Abstract
According to the Developmental Origin of Health and Disease (DOHaD) concept, alterations of nutrient supply in the fetus or neonate result in long-term programming of individual body weight (BW) setpoint. In particular, maternal obesity, excessive nutrition, and accelerated growth in neonates have been shown to sensitize offspring to obesity. The white adipose tissue may represent a prime target of metabolic programming induced by maternal obesity. In order to unravel the underlying mechanisms, we have developed a rat model of maternal obesity using a high-fat (HF) diet (containing 60% lipids) before and during gestation and lactation. At birth, newborns from obese dams (called HF) were normotrophs. However, HF neonates exhibited a rapid weight gain during lactation, a key period of adipose tissue development in rodents. In males, increased BW at weaning (+30%) persists until 3months of age. Nine-month-old HF male offspring was normoglycemic but showed mild glucose intolerance, hyperinsulinemia, and hypercorticosteronemia. Despite no difference in BW and energy intake, HF adult male offspring was predisposed to fat accumulation showing increased visceral (gonadal and perirenal) depots weights and hyperleptinemia. However, only perirenal adipose tissue depot exhibited marked adipocyte hypertrophy and hyperplasia with elevated lipogenic (i.e. sterol-regulated element binding protein 1 (Srebp1), fatty acid synthase (Fas), and leptin) and diminished adipogenic (i.e. peroxisome proliferator-activated receptor gamma (Pparγ), 11β-hydroxysteroid dehydrogenase type 1 (11β-Hds1)) mRNA levels. By contrast, very few metabolic variations were observed in HF female offspring. Thus, maternal obesity and accelerated growth during lactation program offspring for higher adiposity via transcriptional alterations of visceral adipose tissue in a depot- and sex-specific manner.
Introduction
The rising prevalence of obesity in the world is considered a global epidemic (Popkin et al. 2012). Obesity is characterized by accumulation and functional alterations of white adipose tissue (WAT) predisposing the individuals to increased risk of metabolic pathologies (Sun et al. 2011). The expansion of WAT results from hyperplasia (increase in adipocyte number) and/or hypertrophy (increase in adipocyte size) along with modifications of tissue sensitivity to circulating hormones (Björntorp & Sjöström 1971). Obesity is the result of a complex interaction between genetic and environmental factors (Bouchard 2009). According to the Developmental Origin of Health and Disease (DOHaD) concept also called ‘developmental programming’ or ‘conditioning’ (Barker 2004, Hanson & Gluckman 2014), alterations of nutrient supply in the fetus or neonate result in long-term programming of individual body weight (BW) setpoint. Epidemiological studies initially showed that maternal undernutrition leading to fetal growth restriction was associated with higher adiposity in adulthood (Ravelli et al. 1999). Clinical studies have also shown that maternal obesity, excessive nutrition, and accelerated growth in neonates sensitize offspring to obesity (Leddy et al. 2008).
Thus, WAT may represent a prime target of metabolic programming induced by maternal obesity. Perturbations to the perinatal nutrient supply may affect adipocyte development, leading to persistent alterations in their number and functional properties (Lukaszewski et al. 2013, Lecoutre & Breton 2014, 2015). Indeed, in fetuses and neonates, adipocyte stem cells are still plastic and potentially sensitive to maternal factors (Tang & Lane 2012). In humans, the number of adipocyte is set early in life and is a major determinant of fat mass in adulthood (Spalding et al. 2008). The timing of adipose tissue development, which differs between species, determines the window of vulnerability to potential adverse environment. In rodents, adipose tissue growth and adipogenesis mainly take place during the last week of gestation and accelerate throughout lactation, whereas in larger mammals, these processes occur before birth. However, there is now convincing evidence that adipogenesis occurs throughout the lifetime (Muhlhausler & Smith 2009).
Little is known about the programming mechanisms that may account for long-lasting perturbation of adipogenesis and WAT metabolism in offspring from obese dams. To unravel the underlying mechanisms, several animal models of maternal obesity have been developed using high-fat (HF) or cafeteria diet applied during the preconception, gestation, and/or lactation periods in dams (Williams et al. 2014). These studies confirmed that maternal obesity has common long-term metabolic consequences sensitizing the offspring to metabolic syndrome features. In particular, maternal obesity at conception programs enhanced adipogenesis and lipogenesis from the fetal period to adulthood resulting in higher WAT mass and larger adipocytes (Muhlhausler & Smith 2009, Borengasser et al. 2013, Murabayashi et al. 2013). Overfeeding during lactation and/or postweaning periods leads to accelerated growth and consistently worsens adipogenesis and lipogenesis programming (Desai & Ross 2011, Guberman et al. 2013, Masuyama & Hiramatsu 2014). Programmed upregulation of the key adipogenic factor peroxisome proliferator-activated receptor gamma (PPARγ) is one of the characteristic features of fat expansion in the offspring of obese dams (Samuelsson et al. 2008, Muhlhausler & Smith 2009, Sen & Simmons 2010, Dahlhoff et al. 2014, Desai et al. 2015). Obesity-prone offspring rats from obese mothers also exhibited modified fatty acid composition within WAT (Benkalfat et al. 2011). However, in rodents, few studies have examined depot- and sex-specific consequences of maternal obesity in offspring’s WAT and there is little agreement among them (Sun et al. 2012, Ornellas et al. 2013, Dahlhoff et al. 2014, Masuyama & Hiramatsu 2014).
In this study, we examined whether maternal obesity differently programs adipocyte number and morphology using a model of maternal obesity in rats fed a HF diet before and during gestation and lactation. We also profiled gene expression in two visceral fat depots (gonadal and perirenal WAT) in adult offspring of both sexes. Here, we demonstrate the maternal obesity and accelerated growth during lactation program offspring for higher adiposity via transcriptional alterations of visceral adipose tissue in a depot- and sex-specific manner.
Materials and methods
Animals
Four-week-old virgin female Wistar rats (1 month) were purchased from Charles River Laboratories (L’Arbresle, France) and were housed in individual cages in a humidity-controlled room with a 12h light:12h darkness cycle. Food and water were available ad libitum. After 2weeks of acclimatization on a control (C) diet (3.85kcal/g with 10% of total calories as fat consisting of soybean oil (5.6%) and lard (4.4%), 70% as carbohydrate, and 20% as protein; D12450J, Research Diets, New Brunswick, NJ, USA), female rats were fed either a HF diet (5.24kcal/g with 60% of total calories as fat consisting of soybean oil (5.6%) and lard (54.4%), 20% as carbohydrate, and 20% as protein; D12492, Research Diets, New Brunswick, NJ, USA) or a C diet for 16weeks (n=12 per group). After 14 weeks of HF diet, 20-week-old female (5months) rats were subjected to an oral glucose tolerance test (OGTT). Plasma levels of insulin, leptin, and corticosterone were also measured after 16h overnight fasting in both groups. After mating with a male rat fed a C diet, 22-week-old pregnant females were transferred into individual cages with free access to water and continued on their respective diets (C or HF diet) throughout gestation and lactation. Maternal BW was measured weekly until delivery. At parturition, pups were weighed and sexed. Litter size was adjusted to eight pups per dam (four males and four females). During lactation, BWs of dams and pups were assessed on postnatal days (PND) 1, 4, 7, 11, 14, 17, and 21. At weaning (PND21), dams were killed and glycemia as well as plasma levels of leptin insulin and corticosterone were determined after 16h overnight fasting. To obviate any litter effects, animals used for each experiment were randomly chosen in different litters and only a limited number of animals (one to two males and females) were used from each litter. After weaning, male (M) and female (F) offspring from C or HF dams were housed individually with free access to water and C diet, divided into four groups (CM, CF, HFM, and HFF (n=16 per group) and weighed weekly until 9months of age. Animal use authorization by the French Ministry of Agriculture (No. 04860) has been granted to our laboratory for experimentation with rats. Experiments were conducted in accordance with the principles of laboratory animal care (European Communities Council Directive of 1986, 86/609/EEC).
