Abstract
Early life nutrition is important in the regulation of metabolism in adulthood. We studied the effects of different fatty acid composition diets on adiposity measures, glucose tolerance, and peripheral glucocorticoid (GC) metabolism in overfed neonatal rats. Rat litters were adjusted to a litter size of three (small litters (SLs)) or ten (normal litters (NLs)) on postnatal day 3 to induce overfeeding or normal feeding respectively. After weaning, SL and NL rats were fed a ω6 polyunsaturated fatty acid (PUFA) diet (14% calories as fat, soybean oil) or high-saturated fatty acid (high-fat; 31% calories as fat, lard) diet until postnatal week 16 respectively. SL rats were also divided into the third group fed a ω3 PUFA diet (14% calories as fat, fish oil). A high-fat diet induced earlier and/or more pronounced weight gain, hyperphagia, glucose intolerance, and hyperlipidemia in SL rats compared with NL rats. In addition, a high-fat diet increased 11β-hsd1 (Hsd11b1) mRNA expression and activity in the retroperitoneal adipose tissue of both litter groups compared with standard chow counterparts, whereas high-fat feeding increased hepatic 11β-hsd1 mRNA expression and activity only in SL rats. SL and a high-fat diet exhibited significant interactions in both retroperitoneal adipose tissue and hepatic 11β-HSD1 activity. Dietary ω3 PUFA offered protection against glucose intolerance and elevated GC exposure in the retroperitoneal adipose tissue and liver of SL rats. Taken together, the results suggest that dietary fatty acid composition in the post-sucking period may interact with neonatal feeding and codetermine metabolic alterations in adulthood.
Introduction
Obesity, particularly visceral obesity, is related to metabolic diseases such as insulin resistance, hyperlipidemia, hypertension, type 2 diabetes, and nonalcoholic fatty liver disease (Boden 2011). Numerous clinical and experimental evidence indicates that the early nutritional status of an organism may be a programming factor for the development of obesity and metabolic syndrome later in life (Ong & Loos 2006, Symonds et al. 2009), particularly malnutrition or overnutrition during the prenatal period. Elevated glucocorticoid (GC) activity has been proposed as a possible mediator of permanent obesity and metabolic syndrome programming in adults (Metges 2009).
The enzyme 11β-hydroxysteroid dehydrogenase type 1 (11β-HSD1 (HSD11B1)) provides sufficient intracellular GC levels by converting inactive GC to active GC (corticosterone or cortisol), amplifying local GC action in the adipose tissue and liver (Odermatt & Nashev 2010). Evidence from animal models and clinical studies confirmed that overexpression of 11β-HSD1 in the adipose tissue or liver is associated with metabolic disorders (Masuzaki et al. 2001, Morton et al. 2004b). By contrast, A-ring reductases (5α- and 5β-reductase) inactivate GCs in the liver (Livingstone et al. 2000), and elevated hepatic 5α reductase type 1 (5αR1) and 5βR activity is associated with insulin resistance and obesity (Westerbacka et al. 2003, Tomlinson et al. 2008). Therefore, the activities of these enzyme systems may codetermine GC availability. When studying postnatal overfeeding induced by small litter (SL) rearing in rats, Boullu-Ciocca et al. (2005) found that these rats developed hypothalamic-pituitary-adrenal (HPA) axis hyperactivity and higher 11β-hsd1 mRNA expression in the visceral adipose tissue, accompanied by obesity, insulin resistance, and hyperleptinemia in adulthood. Moreover, we (Hou et al. 2011) confirmed that a similar experimental setup induced peripheral tissue-specific alterations in 11β-hsd1 mRNA expression and activity and 5αR1 and 5βR mRNA expression at puberty, which could contribute to elevated tissue-specific GC exposure and aggravate the development of metabolic dysregulation in adults. Importantly, maturation of the rodent neuroendocrine system occurs mainly during the postnatal period, in contrast to primates, in which it occurs during the third trimester (Matthews 2002).
Dietary fatty acids are important regulators of gene expression, acting as intracellular messengers or ligands for transcription factors (Lichtenstein & Schwab 2000, Riccardi et al. 2004, Tierney et al. 2011) and are implicated in the development of metabolic syndrome. Studies have shown that local GC metabolism is influenced not only by cytokines and hormones but also by different dietary fatty acid approaches, such as composition, dose, and duration of the treatment applied in different studies (Bermudez & Sinclair 2009, Donahue et al. 2011). A diet high in saturated fatty acids increases 11β-hsd1 gene expression in the retroperitoneal adipose tissue of rats (Vara et al. 2010) and whole-body 11β-HSD1 activity in humans (Stimson et al. 2007). By contrast, a safflower oil diet (rich in polyunsaturated fatty acids (PUFAs)) reduces plasma corticosterone and adipose tissue 11β-HSD1 activity (Man et al. 2011). Dietary fish oil (rich in ω3 PUFAs) for 6 months after birth prevents hypertension and hyperleptinemia in rats programmed by fetal GC exposure (Wyrwoll et al. 2006). Taken together, these observations strongly suggest the existence of an association between local GC metabolism and nutritional status or fatty acid composition.
In this study, we investigated the effects of three diets with different fatty acid compositions on adiposity measures, glucose tolerance, and peripheral GC metabolism in neonatal rats induced to overfeed by rearing in SLs.
Materials and Methods
Animals
All the studies were approved by the University Committee on Use and Care of Animals and overseen by the Unit for Laboratory Animal Medicine at Nanjing Medical University (ID: 2008031801). Sprague Dawley rats (Nanjing, Jiangsu, China) were maintained under controlled light (0600–1800 h) and temperature (22±2 °C) conditions with free access to tap water.
