Molecular Human Reproduction, Vol. 5, No. 4, 291-298,
April 1999
© 1999 European Society of Human Reproduction and Embryology
Steroidogenic enzyme expression in human corpora lutea in the presence and absence of exogenous human chorionic gonadotrophin (HCG)
1 MRC Reproductive Biology Unit, Centre for Reproductive Biology, 37 Chalmers Street, Edinburgh EH3 9EW, UK
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In a human conception cycle, the expected decline in progesterone production by the corpus luteum during the late luteal phase is prevented by human chorionic gonadotrophin (HCG) secreted by the implanting blastocyst. This study investigated the expression of components of the synthetic pathway for progesterone in human corpora lutea in the presence and absence of HCG in vivo. Corpora lutea were obtained from: (i) normally cycling women at the time of hysterectomy and classified on the basis of the urinary luteinizing hormone (LH) surge as early (n = 3), mid- (n = 3), or late luteal (n = 3); or (ii) women who had received daily doubling doses of HCG (n = 3) to `rescue' the corpus luteum. Expression patterns of steroidogenic acute regulatory protein (StAR), cytochrome P450 cholesterol side-chain cleavage (P450scc) and 3ß-hydroxysteroid dehydrogenase (3ß-HSD) were investigated by Northern blotting, in-situ hybridization and immunohistochemistry. Luteal `rescue' with HCG was associated with the continued expression of these components. In the late luteal phase, in the absence of HCG, expression remained but was more variable. The expression of 3ß-HSD mRNA was significantly reduced during the luteal phase (P < 0.01). In conclusion, during luteal `rescue', HCG acts to maintain the steroidogenic pathway. In the absence of HCG, the decline in progesterone production begins in the presence of the main components of the steroidogenic pathway. While unlikely to initiate this decline, the altered expression levels of these components, particularly that of 3ß-HSD, may contribute to the continued reduction in progesterone production.
corpus luteum/enzymes/HCG/steroidogenesis/steroidogenic acute regulatory protein
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In non-conception cycles, the human corpus luteum will undergo luteolysis, with loss of functional and structural integrity after 14 days. In the presence of human chorionic gonadotrophin (HCG) from the implanting blastocyst, the corpus luteum is `rescued' and its function and structure are maintained (Behrman et al., 1993
). The molecular mechanisms of luteolysis and how they are prevented by exposure to HCG are still not clear in the primate.
Progesterone production by the primate corpus luteum is dependent on luteinizing hormone (LH) from the anterior pituitary gland (Hutchison and Zeleznik, 1984; Fraser et al., 1986). LH binds to, and activates, a specific glycoprotein LH/HCG receptor present on the membranes of steroidogenic luteal cells (McNeilly et al., 1980; Bramley et al., 1987; Segaloff and Ascoli, 1993). The LH receptor is a G protein coupled receptor with a large glycosylated extracellular domain, seven transmembrane loops, and a smaller intracellular C-terminal domain (Segaloff and Ascoli, 1993). Specific ligand binding activates second messenger systems, notably cyclic AMP (cAMP) (Segaloff and Ascoli, 1993). This stimulates the uptake of cholesterol into mitochondria by the action of steroidogenic acute regulatory protein (StAR) (Stocco and Clark, 1996), its conversion into pregnenolone by the enzyme cytochrome P450 cholesterol side-chain cleavage (P450scc) (Simpson and Boyd, 1967) and the further conversion of pregnenolone into progesterone by 3ß-hydroxysteroid dehydrogenase (3ß-HSD) (Strauss and Miller, 1991).
Luteolysis can be induced by the withdrawal of LH, by inhibition of gonadotrophin releasing hormone (GnRH) (Hutchison and Zeleznik, 1984; Fraser et al., 1986). However, LH withdrawal is not the cause of luteolysis in natural cycles, as luteolysis still occurs in the presence of continued exposure to LH in the late luteal phase (Hutchison et al., 1986). This suggests that functional luteolysis is associated with an increasing block to LH action within the corpus luteum. Therefore, we recently investigated the expression of LH/HCG receptors in the human corpus luteum (Duncan et al., 1996a). We, and others, have shown that the primate corpus luteum continues to express LH receptors at a time when progesterone synthesis is declining (Ravindranath et al., 1992a; Nishimori et al., 1995; Duncan et al., 1996a). Recent reports have suggested that the rate-limiting step in steroidogenesis may be the transport of cholesterol to the inner mitochondrial membrane under the action of StAR (Clark et al., 1994; Stocco and Clark, 1996). Therefore, in this study we aimed to investigate the expression of components of the steroidogenic pathway beyond the LH/HCG receptor, notably StAR, P450scc and 3ß-HSD, in the human corpus luteum throughout the luteal phase and after luteal `rescue' with HCG, in vivo, to simulate early pregnancy.
