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Endocrinology Vol. 140, No. 12 5587-5597
Copyright © 1999 by The Endocrine Society

ARTICLES

Expression of Growth Hormone and Its Receptor in the Placental and Feto-Maternal Environment during Early Pregnancy in Sheep

Marie-Christine Lacroix, Eve Devinoy, Sandrine Cassy, Jean-Luc Servely, Michel Vidaud and Guy Kann

Unité de Biologie Cellulaire et Moléculaire, Institut National de la Recherche Agronomique (M.C.L., E.D., S.C., J.L.S., G.K.), 78352 Jouy en Josas, France; Laboratoire de Génétique Moléculaire, Faculté de Pharmacie (M.V.), 75006 Paris, France

Address all correspondence and requests for reprints to: Dr. Marie-Christine Lacroix, Unité de Biologie Cellulaire et Moléculaire, Institut National de la Recherche Agronomique, 78352 Jouy en Josas, France. E-mail: lacroix{at}jouy.inra.fr


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
In a previous study we showed the existence of GH in the ovine placenta. We now supplement the information available on placental GH and describe the presence and distribution of GH receptor (GH-R) messenger RNA (mRNA) in uterine, fetal, and placental tissues during early pregnancy. GH mRNA was not detected in the placenta before day 27 (d27). Its expression peaked between d40 and d45 and fell after d55. GH mRNA was localized in the trophectoderm and syncytium. During the d35-d50 period, concentrations of GH in the maternal circulation were not increased. In umbilical blood, however, GH was detected from d35 and was presumed to be of placental origin, because GH mRNA was not detected in the fetal pituitary gland on d40. We report on GH-R mRNA expression in the placenta between d20-d120. The relative abundance of GH-R transcripts increased significantly between d25-d43. In the endometrium, GH-R mRNA was detected from d8-d120 of pregnancy and from d4-d16 of the cycle. GH-R mRNA was localized in the trophectoderm, fetal mesoderm, and maternal uterine stroma. In the fetal liver, GH-R mRNA was first detectable on d35. The results of this study indicate that between d35-d50 of pregnancy, the endometrium, placenta, and fetus are all potential targets for the placental GH.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
THE PLACENTA PERFORMS many important functions during pregnancy, and its normal development is essential for fetal growth. It is no longer considered as a passive tissue only involved in the transport of nutrients and gases to the fetus, because recent studies have shown that the placenta is an endocrine organ producing many hormones, including members of the GH/PRL family, in several species (1). GH is expressed by human (2) and monkey (3) placentas, and we reported in a previous study that between days 35 (d35) and d50 of pregnancy, the ovine placenta exhibits GH immunoreactivity that corresponds to a 22-kDa protein. This protein is localized to the trophectoderm and syncytium and is most likely produced by these tissues, as GH messenger RNA (mRNA) has been detected in placenta collected between d40-d50 (4).

Available evidence suggests different roles for these placental GH/PL hormones at different stages of pregnancy (5), but their role(s) in fetal and placental growth remains unclear. The authors of recent studies have argued in favor of a role for GH in implantation, blastocyst development, and fetal growth (6, 7, 8). In sheep, no data are available concerning the presence of GH in fetal-placental tissues, either at preimplantation stages or during the early placentation period. The earliest stage at which GH transcripts are detected in ovine placenta and the type of cells expressing them still require clarification.

The placental GH transcripts described in sheep between d40-d50 are all presumed to encode for GH proteins that bind to GH receptors (4). Their earlier expression may influence fetal-placental development before d50, when the fetal pituitary starts to produce GH (9). The GH receptor (GH-R) has been described in several tissues during pregnancy, including blastocyst (7, 10), fetal tissues (10, 11, 12, 13, 14, 15), placenta (10, 12, 16, 17), and uterine epithelium (12, 18, 19). These findings suggest that these tissues are potential targets for GH. In sheep, GH-R mRNA is expressed in fetal liver and muscle and in the placenta, but only from d51 of pregnancy (20, 21), when placental GH production has decreased (4). The aim of this study was therefore to obtain additional information on the expression of GH and GH-R in maternal uterus and fetal-placental tissues from ewes before d50 of pregnancy to gain a better understanding of the role(s) of placental GH during early pregnancy.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animals and tissue collection
All cyclic and pregnant ewes were of the Préalpes du Sud breed. They were fed according to French nutrition recommendations, and all experiments were conducted in accordance with French guidelines concerning the use of experimental animals, including animal welfare and conditions for animal handling before slaughter (22). Blood samples were collected twice daily from one group of four pregnant ewes at 0900 and 1600 h between d6-d60 of pregnancy. In a second group of 33 pregnant ewes, the uteri were collected between d8-d120 (term = 145 days) at the local abattoir just after slaughter. Allantoic and amniotic fluids were collected between d20-d120. When possible, blood was collected from the umbilical artery from d35 and also from the umbilical vein on d41, d42, d43, and d44. Fetal livers were collected on d31, d35, d42, d46, d50, and d83, and fetal heads were collected on d40 and d60. From d20, placentas were collected after removal of the fetus and weighed. On d20, d25, and d30, the placental chorio-allantois was separated from the endometrium. At later stages, the cotyledon (fetal side of each placentome) was manually separated from the caruncle (maternal side of each placentome). After removal of the placenta, the endometria were collected. On d8, d10, d12, d14, and d17, the whole endometrium was collected, but between d25-d120, caruncular and intercaruncular endometria were collected separately. On d40, in seven ewes with a single conceptus, intercaruncular endometrium was collected from both the pregnant uterine horn containing the fetus and the chorio-allantois, and the contralateral uterine horn that had not yet been invaded by the chorio-allantois. Endometrium was collected from cyclic ewes (six animals) on d4, d6, d8, d11, d14, and d17. Pituitary glands were obtained from pregnant ewes, and liver was obtained from a castrated male (21) for use as positive control tissues. For in situ hybridization, the whole placentome was collected to include both the cotyledon and the caruncle. All tissues were frozen in liquid nitrogen and stored at -80 C until processing.