Experimental design
The experimental setup was similar to Boullu-Ciocca's study (Boullu-Ciocca et al. 2008). Briefly, female rats were time-mated, and at postnatal day 3, male pups were redistributed to litter sizes of three (SLs) or ten (normal litters (NLs)) to induce early postnatal overfeeding or normal feeding respectively (Velkoska et al. 2008, Rodrigues et al. 2009). After weaning (postnatal day 21), the rats from the NLs were fed a soybean oil diet (dietary fat was soybean oil rich in ω6 PUFAs; NL group) or a diet high in saturated fats (NL-HF group), and the rats from the SLs were fed a soybean oil diet (SL group), a diet high in saturated fat (SL-HF group), or a fish oil-rich diet (dietary fat was fish oil rich in ω3 PUFAs; SL-FO group) until postnatal week 16 (W16). The diets (Slac, Shanghai, China) and their nutriment composition are shown in Table 1 (Note: the soybean oil diet was often referred to as ‘standard chow’ in previous studies (Reeves et al. 1993, Bringhenti et al. 2011)). All animals were housed three per cage postweaning and fed ad libitum. Body weight, body length, tail length, and food intake were monitored throughout life. The animals were killed at W3, W6, and W16 between 0830 and 1000 h after fasting overnight.
Purified diet formula and composition (weight (%))
| Soybean oil diet (%) | Fish oil diet (%) | High-fat diet (%) | |
|---|---|---|---|
| Casein | 18.9 | 18.9 | 18.9 |
| l-Cystine | 0.3 | 0.3 | 0.3 |
| Corn starch | 48.3 | 48.3 | 39.3 |
| Maltodextrin | 3.3 | 3.3 | 3.3 |
| Sucrose | 13.0 | 13.0 | 13.0 |
| Cellulose | 4.7 | 4.7 | 4.7 |
| Mineral mix | 4.3 | 4.3 | 4.3 |
| Vitamin mix | 1.1 | 1.1 | 1.1 |
| Soybean oil | 6.0 | 0.0 | 6.0 |
| Fish oil | 0.0 | 6.0 | 0.0 |
| Lard | 0.0 | 0.0 | 9.0 |
| Total | 100.0 | 100.0 | 100.0 |
| Energy (kcal/100 g) | 392.6 | 392.6 | 438.2 |
Serum and tissue collection
Rats were anesthetized with chloral hydrate (300 mg/kg body weight, i.p.) at 0830 h after overnight fasting (12 h) and blood samples were obtained from the right ventricle. The blood was centrifuged (2000 g, 4 °C, 15 min) and the separated serum stored at −70 °C for subsequent determination of biochemical parameters, serum insulin, corticosterone, and leptin. Kidney, liver, and epididymal and retroperitoneal fat pads were dissected immediately. The kidney and fat pads were weighed, and all tissues were snap-frozen in liquid nitrogen and kept at −80 °C until gene expression and activity analysis.
Biochemical analysis
Total triglycerides (TGs) and total cholesterol (TC) in the serum were measured using an Olympus AU400 analyzer with enzymatic reagents (Olympus America, New York, NY, USA). Serum corticosterone (Cayman, Ann Arbor, MI, USA) and leptin (Millipore, Billerica, MA, USA) were measured by ELISAs. The limit of sensitivity was 40 pg/ml for corticosterone and 0.04 ng/ml for leptin, and the intra-assay coefficients of variation for corticosterone and leptin were 7–9 and 2–3% respectively.
Intraperitoneal glucose tolerance test
The intraperitoneal glucose tolerance test (IPGTT) was performed as described previously (Chen et al. 2008). Briefly, at W6 and W16, rats were fasted overnight. A blood sample was then taken from a tail vein and the rats were injected i.p. with 2.0 g d-glucose (50% stock solution in saline)/kg body weight. Blood samples were drawn at 30-, 60-, and 120-min intervals after the glucose injection, and glucose levels were measured by a glucose meter (Accu-Chek; Roche).
Quantitative real-time PCR
Total RNA was isolated from the adipose tissue and liver using TRIzol (Invitrogen) according to the manufacturer's instructions and quantified spectrophotometrically by the OD260. The integrity of total RNA was assessed using agarose gel electrophoresis, and cDNA was synthesized using M-MLV reverse transcriptase (Promega) with 1.0 μg of the RNA sample as recommended by the manufacturer. PCR amplification using glyceraldehyde-3-phosphate dehydrogenase (GAPDH) primers with a subset of the cDNA samples confirmed successful reverse transcription. Real-time PCR was performed using the SYBR GREEN ABI Prism 7500 sequence detector with the following program: 50 °C for 2 min, 95 °C for 10 min, 40 cycles of 95 °C for 15 s, and 60 °C for 1 min. The mRNA levels were normalized to GAPDH. Data were analyzed using the 2−ΔΔct method (Pfaffl 2001). Primer sequences were as follows: 11β-HSD1, forward 5′-GAA GAA GCA TGG AGG TCA AC-3′, reverse 5′-GCA ATC AGA GGT TGG GTC AT-3′; 5αR1, forward 5′-CTG TTT CCT GAC AGG CTT TGC-3′, reverse 5′-GCC TCC CCT GGG TAT CTT GT-3′; 5βR, forward 5′-GCC TTT AAG CCT GGA GAG GAA-3′, reverse 5′-ACG TGG CAC ACA GAT TTG ATT-3′; GAPDH, forward 5′-CAA GTT CAA CGG CAC AGT CAA-3′, reverse 5′-TGG TGA AGA CGC CAG TAG ACT C-3′.