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Source of reagents
All reagents were obtained from Sigma Chemical (Poole, Dorset, UK), unless otherwise stated. A 1.2 kb cDNA construct of human 3ß-HSD in pIBI25 was supplied by Professor J.I.Mason, Dept of Clinical Biochemistry, University of Edinburgh, Edinburgh, UK, and a polyclonal antibody to human type I 3ß-HSD was kindly provided by Prof. Van Luu-The (CHUL Research Centre, Quebec, Canada). A 600 bp cDNA fragment of bovine P450scc was obtained from Professor A.S.McNeilly, MRC Reproductive Biology Unit, Edinburgh, UK.
Collection of tissue
Corpora lutea were enucleated at the time of hysterectomy in 12 women undergoing surgery for benign conditions as previously described (Duncan et al., 1996a). All women were healthy, aged 32–45 years, with regular menstrual cycles and had not received any form of hormonal treatment for at least 3 months prior to taking part in the study. The corpora lutea were dated on the basis of serial urinary LH measurements on samples collected daily prior to operation (Djahanbakhch et al., 1981a). On this basis, three corpora lutea were classified as early (LH+1 to LH+5), three as mid (LH+6 to LH+10) and three as late (LH+11 to LH+14). In addition, three women received daily i.m. injections of HCG (Profasi, Serono Laboratories, Welwyn Garden City, UK) from LH+7 in daily doubling doses, starting at 125 IU, for 5–8 days until surgery. This regimen has been shown to reproduce the hormonal changes of early pregnancy (Illingworth et al., 1990).
The whole corpus luteum was enucleated from the ovary by blunt dissection and the ovary oversewn as described by Duncan et al. (1996a,b). The tissue was immediately divided into radial blocks in order to ensure that the whole thickness of the gland was represented in any piece. One piece was fixed in 4% paraformaldehyde for 24 h and embedded in paraffin wax for subsequent immunohistochemistry, and another piece was frozen in embedding medium (Tissue-Tek OCT compound; Miles Inc, Elkhart, USA) and stored at –70°C. Frozen sections (5 µm) were cut from this block onto poly-L-lysine (50 µg/l)-coated slides and stored at –70°C until use. A third piece was snap-frozen in liquid nitrogen and stored at –70°C for subsequent RNA extraction. In each case an endometrial biopsy was also fixed in paraformaldehyde and processed into paraffin wax for luteal phase dating by tissue morphometry (Li et al., 1988). Plasma was taken before surgery and progesterone concentration was measured by a standard radioimmunoassay (Djahanbakhch et al., 1981b). This study was approved by the Reproductive Medicine Subcommittee of the South East Scotland Research Ethics Committee, and informed consent was obtained from all patients prior to tissue collection.
Cloning of StAR from the human corpus luteum
Total cellular RNA was extracted from a mid-luteal human corpus luteum by the method of Chomczynski and Sacchi (1987) and reverse transcribed into cDNA using a commercial kit (Promega, Southampton, UK). Specific oligonucleotide primers for StAR were synthesized by Oswell DNA Services (Southampton, UK). The primer sequences, 5'-AACCAGGAAGGCTGGAA-3' and 5'-CCATGCAGGTGGGGCCGTGTTCAGC-3' (Clark et al., 1994) were used to amplify the expected 400 bp fragment of StAR cDNA using the polymerase chain reaction conditions as described previously (Clark et al., 1994). This was cloned into a plasmid PCR II vector using a commercial kit (TA Cloning Kit; Invitrogen NV, Leek, The Netherlands) and sequenced using a Taq dye deoxy termination cycle sequencing kit (Applied Biosystems, Warrington, UK) using an automatic sequencer (373A; Applied Biosystems). Data were analysed using a commercial computer program (GeneJockey II; Biosoft, Cambridge, UK).
