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Original Articles: Hormones and Reproductive Health |
Service de Gynécologie-Obstétrique (J.L.), Hôpital Cochin–Saint Vincent-de-Paul, 75674 Paris; Université P. & M. Curie, Upres EA2396 (J.-C.C.), 75005 Paris; INSERM U 465 (M.G.-M., 75006 Paris; Centre National de la Recherche Scientifique UPR 1524 (M.C., S.H.-d.M.), 92120 Meudon; and INSERM U 449 (H.V.), 69372 Lyon, France
Address correspondence and requests for reprints to: Jacques Lepercq, M.D., Service de Gynécologie-Obstétrique, Hôpital Cochin–Saint Vincent-de-Paul, 82 avenue Denfert-Rochereau, 75674 Paris Cedex 14, France. E-mail: j.lepercq{at}svp.ap-hop-paris.fr
Abstract
In the adult, circulating leptin is highly correlated to adipose tissue mass. Whether such a relationship exists prenatally is unknown, because the actual source of fetal leptin has not been determined. In the present study, we have assessed the placental contribution to fetal and maternal circulating leptin concentrations and determined whether fetal adipose tissue produces leptin. The rate of leptin production in dually perfused human placenta was 0.036 ng/min·g. Ninety-five percent of the leptin released was delivered into the maternal circulation, vs. only 5% on the fetal side. Leptin messenger RNA and protein were detected in adipose tissue biopsies of 20–38 week human fetuses. However, leptin concentration was twice lower in fetal (0.22 ± 0.11 ng/mg protein, n = 6) than in adult (0.49 ± 0.12 ng/mg protein, n = 8) adipose tissue. Umbilical leptin levels closely reflected ponderal index at birth over a wide range of birth weights (1.6–4.1 kg). In sharp contrast, maternal and placental leptin concentrations were increased in pregnancies associated with fetal growth retardation.
We conclude that umbilical leptin levels are independent of placental leptin production and can be taken as a marker of fat mass in human fetuses. By contrast, placental leptin production makes a substantial contribution to maternal circulating leptin levels during pregnancy.
THE BEST DOCUMENTED function of leptin is the regulation of feeding and energy balance, through multiple hypothalamic pathways ending to the control of food intake and body fat mass (1). Leptin, which was discovered as an antiobesity hormone in ob/ob mice (2), now seems to serve a variety of roles in reproduction (3), puberty (4), early human development (5), and pregnancy (6). Following its original discovery in adipose tissue (2), leptin has been detected in the placenta (6), the amniotic fluid (7), and in fetal plasma as early as week 18 of gestation (8). The actual source of fetal circulating leptin and the role(s) of prenatal leptin have yet to be elucidated.
In the adult, white adipose tissue is the main source of leptin, and circulating concentrations are positively correlated with body mass index and fat mass (9, 10). During pregnancy, leptin increases out of proportion with change in adipose tissue mass and returns to normal after delivery. This suggests that leptin production by the placenta contributes to maternal leptinemia. In the umbilical cord blood, leptin concentrations are closely related to birth weight (8, 11). Moreover, high serum leptin concentrations are systematically found in macrosomic newborns from normal and pathological pregnancies (12, 13). These observations have led to the hypothesis that fetal fat mass determines fetal circulating leptin, as in the adult (13, 14). Alternatively, placental leptin may be delivered in the umbilical circulation and contributes to fetal leptin (6, 15).
The present study was designed to establish the respective contribution of fetal adipose tissue and placenta to fetal circulating leptin. We present direct evidences that human fetal adipose tissue produces leptin and that placental contribution to fetal blood leptin is minimal.
Materials and Methods
Placental perfusions
Human placentas from uncomplicated pregnancy were collected after cesarean sections. Perfusion of a suitable placental lobule was carried out as described previously (16). Briefly, after a washout period, perfusion medium was recirculated in the fetal and maternal circulation during 120 min. The media were continuously gassed with 5% O2 + 5% CO2 (fetal) or 95% O2 + 5% CO2 (maternal). The volumes of fetal (Vf) and maternal (Vm) perfusion media collected at the end of the recirculation period were carefully recorded, and the media were stored at -20 C until use. Leptin concentrations were measured in fetal (Cf) and maternal (Cm) media, allowing to calculate the amounts of leptin released in fetal (Qf = Cf x Vf) and maternal (Qm = Cm x Vm) circulation. These values were divided by the time length of perfusion (t = 120 min) to give the rates of leptin release toward the fetal and maternal side of the placental circulation. Leptin was also assessed in the placenta before (Qti) and by the end (Qte) of the perfusion, to evaluate change in placental leptin (Q = Qte - Qti). The rate of placental leptin synthesis (PLS) was estimated as PLS = Qf + Qm + Q divided by the time length of perfusion for one cotyledon, and as TPS = PLS x P/L, where P/L is the placenta to lobule weight ratio, for the whole placenta. The mean lobule and placental weights were 20.8 ± 4.3 g and 464.6 ± 72.5 g, respectively.