Food intake and metabolic parameters
Food consumption was recorded weekly from weaning to adulthood until killing the animal in the four groups. Food intake of rats was measured once a day at the beginning of the light phase (09:00h) by subtracting the uneaten food from the initial amount. Weight-related energy intake is defined as the energy content of the food ingested (kcal) expressed relative to BW (g). Twenty-four-week-old (6months) offspring was placed in metabolic cage (Bioseb, Vitrolles, France). After an acclimatization period, food intake was recorded for each 24h period during 1week to investigate light/dark phase food intake rhythm.
Oral glucose tolerance test
For OGTT, rats were fasted overnight. Basal blood glucose level defined as T0 was determined using a glucometer (Glucotrend 2, Roche Diagnostics) before oral glucose administration (2g/kg BW). Tail vein blood glucose was then measured at 0, 30, 60, 90, and 120min after administration.
Endocrine parameters
Plasma hormone levels were evaluated in 30-week-old (7months) and 36-week-old offspring (9months) while killing. Blood glucose was determined as described above. Plasma leptin and adiponectin concentrations were measured using murine ELISA kits (Diagnostic Systems Laboratories, Inc. Webster TX, USA and Adipogen Inc, Korea, respectively). Plasma corticosterone levels were determined by a competitive enzyme immunoassay (Immunodiagnostic Systems Ltd, Boldon, UK). Plasma insulin concentrations were measured by ELISA (DRG International, Inc, Springfield Township, NJ, USA). Plasma apelin content was determined by ELISA (Phoenix Pharmaceuticals, Burlingame, CA, USA). The assay sensitivity was 0.07ng/mL (insulin), 0.04ng/mL (leptin), 0.1ng/mL (adiponectin), 0.55ng/mL (corticosterone), and the intra- and inter-assay coefficients of variation were 4 and 9.1% (insulin), 5.4 and 7.3% (leptin), 4.4 and 6.1% (adiponectin), 4.9 and 7.8% (corticosterone), respectively. Assay kits were used to determine the contents of plasma triglycerides and total cholesterol (61238 Triglyceride Enzymatique PAP100, 61218 Cholesterol Liquide, BioMérieux, France) as well as free cholesterol and free fatty acid (FFA) (references 279-47106 and 999-75406, Wako Chemicals, Neuss, Germany). Each sample was measured in duplicate.
Plasma and tissue collections
Thirty-six-week-old rats (9months) were rapidly weighed and killed by decapitation between 09:00 and 10:00h after 16h overnight fasting. Trunk blood samples were collected into prechilled tubes containing EDTA (20µL of a 5% solution) and centrifuged at 4000g for 10min at 4°C. Plasma was stored at −20°C. Several tissues (brown adipose tissue, liver, heart, kidney, and adrenal gland) as well as gonadal (GWAT) and perirenal (PWAT) fat pads were weighed, frozen in liquid nitrogen and stored at −80°C. For histology experiments, animals were fixed by intracardiac perfusion using buffered 4% paraformaldehyde solution.
Gene expression analysis
GWAT and PWAT gene expression levels were determined in the four groups using RT-quantitative PCR as previously validated (Lukaszewski et al. 2011). Briefly, total RNA was extracted and purified using RNeasy lipid tissue kit (Qiagen) according to the manufacturer’s recommendations. The yield of total RNA was quantified on a Multiskan GO Microplate Spectrophotometer (Thermo Scientific). The quality of total RNA was assessed by determining the 260/280 and the 260/230 absorbance ratio and by agarose gel electrophoresis. First-strand cDNAs were synthesized using ThermoScript RT Kit (Invitrogen and Life Technologies). Relative expression levels of RNA per sample were quantified by SYBR Green assay on a Roche Light Cycler 480 sequence detection assay (Roche). Primer sequences are presented in Table 1. For each transcript, PCR was performed in duplicate with 10µL final reaction volumes with 1µL of cDNA, 8μL of QuantiTect SYBR Green Master mix (Qiagen), and 0.5 µL of each primer set (Table 1). PCR was conducted using the following cycle parameters: 10min at 95°C, and 40 three-step cycles of 15 s at 95°C, 20 s at 60°C, and 30 s at 72°C. A pool of cDNA from control tissues was used as a standard for quantitative correction. All cDNA samples were applied in dilution of 1:10 to obtain results within the range of the standard. Analysis of transcript level was carried out using the determination of the threshold cycle Ct for each reaction corrected by the efficiency. The level of gene expression was normalized to the reference gene transcript cyclophilin A RNA.
Primers used for RT-quantitative PCR analysis.
| Target | Sequence | NCBI detected transcript no. | Length of amplicon (bp) |
|---|---|---|---|
| Leptin | |||
| Forward | 5’-GTTCCTGTGGCTTTGGTCCT-3’ | NM_013076 | 99 |
| Reverse | 5’-CTGGTGACMTGGTCTTGATGA-3’ | ||
| Ob-Rb | |||
| Forward | 5’-GCATGCAGAATCAGTGATATTTGG-3’ | NM_012596 | 81 |
| Reverse | 5’-CAAGCTGTATCGACACTGATTTCTTC-3’ | ||
| Adiponectin | |||
| Forward | 5’-CAGGAGATGCTGGAATGACA-3’ | NM_144744 | 114 |
| Reverse | 5’-CTACGCTGAATGCTGAGTGAT-3’ | ||
| Fas | |||
| Forward | 5’-GGACATGGTCACAGACGATGAC-3’ | NM_017332 | 94 |
| Reverse | 5’-GTCGAACTTGGACAGATCCTTCA-3’ | ||
| C/Ebpα | |||
| Forward | 5’-AGTTGACCAGTGACAATGACCG-3’ | NM_012524 | 94 |
| Reverse | 5’-TCAGGCAGCTGGCGGAAGAT-3’ | ||
| Pparγ | |||
| Forward | 5’-TTGGCCATATTTATAGCTGTCATTATT-3’ | NM_001138838 | 76 |
| Reverse | 5’-TGTCCTCGATGGGCTTCA-3’ | ||
| Srebp1 | |||
| Forward | 5’-CATCGACTACATCCGCTTCTTACA-3’ | NC_005109 | 97 |
| Reverse | 5’-GTCTTTCAGTGATTTGCTTTTGTGA-3’ | ||
| Gr | |||
| Forward | 5’-GTGAAATGGGCAAAGGCGATAC-3’ | NM_012576 | 98 |
| Reverse | 5’-GCAAATGCCATGAGAAACATCC-3’ | ||
| Mr | |||
| Forward | 5’-CTGTGCCCGGCTTTGATGGTAG-3’ | NM_013131 | 100 |
| Reverse | 5’-GGGATGCTGGCTTCAGGGTAAT-3’ | ||
| 11β-Hsd1 | |||
| Forward | 5’-GGACTGGACATGCTCATTCTCA-3’ | NM_017080 | 84 |
| Reverse | 5’-GCTTCTTCGCACAGAGTGGATA-3’ | ||
| 11β-Hsd2 | |||
| Forward | 5’-ATGTGAACCTCTGGGAGAAGCG-3’ | NM_012524 | 100 |
| Reverse | 5’-CCATGCAAGTGCTCAATGTAGTC-3’ | ||
| Housekeeping gene Ppia (Cyclophilin A) | |||
| Forward | 5’-ATTCATGTGCCAGGGTGGT-3’ | NM_017101 | 115 |
| Reverse | 5’-GATGCCAGGACCTGTATGCT-3’ |
See text for definitions. Accession numbers, primers sequences, and size of PCR products are indicated. Accession numbers correspond to the mRNA sequences.