11β-HSD activity assay
To estimate the 11β-HSD1 protein levels, 11β-HSD1 activity was measured in adipose and hepatic tissues in the direction of the dehydrogenase reaction, as it is more stable in vitro than in the reductase direction (Lakshmi & Monder 1988). 11β-HSD1 enzyme activity was measured as described previously (Buren et al. 2007, Hou et al. 2011). Briefly, tissue was homogenized in homogenization buffer (10% glycerol, 300 mM NaCl, 1 mM EDTA, and 50 mM Tris, pH 7.7) containing 1 mM dithiothreitol and then centrifuged at 4 °C. The protein concentration was determined using a Pierce BCA protein assay kit with BSA as the standard (Thermo Fisher Scientific, Rockford, IL, USA). Each sample was analyzed in duplicate using an internal control. Samples of adipose tissue (0.5 mg protein/ml) or liver (10 μg protein/ml) homogenate were incubated with 2 mM NADP and 100 nM 1,2,6,7-[3H]4-corticosterone (Amersham). After incubation in a shaking water bath at 37 °C for 1 or 2 h (liver and adipose tissues respectively), the reaction was interrupted and the steroids extracted with ethyl acetate and dried, dissolved in ethanol, separated by thin-layer chromatography (TLC; mobile phase chloroform:ethanol (92:8)), and exposed to a phosphorimager tritium screen (GE Healthcare Europe, Freiburg, Germany). The TLC plates were then scanned and quantified using a Typhoon scanner (GE Healthcare Europe). 11β-HSD1 activity was expressed as the percentage conversion of corticosterone into 11-dehydrocorticosterone.
Statistical analysis
Results are expressed as mean±s.e.m. Significant differences between SLs and NLs of rats at W3 were analyzed by unpaired t-tests. Significant differences between groups of rats at W6 and W16 were analyzed by two-way ANOVA followed by a post hoc Bonferroni test. Body weight, food energy intake, and serum glucose during IPGTT were analyzed using two-way ANOVA with repeated measures, followed by a post hoc Bonferroni test. P<0.05 was considered significant.
Results
Body weight gain, energy intake, and adipose tissue weight
SL rearing induced a significant increase in weight gain compared with NLs by postnatal W2 (41.3±2.3 vs 27.8±2.3 g, P<0.01). The difference in body (Fig. 1) and fat pad (retroperitoneal and epididymal) weight persisted until W16 (Table 2). After 3 weeks on a high-fat diet, body weight and retroperitoneal fat pad weight of SL-HF rats were higher than that of SL rats, and these effects persisted until W16. SLs showed significant interactions with high-fat diet with regard to body weight (F=5.033, P<0.05). By contrast, the weight gain of SL-FO rats, as well as their fat pad weight, was less than that of SL rats, and from W12 to W16, the SL-FO rats did not differ significantly from NL rats. NL-HF rats had greater body weight and fat pad weight than NL rats at W15 and W16, but they weighed less than SL-HF rats during the experimental period. No differences were found in body and tail length or kidney weight among the groups (data not shown).
Effect of different diets on body weight from W3 to W16. Data are mean±s.e.m., n=6–9 in each group. aP<0.05 vs NL at every time point, bP<0.05 vs SL from W6 to W16, cP<0.05 vs NL-HF at every time point, and dP<0.05 for SL vs NL at every time point. The body weight of NL-HF rats were higher than NL rats at W15 and W16; there was no difference between SL-FO rats' body weight and NL rats from W12 to W16.
Citation: Journal of Endocrinology 215, 1; 10.1530/JOE-12-0191
Adiposity measurements at W3, W6, and W16 (g)
| NL | SL | NL-HF | SL-HF | SL-FO | |
|---|---|---|---|---|---|
| W3a | |||||
| RAT weight | 0.13±0.01 | 0.23±0.01* | – | – | – |
| EAT weight | 0.14±0.02 | 0.20±0.02* | – | – | – |
| W6 | |||||
| RAT weight | 1.55±0.22 | 1.98±0.44* | 1.57±0.11† | 2.26±0.14*,†,‡ | 1.90±0.87*,‡ |
| EAT weight | 1.55±0.13 | 1.80±0.21* | 1.54±0.10*,† | 1.81±0.06*,‡ | 1.77±0.23*,‡ |
| W16 | |||||
| RAT weight | 6.2±0.6 | 8.9±0.9* | 9.9±0.9* | 11.4±1.4*,†,‡ | 7.4±1.3†,‡ |
| EAT weight | 6.0±0.3 | 8.1±1.1* | 9.2±0.8* | 10.9±1.1*,†,‡ | 6.1±1.3†,‡ |
RAT, retroperitoneal adipose tissue; EAT, epididymal adipose tissue. Data are expressed as mean±s.e.m., n=6–9 in each group. *P<0.05 vs NL at the same time point, †P<0.05 vs SL at the same time point, and ‡P<0.05 vs NL-HF at the same time point.
Measured before randomization to feeding groups.
Both SL-HF and NL-HF rats exhibited higher energy intake compared with their counterparts from W3 to W16, and SL-HF rats had a higher energy intake than NL-HF rats throughout the study period. SLs and a high-fat diet showed a significant interaction for general energy intake (F=84.11, P<0.01). Although SL and SL-FO rats exhibited a higher energy intake than NL rats at W3 and W4, no difference was found in food energy intake among NL, SL, and SL-FO rats from W5 to W16 (Fig. 2).
Effect of different diets on energy intake from W3 to W16. Data are mean±s.e.m., n=6–9 in each group. aP<0.05 vs NL at every time point, bP<0.05 vs SL at every time point, and cP<0.05 vs NL-HF at every time point. *P<0.05 vs SL and SL-FO at the corresponding time point.
Citation: Journal of Endocrinology 215, 1; 10.1530/JOE-12-0191
Serum lipids
No difference was observed in serum TGs (Fig. 3A) and TC (Fig. 3B) between NL and SL rats from W6 to W16. Serum TG levels were increased in SL-HF rats compared with SL rats from W6, and a high-fat diet and SLs showed a significant interaction with regard to TGs (F=4.899, P<0.05). TG levels in NL-HF rats were increased at W16, whereas TG levels in SL-FO rats were unchanged compared with NL or SL during the experimental period (Fig. 3A). The TC levels of NL-HF and SL-HF rats were higher than the levels in their counterparts at W6 and W16 respectively, whereas fish oil decreased TC levels compared with NL and SL rats during the experimental period (Fig. 3B).
