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Regular Article |
a Departments of Physiologyand
b Obstetrics and Gynaecology, Adelaide University, South Australia 5005, Australia
c Department of Physiology, University of New England, Armidale, New South Wales 2350, Australia
d Academic Division of Child Health, School of Human Development, University Hospital, Nottingham NG7 2UH, United Kingdom
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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50% of the control diet until 144–147 days of gestation. Maternal plasma glucose, but not leptin, concentrations were lower in the undernourished ewes. A significant correlation was found, however, between mean maternal plasma leptin (y) and glucose (x) concentrations (y = 2.9x - 2.4; r = 0.51, P < 0.02) when the control and undernourished groups were combined. Fetal plasma glucose and insulin, but not fetal leptin, concentrations were lower in the undernourished ewes, and no correlation was found between mean fetal leptin concentrations and either mean fetal glucose or insulin concentrations. A positive relationship, however, was found between mean fetal (y) and maternal (x) plasma leptin concentrations (y = 0.18x + 0.45; r = 0.66, P < 0.003). No significant difference was found in the relative abundance of leptin mRNA in fetal perirenal fat between the undernourished (0.60 ± 0.09, n = 10) and control (0.70 ± 0.08, n = 10) groups. Fetal plasma concentrations of leptin (y) and leptin mRNA levels (x) in perirenal adipose tissue were significantly correlated (y = 1.5x ± 0.3; r = 0.69, P < 0.05). In summary, the capacity of leptin to act as a signal of moderate maternal undernutrition may be limited before birth in the sheep.
insulin, leptin, pregnancy
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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All procedures were approved by the Adelaide University Animal Ethics Committee. Surgery was performed on 29 pregnant Border-Leicester Merino cross-bred ewes under aseptic conditions between 109 and 113 days of gestation (term = 147 ± 3 days [mean ± SEM]) with general anesthesia induced by sodium thiopentone (1.25 g i.v.; Pentothal; Rhone Merieux, Pinkenba, Qld, Australia) and maintained with 2.5–4% (v/v) halothane (Fluothane; ICI, Melbourne, Vic, Australia) in oxygen. Vascular catheters were implanted in a maternal jugular vein, a fetal carotid artery and jugular vein, and the amniotic cavity as previously described [11]. Catheters were filled with heparinized saline, and the fetal catheters were exteriorized through an incision made in the ewes' flank. During surgery, ewes and fetuses received a 2-ml i.m. injection of antibiotics (procaine penicillin [250 mg/ml], dihydrostreptomycin [250 mg/ml], and procaine hydrochloride [20 mg/ml]; Penstrep Illium; Troy Laboratories, Smithfield, NSW, Australia). Ewes were housed in individual pens in rooms with a 12L:12D photoperiod and fed once daily at 1100 h with water provided ad libitum. Animals were allowed to recover from surgery for at least 4 days before collection of fetal and maternal blood samples commenced.
Feeding Regime
Pregnant ewes were randomly assigned at 115 days either to a control group weighing 56.7 ± 1.9 kg (n = 13) that received 19.8 ± 0.2 g/kg of lucerne and 3.0 ± 0.1 g/kg of oats per day or to an undernourished group weighing 53.5 ± 2.3 kg (n = 16) that received 10.3 ± 0.1 g/kg of lucerne and 1.6 ± 0.1 g/kg of oats per day. Maternal food allocation was increased in both the control and undernourished groups (lucerne by 15%, oats by 10%) every 10 days until postmortem at 144–147 days of pregnancy [11].
Blood Sampling Protocol
Maternal venous (5 ml) and fetal arterial (3.5 ml) blood samples were collected between 0800 and 1100 h, before the ewes were fed, three times each week between 116 and 140 days of gestation. Blood samples were centrifuged at 1500 x g for 10 min, and plasma was separated into aliquots and stored at -20°C for subsequent glucose and hormone assay. At times during the 25-day protocol, blood samples could not be collected due to technical problems (primarily related to blocked vascular catheters). The number of maternal and fetal blood samples that were available for glucose, insulin, and leptin determination are detailed in the subsequent assay sections. Fetal arterial blood (0.5 ml) samples were also collected for the measurement of arterial blood gas status (ABL 520 blood gas analyzer; Radiometer, Copenhagen, Denmark).
Tissue Collection
Ewes were killed between 144 and 147 days of pregnancy with a lethal overdose of sodium pentobarbitone (Virbac Pty Ltd.; Peakhurst, NSW, Australia). Fetuses were delivered by hysterotomy, weighed, and killed by decapitation (control group, 12 singletons and 2 twins; undernutrition group, 15 singletons and 1 twin). Fetal perirenal adipose tissue was collected and weighed, and a sample was frozen in liquid nitrogen and stored at -80°C.