Morphometric analysis of adipose tissue
Fat pad mass as well as cell-size distributions were measured. GWAT and PWAT from the four groups (n=6 per group) were postfixed for 24h in 4% paraformaldehyde in phosphate buffer saline and embedded in paraffin. Fixed tissues were then cut into serial 10 µm sections, mounted on gelatin-coated slides and stained with Groat’s hematoxylin and phloxin (2%), according to standard laboratory protocols. Sections were examined using light microscopy (Leica DM IRE2) and photomicrographs were captured at ×20 magnification. The surface of adipocytes was evaluated in ten randomly selected fields of vision for a total of at least 250 adipocytes using ImageJ software (NIH). Total cell number is a direct measure reflecting hyperplasia. The number of cells was estimated using the formula as described previously (Lemonnier 1972).
Statistical analysis
All data are expressed as mean±s.e.m. Statistical analysis was carried out using GraphPad Prism5 (GraphPad). A direct comparison between a pair of groups was made using an unpaired Student’s t-test or a two-way ANOVA for repeated measures followed by a Bonferroni post hoc test, where appropriate. A P value of <0.05 was considered to be statistically significant.
Results
Effects of HF diet on maternal parameters
HF-fed female rats gained more weight than C females (ANOVA P<0.0001) (Fig. 1A). After 14weeks of HF diet, females had about two-fold higher plasma leptin concentration (4.3±0.6 vs 2.2±0.4ng/mL, P<0.05) compared with C females (Table 2). No difference was observed in basal glycemia, plasma insulin, and corticosterone levels. However, HF-fed females showed a more pronounced increase in glucose levels during OGTT with a higher area under the curve (AUC) (Fig. 1B), reflecting impaired glucose tolerance (ANOVA, P<0.0001). HF-fed dams displayed a 20% increase in BW at the end of the gestation (Fig. 1A). At weaning, HF-fed dams exhibited about three-fold higher plasma leptin (2.5±0.4 vs 0.9±0.2ng/mL, P<0.05) and corticosterone levels (796.2±187 vs 278.1±86.8ng/mL, P<0.05) (Table 2), while showing a marked increase in all fat pads weights (data not shown). No variation was observed in glycemia or plasma insulin levels.
Maternal phenotype. BW of dams fed either a C or a HF diet during preconception, gestation, and lactation (A). Concentration of blood glucose during OGTT after 14 weeks of diet and area under the curve (B) (n=12 per group). *Effect of maternal HF diet vs maternal C diet (**P<0.01; ***P<0.001).
Citation: Journal of Endocrinology 230, 1; 10.1530/JOE-16-0037
Plasma parameters of dams fed either a control (C) or a high-fat (HF) diet.
| Dams after 14 weeks of diet | Dams at weaning | |||||
|---|---|---|---|---|---|---|
| Variable | C | HF | Diet effect P-value | C | HF | Diet effect P-value |
| Glucose (dg/mL) | 95.1 ± 4.7 | 96±2.2 | NS | 107±8.7 | 114.3±21 | NS |
| Insulin (µU/mL) | 8.6 ± 0.4 | 9.3±1.3 | NS | 10.4±1 | 12.4±0.6 | NS |
| Leptin (ng/mL) | 2.2 ± 0.4 | 4.3±0.6 | <0.05 | 0.9±0.2 | 2.5±0.4 | <0.05 |
| Corticosterone (ng/mL) | 756.4 ± 206.6 | 1143±317.6 | NS | 278.1±86.8 | 796.2±187.7 | <0.05 |
Values are means±s.e.m. (n=12 females per group).
Effects of maternal obesity on offspring growth during lactation
Maternal obesity did not impact the litter size (C: 10±2 pups vs HF: 9±3 pups) or the birth weight of offspring (C: 6.2±0.1g vs HF: 6.3±0.1g). However, both sexes of HF neonates exhibited rapid weight gain during lactation (ANOVA, P<0.0001) (Fig. 2). Post hoc analysis revealed difference in BW from PND10 in male offspring (Fig. 2A) and from PND17 in female offspring (Fig. 2B). At weaning, HF male offspring exhibited a 30% increase, whereas HF female offspring only showed a 10% increase in BW (Fig. 2A and B).
Rat offspring growth curves from birth to weaning. BW of male (A) and female (B) from C or HF dams (n=16 per group). *Effect of maternal HF diet vs maternal C diet (*P<0.05; **P<0.01; ***P<0.001).
Citation: Journal of Endocrinology 230, 1; 10.1530/JOE-16-0037
Effects of maternal obesity on offspring growth and energy intake from weaning to adulthood
Weaned offspring was fed a C diet until 9months of age. The BW difference between C and HF male rats persisted until 12weeks of age (Fig. 3A). Among females, BW equalized between C and HF rats as early as 1week after weaning (Fig. 3B). In both sexes, HF rats exhibited similar weight-normalized energy intake during adulthood, suggesting that they were not hyperphagic (Fig. 3C and D). However, HF 4-week-old female offspring exhibited significantly increased weight-normalized energy intake as compared with C rats (Fig. 3D). This transient postweaning hyperphagia may reflect female-specific regulations of food intake from milk to solid diet. HF 6-month-old rats placed in metabolic cages exhibited a lower food intake during the dark phase and a higher food intake during the light phase, resulting in modified light/dark phase food intake rhythm (data not shown).
Rat offspring growth curves and food intake from weaning to adulthood. BWs of male (A) and female (B) from C or HF dams. Weekly weight-related energy intake of male (C) and female (D) from C or HF dams (n=16 per group). Weight-related energy intake is defined as the energy content of the food ingested (kcal) expressed relative to BW (g). *Effect of maternal HF diet vs maternal C diet (*P<0.05; **P<0.01; ***P<0.001).
Citation: Journal of Endocrinology 230, 1; 10.1530/JOE-16-0037
Effects of maternal obesity on adult offspring metabolic parameters
At 7months of age, C and HF offspring had comparable fasting blood glucose concentrations. During OGTT, HF males displayed increased glucose levels at 30min (P<0.01) (Fig. 4A) with a trend toward higher AUC (P=0.09) (Fig. 4B), reflecting mild glucose intolerance. No difference was observed in HF female rats (Fig. 4C and D). In HF male rats, serum corticosterone concentration was about two-fold higher than that in C animals after fasting (188.7±27.94 vs 90.42±14.18ng/mL, P<0.01) and feeding (57.4±9.4 vs 23±3.4ng/mL, P<0.05) conditions (Fig. 5A). No difference was observed in HF female rats (Fig. 5B).
Oral glucose tolerance test in 7-month-old adult offspring. Time course of plasma glucose in male (A) and female (C) from C or HF dams (n=16 per group). The respective AUCs are shown (B, D). *Effect of maternal HF diet vs maternal C diet (**P<0.01).
Citation: Journal of Endocrinology 230, 1; 10.1530/JOE-16-0037
Plasma corticosterone concentrations in fasted and fed conditions in 7-month-old adult offspring. Plasma levels in male (A) and female (B) from C or HF dams (n=16 per group). Effect of maternal HF diet vs maternal C diet (***P<0.001). #Effect of fasted condition vs fed condition (#P<0.001).