Study population
The study population included 74 nonobese, nondiabetic pregnant women (mean body mass index, 22.4 ± 1.3 kg/m2) and their infants classified into three groups: large for gestational age (LGA; n = 12), average for gestational age (AGA; n = 38), and small for gestational age (SGA; n = 24). Placental and neonatal anthropometric measurements at birth included weight on a calibrated scale and length on a measuring board. Gestational age at delivery was determined from the date of the last menstrual period and was confirmed by first-trimester ultrasonography. Birth weight according to gestational age was used to define SGA, AGA, and LGA infants based on French growth standard curves (17); the ponderal index was expressed as weight (g) x 100/length3 (cm3) (13); and birth weights (g) and mean gestational age (weeks) were 1620 ± 11 and 35.2 ± 1 in SGA, 3295 ± 21 and 39.8 ± 0.3 in AGA, and 4192 ± 77 and 39.60 ± 0.5 in LGA infants, respectively.
Blood and tissue sampling
Venous blood samples were collected from the mother and umbilical cord at delivery. The blood was immediately centrifuged, and plasma was stored at -20 C until analysis. Biopsies of placenta were obtained at delivery. Fetal adipose tissue was sampled during autopsies performed within 1–36 h in six cases of in utero fetal death at 20, 33, 34, 37, 37, and 38 weeks of gestation. The cause of death was placental insufficiency in two cases, diabetes in one case, and umbilical cord abnormality in three cases. Fetal body weights were, respectively, 430, 1020, 1600, 4770, 3220, and 2650 g, and fetuses were not macerated. Maternal white adipose tissue was sampled from abdominal sc depot at the time of elective cesarean section in uncomplicated pregnancies with AGA infants at weeks 38–39 of gestation. Liver biopsy was obtained from one nonpregnant woman at the time of surgical resection for severe cirrhosis. All tissue biopsies were snap-frozen in liquid nitrogen and stored at -80 C.
Biochemical assays
Serum leptin levels were measured by RIA (Linco Research, Inc., St.Charles, MO), with a detection limit of 0.1 ng/mL. Inter- and intra-assay variations were 4.6% and 3.6%, respectively. Leptin concentrations in fat and placental biopsies, including those from perfused placentas, were determined on clarified supernatant (10 min, 2500 g) of whole tissue homogenates [10% v/w in 250 mM sucrose, 25 mM HEPES buffer (pH 7.4) containing 1 mM phenylmethylsulfonyl fluoride, 2 mM aprotinin, and 2 mM dithiothreitol]. Measurements were performed using the same RIA and linearity was established using increasing volumes (50–300 µL) of supernatant as previously described (15).
RNA isolation and RT-PCR
Total RNA was extracted from control liver, placenta, and maternal and fetal adipose tissue as described by Chirgwin et al. (18). The RNA concentration was determined by measuring absorbance at 260 nm. RNA (100 ng) was reverse transcribed and amplified by PCR in the presence of specific primers for leptin, Ob receptor (Ob-R), and the transcription factor Sp1, which is ubiquitously expressed and was used as an internal standard. The primer pair derived from the human leptin complementary DNA sequence (2) was: upstream primer 3'-CATTGGGGAACCCTGTGCGGATTC-5'; downstream primer 5'-TGGCAGCTCTTAGAGAAGGCCAGC-3'. The primer pair derived from the human Ob-R (19) was designed to detect the long form (3565 bp) of Ob-R: downstream primer 5'-TACAAAGTGTGAGCAACTGT-3' and upstream primer 5'-GGGTTCTGTTTGTATTAGTG-3'. Amplifications consisted of one cycle at 94 C for 2 min, followed by 30 cycles at 94 C for 30 sec, 60 C for 1 min, and 68 C for 2 min, with a final extension step at 68 C for 7 min. To control the quality of the RNA and the intensity of the depot, we measured Sp1 expression, using the same amplification conditions and the following oligonucleotide pair: 5'-GAGAGTGGCTCACAGCCTGTC-3' upstream and 5'-GTTCAGAGCATCAGACCCCTG-3' downstream (20). PCR products were analyzed on a 1.2% agarose gel and exposed to an ultraviolet transilluminator for photography.