Ovine (o) GH RIAs
Concentrations of oGH in serum from maternal and umbilical blood were determined using an oGH RIA kit (antiserum AFP-C0123080 and oGH I-4; National Hormone and Pituitary Program, Bethesda, MD) supplied by the NIDDK as described previously (4). Briefly, oGH was iodinated to a specific activity of 100 µCi/µg, using a modified chloramine-T method described previously (23). Serum or standards (100 µl) were incubated with antiserum AFP-C0123080 (final dilution, 1:157,000) and [125I]oGH (25,000 cpm) in a final volume of 500 µl for 4 days at 4 C. Bound and free hormones were separated by the addition of 100 µl sheep antirabbit IgG prepared in our laboratory (1:5 dilution; 1-h incubation at 20 C) and 2 ml polyethylene glycol (4.6%; Prolabo, Paris, France) and then centrifugation (30 min, 3,000 x g, 4 C). Assay specificity, defined by the NIDDK, showed cross-reactivity with bovine GH only. Different ovine pituitary hormones and the ovine chorionic somatomammotropin (oCS) did not cross-react at up to 2 µg/tube. The inhibition curves of serial dilutions of serum were parallel to the standard curve (data not shown). Intra- and interassay variation coefficients were 5.8% and 11%, respectively. The sensitivity of the assay was 0.62 ng GH/ml.

Probes
oGH. For GH mRNA detection using Northern blot, a 125-bp PstI/SmaI fragment of the pbGH7 plasmid (24) (GH probe) was used as described previously (4). This probe exhibited 98% homology with oGH complementary DNA (cDNA) and no significant homology with oCS cDNA. For the detection of GH mRNA using in situ hybridization, two riboprobes, T3 antisense and T7 sense, were generated. Two primers were designed according to the method of Logel et al. (25): 5'-CCAAGCTTCATTAACCCTCACTAAAGGGAGACCTCAGGTACGTCTC-3' and 5'-CCAAGCTTCTAATACGACTCACTATAGGGAGACGCATCTCACTGCTC-3'. The specific oGH sequence is double underlined, and it is preceded by T3 or T7 consensus sequence, respectively, and a 9-bp leader sequence, which is underlined. The T3/T7 oGH template was generated using RT-PCR of ovine pituitary total RNA. The resulting PCR product (364 bp) was used as a template for digoxigenin (DIG)-riboprobe synthesis (DIG labeling mix, Boehringer Mannheim, Mannheim, Germany) in accordance with the Promega Corp. protocol (labeling kit, Promega Corp., Madison, WI).

GH-R. For GH-R mRNA analysis using Northern blot, two fragments of ovine GH-R cDNA (26) were used simultaneously: a 1186-bp EcoRI fragment (from positions 1–1186) and a 1057-bp EcoRI/XbaI fragment (from positions 1187–2230), which included 14 nucleotides of the SP72 polylinker sequence. For in situ hybridization, the same plasmid was digested by ClaI or BamHI in the polylinker sequence at the 5'- or 3'-end of the GH-R insert, respectively. Each of these linearized DNAs was used as a template to generate SP6 antisense or T7 sense [{alpha}-32P]UTP-labeled (Amersham Pharmacia Biotech, Les Ulis, France) riboprobes (Promega Corp.).