Citation: Journal of Endocrinology 230, 1; 10.1530/JOE-16-0037
Unlike females, HF 9-month-old male rats displayed increased PWAT (45.8±2.4 vs 39.2±2.6mg/g BW, P<0.05) and GWAT weights, when normalized to BW (35±1.6 vs 30±1.7mg/g BW, P<0.05) compared with C rats (Table 3). These findings were consistent with an increase in plasma leptin levels (Table 4). HF females exhibited a decrease in interscapular brown fat pad weight, but this was not observed in HF males (1.54±0.16 vs 1.97±0.1mg/g BW, P<0.05) (Table 3).
Body composition of 9-month-old offspring from control (C) or obese (HF) dams.
| Male offspring | Female offspring | ||||||||
|---|---|---|---|---|---|---|---|---|---|
| Variable | C | HF | Diet effect P-value | C | HF | Diet effect P-value | Effect of sex P | Effect of maternal diet P | Interaction P |
| Body weight (g) | 563.7±10.4 | 581.3±9.9 | NS | 291.06±6.38 | 282.89±8.34 | NS | <0.0001 | 0.65 | 0.22 |
| Perirenal adipose tissue (mg/g BW) | 39.2±2.6 | 45.8±2.4 | <0.05 | 38±4.72 | 37.94±2.04 | NS | 0.16 | 0.14 | 0.12 |
| Gonadal adipose tissue (mg/g BW) | 30±1.7 | 35±1.6 | <0.01 | 28.88±3.43 | 26.99±2.15 | NS | 0.03 | 0.17 | 0.03 |
| Brown fat pad (mg/g BW) | 1.2±0.05 | 1.2±0.08 | NS | 1.97±0.1 | 1.54±0.16 | <0.05 | <0.0001 | 0.02 | 0.02 |
| Liver (mg/g BW) | 22.43±0.4 | 23.37±0.4 | NS | 23.43±0.64 | 23.11±0.48 | NS | 0.57 | 0.66 | 0.28 |
| Heart (mg/g BW) | 2.38±0.04 | 2.28±0.04 | NS | 2.75±0.19 | 3.16±0.11 | <0.05 | <0.0001 | 0.15 | 0.02 |
| Kidney (mg/g BW) | 4.8±0.2 | 4.9±0.09 | NS | 6.16±0.143 | 6.17±0.13 | NS | <0.0001 | 0.35 | 0.39 |
| Adrenal (mg/g BW) | 0.1±0.01 | 0.1±0.01 | NS | 0.23±0.013 | 0.19±0.02 | <0.05 | <0.0001 | 0.03 | 0.1 |
Values are mean±s.e.m. (n=16 rats per group).
Plasmaparametersof9-month-oldoffspringfromcontrol(C)orobese(HF)dams.
| Maleoffspring | Femaleoffspring | ||||||||
|---|---|---|---|---|---|---|---|---|---|
| Variable | C | HF | Diet effect P-value | C | HF | Diet effect P-value | Effectofsex P | Effectofmaternaldiet P | Interaction P |
| Glucose(dg/mL) | 105.7±2.76 | 104±1.63 | NS | 99±3.7 | 87.89±2.84 | NS | 0.0003 | 0.04 | 0.12 |
| NEFA(mM) | 0.63±0.03 | 0.62±0.03 | NS | 0.72±0.04 | 0.74±0.09 | NS | 0.02 | 0.77 | 0.74 |
| Triacylglycerol(mg/dL) | 198.9±16.94 | 216.7±16.08 | NS | 91.77±10.94 | 76.01±18.41 | NS | <0.0001 | 0.95 | 0.73 |
| Totalcholesterol(mM) | 0.99±0.04 | 1.01±0.04 | NS | 0.66±0.04 | 0.59±0.04 | NS | <0.0001 | 0.55 | 0.29 |
| Insulin(µU/mL) | 33.47±3.2 | 43.63±3.79 | <0.05 | 10.79±2.07 | 10.5±1.86 | NS | <0.0001 | 0.16 | 0.13 |
| Leptin(ng/mL) | 10.15±1.11 | 15.06±1.25 | <0.01 | 3.27±0.51 | 5.76±1.35 | NS | <0.0001 | 0.002 | 0.3 |
| Adiponectin(µg/mL) | 13.73±1.1 | 14.5±1.12 | NS | 22.12±1.56 | 21.93±3.93 | NS | <0.0001 | 0.85 | 0.76 |
| Apelin(ng/mL) | 7.1±0.86 | 6.21±1.08 | NS | 3.99±0.25 | 2.97±0.2 | NS | 0.001 | 0.29 | 0.93 |
Valuesaremean±s.e.m.(n=16ratspergroup).
At 9months of age, HF male rats had about 1.5-fold higher plasma insulin (43.63±3.79 vs 33.47±3.23µ/mL, P<0.05) and leptin concentrations (15.06±1.25 vs 10.15±1.11ng/mL, P<0.05) compared with C rats, whereas no difference was observed in HF females (Table 4). The increased Homeostasis Model Assessment-insulin resistance (HOMA-IR) index (+38.5%, P<0.05) suggests that HF male rats had decreased insulin sensitivity compared with C rats.
Effects of maternal obesity on adult offspring adipose tissue morphometric parameters
As shown in representative photographs (Fig. 6A), PWAT of HF male offspring exhibited an increase in average adipocyte area (Fig. 6B) and total cell number (Fig. 6C) compared with C rats. Adipocytes measuring 7500µm2 or less represented 65% of all adipocytes in C male offspring, whereas they represented only 40% of all adipocytes in HF male offspring. This indicates that maternal HF diet decreased the frequency of small-sized adipocytes (Fig. 6D). In particular, the proportion of adipocytes measuring 2500µm2 or less displayed a marked sixfold decrease in HF male offspring. In addition, these animals showed greater percentage of large-sized adipocytes (7500–40,000µm2) (60% vs 35%) compared with C rats. Although no changes in average adipocyte area (Fig. 7A and B) and total cell number (Fig. 7C) were observed in GWAT of HF males, a marked five-fold reduction in frequency of 0–2500 size adipocytes similarly occurred (Fig. 7D). By contrast, both fat pads of HF female offspring (Figs 6 and 7) showed no major changes.
Morphometric analysis of perirenal white adipose tissue in 9-month-old offspring. Representative photomicrographs of paraffin-embedded sections (scale bars=100µm) (A), average area (B), total cell number (C), and percentage of adipocytes in a given size range (area in µm2) (D) in male and female offspring from C or HF dams (n=6 per group). *Effect of maternal HF diet vs maternal C diet (*P<0.05; **P<0.01). #Effect of male vs female (#P<0.05; ##P<0.01).
Citation: Journal of Endocrinology 230, 1; 10.1530/JOE-16-0037
Morphometric analysis of gonadal white adipose tissue in 9-month-old offspring. Representative photomicrographs of paraffin-embedded sections (scale bars=100µm) (A), average area (B), total cell number (C), and percentage of adipocytes in a given size range (area in µm2) (D) in male and female offspring from C or HF dams (n=6 per group). *Effect of maternal HF diet vs maternal C diet (*P<0.05; **P<0.01). #Effect of male vs female (#P<0.05; ##P<0.01).