Glucose Assay
Plasma glucose concentrations were determined in 234 maternal plasma samples (control group, 90 samples, n = 8 sheep; undernutrition group, 144 samples, n = 13 sheep) and 348 fetal plasma samples (control group, 160 samples, n = 13 sheep; undernutrition group, 188 samples, n = 16 sheep) by enzymatic analysis using hexokinase and glucose-6-phosphate dehydrogenase and measuring the formation of NADH spectrophotometrically at 340 nm (COBAS MIRA automated analysis system; Roche Diagnostic, Basel, Switzerland) [11]. The intra- and interassay coefficients of variation were both <5%.
Insulin Radioimmunoassay
Fetal plasma insulin concentrations were measured in 196 samples (control group, 88 samples, n = 12 sheep; undernutrition, 108 samples, n = 13 sheep) using a commercial kit (Phadaseph radioimmunoassay kit; Pharmacia & Upjohn, Uppsala, Sweden). The detection range of the assay was 1.5–240 µU ml-1. Guinea pig anti-insulin antisera and [125I]human insulin (100 µl) were added to plasma samples (100 µl), which were then incubated for 2 h at room temperature before the addition of 2 ml of sheep anti-guinea pig immunoglobulin G. Samples were allowed to stand at room temperature for a further 30 min before being centrifuged at 1500 x g for 10 min as described previously [11]. The inter- and intraassay coefficients of variation were <10%.
Leptin Assay
Plasma leptin concentrations were determined in 119 maternal plasma samples (control group, 50 samples, n = 10 sheep; undernutrition group, 69 samples, n = 15 sheep) and 99 fetal plasma samples (control group, 44 samples, n = 9 sheep; undernutrition group, 55 samples, n = 12 sheep) using a competitive ELISA previously validated for sheep plasma [12]. The ELISA plate was coated with 6 ng of recombinant bovine leptin in 50 µl of 0.1 M bicarbonate buffer (pH 9.0) overnight at 37°C. The plate was blocked with 200 µl of 5% skim milk in ELISA buffer for 1 h at 37°C. Samples (100 µl) were assayed in duplicate and added to wells containing 50 µl chicken antirecombinant bovine leptin antisera in 100% Triton-X 100, 0.5% SDS, and 5% sodium deoxycholate, and the plate was incubated overnight at 37°C. Strepavidin conjugated to alkaline phosphatase (Amrad Biotech; Boronia, Vic, Australia) was added, and after incubation for 1 h, the plate was developed with p-nitrophenylphosphate disodium salt hexahydrate. The sensitivity of the assay was 0.25 ng/ml, and the inter- and intraassay coefficients of variation were 15.7% and 11%, respectively.
Leptin Reverse Transcription-Polymerase Chain Reaction
Perirenal adipose tissue was collected from 20 (control group, n = 10; undernourished group, n = 10) of the 29 fetal sheep, and total RNA was extracted as previously described [8]. Briefly, approximately 100 mg of fetal adipose tissue were homogenized with 1 ml of Sigma Trireagent (Sigma Chemical Co., St. Louis, MO) and allowed to stand at room temperature for 5 min. This was then mixed with 1-bromo-3-chloro-propane (100 µl), left standing at room temperature for 10 min, and then centrifuged at 4°C at 3500 x g for 10 min. An aliquot of the aqueous layer (500 µl) was recovered and mixed with isopropanol (500 µl). The RNA was precipitated by centrifugation at 3500 x g for 5 min at 4°C. The pellet was washed in 70% ethanol and allowed to air dry. The RNA pellet was then dissolved in sterile water (20 µl), and 1 µl of the solution was diluted in sterile water (500 µl) for determination of the spectrophotometric absorbance at 260 and 280 nm. The ratio of nucleic acid to protein was >1.6, and the RNA yield was 0.44 ± 0.02 µg/mg adipose tissue. Integrity of RNA preparations was evaluated by agarose gel electrophoresis, followed by ethidium bromide staining and identification of ribosomal RNA.