Northern blotting
Total cellular RNA was isolated by the method of Chomczynski and Sacchi (1987) using a commercial kit, and its concentration was determined by absorption at 260 nm. Total RNA (20 µg) was denatured, electrophoresed in a 1.5% formaldehyde–agarose gel and transferred to a nylon membrane (Amersham International plc, Aylesbury, Bucks, UK) by capillary action in 20x SSC (1x SSC is 150 mM NaCl, 15 mM sodium citrate, pH 7). RNA was then fixed onto the membranes by UV cross-linkage (Spectronics Corporation, New York, USA). Northern blot analysis was conducted as described previously (Duncan et al., 1996a).
Briefly, membranes were pre-hybridized for 5 h in 15 ml hybridization buffer [0.5M sodium phosphate, 1 mM EDTA, 1% (w/v) bovine serum albumin (BSA), 7% (w/v) sodium dodecyl sulphate (SDS), 6.7% (v/v) deionized formamide] at 65°C. The cDNA probes were labelled with 50 µCi [32P]-dCTP by the random priming method using a commercial kit (Amersham International plc) and added to the hybridization buffer for 20 h at 65°C. The membranes were washed twice at 65°C with 2x SSC for 15 min and once more with 2x SSC/0.1% SDS at 65°C for 15 min. The blots were then laid down to a phosphor screen for 48 h and visualized using a phosphorimager computer (Molecular Dynamics, Maidstone, Kent, UK). The blots were then stripped and then re-probed with a [32P]-end-labelled oligonucleotide which hybridizes to 18S RNA as described previously (Brooks et al., 1992).
In-situ hybridization
Isotopic in-situ hybridization was performed on frozen sections using [35S]-labelled riboprobes using the technique described previously (Duncan et al., 1996a). The antisense 3ß-HSD riboprobe, incorporating [35S]-labelled UTP (Amersham International plc), was generated by T7 RNA polymerase (Promega) using a commercial kit (Promega). Frozen sections (5 µm) on poly-L-lysine (50 µg/l)-coated slides were quickly thawed and fixed in 4% paraformaldehyde for 5 min at room temperature. After washing in 0.1 M sodium phosphate, slides were rinsed firstly in water and then in 0.1 M triethanolamine (TEA) pH 8. The slides were then acetylated in 0.25% (v/v) acetic anhydride (BDH Laboratory Supplies, Poole, UK) in TEA. After acetylation, the slides were washed in 2x SSC pH 7, and dehydrated through graded alcohols. The slides were then dried under vacuum in a desiccator for 1 h at room temperature.
Hybridization buffer (100 µl; 50% deionized formamide, 10% dextran sulphate, 1x Denhardt's solution, 0.5 mg/ml yeast tRNA, 10 mM dithiothreitol (DTT), 0.3 M NaCl, 10 mM Tris, 1 mM EDTA pH 8) containing 1x106 c.p.m. radiolabelled probe was added to each section. The slides were covered with a hydrophobic coverslip (Gel Bond; ICN Biomedical Ltd, High Wycombe, UK) and incubated overnight at 55°C in a moist chamber. The following day the coverslips were washed off in 4x SSC. After several rinses in 4x SSC, the slides were treated with RNAse A (20 µg/ml) in RNAse buffer (10 mM Tris, 1 mM EDTA, 0.5 M NaCl, pH 8) for 30 min at 37°C. The sections were de-salted by rinsing in 2x SSC/1 mM DTT, followed by 1x SSC/1 mM DTT and 0.5x SSC/1 mM DTT at room temperature. The slides were then washed for 30 min in 0.1x SSC at 70°C in a shaking water bath. After rinsing in 0.1x SSC/1 mM DTT at room temperature, the sections were dehydrated through graded alcohols containing 1 mM DTT and 0.08 x SSC, washed in pure ethanol and allowed to dry.
The slides were then dipped in photographic emulsion (Kodak NTB-2; IBI Ltd, Cambridge, UK) and stored at 4°C for 18 days in the dark. After developing (Kodak D-19) and fixing (Kodak Unifix) at 15 °C in the dark, the slides were washed in water, counterstained in haematoxylin, dehydrated through graded alcohols and mounted in Pertex mounting medium (Cellpath, Hemel Hempstead, UK). The sections were viewed and photographed under dark-field illumination. The localization of the grains was determined by reference to the section viewed under bright-field.