RNA isolation and Northern blot analysis
Total RNA from chorio-allantois, cotyledons, endometrium, maternal pituitary (GH hybridization control), fetal liver, and liver from a castrated male (GH-R hybridization control) was extracted according to a modified guanidium-thiocyanate-phenol-chloroform procedure and analyzed using Northern blot, as previously described (4). Total RNA was denatured and separated by gel electrophoresis using a 1% (wt/vol) agarose gel containing 2.2 M formaldehyde (27). Fractionated RNA was transferred to a nylon membrane (Nytran NY 13N, Schleicher & Schuell, Inc., Equevilly, France) according to the manufacturer’s instructions. Equal loading and transfer were checked by ethidium bromide staining. Blots were prehybridized for at least 2 h and were hybridized overnight at 65 C in the presence of 0.5 M sodium phosphate buffer, 7% SDS (28), and the appropriate probe labeled by random priming using [{alpha}-32P]deoxy-CTP (3000 Ci/mmol; Amersham Pharmacia Biotech). Blots were washed twice for 10 min each time in 4 x SSC (20 x SSC = 3.0 M NaCl, 0.3 M citrate, pH 7) 0.5% SDS at 65 C for the GH probe or in 1 x SSC-0.1% SDS at 65 C once at room temperature and twice at 65 C, 5 min each time, for GH-R probe. Blots were exposed to x-ray film (MP film, Amersham Pharmacia Biotech). Differences in RNA loading and quality were checked using a radiolabeled 18S ribosomal (29) or glyceraldehyde-3-phosphate dehydrogenase (GAPDH) (30) cDNA probe. RNA levels were determined using Storm and ImageQuant software (Molecular Dynamics, Inc., Sunnyvale, CA). The relative abundance of GH and GH-R mRNA was determined by calculating the ratio between the intensity of GH or GH-R bands and the intensity of the 18S or GAPDH RNA band.

RT-PCR and sequencing of PCR products
Placental and endometrial RNA were reverse transcribed, and RT products were amplified as previously described (31) using GH-R-specific primers (5'-TCACAGGCACTTCATACTCCT-3' and 5'-GGCTCCAGTGATGCTTTTCT-3', corresponding to nucleotides 854–874 and 235–255, respectively, of the GH-R cDNA) (26). Amplification was performed for 25 cycles at 95 C for 1 min, 67 C for 1 min, and 72 C for 1 min.

Amplification products were sequenced using ABI PRISM model 377 (Perkin Elmer Corp., PE Applied Biosystems, Foster City, CA) and a Big Dye terminator kit (Perkin Elmer Corp.) after purification with QIAquick PCR purification kit (QIAGEN, Valencia, CA).

Real-time quantitative RT-PCR
Real-time quantitative PCR analyses for oGH were performed on placenta and pituitary total RNA, using an ABI PRISM 7700 Sequence Detection System instrument and software (Perkin Elmer Corp.). The theoretical bases of the method have been described previously (32, 33). oGH real-time PCR was performed using specific forward (5'-CCCAGGTTGCCTTCTGCTTC-3) and reverse (5'-GCGAAGCAGCTCCAAGCTG-3') primers. In addition, the level of transcripts for the constitutive housekeeping gene product 36B4 coding for acidic ribosomal phosphoprotein (PO) (34) was measured for each sample to control for sample to sample differences in RNA concentration. In each case, 1 µg total RNA was reverse transcribed for 30 min at 42 C with 1.5 mM random hexamers (Pharmacia Biotech, Les Ulis, France), 3 mM MgCl2, 75 mM KCl, 50 mM Tris-HCl buffer (pH 8.3), 500 mM of each deoxy (d)-NTP, 10 mM dithiothreitol, 10 U RNasin ribonuclease inhibitor (Promega Corp.) and 50 U Moloney virus reverse transcriptase (Superscript II, Life Technologies, Inc., Grand Island, NY) in a total volume of 20 µl. Amplification reactions were then set up in a reaction volume of 50 µl using a SYBR Green DNA PCR Core Reagent Kit (PE Biosystems). One microliter of the RT reaction was used for real-time PCR, which consisted of one-step denaturation at 95 C for 10 min, followed by 35 cycles of amplification with denaturation at 95 C for 15 sec and annealing at 65 C for 1 min in the presence of 200 nM specific forward and reverse primers; 5 mM MgCl2; 50 mM KCl; 10 mM Tris buffer (pH 8.3); 200 mM each of dATP, dGTP, and dCTP; 400 mM dUTP; and 1.25 U AmpliTaq Gold. Each sample was analyzed in duplicate. In parallel, 10-fold serial dilutions of total pituitary RNA were run with each analysis as a calibration curve. For each sample, the amounts of oGH mRNA and PO mRNA were determined with relation to the standard curves. Levels of oGH mRNA were then expressed as a ratio to PO mRNA values.