Ovine leptin and ß-actin cDNA were amplified by reverse transcription-polymerase chain reaction (RT-PCR) as previously described [8]. Briefly, cDNA was obtained by RT of 2 µg of total RNA with random hexamer oligonucleotides (GeneWorks, Adelaide, SA, Australia) and Super-Script RNase H- (Gibco BRL, Gaithersburg, MD). A fragment of ovine leptin cDNA was amplified through 26 cycles of 60 sec at 94°C, 15 sec at 53°C, and 60 sec at 72°C (Hybaid PCR Express, Teddington, U.K.) from 5 µl of RT product using Taq DNA polymerase (Biotech International, Bently, WA, Australia) according to the manufacturer's instructions with 5'-GACATCTCACACACGCAG-3' and 5'-GAGGTTCTCCAGGTCATT-3' (GeneWorks) as primers. This produced a double-stranded fragment of ovine leptin of 183 base pairs (bp) whose sequence was confirmed. A fragment of ovine ß-actin cDNA was similarly amplified separately by PCR of the same RT product used for amplification of leptin cDNA with 5'-TGGATGGTGGGTATATGGGTC-3' and 5'-TAGATGGGCACAGTGTGGGT-3', and the identity of the 349-bp product was confirmed by sequencing. Both cDNA products from RT-PCR (8 µl) were electrophoresed through a 2.0% (w/v) agarose gel, stained with ethidium bromide, visualized by ultraviolet transillumination, photographed using a digital camera, and quantified using 1D Image Analysis Software Electrophoresis Documentation and Analysis System 120 (Kodak dS Digital Science, Rochester, NY).
Statistical Analysis
Data are presented as the mean ± SEM. The effects of maternal nutrition on fetal body weight, total perirenal fat mass, mean gestational arterial PO2, and the relative abundance of leptin mRNA (ratio of leptin mRNA to ß-actin mRNA) in fetal perirenal adipose tissue were determined using unpaired Student t-test. The effects of maternal nutrition on maternal plasma glucose and leptin concentrations were determined by multifactorial ANOVA with repeated measures using feeding group (control vs. undernutrition) and gestational age (in 5-day blocks) as the specified factors. Similarly, the effects of maternal nutrition and gestation on fetal plasma glucose, insulin, and leptin concentrations were also determined using multifactorial ANOVA with repeated measures. Data were transformed when required to reduce heterogeneity of variance and overcome nonnormal distributions. The Duncan new multiple-range test was used after ANOVA to identify significant differences between mean values. Linear regression analysis was used to assess the relationship between the mean plasma leptin and the mean plasma glucose concentrations measured in each ewe and fetus from 116 to 140 days of pregnancy. Similarly, linear regression analysis was also used to assess relationships between mean plasma leptin and mean plasma glucose or insulin concentrations measured in each fetus from 116 to 140 days of pregnancy. Relationships between mean fetal plasma leptin concentrations and fetal body weight, fetal fat mass, and maternal plasma leptin concentrations were similarly determined. A probability of 5% (P < 0.05) was taken as the level of significance in all analyses.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The mean fetal arterial PO2 throughout late gestation was not different between the control (21.9 ± 0.5 mm Hg) and undernourished (23.4 ± 0.6 mm Hg) groups. No difference was found in fetal body weights (control, 5.02 ± 0.12 kg; undernourished, 4.70 ± 0.16 kg) or relative fat mass (control, 3.89 ± 0.15 g/kg; undernourished, 4.13 ± 0.29 g/kg) between the two groups.
Maternal Plasma Glucose and Leptin Concentrations
Maternal plasma concentrations of glucose were significantly lower (F = 5.13, P < 0.05) in undernourished ewes throughout late pregnancy (Fig. 1A). Plasma glucose concentrations were also lower (F = 4.88, P < 0.002) in both control and undernourished ewes after 120 days when compared with earlier in pregnancy (Fig. 1A). No significant effect of undernutrition, however, was found on maternal plasma concentrations of leptin, and no significant change in maternal leptin concentrations was found between 116 and 140 days of pregnancy in either the control or the undernourished ewes (Fig. 1B). Mean maternal plasma leptin (y) and glucose (x) concentrations were not correlated within each separate feeding group; however, they were significantly correlated when data from the control and undernourished groups were combined (y = 2.9x - 2.4; r = 0.51, P < 0.02, n = 20) (Fig. 2A). No relationship was observed between the mean maternal plasma concentrations of leptin and either maternal body weight at 110–115 days of gestation or fetal body weight at 144–147 days of gestation.
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Fetal Plasma Glucose, Insulin, and Leptin Concentrations
Fetal plasma concentrations of glucose (F = 10.13, P < 0.005) and insulin (F = 6.64, P < 0.02) were significantly lower in the undernourished group (Fig. 3, A and B). Fetal plasma concentrations of insulin were lowest (P < 0.003) between 131 and 135 days compared with other gestational periods. No significant effect of maternal undernutrition on fetal leptin concentrations was found, however, and no significant change in fetal plasma leptin concentrations was observed between 116 and 140 days of gestation in either the undernourished or the control group (Fig. 3C). Also, no difference was observed between plasma leptin concentrations in male and female fetuses.