Immunohistochemistry
Paraffin wax sections (5 µm) were cut onto poly-L-lysine (50 µg/l)-coated slides, then de-waxed and rehydrated. Endogenous peroxidase activity was blocked with 2% (v/v) hydrogen peroxide in 60% methanol for 30 min at room temperature. This tissue was then permeabilized with 0.1% Triton-X100 in 0.05M Tris-buffered saline, pH 8.0 (TBS) and rinsed in TBS prior to blocking with 20% (v/v) normal goat serum (NGS) (SAPU; Carluke, Scotland, UK) in TBS with 4% (w/v) BSA for 20 min. Sections were incubated overnight at 4°C with the polyclonal rabbit anti-human 3ß-HSD antiserum diluted 1:1000 in 20% NGS in TBS. The following day, the sections were washed with TBS and then incubated with biotinylated goat anti-rabbit immunoglobulins (Ig) (Dako Ltd, High Wycombe, Bucks, UK) diluted 1:500 in TBS, for 30 min at room temperature. After being washed with TBS, sections were incubated with avidin–biotin horseradish peroxidase complex (Dako Ltd) for 30 min at room temperature, then washed again with TBS and developed with diamino-benzidine (DAB) to give a brown end product (Vector Laboratories, Peterborough, UK). Sections were counterstained with haematoxylin, dehydrated through graded alcohols and cleared in xylene prior to mounting. Polyclonal rabbit IgG (Dako Ltd) at the same antibody concentration was used in place of the primary antibody, in serial sections, as a negative control.
Analysis of results
Northern blot band intensity was measured using the phosphorimager computer. To correct for minor differences in loading, the ratio of the relative band intensity to the 18S band intensity was used for data analysis. One-way analysis of variance was used to investigate differences in expression throughout the luteal phase. The early luteal corpora lutea were compared to the mid- and late luteal corpora lutea using an unpaired t-test. A commercial software package was used for statistical analysis (StatView 4.0; Abacus Concepts Inc., Berkeley, CA, USA). Immunohistochemical staining for 3ß-HSD was assessed in sections from the same run performed under carefully controlled conditions by a observer blinded to tissue identity. This was repeated two weeks later to confirm consistency of scoring. The staining intensity in the granulosa–lutein cell layer was graded as absent (–), faint (+), moderate (++), intense (+++) or very intense (++++) for each section. Where the sections differed between each scoring session an intermediate value was given.
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Progesterone concentrations
The classification of the corpora lutea by serial urinary LH measurement agreed with the luteal-phase dating of endometrial biopsies using the method of Li et al. (1988). The plasma progesterone concentrations were 35.3 ± 9.8 nmol/l in the early luteal samples, 41.0 ± 9.9 nmol/l in the mid-luteal samples and 19.2 ± 12.9 nmol/l in the late luteal samples. After luteal rescue by exogenous HCG the plasma progesterone concentrations had increased to 52.6 ± 1.5 nmol/l.
Cloning of human StAR
The specific oligonucleotide primers for StAR amplified the expected 400 bp fragment (Clark et al., 1994) from cDNA transcribed from mRNA extracted from the mid-luteal human corpus luteum. When this fragment was sequenced, the sequence was 97% identical to the sequence for human StAR in the gene sequence databases. This StAR cDNA sequence was used to study the expression of StAR mRNA in the human corpus luteum.
Expression of steroidogenic enzymes in human corpora lutea
Specific mRNA transcripts for StAR, P450scc and 3ß-HSD could be detected in corpora lutea from different stages of the luteal phase and after luteal `rescue' with exogenous HCG (Figure 1). Two major StAR mRNA transcripts of 1.7 and 4.8 kb were detected. These are consistent with the size of human StAR transcripts already published (Kiriakidou et al., 1996). Major transcripts of 2.0 and 1.7 kb were detected in corpora lutea after northern blotting for P450scc and 3ß-HSD respectively (Figure 1) These are consistent with the expected size of the mRNAs for these steroidogenic enzymes in the primate (Doody et al., 1990; Bassett et al., 1991).
