis. Whether the pre-ovulatory LH surge is involved in this event directly or through another mediator remains to be established. The presence of PRL-R in preovulatory granulosa cells may also provide an attractive mechanism for the coordination between granulosa cells and follicular macrophages in the process of ovulation. Further studies are needed to elucidate the pattern of temporal expression of PRL-R within the human follicle.

References Ali, S., Pellegrini, I. and Kelly, P.A. (1991) A prolactin-dependent immune cell line (Nb2) expresses a mutant form of prolactin receptor. J. Biol. Chem., 266, 20110–20117. Alila, H.W., Rogo, K.O. and Gombe, S. (1987) Effects of prolactin on steroidogenesis by human luteal cells in culture. Fertil. Steril., 47, 947–955. Boutin, J.M., Jolicoeur, C., Okamura, H. et al. (1988) Cloning and expression of the rat prolactin receptor, a member of the growth hormone/prolactin receptor gene family. Cell, 53, 69–77. Boutin, J.M., Edery, M., Shirota, M. et al. (1989) Identification of a cDNA encoding a long form of prolactin receptor in human hepatoma and breast cancer cells. Mol. Endocrinol., 3, 1455–1461. Clarke, D.L., Arey, B.J. and Linzer, D.I. (1993) Prolactin receptor messenger ribonucleic acid expression in the ovary during the rat estrous cycle. Endocrinology, 133, 2594–603. Clevenger, C.V. and Plank, T.L. (1997) Prolactin as an autocrine/paracrine factor in breast tissue. J. Mamm. Gland Biol. Neoplasia, 2, 59–68. Clevenger, C.V., Freier, D.O. and Kline, J.B. (1998) Prolactin receptor signal transduction in cells of the immune system. J. Endocrinol., 157, 187–97. Cox, D.B., Kent, J.C., Casey, T.M. et al. (1999) Breast growth and the urinary excretion of lactose during human pregnancy and early lactation: endocrine relationships. Exp. Physiol., 84, 421–34.

1038

Dardenne, M., de Moraes, M.d.C., Kelly, P.A. et al. (1994) Prolactin receptor expression in human hematopoietic tissues analyzed by flow cytofluorometry. Endocrinology, 134, 2108–21014. Dunaif, A.E., Zimmerman, E.A., Friesen, H.G. et al. (1982) Intracellular localization of prolactin receptor and prolactin in the rat ovary by immunocytochemistry. Endocrinology, 110, 1465–1471. Garcia, J.E., Padilla, S.L., Bayati, J. et al. (1990) Follicular phase gonadotropinreleasing hormone agonist and human gonadotropins: a better alternative for ovulation induction in in vitro fertilization. Fertil. Steril., 53, 302–305. Glasier, A. and McNeilly, A.S. (1990) Physiology of lactation. Baillie`res Clin. Endocrinol. Metab., 4, 379–395. Horseman, N.D. (1999) Prolactin and mammary gland development. J. Mamm. Gland Biol. Neoplasia, 4, 179–88. Jabbour, H.N., Critchley, H.O. and Boddy, S.C. (1998) Expression of functional prolactin receptors in nonpregnant human endometrium: janus kinase-2, signal transducer and activator of transcription-1 (STAT1), and STAT5 proteins are phosphorylated after stimulation with prolactin. J. Clin. Endocrinol. Metab., 83, 2545–2553. Jones, R.L., Critchley, H.O., Brooks, J. et al. (1998) Localization and temporal expression of prolactin receptor in human endometrium. J. Clin. Endocrinol. Metab., 83, 258–262. Kline, J.B., Roehrs, H. and Clevenger, C.V. (1999) Functional characterization of the intermediate isoform of the human prolactin receptor. J. Biol. Chem., 274, 35461–35468. Laufer, N., Botero-Ruiz, W., DeCherney, A.H. et al. (1984) Gonadotropin and prolactin levels in follicular fluid of human ova successfully fertilized in vitro. J. Clin. Endocrinol. Metab., 58, 430–4. Machelon, V., Nome, F., Durand-Gasselin, I. et al. (1995) Macrophage and granulosa interleukin-1 beta mRNA in human ovulatory follicles. Hum. Reprod., 10, 2198–203. McNatty, K.P., Sawers, R.S. and McNeilly, A.S. (1974) A possible role for prolactin in control of steroid secretion by the human Graafian follicle. Nature, 250, 468, 653–655. Pellegrini, I., Lebrun, J.J., Ali, S. et al. (1992) Expression of prolactin and its receptor in human lymphoid cells. Mol. Endocrinol., 6, 1023–1031. Roy, S.K., Wang, S.C. and Greenwald, G.S. (1987) Radioreceptor and autoradiographic analysis of FSH, hCG and prolactin binding sites in primary to antral hamster follicles during the periovulatory period. J. Reprod. Fertil., 79, 307–313. Russell, D.L. and Richards, J.S. (1999) Differentiation-dependent prolactin responsiveness and stat (signal transducers and activators of transcription) signaling in rat ovarian cells. Mol. Endocrinol., 13, 2049–2064. Schwarzler, P., Untergasser, G., Hermann, M. et al. (1997) Prolactin gene expression and prolactin protein in premenopausal and postmenopausal human ovaries. Fertil. Steril., 68, 696–701. Shirota, M., Kurohmaru, M., Hayashi, Y. et al. (1995) Detection of in situ localization of long form prolactin receptor messenger RNA in lactating rats by biotin-labeled riboprobe. Endocr. J., 42, 69–76. Short, R.V. (1993) Lactational infertility in family planning. Ann. Med., 25, 175–80. Takehara, Y., Dharmarajan, A.M., Kaufman, G. et al. (1994) Effect of interleukin-1 beta on ovulation in the in vitro perfused rabbit ovary. Endocrinology, 134, 1788–1793. Telleria, C.M., Parmer, T.G., Zhong, L. et al. (1997) The different forms of the prolactin receptor in the rat corpus luteum: developmental expression and hormonal regulation in pregnancy. Endocrinology, 138, 4812–4820. Tyson, J.E., Mowat, G.S. and McCoshen, J.A. (1984) Simulation of a probable biologic action of decidual prolactin on fetal membranes. Am. J. Obstet. Gynecol., 148, 296–300. Wallach, E.E. and Vlahos, N.P. (1998) The mechanism of mammalian ovulation. Israel J. Obstet. Gynecol., 9, 127–131. Received on April 30, 2001; accepted on August 8, 2001