Effect of different diets on serum levels of total triglycerides (A) and total cholesterol (B) at W6 and W16. Data are mean±s.e.m., n=6–9 in each group. aP<0.05 vs NL, bP<0.05 vs SL, and cP<0.05 vs NL-HF.
Citation: Journal of Endocrinology 215, 1; 10.1530/JOE-12-0191
Glucose homeostasis
IPGTT confirmed that SLs and the high-fat diet induced glucose intolerance with a pronounced interaction at W6 (F=5.365, P<0.05), and this effect persisted to W16 (F=4.457, P<0.05). The area under the curve (AUC) for plasma glucose increased in SL-HF rats already at W6, which indicated higher insulin resistance early in development (Fig. 4C). At W16, the AUC was increased in SL, SL-HF, and NL-HF rats compared with NL rats and was highest in the SL-HF rats (Fig. 4D). By contrast, the AUC in SL-FO rats was decreased compared with SL rats and did not differ from that of NL rats.
Effect of different diets on intraperitoneal glucose tolerance test (A and B) and areas under the curve (C and D) at W6 (A and C) and W16 (B and D). Data are mean±s.e.m., n=6–9 in each group. aP<0.05 vs NL, bP<0.05 vs SL, and cP<0.05 vs NL-HF.
Citation: Journal of Endocrinology 215, 1; 10.1530/JOE-12-0191
Serum leptin and corticosterone
As shown in Table 3, serum leptin mirrored adiposity measures very well. Both SL and the high-fat diet resulted in elevated serum leptin levels. By contrast, the fish oil diet decreased serum leptin compared with SL and NL-HF rats, but serum leptin was elevated compared with NL rats at W16.
Serum leptin and corticosterone levels at W16
| NL | SL | NL-HF | SL-HF | SL-FO | |
|---|---|---|---|---|---|
| Leptin (ng/ml) | 13.0±1.0 | 18.0±0.2* | 18.8±1.8* | 21.9±1.1*,†,‡ | 15.1±1.5*,†,‡ |
| Corticosterone (ng/ml) | 10.2±0.8 | 14.0±0.7* | 10.2±1.0† | 15.5±0.6*,‡ | 12.6±0.8* |
Data are expressed as mean±s.e.m., n=6–9 in each group. *P<0.05 vs NL, †P<0.05 vs SL, and ‡P<0.05 vs NL-HF.
Serum corticosterone levels were elevated in SL rats compared with NL rats at W16. A high-fat diet had no effect on serum corticosterone levels in normal or SL rats. Moreover, we found a nonsignificant tendency for decreased serum corticosterone levels in SL-FO rats compared with SL rats (Table 3).
Hepatic 11β-hsd1 mRNA expression and enzyme activity
Hepatic 11β-hsd1 mRNA expression in rats in SLs was higher than the expression in rats from NLs at W3 (15.5±1.8 vs 8.5±1.8, P<0.05) and a similar difference persisted until W16 (Fig. 5A). Compared with SL rats, 11β-hsd1 mRNA expression was increased in SL-HF rats but decreased in SL-FO rats at W16. No difference in 11β-hsd1 mRNA expression was found between NL-HF rats and NL rats (Fig. 5A). Moreover, the pattern of hepatic 11β-HSD1 enzyme activity paralleled 11β-hsd1 gene expression in all groups, and SL and the high-fat diet exhibited a significant interaction with regard to hepatic 11β-HSD1 enzyme activity (F=22.49, P<0.01; Fig. 5B).
Effect of different diets on 11β-hsd1 mRNA expression (A) and enzyme activity (B) in the liver at W16. Data are mean±s.e.m., n=6–9 in each group. aP<0.05 vs NL, bP<0.05 vs SL, and cP<0.05 vs NL-HF.
Citation: Journal of Endocrinology 215, 1; 10.1530/JOE-12-0191
11β-Hsd1 mRNA expression and enzyme activity in adipose tissue
As shown in Fig. 6A, 11β-hsd1 mRNA expression was increased in SL rats compared with NL rats. Feeding rats a high-fat diet induced a pronounced increase in 11β-hsd1 mRNA expression in the retroperitoneal adipose tissue of both SL-HF and NL-HF rats compared with their counterparts fed standard chow. 11β-HSD1 activity was similar to gene expression data, and SL and the high-fat diet showed a significant interaction with regard to adipose tissue 11β-HSD1 activity (F=18.45, P<0.01). Similar to the liver, 11β-hsd1 mRNA expression and enzyme activity in adipose tissue was decreased in SL-FO rats compared with SL rats (Fig. 6B). In epididymal adipose tissue, 11β-hsd1 mRNA expression and enzyme activity did not differ significantly between different diet groups (data not shown).
Effect of different diets on 11β-hsd1 mRNA expression (A) and enzyme activity (B) in retroperitoneal adipose tissue at W16. Data are mean±s.e.m., n=6–9 in each group. aP<0.05 vs NL, bP<0.05 vs SL and cP<0.05 vs NL-HF.
Citation: Journal of Endocrinology 215, 1; 10.1530/JOE-12-0191
Hepatic A-ring reductase mRNA expression
No group difference was found in hepatic 5αR1 mRNA expression (data not shown). Hepatic 5βR mRNA expression was 27% lower in SL rats than in NL rats (P<0.05) but increased in SL-HF (50%, P<0.05) and NL-HF rats (62%, P<0.05) compared with SL and NL respectively. For SL-FO rats, hepatic 5βR mRNA expression was not significantly altered compared with NL or SL rats at W16 (data not shown).