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Changes in steroidogenic enzyme expression during the luteal phase
As can be seen in Figure 1
expression the steroidogenic enzymes did not appear to be constant throughout the luteal phase. In the late luteal phase, band intensity was more variable than during other stages of the luteal phase. Band intensity was measured and corrected for 18S RNA expression for each steroidogenic enzyme (Figure 2). It can be seen that expression of enzymes tended to be lower in the late luteal phase. However, this trend did not reach statistical significance for any of the enzymes. However, when the mid- and late luteal phases were combined, the expression of 3ß-HSD was markedly lower (P < 0.01) than in the early luteal phase or after luteal `rescue' with HCG. The most marked change in steroidogenic enzyme expression throughout the luteal phase was that of 3ß-HSD.
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Localization and expression of 3ß-HSD in human corpora lutea
As Northern blotting revealed that 3ß-HSD changed most during the luteal phase, its expression and localization were further investigated by isotopic in-situ hybridization and immunohistochemistry. mRNA could be detected by in-situ hybridization in corpora lutea at all stages of the luteal phase. However, during the late luteal phase, detection of 3ß-HSD mRNA was variable. The hybridization signal at the beginning of the late luteal phase (Figure 3a
) was similar to that seen in the mid-luteal phase. However, towards the end of the late luteal phase much less 3ß-HSD mRNA could be detected by in-situ hybridization (Figure 3c). In contrast, 3ß-HSD protein, as detected by immunohistochemistry, could be detected throughout the late luteal phase (Figure 3b,d). When 3ß-HSD immunostaining was analysed, the intensity did not vary across the luteal phase or after luteal `rescue' with exogenous HCG (Figure 4).
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We have previously reported the expression of LH/HCG receptors in human corpora lutea throughout the luteal phase and after luteal `rescue' with HCG (Duncan et al., 1996a
). We found that LH/HCG receptor mRNA and binding activity were retained in corpora lutea during the late luteal phase when progesterone production is falling. There is a decline in LH secretion during the luteal phase (Ellinwood et al., 1984), but this is not wholly responsible for the falling luteal progesterone production (Hutchison et al., 1986). The reasons for reduced progesterone synthesis in the late luteal phase in the presence of LH and LH receptors are, therefore, not yet clear. The stage of steroidogenesis which is generally thought to be rate-limiting is the conversion of cholesterol to pregnenolone (Strauss and Miller, 1991; Stocco and Clark, 1996). This stage involves the actions of both StAR (Stocco and Clark, 1996) and P450scc (Simpson and Boyd, 1967). As the tools are now available to investigate StAR expression at a molecular level, in this follow-up study, we looked at its expression during the luteal phase and compared its expression with that of the steroidogenic enzymes involved in progesterone synthesis, P450ssc and 3ß-HSD.
We have cloned StAR from the human corpus luteum and shown it to be expressed throughout the luteal phase and after luteal `rescue' with exogenous HCG. Since it was first characterized at a molecular level in 1994 (Clark et al., 1994), it has become increasingly clear that StAR has a fundamental role in the control of steroidogenesis (King et al., 1995; Lin et al., 1995). It has been shown to be involved in steroidogenesis in ovine (Juengel et al., 1995) and bovine (Hartung et al., 1995) corpora lutea. Recently, Kiriakidou et al. (1996) reported the expression of StAR in the human ovary. We have confirmed that StAR is expressed by the human corpus luteum during the functional luteal phase after luteal `rescue' in simulated early pregnancy. This suggests that expression of StAR may be a major factor in the control of luteal steroidogenesis.
StAR is a good candidate molecule to explain how progesterone production falls in the late luteal phase and is increased by HCG during luteal `rescue'. Expression of StAR has been shown to be regulated by LH and cAMP in granulosa–luteal cells (Sugawara et al., 1995; Kiriakidou et al., 1996) and it is the expression of StAR that is now thought to be the major rate-limiting step in the steroidogenic pathway (Stocco and Clark, 1996). However, we have found that StAR expression continues across the luteal phase in a similar fashion to P450scc and the LH receptor (Duncan et al., 1996a). Clearly the elements steroidogenic pathway must decline in the late luteal phase as LH receptor and steroidogenic enzyme expression is absent from corpora lutea collected during menstruation (Doody et al., 1990; Bassett et al., 1991; Ravindranath et al., 1992b; Nishimori et al., 1995). In our study, there was more heterogeneity of expression in the late luteal phase but this did not reach statistical significance. It appears that the fall in progesterone production occurs in the continued expression of the main elements of the steroidogenic pathway including StAR. It is unlikely that differential regulation of StAR expression initiates functional luteolysis.