In situ hybridization and detection procedures
Frozen sections were air-dried for 1 h and rehydrated in PBS for 10 min. Subsequently, the sections were fixed for 10 min in 4% paraformaldehyde, washed twice in 4 x SSC, and then treated for 10 min with 0.25% acetic anhydride in 0.1 M triethanolamine. The slides were prehybridized for 1 h at 42 C in prehybridization buffer (4 x SSC, 50% formamide, and 125 µg/ml yeast transfer RNA). Hybridization was carried out in a 4 x SSC-50% formamide humid atmosphere chamber overnight or for 48 h at 42 C in prehybridization buffer (50 µl by slides) supplemented with 10% dextran sulfate, 1 x Denhardt’s solution (50 x = 1% BSA, 1% Ficoll 400, and 1% polyvinylpyrrolidone), and the appropriate probe (2 ng/µl for DIG-labeled probes or 106 cpm for {alpha}-32P-labeled probe) after denaturation for 5 min at 80 C. After hybridization, the slides were washed at 37 C successively in 2 x SSC and 1 x SSC for 15 min each time, then treated for 5 min by RNase A (20 µg/ml) at room temperature and washed twice again in 0.1 x SSC for 30 min each time. For DIG labeling, the sections were rinsed with 100 mM Tris-HCl and 150 mM NaCl, pH 7.5 (TE1), and mRNA were visualized following the procedures described by Boehringer Mannheim. Sections were successively incubated at room temperature 1) for 1 h with Boehringer blocking reagent; 2) for 2 h with sheep anti-DIG-alkaline phosphatase; 3) in TE1, twice for 10 min each time; 4) in 100 mM Tris-HCl, 100 mM NaCl, and 50 mM MgCl2, pH 9.5; and 5) in nitro blue tetrazolium/5-bromo-4-chloro-3-indoyl-phosphate for 10 min, for color reaction revelation. Sections were then counterstained with 0.1% Nuclear Red. For the revelation of radioactive labeling, slides were dehydrated and dipped in NTB2 nuclear emulsion (Eastman Kodak Co., Paris, France), exposed for 3 weeks at room temperature, and then developed in D19 (Kodak) and stained with 1% toluidine blue.

Statistical analysis
The data are expressed as the mean ± SEM. The effects of the stage of gestation on GH concentrations in the maternal circulation were assessed using ANOVA with repeated measures. Variations in concentrations of GH in umbilical cord blood at different stages of pregnancy were analyzed by ANOVA, and pairwise comparisons were tested using Scheffe’s method. The correlation between GH concentrations in umbilical blood and placental weight was analyzed using Kendall’s Tau coefficient method. This test was also used to determine changes in GH mRNA in different tissues at different stages of pregnancy. Statistical analyses were performed using StatView 4.5 (Abacus Concepts, Inc., Berkeley, CA). Probabilities less than 5% were considered significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Expression of GH mRNA in the cotyledon
Total RNA was prepared from chorio-allantois on d20 and d30 and from cotyledons for the other stages of pregnancy, and GH mRNA levels were analyzed using Northern blot. No oGH mRNA hybridization signals were observed on d20 or d30. Such signals were observed between d35 and d50 (Fig. 1Go, A and B). Despite individual variations, the mean GH mRNA signal intensities rose between d35 (5,9 ± 0.52; n = 4) and d40 (18.1 ± 6; n = 4; Fig. 1CGo, left panel; P < 0.05). Peak values were observed between d40-d45 and then tended to fall between d45-d50. GH mRNA levels were very low after d55 (Fig. 1BGo).



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  		Figure 1. Ontogeny of GH mRNA expression in total RNA (18 independent samples) from chorio-allantois (d20 and d30) and cotyledons (d35-d75). Twenty micrograms of each sample were loaded per lane. Adult ovine hypophysis (H; 0.1 and 0.05 µg loaded) was used as a control. A, Hybridization with a 32P-labeled fragment of oGH cDNA (upper panel) and ethidium bromide staining of the gel (lower panel). B, Same blot hybridized with the 32P-labeled 18S cDNA probe. The intensity of the transcript was expressed as a ratio of oGH signal to 18S signal (arbitrary unit). C, Left panel, Comparison of the mean levels of GH transcript expression in cotyledons collected on d35 (n = 4) and d40 (n = 4); right panel, expression of GH transcript analyzed using real-time quantitative RT-PCR. Together with the samples studied above, three more samples, collected on d25, d27, and d63, were analyzed. The level of expression was expressed as a ratio to PO housekeeping gene.

 
To investigate oGH mRNA expression before d35, RNA were analyzed further, using real-time quantitative PCR, a more sensitive method. Together with the samples studied above, three more samples collected on d25, d27, and d63 were analyzed. Expression of the oGH transcript was first detected at a very low level on d27 (4 x 105 arbitrary units). As with Northern analysis, the peak levels of expression were observed on d40 and d45 (8900 x 105 and 7090 x 105 arbitrary units, respectively), and a weak signal was still detected between d60-d75 (95 x 105, 38 x 105, and 81 x 105 arbitrary units, respectively; Fig. 1CGo, right panel).