Discussion
Previous studies have shown that, in rats, neonatal overfeeding induced by SL rearing results in adulthood onset of obesity, insulin resistance, hyperlipidemia, hyperleptinemia, and GC metabolism abnormalities (Boullu-Ciocca et al. 2005, Plagemann et al. 2009, Prior & Armitage 2009, Hou et al. 2011). Our study aimed to investigate whether postweaning diets with different fatty acid compositions could aggravate or ameliorate peripheral GC metabolism abnormalities and systemic metabolic disturbances in neonatally overfed rats. We found that a high-fat diet induced early-onset hyperphagia, obesity, insulin resistance, hyperlipidemia, and high circulating leptin. The effect of the high-fat diet in SL rats was earlier and more pronounced compared with NL rats. In SL-FO rats, the ω3 PUFA-rich diet prevented the increase in body weight and adipose tissue weight, lowered cholesterol, and improved glucose metabolism without reducing energy intake. The novel finding in this study was that 11β-hsd1 mRNA expression and enzyme activity were increased in both the hepatic and the visceral adipose tissue of SL-HF rats but decreased in SL-FO rats compared with SL rats. These data suggest that postweaning dietary fatty acid composition may potentially interact with metabolic and hormonal axes maturation and formation and codetermine the metabolic outcome, partly by regulating local GC action.
A high-fat diet may induce obesity in mice (Kozawa et al. 2011), rats (Mullen et al. 2009), and humans (Boden 2011). Consistent with previous reports, the high-fat diet induced visceral adipose gain and obesity in both NL-HF and SL-HF rats. However, SL-HF rats exhibited a greater rate of fat gain as well as hyperlipidemia and insulin resistance after 3 weeks on the diet, and the body weight and visceral adipose tissue weight measured for SL-HF rats were much higher than in other groups due to a significant interaction between SLs and the high-fat diet in the upregulation of serum TGs, body weight, and adipose tissue weight gain. These data suggest that rats that are overfed during lactation are more susceptible to high-fat stimulation after weaning, and this may lead to abnormal metabolic dysregulation compared with normally fed rats. Such divergent nutritional exposure during early development can reprogram the response to a particular nutritional environment, as reported previously in rat offspring exposed to gestation stress (Tamashiro & Moran 2010) and maternal malnutrition (Minana-Solis & Escobar 2007). By contrast, perinatal mice fed a normal diet are resistant to high-fat diet-induced hyperphagia, obesity, and insulin resistance (Gallou-Kabani et al. 2007). These observations indicate that the perinatal period represents a critical time frame during which metabolic regulatory set points may be modified and that persistent overnutrition (e.g. high-fat diet) may further increase the incidence of obesity in rats exposed to overfeeding during lactation.
Dietary fatty acids are important regulators of gene expression, acting as intracellular messengers or ligands for transcription factors (Lichtenstein & Schwab 2000, Riccardi et al. 2004, Tierney et al. 2011) and cause a pronounced tissue-specific regulation of 11β-HSD1 in rodents (Morton et al. 2004a, Drake et al. 2005, Vara et al. 2010). In agreement with previous findings (Shin et al. 2011), 11β-hsd1 mRNA expression and enzyme activity was increased in retroperitoneal adipose tissue from rats fed a high-fat diet, regardless of litter size. Notably, hepatic 11β-HSD1 activity was persistently higher in SL rats compared with NL rats, and SL and the high-fat diet exhibited a significant interaction with regard to 11β-HSD1 activity, whereas no changes in hepatic 11β-HSD1 expression and enzyme activity was seen in NL-HF rats, which are similar to the majority of animal models of obesity (Morton et al. 2004b, Drake et al. 2005). This inconsistency may be explained by different time frames of exposure to overnutrition (Chen et al. 2009) and implies that the key set point of 11β-HSD1 in the liver is the perinatal period. In addition, nutritional experience in early development may determine the gene response to circumstances later in life (Bouchard 2008). The overexpression of 11β-HSD1 in the liver of transgenic mice results in insulin resistance, dyslipidemia, hypertension, and fatty liver. Therefore, the persistent increase in hepatic 11β-HSD1 activity in SL and SL-HF rats may induce early-onset and more severe obesity, as well as other features of metabolic dysregulation.
Although both ω3 PUFAs and ω6 PUFAs exhibit antiobesity effects, products derived from ω6 PUFAs are more potent mediators of inflammation than similar products derived from ω3 PUFAs (Kang 2003) and a higher dietary ω6:ω3 PUFA ratio increases the risk of obesity (Ailhaud et al. 2008). Thus, we selected a fish oil diet rich in ω3 PUFAs to test the possibility of preventing the programming outcome in overfed neonatal rats. We found that ω3 PUFAs in the diet downregulated 11β-HSD1 expression and enzyme activity in the adipose tissue and liver, which is in line with ameliorated glucose tolerance and the prevention of excess adipose tissue accumulation. This finding is consistent with the well-documented effect of ω3 PUFA-rich diets against body fat gain in rats (Belzung et al. 1993) and mice (Ruzickova et al. 2004). We found that the fish oil diet prevented the development of metabolic disorders without reducing energy intake and body growth and development. Thus, ω3 PUFAs have an effect on tissue function that is not dependant on calorie restriction. Reduced local GC activation by ω3 PUFAs may play a key role in reducing adverse programming outcomes later in life, which might be a useful strategy for improving early programming effects following GC excess.
It is worth noting that we did not include a group of rats from NLs fed a fish oil diet in this study. Interestingly, there is at least one study reporting that a fish oil diet may reduce adipose weight gain in normal fed rats (Wyrwoll et al. 2006), and it is plausible that 11β-HSD1 enzyme activity levels are altered in those rats as well. However, the effect of ω3 PUFAs on GC metabolism in normal size litter fed rats still needs to be studied.