While expression of StAR seems to vary in a similar fashion to P450scc and LH receptors (Duncan et al., 1996a), the changes in expression of 3ß-HSD were more marked. Of the components of the steroidogenic pathway studied, we found the expression of 3ß-HSD mRNA to vary most throughout the luteal phase. Previous studies of steroidogenic enzyme expression in primate corpora lutea have shown similar results. Doody et al. (1990) reported expression of P450scc and 3ß-HSD in a mid-luteal and a late luteal human corpus luteum. Whereas P450scc was easily detected in the late luteal phase, this was not true of 3ß-HSD. In a larger study of the primate corpus luteum, Bassett et al. (1991) found no change in the expression of P450scc mRNA but a marked reduction in 3ß-HSD mRNA during the functional luteal phase. Although 3ß-HSD is not classically thought to be rate-limiting in steroidogenesis (Strauss and Miller, 1991), its expression appears to be more tightly regulated than other steroidogenic components in the primate corpus luteum.
We found the steroidogenic components to be expressed in high levels in the early luteal corpus luteum when progesterone production is increasing. This agrees with previous studies on the primate corpus luteum (Doody et al., 1990; Bassett et al., 1991). It is possible that the increase in progesterone production in the early luteal phase is due to the increasing translation of steroidogenic component mRNA into functional enzymes. However, it is thought that the steroidogenic capacity of the corpus luteum is laid-down during normal late follicular development (Lobb et al., 1998). Indeed, we could immunolocalize 3ß-HSD at this stage, and other groups have reported no differences in immunodetectable steroidogenic enzymes in the early luteal phase (Suzuki et al., 1993; Conley et al., 1995; Sanders and Stouffer, 1997). The likely explanation for the apparent difference in steroidogenic capacity in the early luteal phase and the circulating progesterone concentrations is substrate availability (Carr et al., 1982; Bassett et al., 1991).
It is uncertain whether the changes in 3ß-HSD expression are involved in the initiation of functional luteolysis. We and others (Bassett et al., 1991) have provided evidence that the fall in 3ß-HSD mRNA expression begins in the mid-luteal phase at a time of maximal progesterone output. In vivo, maximal simulation of progesterone production from luteal cells occurs in the early luteal phase and declines throughout the rest of the luteal phase (Fisch et al., 1990). Although there may be changes in the distribution of 3ß-HSD immunostaining in the late luteal stage (Sanders and Stouffer, 1997), we and others have shown that 3ß-HSD protein continues to be present, and presumably functional, throughout the luteal phase (Suzuki et al., 1993; Hild-Petito and Fazleabas, 1997). Inhibition of 3ß-HSD, using trilostane, inhibits progesterone function but the non-steroidogenic functions of the corpus luteum, such as relaxin production continue normally (Duffy et al., 1994, 1995). Other factors must be involved in controlling the lifespan of the corpus luteum. A fall in 3ß-HSD activity does not appear to be able to initiate normal luteolysis by itself.
The mechanism for the apparent differential control of 3ß-HSD and P450scc expression is not known. It remains possible that it is a function of mRNA stability and different half-lives of mRNA species. There is some preliminary evidence for differential control of expression of these enzymes. In rats (Oonk et al., 1989), and humans (Voultilainen et al., 1986) it appears that after the LH surge, expression of P450scc is constitutive and not augmented by gonadotrophin stimulation. In contrast, gonadotrophins have been shown to have a slight stimulatory effect of 3ß-HSD expression in vitro (Chedrese et al., 1990). However, in the primate corpus luteum, Ravindranath et al. (1992b) reported similarities in the control of P450scc and 3ß-HSD expression by showing LH is required for the continued expression of both. Further work is required to study the control of 3ß-HSD expression in the corpus luteum.