Localization of GH mRNA in the trophectoderm and syncytium
Day 40 placentome sections were hybridized with T3 DIG oGH antisense probe. No hybridization signals for oGH mRNA were observed in maternal endometrium stroma or in the fetal mesoderm of the placentome (Fig. 2AGo). GH mRNA were detected as spots localized to the trophectoderm and also in an area between the trophectoderm and maternal stroma, which was presumed to be the syncytium formed by the fusion of the binucleated cells of the trophectoderm and uterine epithelial cells (Fig. 2CGo). No positive reaction was observed using the T7 sense probe (Fig. 2BGo). Pituitaries from adult ewes (positive control for DIG probes) produced, as expected, positive hybridization signals with the T3 antisense probe, but only in the anterior pituitary. No hybridization signal was detected using the T7 sense probe (data not shown).



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  		Figure 2. Localization of GH mRNA in d40 placentome sections using in situ hybridization. Sections (magnification, x12) were hybridized with DIG-labeled antisense (A) and sense (B) mRNA oGH probes. The signal was localized to the trophectoderm (arrow). No signal was detected in the fetal mesoderm (M) and maternal stroma (S) of the caruncle or in the deep endometrium (E). At a higher magnification (x50), the section indicated by an arrow in A, shows more clearly that the hybridization signal was present in the trophectoderm (T) and the area (Y) between the trophectoderm and maternal stroma, presumed to be the syncytium (C).

 
Concentrations of GH in maternal and umbilical cord blood and in amniotic and allantois fluids during early pregnancy
The GH produced by the fetal part of the placenta and syncytium of feto-maternal origin may be released into maternal or fetal compartments. Consequently, GH concentrations were assessed in blood collected from the maternal jugular vein, umbilical artery, and umbilical vein, when possible.

GH concentrations in maternal serum were low during early pregnancy and did not rise significantly between d30-d50, when GH is produced by the placenta (Fig. 3AGo). Mean maternal GH concentrations between d6-d27 (before placental GH production) and between d30-d60 (during the placental GH production period) were 2.22 ± 0.09 and 2.42 ± 0.09 ng/ml (two samples each day, four ewes), respectively. Arterial cord blood collected between d35-d120 contained GH from d35 (Fig. 3BGo). GH concentrations in cord blood did not differ between d35-d47 (4.7 ± 0.41 ng/ml; n = 60), but had risen (P < 0.002) by d50 to 28 ± 3.5 ng/ml (n = 12). Concentrations of GH in the umbilical vein on d41 (n = 10), d42 (n = 3), and d50 (n = 5) were comparable with those found in the umbilical artery on the same days (data not shown). At later stages of pregnancy, arterial cord blood concentrations of GH were analyzed each week (Fig. 3CGo). The values rose (P < 0.05) from week 6 to reach their peak values at week 11 (157.4 ± 18.6 ng/ml; n = 18); they then fell (P < 0.05) at week 12 and remained unchanged until near the end of pregnancy (73.59 ± 5 ng/ml; n = 22). In ewes with a single fetal-placental unit, correlations between placental weight and arterial cord blood GH levels were analyzed between d35-d120. During this period, placental weight increased until d90, and this was positively correlated with GH levels in cord blood between d50-d90 (P < 0.01; n = 13). Between d35-d50 and after d90, correlations between GH levels and placental weight were not significant (n = 19). GH was detected in neither amniotic nor allantoic fluid at any stage of pregnancy.



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  		Figure 3. oGH concentrations (mean ± SEM) in the maternal and umbilical circulation assessed using RIA. A, Changes in oGH levels in maternal blood between d6-d60 of pregnancy in four pregnant ewes. Samples were collected twice daily at 0900 and 1600 h. The black line under the graph delimits the period of placental GH production, showing that there was no significant increase in maternal oGH levels during this period. B and C, oGH concentrations in samples from umbilical artery between d35-d50 (B; d35, n = 5; d40, n = 8; d41, n = 18; d42, n = 15; d43, n = 3; d45, n = 7; d47, n = 4; d50, n = 12) and between d40-d120 (C; n = 23). The results of statistical analysis of the data presented in B and C are detailed in Results.

 
Absence of GH mRNA from the fetal pituitary on d40
The presence of GH in the umbilical cord circulation as early as d35 could derive from GH produced by the fetal pituitary; however, in situ hybridization failed to detect the presence of GH mRNA on d40 in two fetal pituitaries. On d60 the anterior pituitary of the fetus had a strong signal using the antisense probe, but no signal was detected by the T7 sense probe in either case (Fig. 4Go).



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  		Figure 4. oGH transcript in the fetal pituitary on d40 and d60. Frozen sections of fetal pituitary collected on d40 and d60 of pregnancy were hybridized with DIG-labeled antisense and sense mRNA oGH probes. oGH mRNA was not detected in the d40 fetal pituitary, but was detected in the fetal pituitary at d60 using the DIG-labeled antisense probe. No signal was detected with the sense probe on either d40 or d60 of fetal development.