The regulation of 11β-HSD1 by dietary fatty acids may involve peroxisome proliferator-activated receptor-γ (PPARγ) and PPARα and CCAAT enhancer binding protein α (C/EBPα) (Vara et al. 2010, Vagnerova et al. 2011). PPARs are known to downregulate the expression of 11β-HSD1 (Berger et al. 2001, Srivastava 2009) and have a high affinity for PUFAs compared with saturated fatty acids, whereas a high-fat diet may increase the expression of C/EBPα, a key transcription factor for 11β-HSD1 (Kim & Park 2008). However, the details of dietary fatty acid regulation of 11β-HSD1 need to be further studied in vitro.
The A-ring reductases are the main inactivating enzymes for GC in the liver (Livingstone et al. 2000). Studies of obese or insulin-resistant animals (Livingstone et al. 2000) and humans (Tsilchorozidou et al. 2003, Baudrand et al. 2011) have revealed higher expression of hepatic 5α-reductase. In our study, 5βR mRNA expression was increased following a high-fat diet, independent of litter size. This alteration might be due to compensatory activation following a high-fat diet, independent of nutritional status, similar to Drake's report (Drake et al. 2005). The compensation may prevent circulating GC levels from increasing with a high-fat diet. Notably, 5βR mRNA expression seemed to be maintained in SL-FO rats compared with NL rats, with decreased circulating GC levels, though this was not significant. These data suggest a recovery of the GC conversion enzyme in SL-FO rats, which could be one of the mechanisms by which ω3 PUFAs prevent metabolic disease after postnatal overfeeding. However, little is known about the regulation of A-ring reductases by ω3 PUFAs until now, and the specific mechanisms involved in this process require additional study.
In conclusion, overfed neonatal rats had increased sensitivity to a high-fat diet from post-suckling to adulthood and presented early-onset hyperphagia, obesity, insulin resistance, and hyperlipidemia. By contrast, a ω3 PUFA-rich diet prevented programmed excess adipose accumulation and insulin resistance in early overfed rats. Tissue-specific regulation of GC-converting enzymes may underlie this process. This finding raises the possibility that dietary supplementation with ω3 PUFAs may provide a viable therapeutic option in humans for preventing and/or reducing adverse programming outcomes later in life.
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 work was supported by the Natural Science Foundation of China (8127304), Jiangsu Health Department of China (H2009) and the Scientific Research Foundation for the Returned Overseas Chinese Scholars, State Education Ministry (2009).
References
Ailhaud G, Guesnet P & Cunnane SC 2008 An emerging risk factor for obesity: does disequilibrium of polyunsaturated fatty acid metabolism contribute to excessive adipose tissue development? British Journal of Nutrition 100 461–470. (doi:10.1017/S0007114508911569)
Baudrand R, Dominguez JM, Carvajal CA, Riquelme A, Campino C, Macchiavello S, Bozinovic M, Morales M, Pizarro M & Solis N et al. 2011 Overexpression of hepatic 5α-reductase and 11β-hydroxysteroid dehydrogenase type 1 in visceral adipose tissue is associated with hyperinsulinemia in morbidly obese patients. Metabolism: Clinical and Experimental 60 1775–1780. (doi:10.1016/j.metabol.2011.05.001)
Belzung F, Raclot T & Groscolas R 1993 Fish oil n-3 fatty acids selectively limit the hypertrophy of abdominal fat depots in growing rats fed high-fat diets. American Journal of Physiology 264 R1111–R1118.
Berger J, Tanen M, Elbrecht A, Hermanowski-Vosatka A, Moller DE, Wright SD & Thieringer R 2001 Peroxisome proliferator-activated receptor-γ ligands inhibit adipocyte 11β -hydroxysteroid dehydrogenase type 1 expression and activity. Journal of Biological Chemistry 276 12629–12635. (doi:10.1074/jbc.M003592200)
Bermudez MDLG & Sinclair AJ 2009 Fatty acids and obesity. Current Pharmaceutical Design 15 4117–4125. (doi:10.2174/138161209789909674)
Boden G 2011 Obesity, insulin resistance and free fatty acids. Current Opinion in Endocrinology, Diabetes, and Obesity 18 139–143. (doi:10.1097/MED.0b013e3283444b09)
Bouchard C 2008 Gene–environment interactions in the etiology of obesity: defining the fundamentals. Obesity 16 (Suppl 3) S5–S10. (doi:10.1038/oby.2008.528)
Boullu-Ciocca S, Dutour A, Guillaume V, Achard V, Oliver C & Grino M 2005 Postnatal diet-induced obesity in rats upregulates systemic and adipose tissue glucocorticoid metabolism during development and in adulthood: its relationship with the metabolic syndrome. Diabetes 54 197–203. (doi:10.2337/diabetes.54.1.197)
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)
Bringhenti I, Schultz A, Rachid T, Bomfim MA, Mandarim-de-Lacerda CA & Aguila MB 2011 An early fish oil-enriched diet reverses biochemical, liver and adipose tissue alterations in male offspring from maternal protein restriction in mice. Journal of Nutritional Biochemistry 22 1009–1014. (doi:10.1016/j.jnutbio.2010.08.013)
Buren J, Bergstrom SA, Loh E, Soderstrom I, Olsson T & Mattsson C 2007 Hippocampal 11β-hydroxysteroid dehydrogenase type 1 messenger ribonucleic acid expression has a diurnal variability that is lost in the obese Zucker rat. Endocrinology 148 2716–2722. (doi:10.1210/en.2006-0897)
Chen H, Simar D, Lambert K, Mercier J & Morris MJ 2008 Maternal and postnatal overnutrition differentially impact appetite regulators and fuel metabolism. Endocrinology 149 5348–5356. (doi:10.1210/en.2008-0582)
Chen H, Simar D & Morris MJ 2009 Hypothalamic neuroendocrine circuitry is programmed by maternal obesity: interaction with postnatal nutritional environment. PLoS ONE 4 e6259 doi:10.1371/journal.pone.0006259)
Donahue SM, Rifas-Shiman SL, Gold DR, Jouni ZE, Gillman MW & Oken E 2011 Prenatal fatty acid status and child adiposity at age 3 y: results from a US pregnancy cohort. American Journal of Clinical Nutrition 93 780–788. (doi:10.3945/ajcn.110.005801)
Drake AJ, Livingstone DE, Andrew R, Seckl JR, Morton NM & Walker BR 2005 Reduced adipose glucocorticoid reactivation and increased hepatic glucocorticoid clearance as an early adaptation to high-fat feeding in Wistar rats. Endocrinology 146 913–919. (doi:10.1210/en.2004-1063)
Gallou-Kabani C, Vige A, Gross MS, Boileau C, Rabes JP, Fruchart-Najib J, Jais JP & Junien C 2007 Resistance to high-fat diet in the female progeny of obese mice fed a control diet during the periconceptual, gestation, and lactation periods. American Journal of Physiology. Endocrinology and Metabolism 292 E1095–E1100. (doi:10.1152/ajpendo.00390.2006)
Hou M, Liu Y, Zhu L, Sun B, Guo M, Buren J & Li X 2011 Neonatal overfeeding induced by small litter rearing causes altered glucocorticoid metabolism in rats. PLoS ONE 6 e25726 doi:10.1371/journal.pone.0025726)
Kang JX 2003 The importance of ω-6/ω-3 fatty acid ratio in cell function. The gene transfer of ω-3 fatty acid desaturase. World Review of Nutrition and Dietetics 92 23–36.
