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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
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Introduction |
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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.
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Materials and Methods |
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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 [
-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
[-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 -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.
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Results |
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, 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. 1C, 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. 1B).
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, 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. 2A). 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. 2C). No positive
reaction was observed using the T7 sense probe (Fig. 2B). 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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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. 3A). 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. 3B). 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. 3C). 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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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. 5), 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. 6A). The relative abundance
of the GH-R transcript (corrected for GAPDH RNA signal intensity)
increased significantly (P < 0.01) between d25 and d43
(Fig. 6, B and C).
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). GH-R mRNA expression did not vary
between d25-d120 of pregnancy, and it was comparable in caruncular and
intercaruncular endometria (Fig. 7B). 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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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. 9, 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. 9, C and D) and liver (data not
shown) tissues.
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Discussion |
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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.
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Acknowledgments |
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Received July 6, 1999.
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