Citation: Journal of Endocrinology 230, 1; 10.1530/JOE-16-0037
Effects of maternal obesity on adult offspring adipose tissue gene expression profile
Maternal obesity led to pronounced changes in PWAT gene expression in HF adult male offspring. RT-quantitative PCR data showed that leptin mRNA content was increased (+1.5-fold) in HF male rats compared with C animals (Fig. 8A). This is in agreement with the increased serum leptin levels (Table 4). In PWAT, maternal obesity resulted in increased mRNA levels for genes promoting de novo lipogenesis such as fatty acid synthase (Fas, +1.6-fold) and sterol regulatory element-binding protein-1 (Srebp1, +1.7-fold) in HF male offspring. In addition, mRNA expression levels of genes involved in adipogenesis such as Pparγ and 11β-hydroxysteroid dehydrogenase type 1 (11β-Hsd1) were decreased (–1.3- and –1.5-fold, respectively) in HF male offspring vs C rats (Fig. 8A). By contrast, maternal obesity did not affect GWAT mRNA expression profiles to the same extent in HF male rats, except for a downregulation of Ob-Rb (Fig. 9A). Compared with HF males, less change in gene expression profile was observed in HF female adult offspring in both fat pads. HF female offspring still showed increased CCAAT/enhancer binding proteinα(C/Eepα) mRNA expression levels (+2-fold) in PWAT (Fig. 8B) and lower adiponectin (–1.4-fold) and glucocorticoid receptor (Gr) (–1.4-fold) mRNA expression levels in GWAT (Fig. 9B). This was consistent with differences in fat depot weights and plasma leptin concentrations observed in HF male vs female offspring (Tables 3 and 4).
mRNA expression levels of perirenal white adipose tissue in 9-month-old offspring. Gene expression was measured in male (A) and female (B) by quantitative real-time PCR, corrected for the mean expression of reference gene (cyclophilin A) (n = 10 per group). Gene symbols are detailed in Table 1. *Effect of maternal HF diet vs maternal C diet (*P < 0.05; **P < 0.01).
Citation: Journal of Endocrinology 230, 1; 10.1530/JOE-16-0037
mRNA expression levels of gonadal white adipose tissue in 9-month-old offspring. Gene expression was measured in male (A) and female (B) by quantitative real-time PCR, corrected for the mean expression of reference gene (cyclophilin A) (n=10 per group). Gene symbols are detailed Table 1. *Effect of maternal HF diet vs maternal C diet (*P<0.05).
Citation: Journal of Endocrinology 230, 1; 10.1530/JOE-16-0037
Discussion
The main finding of this study is that maternal obesity has long-lasting consequences on visceral WAT of adult rat offspring in a depot- and sex-specific manner. In particular, we showed that HF adult male offspring exhibit greater visceral fat pad weights with adipocyte hypertrophy and hyperplasia, despite no difference in BW and energy intake. Our findings disagree with other studies (Kirk et al. 2009, Nivoit et al. 2009, Desai et al. 2014), reporting that maternal obesity prior and throughout pregnancy and lactation programs hyperphagia and marked increased BW in adult rat offspring. The discrepancy between our results and those of others may reflect differences in fetal and/or postnatal programming. These differences may depend on the duration of maternal HF feeding, the dietary lipid content (percentage of lipids vs carbohydrates), the lipid composition (saturated vs unsaturated), and the palatability (presence of sweetened condensed milk) of the diet and therefore the severity of maternal obesity (i.e. weight gain, hormonal status, inflammation grade, and so on). It may also depend on the difference of genetic background of rat strain used (Wistar vs Sprague–Dawley) (Zambrano et al. 2010).
In our model, HF adult offspring might have developed a modification of WAT’s sensitivity to circulating hormones. Indeed, perirenal WAT depot exhibited changes in lipogenic and adipogenic pathways that may favor triglyceride storage in mature adipocytes. Consistent with increased fat deposition, HF adult male offspring displayed higher serum leptin concentration. Hyperleptinemia may be interpreted as a leptin-resistant state (Kirk et al. 2009, Sun et al. 2012). First, leptin is known to activate adipogenesis by promoting preadipocyte differentiation (Bol et al. 2008, Guo et al. 2009). However, despite hyperleptinemia, the number of small adipocytes was markedly decreased in both depots of HF adult male offspring. In agreement with these findings, we observed that Pparγ gene expression was downregulated. Our findings disagree with other studies that reported an upregulation of Pparγ contents in WAT along with enhanced adipogenesis in obese-prone offspring from malnourished dams (Samuelsson et al. 2008, Muhlhausler & Smith 2009, Sen & Simmons 2010, Desai et al. 2015). This discrepancy may be due to differences in the establishment of epigenetic marks during adipogenesis and/or hormonal environment, tissue sensitivity, as well as inflammatory status in adipose tissue of adult offspring (Breton 2013). The decrease in gene expression might be seen as an adaptive mechanism to limit fat accumulation (Lukaszewski et al. 2011). Indeed, an increase in the lipogenic capacity of adipose tissue is expected during the ‘dynamic phase of obesity’, when fat stores are rapidly expanding. However, during long-lasting and stable obesity, the decreased expression of lipogenic genes may prevent a further development of fat mass (Ortega et al. 2010). Further experiments on the kinetic of fat deposition and the transcriptional profile of lipogenic genes during the development of WAT are needed to address this question. In agreement with this hypothesis, several studies described a relationship between obesity and lower expression and/or activity of Pparγ in visceral WAT. These modifications appear to be strongly associated with the pathogenesis of metabolic syndrome (Zhang et al. 1996, Fujiki et al. 2009). Second, given the antilipogenic leptin action on mature adipocytes (Huan et al. 2003, Jiang et al. 2009), HF adult male offspring had a trend toward reduced leptin receptor contents and did not show any suppression of Fas gene expression, but instead a markedly increased expression. In accordance with these findings, we reported that hyperleptinemic rat offspring from undernourished dams exhibited impaired leptin sensitivity with reduced pSTAT3 in WAT (Lukaszewski et al. 2011).
Moreover, HF adult male offspring exhibited hyperinsulinemia with elevated Srebp1 and Fas mRNA levels in PWAT, two genes that are known to be upregulated by insulin levels. This suggests that adipose tissue remains sensitive to insulin. This is in agreement with a greater insulin-induced AKT phosphorylation and the upregulation of lipogenic pathways observed in WAT of HF offspring (Borengasser et al. 2013). HF adult male offspring also displayed hypercorticosteronemia. Several lines of evidence prompted us to explore whether sensitivity of WAT to glucocorticoid (GC) was modified in offspring. First, GC alone or in interaction with insulin regulates the differentiation of preadipocytes and lipogenic genes (Campbell et al. 2011). Second, a close link between chronic excess of plasma GC levels and/or increased GC sensitivity within WAT (i.e. modifications of GR, MR, 11β-hydroxysteroid dehydrogenase type 1 (11β-HSD1), and 11β-hydroxysteroid dehydrogenase type 2 (11β-HSD2) contents) and fat expansion has been observed in offspring from malnourished dams (Gnanalingham et al. 2005, Lukaszewski et al. 2011, Guo et al. 2013). Third, increased expression of GR and 11β-HSD1 in visceral adipose tissue has been associated with the development of obesity in rats overfed during lactation (Boullu-Ciocca et al. 2008). We observed a depot-specific downregulation of 11β-Hsd1 mRNA in PWAT in HF adult offspring as previously reported in 3-month-old offspring of obese mice (Samuelsson et al. 2008). We also showed that the ratio between 11β-Hsd1 and 11β-Hsd2 expression that controls local balance between active and inactive GC metabolites (Lee et al. 2014) was decreased. As described in obesity-prone progeny from undernourished dams, it may diminish intratissular GC responsiveness and represent an adaptive mechanism to counteract excess fat storage (Lukaszewski et al. 2011). We cannot exclude that HF offspring may have decreased energy expenditure. Indeed, additional programming mechanisms such as elevated FFA transport and/or lower lipolysis/β-oxydation activities within WAT might account for increased triglyceride storage (Dahlhoff et al. 2014).