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No significant correlation was found between mean fetal plasma concentrations of leptin and either glucose or insulin when data from the undernourished and control groups were combined. Mean fetal (y) and maternal (x) plasma leptin concentrations were significantly correlated (y = 0.18x + 0.45; r = 0.66, P < 0.003, n = 17) (Fig. 2B). The mean fetal plasma leptin concentrations between 116 and 140 days of gestation were not correlated with either fetal body weight or with absolute or relative fetal fat mass at 144–147 days of gestation.
Leptin mRNA Expression in Fetal Perirenal Adipose Tissue
No significant difference was found in the relative abundance of leptin mRNA in fetal perirenal fat between the undernourished (0.60 ± 0.09, n = 10) and control (0.70 ± 0.08, n = 10) groups. The mean fetal plasma concentrations of leptin (y) and the relative abundance of leptin mRNA (x) in perirenal adipose tissue were significantly correlated (y = 1.5x + 0.3; r = 0.69, P < 0.05, n = 9) (Fig. 4). Leptin mRNA expression in fetal adipose tissue was not related to either fetal weight (P = 0.09), fetal perirenal fat mass, mean fetal glucose, or insulin concentrations.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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We have also found that maternal plasma leptin concentrations varied between 3 and 10 ng/ml throughout late pregnancy and that no significant change occurred in circulating leptin concentrations between 115 days of pregnancy and term in either control or undernourished adult ewes. These circulating leptin concentrations are similar to those reported by Thomas et al. [13] in moderately fed, adolescent pregnant ewes from 50 days of pregnancy until term. Those authors also found no significant change in plasma leptin concentrations throughout late pregnancy. The lack of a change in plasma leptin concentrations toward the end of pregnancy in the sheep is in contrast to the increase in plasma leptin concentrations that occurs during late pregnancy in the human [14], rat [15], and mouse [16]. Adipose tissue is the main source of circulating leptin in all species, but to what extent other tissues, such as the placenta, are also a source of leptin in the maternal circulation during late pregnancy is unclear. Leptin gene expression is relatively high in the human placenta [17] and is also detectable in the rodent placenta [18]. Species-specific differences in the relative level of placental leptin expression may account for differences in the effect of pregnancy on the maternal plasma concentrations of leptin between sheep and other species. The sheep placenta also expresses the leptin receptor gene [13], and it is therefore possible that maternal leptin may interact with leptin receptors within the placenta to impact on fetal growth and development. Overfeeding the adolescent ewe throughout pregnancy increases maternal growth at the expense of the placenta, leading to growth restriction of the fetus [19]. In a cohort of overfed and normally fed adolescent pregnant ewes, a negative association was found between maternal plasma leptin concentrations and birth weight, placental weight, and number of placentomes [13]. In the present study in the mature ewe, however, we found no significant relationship between maternal plasma leptin concentrations and fetal body weight. Clearly, further work is required to define the relative roles that maternal leptin and nutrients play in placental and fetal growth and development at different stages of reproductive maturity.
Plasma concentrations of leptin in the fetus (<0.3–3 ng/ml) were substantially lower than those in the pregnant ewe, and no effect of either maternal undernutrition or gestational age was found on circulating fetal leptin concentrations between 116 and 140 days of gestation. In a previous study [8], we reported that the abundance of leptin mRNA in fetal adipose tissue increased between 125 and 144 days of gestation. It may be that leptin concentrations increase in the fetal circulation after 140 days of gestation. We also found a positive relationship between fetal and maternal plasma concentrations of leptin during late gestation. One possible explanation is that maternal body composition or fatness either at the beginning or during pregnancy determines the leptin synthetic and secretory capacity of both maternal and fetal adipose tissue or the amount of fetal adipose tissue deposited during late gestation. A positive relationship was found between circulating fetal leptin and the relative abundance of leptin mRNA in fetal adipose tissue; however, no relationship was found with either maternal or fetal leptin concentrations or with the absolute or relative fetal fat mass. Thus, any impact of maternal body composition on circulating fetal leptin concentrations is presumably expressed through the leptin synthetic and secretory capacity of the fetal adipose tissue. An alternative explanation for the close correlation between maternal and fetal plasma leptin concentrations is that the placental leptin receptor may mediate the uptake of leptin from the maternal into the fetal circulation. This would be similar to the postulated mode of action for the short isoform of the leptin receptor in the choroid plexus epithelium to transport leptin from plasma into the cerebrospinal fluid [20].