Kim YJ & Park T 2008 Genes are differentially expressed in the epididymal fat of rats rendered obese by a high-fat diet. Nutrition Research 28 414–422. (doi:10.1016/j.nutres.2008.03.015)
Kozawa S, Honda A, Kajiwara N, Takemoto Y, Nagase T, Nikami H, Okano Y, Nakashima S & Shimozawa N 2011 Induction of peroxisomal lipid metabolism in mice fed a high-fat diet. Molecular Medicine Reports 4 1157–1162. (doi:10.3892/mmr.2011.560)
Lakshmi V & Monder C 1988 Purification and characterization of the corticosteroid 11β-dehydrogenase component of the rat liver 11β-hydroxysteroid dehydrogenase complex. Endocrinology 123 2390–2398. (doi:10.1210/endo-123-5-2390)
Lichtenstein AH & Schwab US 2000 Relationship of dietary fat to glucose metabolism. Atherosclerosis 150 227–243. (doi:10.1016/S0021-9150(99)00504-3)
Livingstone DE, Jones GC, Smith K, Jamieson PM, Andrew R, Kenyon CJ & Walker BR 2000 Understanding the role of glucocorticoids in obesity: tissue-specific alterations of corticosterone metabolism in obese Zucker rats. Endocrinology 141 560–563. (doi:10.1210/en.141.2.560)
Man TY, Michailidou Z, Gokcel A, Ramage L, Chapman KE, Kenyon CJ, Seckl JR & Morton NM 2011 Dietary manipulation reveals an unexpected inverse relationship between fat mass and adipose 11β-hydroxysteroid dehydrogenase type 1. American Journal of Physiology. Endocrinology and Metabolism 300 E1076–E1084. (doi:10.1152/ajpendo.00531.2010)
Masuzaki H, Paterson J, Shinyama H, Morton NM, Mullins JJ, Seckl JR & Flier JS 2001 A transgenic model of visceral obesity and the metabolic syndrome. Science 294 2166–2170. (doi:10.1126/science.1066285)
Matthews SG 2002 Early programming of the hypothalamo-pituitary–adrenal axis. Trends in Endocrinology and Metabolism 13 373–380. (doi:10.1016/S1043-2760(02)00690-2)
Metges CC 2009 Early nutrition and later obesity: animal models provide insights into mechanisms. Advances in Experimental Medicine and Biology 646 105–112. (doi:10.1007/978-1-4020-9173-5-11)
Minana-Solis MC & Escobar C 2007 Increased susceptibility to metabolic alterations in young adult females exposed to early malnutrition. International Journal of Biological Sciences 3 12–19. (doi:10.7150/ijbs.3.12)
Morton NM, Ramage L & Seckl JR 2004a Down-regulation of adipose 11β-hydroxysteroid dehydrogenase type 1 by high-fat feeding in mice: a potential adaptive mechanism counteracting metabolic disease. Endocrinology 145 2707–2712. (doi:10.1210/en.2003-1674)
Morton NM, Paterson JM, Masuzaki H, Holmes MC, Staels B, Fievet C, Walker BR, Flier JS, Mullins JJ & Seckl JR 2004b Novel adipose tissue-mediated resistance to diet-induced visceral obesity in 11β-hydroxysteroid dehydrogenase type 1-deficient mice. Diabetes 53 931–938. (doi:10.2337/diabetes.53.4.931)
Mullen KL, Pritchard J, Ritchie I, Snook LA, Chabowski A, Bonen A, Wright D & Dyck DJ 2009 Adiponectin resistance precedes the accumulation of skeletal muscle lipids and insulin resistance in high-fat-fed rats. American Journal of Physiology. Regulatory, Integrative and Comparative Physiology 296 R243–R251. (doi:10.1152/ajpregu.90774.2008)
Odermatt A & Nashev LG 2010 The glucocorticoid-activating enzyme 11β-hydroxysteroid dehydrogenase type 1 has broad substrate specificity: physiological and toxicological considerations. Journal of Steroid Biochemistry and Molecular Biology 119 1–13. (doi:10.1016/j.jsbmb.2010.01.007)
Ong KK & Loos RJ 2006 Rapid infancy weight gain and subsequent obesity: systematic reviews and hopeful suggestions. Acta Paediatrica 95 904–908. (doi:10.1080/08035250600719754)
Pfaffl MW 2001 A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Research 29 e45 doi:10.1093/nar/29.9.e45)
Plagemann A, Harder T, Brunn M, Harder A, Roepke K, Wittrock-Staar M, Ziska T, Schellong K, Rodekamp E & Melchior K et al. 2009 Hypothalamic proopiomelanocortin promoter methylation becomes altered by early overfeeding: an epigenetic model of obesity and the metabolic syndrome. Journal of Physiology 587 4963–4976. (doi:10.1113/jphysiol.2009.176156)
Prior LJ & Armitage JA 2009 Neonatal overfeeding leads to developmental programming of adult obesity: you are what you ate. Journal of Physiology 587 2419 doi:10.1113/jphysiol.2009.174045)
Reeves PG, Nielsen FH & Fahey GJ 1993 AIN-93 purified diets for laboratory rodents: final report of the American Institute of Nutrition ad hoc writing committee on the reformulation of the AIN-76A rodent diet. Journal of Nutrition 123 1939–1951.