At birth, HF offspring had a normal birth weight and, then, exhibited a rapid weight gain during lactation, a key period of adipose tissue development. Adipocyte stem cells are also very sensitive to maternal factors during this developmental period (Tang & Lane 2012). Adipocyte number expansion that is set earlier in obese individual may be a major determinant for increased fat mass in adulthood (Spalding et al. 2008). Obesity may arise from increased lipid storage in mature adipocytes during the perinatal period. In line with these findings, we showed that maternal obesity predisposes adult offspring to adiposity by increasing the number of adipocytes and the average fat cell volume. The accelerated postnatal growth in offspring is frequently associated with persisting adiposity throughout life. Several models have shed light on the importance of energy intake and milk composition during the lactation period for adipose tissue programming. Indeed, pups from mothers exposed to HF diet only during lactation (Sun et al. 2012, Desai et al. 2014, White & Tchoukalova 2014) and neonates reared in small litters, representing a model of postnatal overfeeding (Boullu-Ciocca et al. 2008), also displayed persistent hypertrophic adipocytes with enhanced adipogenic and lipogenic mRNA expression levels. However, maternal obesity before conception and gestation is also able to program similar outcomes during the embryonic period. Indeed, despite normal fetal weight, fetus from mice fed a HF diet prior and throughout pregnancy displayed larger adipocytes (Murabayashi et al. 2013, Umekawa et al. 2015) and increased mRNA expression levels of Zfp423, a key transcriptional factor initiating adipogenic commitment (Yang et al. 2013). Adult mouse offspring also exhibited increased mRNAs levels of several genes involved in de novo lipogenesis and lipid droplet size in visceral WAT (Dahlhoff et al. 2014). Similarly, obesity-prone rat offspring from obese dams that were induced intragastric HF diet feeding displayed an increase in adipogenic and lipogenic pathways (Shankar et al. 2008).
Finally, we showed that maternal obesity sensitizes adult rat offspring to increased visceral adiposity in a depot- and sex-specific manner. Indeed, among GWAT and PWAT, only the latter shows marked programming features in HF male offspring. By contrast, very few variations were observed in WAT of HF female offspring. In line with these findings, studies have previously demonstrated the heterogeneity of the adipose lineage. All adipogenic stem cells and adipocytes do not behave equally during adipogenesis. Indeed, each fat depot has a unique developmental gene expression signature (Yamamoto et al. 2010). Fat stem cells are influenced by the anatomic location of the depot and/or the hormonal microenvironment, as well as aging, gender, and metabolic health (Williams et al. 2014). Thus, intrinsic genetic depot-specific differences in adipose stem cells result in different adipogenic potential, gene expression profile, growth rate, and biological properties (i.e. hormone sensitivity) between visceral and subcutaneous fat pads, but also between each specific visceral fat pad. The fact that different adipocyte precursors might determine the development and the function of specific fat pads led to the notion that each WAT depot could be considered a separate mini-organ (Berry et al. 2013).
Among programming mechanisms, inappropriate hormone levels during the perinatal period, are a key factor leading to persistent deregulation of energy homeostasis in progeny. It may result in long-term fat expansion with permanent changes in plasma hormone levels in adult offspring (Breton 2013). Consistent with this notion, maternal obesity prolonged and amplified the plasma leptin surge in offspring in a sex-specific manner (Kirk et al. 2009, Masuyama & Hiramatsu 2014). Maternal HF diet during lactation was also associated with increased insulin and leptin levels in milk (Vogt et al. 2014). Leptin that displays differential morphogenesis effects on male and female adipocytes (Guo et al. 2009) might account for WAT’s programming dimorphism. However, despite the marked lactation effect in HF male vs female neonates, gender-specific modifications of plasma hormone levels and/or adipose tissue hormonal sensitivity remain to be determined.
Maternal obesity may also affect epigenetic mechanisms during adipogenesis. These modifications might be persistent and have long-term effects on the expression of adipogenic and lipogenic genes. We hypothesize that maternal obesity affects offspring’s energy and hormonal status modifying activity of the enzymatic components of the epigenetic machinery. It may cause epigenetic modifications that reprogram offspring’s adipose tissue. Differences in fat cell embryonic origin, development, genetic, and hormonal sensitivity may result in a depot-specific programming effects that may predispose offspring to higher adiposity (öst & Pospisilik 2015). Indeed, maternal obesity in mice induces increased gene expression of Zfp423 with lower promoter methylation levels in fetal offspring (Yang et al. 2013). Similarly, weanling rats from obese dams display increased Zfp423 and C/Ebpβ mRNA expression levels with alterations in DNA methylation of CpG sites (Borengasser et al. 2013). Maternal HF diet during pregnancy also results in histone modifications within leptin and adiponectin promoter regions with gene expression modifications in mouse offspring (Masuyama & Hiramatsu 2012).
Few studies have reported that maternal obesity programs metabolic alterations and adiposity differently in a sex-dependent manner in progeny (Sun et al. 2012, Ornellas et al. 2013, Dahlhoff et al. 2014, Masuyama & Hiramatsu 2014). The basis of the sex-specific programming effects remains unclear, but could reflect direct interactions between nutritional signals and sex hormones in the tissues of the developing fetus (Aiken & Ozanne 2013). In human, numerous studies suggest that sex differences in fetal growth in response to adverse pregnancy conditions are likely to be mediated by sex-specific adaptation of the placenta (Clifton 2010). Similarly, sex-specific programming effects in rat offspring from obese dams might be due to sex-specific differences in placental response to maternal obesity (Reynolds et al. 2015). Epigenetic mechanisms may also contribute to placental programming in a dimorphic manner. Thus, the consumption of HF diet during pregnancy appears to differently affect placental methylation and placental gene expression patterns in male and female mice offsprings (Gallou-Kabani et al. 2010). Thus, sex-specific differences in terms of epigenetic modulations may be associated with developmentally programmed phenotypes. It is possible that postnatal hormonal milieu, which is different between male and female offspring, modify the programming of adipose tissue induced by maternal obesity. This may result in gender-specific outcomes in relation to different sex-steroids (Dunn et al. 2011). Thus, a better knowledge of the epigenome changes in response to maternal obesity may provide a promising way forward to reverse adverse programming of adiposity.
Declaration of interest
The authors declare that there is no conflict of interest that could be perceived as prejudicing the impartiality of the research reported.
Funding
This study was supported by grants from the French ‘Heart and Arteries’ Foundation and grants of the French Ministry of Education.
Acknowledgments
The authors thank Valérie Montel, Anne Dickes-Coopman, Charlène Pourpe, and Capucine Besengez for their excellent technical assistance.