Quantitative RT-PCR analysis
The absolute messenger RNA (mRNA) concentrations of leptin mRNA were determined in fat and placenta biopsies using a previously described RT-competitive PCR assay (RT-cPCR; Ref. 21). As a control, insulin receptor mRNA levels were also quantified using a similar methodology (22). The RT-cPCR method relies on the use of a specific competitor DNA molecule during the amplification process, after a specific RT reaction. The conditions of the RT-cPCR reactions and the sequences of the primers have been described in detail previously (21, 22). For each mRNA, the specific first-strand complementary DNA was synthesized from 0.2 µg total RNA added in the RT reaction. During the PCR, Cy-5 5'-end labeled sense primers were used, and the PCR products were analyzed with an automated laser fluorescence DNA sequencer (ALFexpress; Pharmacia, Upsala, Sweden) in 4% denaturing polyacrylamide gels (23). The initial concentration of target mRNA was determined at the competition equivalence point, as described previously (21, 22). The interassay variation of the RT-cPCR assay was less than 10% (21).
Statistical analysis
Results are expressed as means ± SEM. Statistical analysis was performed by variance analysis and Student’s t test for unpaired data. Statistical significance was defined as P less than 0.05. Analyses were performed using SAS statistical software (SAS Institute, Inc., Cary, NC).
Ethics
All protocols were approved by the institutional ethics review board of Cochin Hospital, Rene Descartes University Paris V (Paris, France), and all the mothers gave their informed consent.
Results
Placental leptin production
The method of dual perfusion of human placental cotyledon was used
to directly assess the rate of leptin production toward the fetal and
the maternal side of placental circulation. We determined that the rate
of placental leptin production was 0.036 ± 0.015 ng/min·g and
14 ± 4.5 ng/min per placenta (Table 1
). Most of the leptin released in media
(95 ± 5%) was delivered into the maternal circulation, whereas
only 5 ± 2% was released into the fetal circulation. These data
demonstrate unequivocally that placental leptin is secreted
asymmetrically in the maternal and fetal circulation.
|
To further examine the origin of fetal circulating leptin, we
sought to determine whether fetal adipose tissue was capable of
producing leptin. As a first attempt to answer this question, we
assessed the presence of leptin mRNA in fetal adipose tissue biopsies
obtained postmortem from fetuses of various gestational ages. As
expected, leptin mRNA was detected by RT-PCR in adult adipose tissue
and in the placenta, but not in the liver obtained from a nonpregnant
woman (Fig. 1). With the same set of
primers, leptin mRNA was readily detected in fetal adipose tissue. In
addition, Ob-R mRNA encoding the long isoform of the receptor was
detected in fetal and maternal fat, as well as in placenta and
liver.
|
). In contrast to leptin mRNA,
similar concentrations of insulin receptor mRNA were detected in fetal
and maternal tissues, indicating that the low mRNA levels detected in
fetal tissue are likely to be specific for leptin. Placental leptin
mRNA concentrations were in the range of concentrations in fetal
adipose tissue, whereas insulin receptor mRNAs were much higher.
|
). In pregnant women, leptin
concentration per milligram of total protein was in the same range in
placenta and in adipose tissue. Umbilical leptin reflects the infant ponderal index
To evaluate whether umbilical leptin was related to fetal adipose
tissue mass, each infant included in the study was assigned to SGA,
AGA, or LGA group, based on birth weight according to gestational age.
The ponderal index of the infants was chosen as a marker of adiposity.
Umbilical cord leptin concentrations were markedly different between
the three groups and increased in relation to the ponderal index (Fig. 2A). In sharp contrast, maternal plasma
leptin concentrations were twice higher in pregnancies associated with
intrauterine growth retardation (SGA), compared with pregnancies
associated with AGA or LGA infants (Fig. 2B). Moreover, placental
leptin concentrations were also significantly increased in pregnancies
associated with intrauterine growth retardation. Thus, in this
population, fetal leptin was related to ponderal index, but not to
maternal and placental leptin levels.
|
In the present study, we directly assessed placental leptin
release in the fetal and maternal circulation. The finding that 95% of
the leptin released by placenta is delivered into the maternal
circulation supports an earlier report (24). These data
are in line with the observation that leptin levels are increased in
pregnant women and fall rapidly after delivery (6, 25).