All elements of the steroidogenic pathway were maintained in the `rescued' corpus luteum of simulated early pregnancy. In granulosa cells, the LH surge initiates the expression of P450scc and 3ß-HSD (Strauss and Miller, 1991). Recently, it has been shown that granulosa cell StAR expression is also initiated by LH at the time of the LH surge (Kiriakidou et al., 1996). Administration of GnRHant in the mid-luteal phase clearly shows that LH is required for the continued expression of these enzymes in the corpus luteum (Ravindranath et al., 1992b). During luteal `rescue', HCG acts through the LH receptor (Cole et al., 1973) to prevent luteolysis and maintain progesterone production. It appears that LH/HCG has a stimulatory effect on the expression of the key elements of the steroidogenic pathway. One of the effects of HCG in early pregnancy appears to be the facilitation of the expression of the enzymes responsible for progesterone synthesis.
What causes the fall in the expression of the steroidogenic enzymes towards menstruation is not clear. Expression is variable in the late luteal phase and absent from the corpus luteum after menstruation (Bassett et al., 1991; Suzuki et al., 1993; Sanders and Stouffer, 1997). It appears that at the beginning of the late luteal phase when progesterone production is falling that the elements of the steroidogenic pathway are still being expressed. At the end of the luteal phase, expression of these elements, including LH receptors, appears to be reduced (Bassett et al., 1991; Ravindranath et al., 1992b; Suzuki et al., 1993; Nishimori et al., 1995). As the production of progesterone and the maintenance of steroidogenic enzyme expression is dependant on LH (Hutchison and Zeleznik, 1984; Fraser et al., 1986; Ravindranath et al., 1992b), it is possible that, in the late luteal phase, LH action is being diluted at the level of the LH receptor (Zeleznik and Hillier, 1996). As LH receptors are still present (Ravindranath et al., 1992a; Duncan et al., 1996a), this effect may be at the level of coupling to second messenger systems. Studies of the LH receptor have shown that uncoupling does occur in some circumstances (Segaloff and Ascoli, 1993). It is likely that the continued stimulation of the steroidogenic enzymes also stimulates their continued expression.
Several other factors may affect the expression or activity of the steroidogenic enzymes. It is clear that the diverse cytokines and growth factors in the corpus luteum (Behrman et al., 1993; Brännström and Norman, 1993) may be involved in regulating the steroidogenic pathway (Brännström and Norman, 1993; Nappi et al., 1994). It is also possible that local regulation of luteal function may be compartmentalized within the corpus luteum itself (Ottander et al., 1997). Progesterone and other local steroids may be involved in controlling 3ß-HSD activity by retroinhibition via direct non-genomic pathways (Chavatte et al., 1995). Although not fully characterized, the paracrine interactions in the human corpus luteum are likely to intimately interact with the steroidogenic pathway.
In conclusion, the molecular mechanisms of the initiation of functional luteolysis are still unknown. It is likely that a reduction in expression of components of the steroidogenic pathway, particularly 3ß-HSD, are involved in the continued fall in progesterone production pre-menstrually. It is, however, unlikely that changes in their expression initiate the initial reduction in progesterone production. However, in the presence of logarithmically increasing concentrations of HCG in early pregnancy, the steroidogenic pathway is maintained, facilitating the continuing luteal synthesis of progesterone. In the late luteal phase the fall in progesterone production appears to occur in the presence of the major components of the steroidogenic pathway, including StAR. The fall in progesterone production is then associated with alterations in steroidogenic enzyme expression, particularly 3ß-HSD. The mechanisms of the initial drop in progesterone production, and why the expression of 3ß-HSD appears to vary most throughout the luteal phase remains unclear. It appears that further studies on luteal steroidogenesis, focusing on the stimulation of second messenger signals in response to LH receptor ligand binding are required.
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Acknowledgments |
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We thank Dr P.K.Saunders, Mrs W.Struthers, Dr R.M.Shiels and Mr M.R.Millar for technical advice. Mr J.Gaughan helped with the sequencing reactions. Mrs V.Reid-Thomas helped in the recruitment of patients and Professor A.S.McNeilly and Dr H.M.Fraser were involved in helpful discussions. We are once again grateful to Dr G.F.Erickson for providing a copy of his protocol for in-situ hybridization. Dr W.C.Duncan is a clinical training fellow supported by the Wellcome Trust.
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2 Present address: Department of Obstetrics and Gynaecology, University of Sydney, Westmead Hospital, Westmead, Sydney, Australia
3 To whom correspondence should be addressed
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References |
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Submitted on August 8, 1998; accepted on January 22, 1999.
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