References
Aiken CE & Ozanne SE 2013 Sex differences in developmental programming models. Reproduction 145 R1–R13. (doi:10.1530/REP-11-0489)
Barker DJP 2004 Developmental origins of adult health and disease. Journal of Epidemiology and Community Health 58 114–115. (doi:10.1136/jech.58.2.114)
Benkalfat NB, Merzouk H, Bouanane S, Merzouk S-A, Bellenger J, Gresti J, Tessier C & Narce M 2011 Altered adipose tissue metabolism in offspring of dietary obese rat dams. Clinical Science 121 19–28. (doi:10.1042/CS20100534)
Berry DC, Stenesen D, Zeve D & Graff JM 2013 The developmental origins of adipose tissue. Development 140 3939–3949. (doi:10.1242/dev.080549)
Björntorp P & Sjöström L 1971 Number and size of adipose tissue fat cells in relation to metabolism in human obesity. Metabolism: Clinical and Experimental 20 703–713. (doi:10.1016/0026-0495(71)90084-9)
Bol VV, Reusens BM & Remacle CA 2008 Postnatal catch-up growth after fetal protein restriction programs proliferation of rat preadipocytes. Obesity 16 2760–2763. (doi:10.1038/oby.2008.417)
Borengasser SJ, Zhong Y, Kang P, Lindsey F, Ronis MJJ, Badger TM, Gomez-Acevedo H & Shankar K 2013 Maternal obesity enhances white adipose tissue differentiation and alters genome-scale DNA methylation in male rat offspring. Endocrinology 154 4113–4125. (doi:10.1210/en.2012-2255)
Bouchard C 2009 Childhood obesity: are genetic differences involved? American Journal of Clinical Nutrition 89 1494S–1501S. (doi:10.3945/ajcn.2009.27113C)
Boullu-Ciocca S, Achard V, Tassistro V, Dutour A & Grino M 2008 Postnatal programming of glucocorticoid metabolism in rats modulates high-fat diet-induced regulation of visceral adipose tissue glucocorticoid exposure and sensitivity and adiponectin and proinflammatory adipokines gene expression in adulthood. Diabetes 57 669–677. (doi:10.2337/db07-1316)
Breton C 2013 The hypothalamus – adipose axis is a key target of developmental programming by maternal nutritional manipulation. Journal of Endocrinology 216 R19–R31 (doi:10.1530/JOE-12-0157)
Campbell JE, Peckett AJ, D’souza AM, Hawke TJ & Riddell MC 2011 Adipogenic and lipolytic effects of chronic glucocorticoid exposure. American Journal of Physiology: Cell Physiology 300 C198–C209. (doi:10.1152/ajpcell.00045.2010)
Clifton VL 2010 Review: sex and the human placenta: mediating differential strategies of fetal growth and survival. Placenta 31 (Supplement) S33–S39. (doi:10.1016/j.placenta.2009.11.010)
Dahlhoff M, Pfister S, Blutke A, Rozman J, Klingenspor M, Deutsch MJ, Rathkolb B, Fink B, Gimpfl M & Hrabě de Angelis M et al. 2014 Peri-conceptional obesogenic exposure induces sex-specific programming of disease susceptibilities in adult mouse offspring. Biochimica et Biophysica Acta 1842 304–317. (doi:10.1016/j.bbadis.2013.11.021)
Desai M & Ross MG 2011 Fetal programming of adipose tissue: effects of intrauterine growth restriction and maternal obesity/high-fat diet. Seminars in Reproductive Medicine 29 237–245. (doi:10.1055/s-0031-1275517)
Desai M, Jellyman JK, Han G, Beall M, Lane RH & Ross MG 2014 Maternal obesity and high-fat diet program offspring metabolic syndrome. American Journal of Obstetrics and Gynecology 211 237.e1–e237.e13. (doi:10.1016/j.ajog.2014.03.025)
Desai M, Jellyman JK, Han G, Lane RH & Ross MG 2015 Programmed regulation of rat offspring adipogenic transcription factor (PPARγ) by maternal nutrition. Journal of Developmental Origins of Health and Disease 6 530–538. (doi:10.1017/S2040174415001440)
Dunn GA, Morgan CP & Bale TL 2011 Sex-specificity in transgenerational epigenetic programming. Hormones and Behavior 59 290–295. (doi:10.1016/j.yhbeh.2010.05.004)
Fujiki K, Kano F, Shiota K & Murata M 2009 Expression of the peroxisome proliferator activated receptor gamma gene is repressed by DNA methylation in visceral adipose tissue of mouse models of diabetes. BMC Biology 7 38. (doi:10.1186/1741-7007-7-38)
Gallou-Kabani C, Gabory A, Tost J, Karimi M, Mayeur S, Lesage J, Boudadi E, Gross M-S, Taurelle J & Vigé A et al. 2010 Sex- and diet-specific changes of imprinted gene expression and DNA methylation in mouse placenta under a high-fat diet. PLoS ONE 5 e14398. (doi:10.1371/journal.pone.0014398)
Gnanalingham MG, Mostyn A, Symonds ME & Stephenson T 2005 Ontogeny and nutritional programming of adiposity in sheep: potential role of glucocorticoid action and uncoupling protein-2. American Journal of Physiology: Regulatory, Integrative and Comparative Physiology 289 R1407–R1415. (doi:10.1152/ajpregu.00375.2005)
Guberman C, Jellyman JK, Han G, Ross MG & Desai M 2013 Maternal high-fat diet programs rat offspring hypertension and activates the adipose renin-angiotensin system. American Journal of Obstetrics and Gynecology 209 262. e1–e8. (doi:10.1016/j.ajog.2013.05.023)
Guo K, Mogen J, Struzzi S & Zhang Y 2009 Preadipocyte transplantation: an in vivo study of direct leptin signaling on adipocyte morphogenesis and cell size. American Journal of Physiology: Regulatory, Integrative and Comparative Physiology 296 R1339–R1347. (doi:10.1152/ajpregu.90691.2008)
Guo C, Li C, Myatt L, Nathanielsz PW & Sun K 2013 Sexually dimorphic effects of maternal nutrient reduction on expression of genes regulating cortisol metabolism in fetal baboon adipose and liver tissues. Diabetes 62 1175–1185. (doi:10.2337/db12-0561)
Hanson MA & Gluckman PD 2014 Early developmental conditioning of later health and disease: physiology or pathophysiology? Physiological Reviews 94 1027–1076. (doi:10.1152/physrev.00029.2013)
Huan J-N, Li J, Han Y, Chen K, Wu N & Zhao AZ 2003 Adipocyte-selective reduction of the leptin receptors induced by antisense RNA leads to increased adiposity, dyslipidemia, and insulin resistance. Journal of Biological Chemistry 278 45638–45650. (doi:10.1074/jbc.M304165200)
Jiang L, Wang Q, Yu Y, Zhao F, Huang P, Zeng R, Qi RZ, Li W & Liu Y 2009 Leptin contributes to the adaptive responses of mice to high-fat diet intake through suppressing the lipogenic pathway. PLoS ONE 4 e6884. (doi:10.1371/journal.pone.0006884)
Kirk SL, Samuelsson A-M, Argenton M, Dhonye H, Kalamatianos T, Poston L, Taylor PD & Coen CW 2009 Maternal obesity induced by diet in rats permanently influences central processes regulating food intake in offspring. PLoS ONE 4 e5870. (doi:10.1371/journal.pone.0005870)
Lecoutre S & Breton C 2014 The cellularity of offspring’s adipose tissue is programmed by maternal nutritional manipulations. Adipocyte 3 256–262. (doi:10.4161/adip.29806)
Lecoutre S & Breton C 2015 Maternal nutritional manipulations program adipose tissue dysfunction in offspring. Frontiers in Physiology 6 158. (doi:10.3389/fphys.2015.00158)
Leddy MA, Power ML & Schulkin J 2008 The impact of maternal obesity on maternal and fetal health. Reviews in Obstetrics & Gynecology 1 170–178.