Leptin content is of the same order of magnitude in term placenta and
maternal adipose tissue expressed per milligram of protein. This is
also relevant when comparing the rate of leptin production by the
placenta measured in the present study (0.036 ± 0.015 ng/min·g)
and by white adipose tissue (0.032 ± 0.005 ng/min·g), as
estimated in humans by arteriovenous balance (26). In
keeping with observations made in normal and pathological pregnancies
(6, 15), this clearly establishes that the placenta is a
major site for leptin synthesis in the human. High mRNA and leptin
levels in placentas from preeclamptic pregnancies may contribute to
further increase maternal circulating leptin. The stimuli responsible
for leptin synthesis are not known so far, but hypoxia has emerged as a
potential stimulating factor of placental leptin production (Ref.
27 and Guerre-Millo, M., A. Grosfeld, J. Lepercq, J.
Andre, M. Cauzac, and S. Hauguel-de Mouzon, unpublished data). By
contrast, very low circulating leptin levels are found in SGA infants
(Fig. 2 and Ref. 28). This argues against a major
contribution of placental production to umbilical leptin levels.
Detection of leptin in fat biopsies obtained between weeks 20 and 38 of gestation provides evidence that adipose tissue of human fetuses produces leptin, at a developmental stage when it can be delivered into the fetal circulation. The lower expression of the leptin gene and protein in adipose tissue of fetuses compared with that of pregnant women may reflect the wide distribution in the age of the fetuses at the time of fat biopsy. However, in our study population, low and high leptin mRNA concentrations distributed independently of gestational age (data not shown), suggesting that the low rate of leptin gene expression is constitutive in the fetal adipose cells. This could reflect the response of adipose tissue to the specific fetal systemic metabolic and hormonal environment.
At birth, body fat mass represents 3% of total body weight in SGA infants (29), 15% in AGA infants (30), and increases up to 30% in LGA infants born to type 1 diabetic mothers (31). The ponderal index allows a satisfactory evaluation of the amount of fat accumulated in the fetus (32, 33) and correlates with skinfold measurement at birth over a wide range of birth weights (34). Moreover, it has been suggested that the fat cell size varies as a function of adipose tissue mass, being smaller in SGA and larger in LGA infants (35). The high leptin concentration in umbilical blood of LGA infants indicates that the bigger the fat mass is, the more leptin is released into the fetal circulation. This relationship strongly suggests that fat cell size is a main determinant of fetal circulating leptin levels, as in the adult (36).
The role(s) of fetal and placental leptin is still speculative. Fetal adipose tissue itself may be a target for the action of its own product as indicated by the presence of mRNA encoding the long alternatively spliced isoform of the Ob-R. An autocrine action of leptin has also been suggested in adult adipose tissue (37, 38), but the functionality of Ob-R in adipose is not clearly demonstrated. Alternatively, the presence of mature leptin and its receptor in several tissues of the fetus (39) raises the possibility that fetal leptin may be involved in regulating specific endocrine pathways different from those developing after birth. Leptin has been visualized by immunohistochemistry in differentiating preadipose cells of week 6 human embryos (40) and may be active from very early developmental stages. The presence of both leptin and Ob-Rs suggests that leptin can act by autocrine or paracrine pathways in the human placenta as suggested for white adipose tissue.
In conclusion, this study provides the first direct evidence that white adipose tissue of human fetuses produces leptin. Fetal plasma leptin levels can be taken as a valuable index of fat mass in human because it appears independent of placental leptin production. By contrast, placental leptin, which is largely delivered into maternal circulation, may contribute to maternal circulating leptin levels during pregnancy.
Acknowledgments
We thank Jocelyne André, Chantal Truong Cong-Y, and Najiba Lahlou for excellent technical assistance. We thank Prof. C. Smadja (Hôpital Antoine Béclère, Clamart, France) for providing liver biopsies.
Received November 29, 2000.
Revised January 30, 2001.
Accepted February 28, 2001.
References
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