Riccardi G, Giacco R & Rivellese AA 2004 Dietary fat, insulin sensitivity and the metabolic syndrome. Clinical Nutrition 23 447–456. (doi:10.1016/j.clnu.2004.02.006)
Rodrigues AL, de Moura EG, Passos MC, Dutra SC & Lisboa PC 2009 Postnatal early overnutrition changes the leptin signalling pathway in the hypothalamic–pituitary–thyroid axis of young and adult rats. Journal of Physiology 587 2647–2661. (doi:10.1113/jphysiol.2009.169045)
Ruzickova J, Rossmeisl M, Prazak T, Flachs P, Sponarova J, Veck M, Tvrzicka E, Bryhn M & Kopecky J 2004 ω-3 PUFA of marine origin limit diet-induced obesity in mice by reducing cellularity of adipose tissue. Lipids 39 1177–1185. (doi:10.1007/s11745-004-1345-9)
Shin E, Shim KS, Kong H, Lee S, Shin S, Kwon J, Jo TH, Park YI, Lee CK & Kim K 2011 Dietary aloe improves insulin sensitivity via the suppression of obesity-induced inflammation in obese mice. Immune Network 11 59–67. (doi:10.4110/in.2011.11.1.59)
Srivastava RA 2009 Fenofibrate ameliorates diabetic and dyslipidemic profiles in KKAy mice partly via down-regulation of 11β-HSD1, PEPCK and DGAT2. Comparison of PPARα, PPARγ, and liver x receptor agonists. European Journal of Pharmacology 607 258–263. (doi:10.1016/j.ejphar.2009.02.024)
Stimson RH, Johnstone AM, Homer NZ, Wake DJ, Morton NM, Andrew R, Lobley GE & Walker BR 2007 Dietary macronutrient content alters cortisol metabolism independently of body weight changes in obese men. Journal of Clinical Endocrinology and Metabolism 92 4480–4484. (doi:10.1210/jc.2007-0692)
Symonds ME, Sebert SP, Hyatt MA & Budge H 2009 Nutritional programming of the metabolic syndrome. Nature Reviews. Endocrinology 5 604–610. (doi:10.1038/nrendo.2009.195)
Tamashiro KL & Moran TH 2010 Perinatal environment and its influences on metabolic programming of offspring. Physiology & Behavior 100 560–566. (doi:10.1016/j.physbeh.2010.04.008)
Tierney AC, McMonagle J, Shaw DI, Gulseth HL, Helal O, Saris WH, Paniagua JA, Golabek-Leszczynska I, Defoort C & Williams CM et al. 2011 Effects of dietary fat modification on insulin sensitivity and on other risk factors of the metabolic syndrome – LIPGENE: a European randomized dietary intervention study. International Journal of Obesity 35 800–809. (doi:10.1038/ijo.2010.209)
Tomlinson JW, Finney J, Gay C, Hughes BA, Hughes SV & Stewart PM 2008 Impaired glucose tolerance and insulin resistance are associated with increased adipose 11β-hydroxysteroid dehydrogenase type 1 expression and elevated hepatic 5α-reductase activity. Diabetes 57 2652–2660. (doi:10.2337/db08-0495)
Tsilchorozidou T, Honour JW & Conway GS 2003 Altered cortisol metabolism in polycystic ovary syndrome: insulin enhances 5α-reduction but not the elevated adrenal steroid production rates. Journal of Clinical Endocrinology and Metabolism 88 5907–5913. (doi:10.1210/jc.2003-030240)
Vagnerova K, Loukotova J, Ergang P, Musilkova J, Miksik I & Pacha J 2011 Peroxisome proliferator-activated receptor-γ stimulates 11β-hydroxysteroid dehydrogenase type 1 in rat vascular smooth muscle cells. Steroids 76 577–581. (doi:10.1016/j.steroids.2011.02.008)
Vara PS, Jeya KS, Kumar PU, Qadri SS & Vajreswari A 2010 Dietary fatty acid composition alters 11β-hydroxysteroid dehydrogenase type 1 gene expression in rat retroperitoneal white adipose tissue. Lipids in Health and Disease 9 111 doi:10.1186/1476-511X-9-111)
Velkoska E, Cole TJ, Dean RG, Burrell LM & Morris MJ 2008 Early undernutrition leads to long-lasting reductions in body weight and adiposity whereas increased intake increases cardiac fibrosis in male rats. Journal of Nutrition 138 1622–1627.
Westerbacka J, Yki-Jarvinen H, Vehkavaara S, Hakkinen AM, Andrew R, Wake DJ, Seckl JR & Walker BR 2003 Body fat distribution and cortisol metabolism in healthy men: enhanced 5β-reductase and lower cortisol/cortisone metabolite ratios in men with fatty liver. Journal of Clinical Endocrinology and Metabolism 88 4924–4931. (doi:10.1210/jc.2003-030596)
Wyrwoll CS, Mark PJ, Mori TA, Puddey IB & Waddell BJ 2006 Prevention of programmed hyperleptinemia and hypertension by postnatal dietary ω-3 fatty acids. Endocrinology 147 599–606. (doi:10.1210/en.2005-0748)