Lee M-J, Pramyothin P, Karastergiou K & Fried SK 2014 Deconstructing the roles of glucocorticoids in adipose tissue biology and the development of central obesity. Biochimica et Biophysica Acta 1842 473–481. (doi:10.1016/j.bbadis.2013.05.029)
Lemonnier D 1972 Effect of age, sex, and sites on the cellularity of the adipose tissue in mice and rats rendered obese by a high-fat diet. Journal of Clinical Investigation 51 2907–2915. (doi:10.1172/JCI107115)
Lukaszewski M-A, Mayeur S, Fajardy I, Delahaye F, Dutriez-Casteloot I, Montel V, Dickes-Coopman A, Laborie C, Lesage J & Vieau D et al. 2011 Maternal prenatal undernutrition programs adipose tissue gene expression in adult male rat offspring under high-fat diet. American Journal of Physiology: Endocrinology and Metabolism 301 E548–E559. (doi:10.1152/ajpendo.00011.2011)
Lukaszewski M-A, Eberlé D, Vieau D & Breton C 2013 Nutritional manipulations in the perinatal period program adipose tissue in offspring. American Journal of Physiology: Endocrinology and Metabolism 305 E1195–E1207. (doi:10.1152/ajpendo.00231.2013)
Masuyama H & Hiramatsu Y 2012 Effects of a high-fat diet exposure in utero on the metabolic syndrome-like phenomenon in mouse offspring through epigenetic changes in adipocytokine gene expression. Endocrinology 153 2823–2830. (doi:10.1210/en.2011-2161)
Masuyama H & Hiramatsu Y 2014 Additive effects of maternal high fat diet during lactation on mouse offspring. PLoS ONE 9 e92805. (doi:10.1371/journal.pone.0092805)
Muhlhausler B & Smith SR 2009 Early-life origins of metabolic dysfunction: role of the adipocyte. Trends in Endocrinology and Metabolism 20 51–57. (doi:10.1016/j.tem.2008.10.006)
Murabayashi N, Sugiyama T, Zhang L, Kamimoto Y, Umekawa T, Ma N & Sagawa N 2013 Maternal high-fat diets cause insulin resistance through inflammatory changes in fetal adipose tissue. European Journal of Obstetrics, Gynecology, and Reproductive Biology 169 39–44. (doi:10.1016/j.ejogrb.2013.02.003)
Nivoit P, Morens C, Van Assche FA, Jansen E, Poston L, Remacle C & Reusens B 2009 Established diet-induced obesity in female rats leads to offspring hyperphagia, adiposity and insulin resistance. Diabetologia 52 1133–1142. (doi:10.1007/s00125-009-1316-9)
Ornellas F, Mello VS, Mandarim-de-Lacerda CA & Aguila MB 2013 Sexual dimorphism in fat distribution and metabolic profile in mice offspring from diet-induced obese mothers. Life Sciences 93 454–463. (doi:10.1016/j.lfs.2013.08.005)
Ortega FJ, Mayas D, Moreno-Navarrete JM, Catalán V, Gómez-Ambrosi J, Esteve E, Rodriguez-Hermosa JI, Ruiz B, Ricart W & Peral B et al. 2010 The gene expression of the main lipogenic enzymes is downregulated in visceral adipose tissue of obese subjects. Obesity 18 13–20. (doi:10.1038/oby.2009.202)
öst A & Pospisilik JA 2015 Epigenetic modulation of metabolic decisions. Current Opinion in Cell Biology 33 88–94. (doi:10.1016/j.ceb.2014.12.005)
Popkin BM, Adair LS & Ng SW 2012 Global nutrition transition and the pandemic of obesity in developing countries. Nutrition Reviews 70 3–21. (doi:10.1111/j.1753-4887.2011.00456.x)
Ravelli AC, van Der Meulen JH, Osmond C, Barker DJ & Bleker OP 1999 Obesity at the age of 50 y in men and women exposed to famine prenatally. American Journal of Clinical Nutrition 70 811–816.
Reynolds CM, Vickers MH, Harrison CJ, Segovia SA & Gray C 2015 Maternal high fat and/or salt consumption induces sex-specific inflammatory and nutrient transport in the rat placenta. Physiological Reports 3 e12399. (doi:10.14814/phy2.12399)
Samuelsson A-M, Matthews PA, Argenton M, Christie MR, McConnell JM, Jansen EHJM, Piersma AH, Ozanne SE, Twinn DF & Remacle C et al. 2008 Diet-induced obesity in female mice leads to offspring hyperphagia, adiposity, hypertension, and insulin resistance: a novel murine model of developmental programming. Hypertension 51 383–392. (doi:10.1161/HYPERTENSIONAHA.107.101477)
Sen S & Simmons RA 2010 Maternal antioxidant supplementation prevents adiposity in the offspring of Western diet-fed rats. Diabetes 59 3058–3065. (doi:10.2337/db10-0301)
Shankar K, Harrell A, Liu X, Gilchrist JM, Ronis MJJ & Badger TM 2008 Maternal obesity at conception programs obesity in the offspring. American Journal of Physiology: Regulatory, Integrative and Comparative Physiology 294 R528–R538. (doi:10.1152/ajpregu.00316.2007)
Spalding KL, Arner E, Westermark PO, Bernard S, Buchholz BA, Bergmann O, Blomqvist L, Hoffstedt J, Näslund E & Britton T et al. 2008 Dynamics of fat cell turnover in humans. Nature 453 783–787. (doi:10.1038/nature06902)
Sun K, Kusminski CM & Scherer PE 2011 Adipose tissue remodeling and obesity. Journal of Clinical Investigation 121 2094–2101. (doi:10.1172/JCI45887.2094)
Sun B, Purcell RH, Terrillion CE, Yan J, Moran TH & Tamashiro KLK 2012 Maternal high-fat diet during gestation or suckling differentially affects offspring leptin sensitivity and obesity. Diabetes 61 2833–2841. (doi:10.2337/db11-0957)
Tang QQ & Lane MD 2012 Adipogenesis: from stem cell to adipocyte. Annual Review of Biochemistry 81 715–736. (doi:10.1146/annurev-biochem-052110-115718)
Umekawa T, Sugiyama T, Du Q, Murabayashi N, Zhang L, Kamimoto Y, Yoshida T, Sagawa N & Ikeda T 2015 A maternal mouse diet with moderately high-fat levels does not lead to maternal obesity but causes mesenteric adipose tissue dysfunction in male offspring. Journal of Nutritional Biochemistry 26 259–266. (doi:10.1016/j.jnutbio.2014.10.012)
Vogt MC, Paeger L, Hess S, Steculorum SM, Awazawa M, Hampel B, Neupert S, Nicholls HT, Mauer J & Hausen AC et al. 2014 Neonatal insulin action impairs hypothalamic neurocircuit formation in response to maternal high-fat feeding. Cell 156 495–509. (doi:10.1016/j.cell.2014.01.008)
White UA & Tchoukalova YD 2014 Sex dimorphism and depot differences in adipose tissue function. Biochimica et Biophysica Acta: Molecular Basis of Disease 1842 377–392. (doi:10.1016/j.bbadis.2013.05.006)
Williams L, Seki Y, Vuguin PM & Charron MJ 2014 Animal models of in utero exposure to a high fat diet: a review. Biochimica et Biophysica Acta 1842 507–519. (doi:10.1016/j.bbadis.2013.07.006)
Yamamoto Y, Gesta S, Lee KY, Tran TT, Saadatirad P & Kahn CR 2010 Adipose depots possess unique developmental gene signatures. Obesity 18 872–878. (doi:10.1038/oby.2009.512)
Yang Q-Y, Liang J-F, Rogers CJ, Zhao J-X, Zhu M-J & Du M 2013 Maternal obesity induces epigenetic modifications to facilitate Zfp423 expression and enhance adipogenic differentiation in fetal mice. Diabetes 62 3727–3735. (doi:10.2337/db13-0433)
Zambrano E, Martínez-Samayoa PM, Rodríguez-González GL & Nathanielsz PW 2010 Dietary intervention prior to pregnancy reverses metabolic programming in male offspring of obese rats. Journal of Physiology 588 1791–1799. (doi:10.1113/jphysiol.2010.190033)
Zhang B, Berger J, Hu E, Szalkowski D, White-Carrington S, Spiegelman BM & Moller DE 1996 Negative regulation of peroxisome proliferator-activated receptor-gamma gene expression contributes to the antiadipogenic effects of tumor necrosis factor-alpha. Molecular Endocrinology 10 1457–1466. (doi:10.1210/mend.10.11.8923470)