 
Ontogeny of GH-R mRNA in placenta, endometrium, and fetal liver
Placental production of GH and the presence of GH in the fetal circulation from d35 led to the assessment of GH-R expression in placenta, endometrium, and fetal liver.

Placenta. GH-R mRNA was detected in chorio-allantois collected on d20, d25, and d30 and in cotyledons between d35-d120. We observed a single, weak band comparable to that detected in liver from a castrated male (4.4 kb; Fig. 5Go), which was shown to correspond to GH-R mRNA after the sequencing of RT-PCR products. The relative level of this transcript tended to increase between d20-d50. To study this period more precisely, five new placental samples were collected on d40 (n = 1), d43 (n = 3), and day 45 (n = 1) and analyzed by comparison with previous samples (Fig. 6AGo). The relative abundance of the GH-R transcript (corrected for GAPDH RNA signal intensity) increased significantly (P < 0.01) between d25 and d43 (Fig. 6Go, B and C).



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  		Figure 5. Presence of oGH-R mRNA transcript in the placenta during pregnancy. Twenty independent samples of total RNA (20 µg) were collected from chorio-allantois on d20 and d30, from placentomes between d35-d120, and from the liver of a castrated male (L; 0.5 and 1 µg) and analyzed using a 32P-labeled oGH-R cDNA probe. The same blot was hybridized with 18S ribosomal cDNA to correct for variations in loading and transfer. The intensity of the hybridization signal was quantified using the ImageQuant system. Transcript levels were expressed as a ratio of the oGH signal to the 18S signal (arbitrary units).

 


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  		Figure 6. Changes in GH-R mRNA expression between d25-d63 of pregnancy in the placentome. A, To study this period in more detail, total RNA prepared from five new placentas collected on d40 (n = 1), d43 (n = 3), and d45 (n = 1) were analyzed in comparison with previous samples (Fig. 5Go; d25, d30, d40, d50, and d63) using a 32P-labeled oGH-R cDNA probe (placenta, 20 µg RNA by lane; L, 0.5 µg RNA castrated male liver as a positive control). The same blot was hybridized with 18S ribosomal cDNA and GAPDH cDNA to correct for variations in loading and transfer. B, The intensity of the hybridization signal was quantified using the ImageQuant system, and oGH-R mRNA levels were expressed as a ratio to the GAPDH signal. C, The relative abundance of oGH-R mRNA increased significantly (P < 0.01) between d25-d43.

 
Endometrium. GH-R mRNA was analyzed using Northern blot in endometrium collected between d4-d16 of the cycle and between d8-d120 of pregnancy. The identity of the transcript was confirmed by the sequencing of RT-PCR products. Levels of expression varied during the estrous cycle and with the stage of pregnancy. GH-R mRNA expression was lower during early pregnancy (before d25), except on d8 and d12. No statistical analyses were performed on the data collected between d4-d16 of the cycle or between d8-d25 of pregnancy, because only one sample at each of those time points was collected and analyzed (Fig. 7AGo). GH-R mRNA expression did not vary between d25-d120 of pregnancy, and it was comparable in caruncular and intercaruncular endometria (Fig. 7BGo). On d40 of pregnancy, RNA prepared from endometrium collected from the nongravid and gravid uterine horns (n = 7) were analyzed, and GH-R mRNA expression was equivalent in both uterine horns (data not shown).



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  		Figure 7. Expression of the oGH-R mRNA transcript in the endometrium during the estrous cycle and pregnancy. Total RNA (20 µg) were prepared from independent samples of endometrium collected A) on d4, d6, d9, d11, d14, and d16 of the cycle and on d8, d10, d12, d14, d17, and d26 of pregnancy (n = 1 for each stage considered); and B) on d30, d75, d83, and d120 (n = 1 for each stage); d25, d35, d45, and d50 (n = 2 for each stage); and d40 (n = 3) of pregnancy. For endometrium collected between d25-d83, total RNA was prepared from either the caruncular (C) or intercaruncular (I) area. Total RNA from the liver of a castrated male (L; 1 and 0.5 µg) was used as a positive control. The two blots were probed with 32P-labeled oGH-R cDNA and 18S ribosomal cDNA to correct for variations due to loading and sample transfer. The intensity of the hybridization signal was quantified using the ImageQuant system. Analysis of the relative abundance of oGH-R mRNA, expressed as a ratio to 18S RNA between d25-d120 of pregnancy, detected no differences in signal intensity between pregnancy stages or between the two types of endometrial tissue (data not shown).

 
Fetal liver. GH-R mRNA expression was first observed on d35 and was present between d35-d83 at comparable intensity (Fig. 8Go).



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  		Figure 8. Ontogeny of oGH-R mRNA expression in fetal liver. Eleven independent samples of total RNA (20 µg) were prepared from fetal liver on d30, d35, d40, d45, and d50 (n = 2 for each day) and d83 (n = 1). Total RNA from the liver of a castrated male (L; 0.5 and 1 µg) was used as a positive control. The blot was probed with 32P-labeled oGH-R cDNA and 18S ribosomal cDNA to correct for variations in loading and sample transfer. The intensity of the hybridization signal was quantified using the ImageQuant system. Results indicate that oGH-R mRNA was first detected on d35.