In the present study, maternal feed availability was reduced by 50% below maintenance for 29–32 days, and this was associated with an 16% fall in maternal glucose concentrations and a 20% fall in fetal plasma glucose and insulin concentrations. No significant effect, however, of this level of maternal undernutrition was found on the fetal plasma concentrations of leptin or on the relative abundance of leptin mRNA in the perirenal adipose tissue. It has recently been reported that continuous infusion of insulin into pregnant ewes for up to 34 days resulted in fetal hypoglycemia and hypoinsulinemia and reduced fetal body weight, but that no change occurred in the expression of leptin mRNA in fetal perirenal fat [21]. Those authors also reported, however, that if the period of continuous insulin infusion was prolonged beyond 36 days (36–76 days), then fetal glucose and insulin concentrations were reduced by 30–50% and leptin mRNA expression was suppressed in fetal perirenal fat [21]. Together, these studies indicate that the synthesis and secretion of leptin in the sheep fetus is resistant to the changes in fetal glucose and insulin concentrations associated with moderate maternal undernutrition. Fetal leptin synthesis is suppressed, however, in the presence of profound fetal hypoglycemia or hypoinsulinemia, which may occur as a consequence of either pharmacological induction of maternal hypoglycemia or severe maternal undernutrition.
In the human, strong positive associations exist between umbilical cord blood leptin concentrations at delivery and infant body weight at birth as well as with other anthropometric markers of fetal growth, including estimates of fetal fat mass [9, 10, 22–25]. We have also previously reported [8] that the abundance of leptin mRNA in fetal adipose tissue was positively correlated with fetal body weight in a cohort of fetuses at an earlier gestational age than those used in the present study. In the present study, however, whereas the relationship between leptin mRNA expression in fetal adipose tissue and fetal weight tended to be positive (P = 0.09), no relationship was found between circulating leptin concentrations and either fetal weight or relative fat mass. These differences between the sheep and the human may be explained, in part, by the different patterns of fat deposition that occur in these species during fetal life. In the sheep fetus, fat is deposited at 0.8 g/kg fetal body weight per day, the proportion of body fat at term is 0.3–2.0%, and the major fat depot is the perirenal adipose tissue, which is comprised predominantly of brown fat cells [26]. Whether leptin is expressed uniformly in all perirenal adipocytes in the sheep fetus before birth is unknown. In contrast, in the human fetus, fat is deposited at a higher rate (3.5 g/kg fetal body weight per day), the proportion of body fat at term is 16%, and subcutaneous fat depots are comprised predominantly of white fat cells [27]. Despite these differences between sheep and human fetuses in the rate of fat deposition, the leptin synthetic capacity of fat stores, and the effect of undernutrition on leptin concentrations during late gestation, it is interesting that perturbations of the intrauterine environment may program the development of postnatal obesity in these and other species. Restricted fetal nutrient supply programs alterations in adiposity or leptin synthesis beyond the postnatal period in the human [28], sheep [29], rat [30], and pig [31]. Further work is required to identify those periods during intrauterine life when changes in the long-term development of the adipocyte and the leptin signaling system are initiated and to clarify the relative importance of maternal body composition and the level of fetal nutrition in the mechanistic pathway that underlies the association between poor intrauterine growth and postnatal obesity.
In summary, we have reported, to our knowledge for the first time, the effect of maternal undernutrition during late pregnancy on maternal and fetal plasma concentrations of leptin and on leptin gene expression in fetal adipose tissue in the sheep. We have found that maternal plasma concentrations of leptin and glucose are positively correlated across the range of circulating glucose concentrations present in well-fed and undernourished pregnant ewes. Interestingly, we have found a positive relationship between the fetal and maternal plasma concentrations of leptin during late gestation, suggesting that maternal body composition during early pregnancy may determine the leptin synthetic and secretory capacity of maternal and fetal adipose tissue. We have found no effect, however, of maternal undernutrition on circulating leptin concentrations or on the abundance of leptin mRNA in adipose tissue in the sheep fetus. The capacity of leptin to act as a signal of moderate maternal undernutrition may, therefore, be limited in this species before birth.
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FOOTNOTES |
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1 We gratefully acknowledge financial support from the National Health and Medical Research Council of Australia. 
2 Correspondence. FAX: 61 8 8 303 3356; caroline.mcmillen{at}adelaide.edu.au
Accepted: April 16, 2002.
Received: January 13, 2002.
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