 
GH-R localization in placentomes
Two monoclonal GH-R antibodies previously described in the literature [MAB 263 (35) and R1-M1-B11 (36), supplied by J. Waters and J. C. Byatt, respectively] were used in an attempt to localize GH-R on d40 placentome sections. A variety of incubation times and temperatures were tested, following the procedure previously described (4), but all attempts to localize GH-R in placentome and castrated male liver sections (positive control) with the two antibodies were unsatisfactory due to a high level of nonspecific background.

GH-R mRNA localization in the placentome
Day 40 placentome sections hybridized with DIG-labeled probe had a very low signal for GH-R mRNA; in situ hybridization was therefore performed using a 35S-labeled GH-R probe. On placentome sections, GH-R transcripts were localized to the trophectoderm, fetal mesoderm, and maternal endometrial stroma of the placentome (Fig. 9Go, A and B). The stronger signal was observed in the trophectoderm. All of these signals were weaker than the very strong signal observed in liver from a castrated male (data not shown). Only low background was observed using the sense probe in placental (Fig. 9Go, C and D) and liver (data not shown) tissues.



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  		Figure 9. Localization of oGH-R mRNA in the placental caruncule. Frozen sections (magnification, x100) of d40 placental caruncle were hybridized with 35S-labeled antisense and sense mRNA oGH-R probes. Brightfield (A) and darkfield (B) views of a section incubated with the antisense probe are shown. The hybridization signal was strongest in the trophectoderm (T) and weaker in the fetal mesoderm (M) and maternal stoma (S) of the caruncule. A hybridization signal equal to the background signal (sense probe; D) was observed in the fetal mesoderm (M). Brightfield (C) and darkfield (D) views are shown of a section incubated with the sense probe. A low level of background signal was observed.

 

   Discussion
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 Abstract
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 Materials and Methods
 Results
 Discussion
References
 
We report here the expression of GH mRNA in the ovine placenta between d27-d75 of pregnancy, and the results of the present study clearly establish that this transcript is not detected in the placenta before d27. oGH mRNA expression lasts for a month, between d27–30 and d63–75. The highest levels of expression were observed between d40-d45, which corresponds to the period of maximum placental GH protein production (4). We have established that placental GH is produced by trophoblastic cells and syncytium, as they contain GH transcripts.

The total GH content per placenta between d40-d50 (calculated using mean placental GH concentrations and weight) (4) ranged from 100-1000 ng. However, there was no increase in maternal serum GH concentrations during this period, strongly suggesting that placental GH is not released in the maternal circulation in significant amounts (<1 ng/ml). Placental GH is probably not implicated in regulation of the maternal metabolism during early pregnancy in sheep, as has been suggested for human placental GH, which is mainly released in the maternal circulation (37, 38). We detected GH as early as d35 in cord blood. We demonstrated that this GH was not of fetal origin, as we could not detect GH mRNA in the fetal pituitary before d40 using in situ hybridization. Some of the GH detected in cord blood before this stage may originate from the transfer of maternal GH across the placenta, but this transfer is presumed to be very low, even nonexistent, in sheep, as we never observed variations in fetal GH concentrations after a 10-fold increase in maternal GH concentrations induced by the treatment with GH-releasing factor of pregnant ewes for 4 days (our unpublished data). An absence of GH transfer from the mother to the fetus had also been reported in humans (39), rabbits (40), and rats (41). Furthermore, mean cord blood GH concentrations between d35-d45 were more than twice those detected in the maternal circulation at the same time points. These data support the hypothesis that the GH detected in umbilical cord blood between d35 and d50, mainly originates from the placenta. However, umbilical GH levels were not correlated with placental weight during the period of placental GH production (d35-d50), probably due to the fact that although placental weight increased at a steady state, placental GH mRNA and protein expressions rose (d35-d45) and then fell (d45 to d50–5d5). The implication of placental GH in the placental growth remains to be established.

In the literature, GH is first reported in the fetal pituitary and circulation between d50-d60 (9, 42, 43). According to these data, we indeed observed a significant rise in GH levels in umbilical cord blood from d50, which reflected the onset of GH production by the fetal pituitary. Between d50-d90, GH concentrations in the umbilical cord and placental weight were correlated. This last observation suggests that fetal GH could be involved in placental growth between d50-d90. Beyond this stage, placental weight stabilizes and is no longer correlated with umbilical GH levels.

The two GH transcripts we had described previously encode two GH proteins, one identical to pituitary GH, and one differing by three amino acids. Both are assumed to bind to GH-R to induce a biological effect. The presence of GH-R in ovine fetal-placental tissues has already been reported, but the earliest stage studied was d50 (21). By d50, placental GH expression has decreased significantly. The results of the present study indicate expression of GH-R mRNA in the placenta from d25-d120 and in the endometrium from d8-d120 of pregnancy. Expression of GH-R mRNA in the placenta and endometrium was not related to that of placental GH or oCS, as it was detected at stages when neither of these hormones was being produced. Nevertheless, placental GH could be involved locally in the up-regulation of GH-R mRNA observed in the cotyledon between d25-d43. A similar regulation was not observed in the endometrium, where GH-R mRNA was expressed weakly during early pregnancy, except on d8 and d12, and then at a consistent and constant levels between d25-d120. Furthermore, there was no significant difference in GH-R mRNA levels in caruncular and intercaruncular endometria. Expression of the GH-R transcript in the endometrium was not regulated locally by a fetal-placental factor, as it was equivalent in the pregnant and nonpregnant uterine horns on d40. Kölle et al. (18) reported comparable data in the bovine. Variations in the levels of GH-R mRNA expression in the endometrium between d8-d17 of pregnancy were not due to RNA degradation or unequal loading. Further samples collected during the first 25 days of pregnancy should be analyzed to determine whether GH-R mRNA expression is enhanced at the time of implantation (d16-d25). However, the expression of GH-R mRNA is not specific to early gestation, and it is also detected in cyclic endometrium. As in the present study, GH-R mRNA is expressed by the endometrium of cyclic rats (19), but not by the endometrium of cyclic cows (18). In sheep, expression in the endometrium was very weak early in the cycle on d4 and d6, fluctuating thereafter. As for the RNA samples collected during early pregnancy, these fluctuations (not due to mRNA degradation or unequal loading) should be analyzed in more detail to ascertain whether there exists a relationship between GH-R mRNA levels and cycle stages. However, our study does establish that in sheep, GH-R mRNA is expressed in the endometrium during both cycles and pregnancy.

In the placenta, GH-R mRNA was localized principally to the trophectoderm and the fetal mesoderm of the cotyledon, with a weaker signal detected in the maternal uterine stroma. These observations differ somewhat from those made by Klempt et al. (21), who did not detect GH-R mRNA in the fetal part of the placenta. The differences in results between these studies could be explained by two facts: the tissue sections were obtained at different stages of pregnancy (d120 vs. d40), and they came from different areas (intercotyledonary characterized by the presence of uterine glands for Klempt et al., and cotyledonary in our study). GH-R transcripts are present in both maternal and fetal components of placentas from humans (14, 15, 44, 45), rabbits (17), cows (12, 18), mice (46), and rats (10, 19). Our results suggest that between d35-d50, the autocrine and paracrine somatotropic effects of placental GH should be considered at both the endometrial and placental levels. After d50, the endometrium and the placenta are both potential target tissues for fetal GH.

The present study confirms the presence of GH-R transcripts in the fetal liver in sheep, as reported by Klempt et al. (21), and that GH-R can first be detected on d35 with the onset of placental GH expression. No expression was detected in fetal liver collected earlier. The role of placental GH in increasing fetal GH-R mRNA expression still requires investigation. In the placenta, endometrium, and fetal liver, only the 4.4-kb full-length receptor transcript was detected. The smaller transcripts (1.7 and 2.5 kb) described in adult liver (21) were not observed. These results agree with those reported by Klempt et al. (21) for ovine placenta and fetal liver. It appears that GH-R transcripts are differentially expressed depending on the type of tissue.

The presence of mRNA coding for GH-R in uterine, fetal, and placental tissues indicates potential roles for placental GH as early as d30 of pregnancy. Placental GH is expressed immediately after the beginning of maternal caruncular stroma invasion by the chorio-allantois (after d20) and during the period when placental growth is at a maximum (mean placenta weight: d30, 10 g; d90, 300 g; our data). In the light of previous reports in sheep (6) and rats (8), we postulate that placental GH influences the placentation process and placental and fetal growth. Its implication could be considered at various levels: regulation of the expression its own receptors (in placenta around d40 and in fetal liver on d30–35), of placental metabolic factors [placental leptin (47) and placental glucose receptors (48)], of placental insulin-like growth factors and the insulin-like growth factor-binding protein system (49), and of placental growth factors or their receptors (such as epidermal growth factor receptors) (50, 51). From d50-d60, fetal GH could act as a relay to placental GH and mediate the same effects.


   Acknowledgments
 
The authors thank D. Durieux, P. Bolifraud, and M. Olivi for technical assistance; Dr. T. E. Adams (University of Melbourne, Melbourne, Australia) for supplying the GH receptor probe; J. P. Furet and C. Giraud-Delville for sequencing PCR products; A. Solari for statistical advice; V. Hawken for correcting the manuscript; and Dr. F. W. Bazer for excellent critical review of the manuscript.

Received July 6, 1999.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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