Low Neonatal Plasma n-6/n-3 PUFA Ratios Regulate Offspring Adipogenic Potential and Condition Adult Obesity Resistance

The possibility that nutrient composition early in life has permanent metabolic effects was recognized >40 years ago (1), a phenomenon later called developmental origins of metabolic disease (2,3). Strong evidence indicates the existence of long-term consequences of early-life nutrients on adult metabolic health, linking perinatal nutrients with predisposition for obesity (4–7). Obesity has doubled in children and quadrupled in adolescents over the past 30 years (8), coincident with a marked increase in maternal intake of refined vegetable oils containing high amounts of omega-6 (n-6) polyunsaturated fatty acids (PUFAs) (4). Infant n-6 PUFA exposures introduced through maternal diet during the perinatal window are believed to stimulate adipogenesis during pre- and postnatal development on the basis of findings in rodent models (4,9–11). In the U.S., n-6 PUFAs in human milk have been threefold greater since the 1950s, whereas n-3 PUFAs have remained constant, thus increasing the infant’s n-6/n-3 PUFA ratio exposure (9,10,12). Infant exposure to a high n-6/n-3 PUFA ratio during gestation and breastfeeding has been associated with increased pediatric adiposity out to 3 years of age (13). We found that the higher the n-6/n-3 PUFA ratio in human milk, the greater the infant adipose deposition by 4 months of age independent of maternal prepregnancy BMI (14). Accordingly, perinatal exposure to a high n-6/n-3 PUFA ratio has been postulated to contribute to the obesity epidemic in developed countries (4,10).

Obesity is defined by excessive adiposity, and regulation of adipose tissue expansion (ATE) occurs through proliferation of adipocyte precursor (AP) cells to increase adipocyte number (hyperplasia) and by adipocyte filling (hypertrophy) (15). In a rat model of maternal obesity and high-fat diet (HFD), cross-fostering techniques identified an offspring’s early-life exposure to maternal HFD-induced obesity as a critical window that conditions adipogenic potential through altered DNA methylation patterning (5,11). In mice, maternal pre- and postnatal consumption of an HFD containing excessive n-6 PUFAs increases the offspring’s n-6/n-3 PUFA ratio exposure and stimulates ATE pathways, resulting in greater offspring adiposity (12,16–18). Moreover, a maternal diet rich in n-6 relative to n-3 PUFAs conditions adipogenic potential in offspring across generations (19), which is believed to occur by regulating expression of master adipogenic transcription factors C/EBPa and peroxisome proliferator–activated receptor-γ (PPARγ) (5,20,21). However, identification of specific molecular targets of n-3 PUFAs, independent of maternal obesity and HFD feeding, is less well characterized.

By using an established transgenic model that overexpresses an n-3 fatty acid desaturase (fat-1) in combination with maternal HFD-induced obesity, we reported that lowering the endogenous maternal n-6/n-3 PUFA ratio reduces placental inflammation and protects adult wild-type (WT) offspring against excessive body fat accumulation and insulin resistance (22). In the current study, we control for confounding maternal variables (HFD and obesity) by comparing offspring born to WT (high n-6/n-3) and fat-1 (low n-6/n-3) mothers provided a standard amount of dietary fat to study outcomes during early-life development independent of maternal obesity or HFD. We tested the hypothesis that lowering n-6/n-3 PUFA ratios in offspring circulation during the preweaning window tempers adipogenesis at the molecular and cellular levels. Of note, a low early-life PUFA ratio imparts long-term metabolic benefit to the adult offspring.

Animal procedures were approved by the institutional animal care and use committee and housed at the University of Colorado Anschutz Campus vivarium. WT C57Bl/6J mice were purchased from The Jackson Laboratory (Bar Harbor, ME). Fat-1 mice were provided by J.X. Kang and genotyped according to his protocol (23). Heterozygous fat-1 females were bred with WT males, generating 50% WT offspring as previously described (22). The study design (Fig. 1) used WT (high n-6/n-3 PUFA ratio) and fat-1 (low n-6/n-3 PUFA ratio) mothers to test PUFA ratio exposures in offspring development. Female mice were bred at 10 weeks old. Dams were provided chow and underwent normal gestation and lactation. At postnatal day 14 (PND14), both dams and their native litters were assessed for body composition, and pups were sacrificed for adipose morphology, cellularity, gene expression, protein levels, and circulating hormone, glucose, and fatty acid composition. Only WT offspring were used for downstream assays, and litters were standardized to six to eight pups per dam. In study 2, long-term effects of early-life PUFA exposure were tested by provision of a western-style diet (high fat, high sucrose [HF/HS]) (Teklad TD.88137; Envigo RMS, Indianapolis, IN). WT offspring with high or low n-6/n-3 PUFA ratio exposure were weaned to chow until 17 weeks old, when adult offspring were assessed by indirect calorimetry during the lead-in period and challenge with HF/HS diet for 1 week. Animals were then maintained on an HF/HS diet for 3 more weeks (Fig. 1).

Contralateral subcutaneous fat pads from one pup were combined and processed for cellularity as described previously (24). Briefly, adipose tissue was enzymatically digested, isolated adipocytes were stained with 0.2% methylene blue for cell integrity, and samples were imaged by using a 0.01-mm stage micrometer. Eight to 12 fields of each sample were quantified by using AdCount software (Mayo Clinic, Rochester, MN) to obtain diameters and cell size frequency distributions.

Total lipids were extracted and fatty acid profiles quantified by gas chromatography–mass spectrometry as previously described (14,25). Data are expressed in micromoles of fatty acid per milliliter of milk or milligram of total plasma protein. The total n-6/n-3 PUFA ratio is the sum of n-6 divided by the sum of n-3 PUFAs; the arachidonic acid/docosahexaenoic acid + eicosapentaenoic acid (AA/DHA + EPA) ratio is the micromoles of 20:4n-6 divided by 22:6n-3 + 20:5n-3; the linoleic/α-linoleic (LA/LNA ratio) is the micromoles of 18:2n-6 (LA) divided by 18:3n-3 (LNA).

PND14 pups were anesthetized by isofluorane and decapitated, and trunk blood was collected in K₂EDTA tubes (BD Microtainer, Franklin Lakes, NJ). Samples were stored on wet ice for 30 min and then centrifuged at 2,000g for 20 min at 4°C. Insulin (#90080; Crystal Chem, Downers Grove, IL), leptin (#22-LEPMS-E01; ALPCO, Salem, NH), and adiponectin (#47-ADPMS-E01; ALPCO) were processed according to manufacturer protocols.

Blood glucose was measured in fasted adults (4 h) by using a Contour blood glucose meter (Ascensia Diabetes Care, Parsippany, NJ) after oral gavage of 50% dextrose (Vet One, Boise, ID) as previously described (26). Pup blood glucose was measured from trunk blood immediately after decapitation.

Total RNA was isolated by using the RNAqueous-Micro Total RNA Isolation Kit (Thermo Fisher Scientific, Waltham, MA), including DNAse digestion, according to the manufacturer’s protocol. Two hundred fifty nanograms of total RNA was used in the Whole-Transcript Expression RNA kit (Life Technologies, Carlsbad, CA), and samples were hybridized to mouse 1.1 ST Gene Arrays (Affymetrix, Santa Clara, CA), washed, stained, and imaged with the Affymetrix GeneAtlas Personal Microarray System. Raw data were GC-RMA (robust multiarray averaging) normalized, log2 transformed, and analyzed by using the Partek Genomic Suite as previously described (27). Quantitative RT-PCR was performed by using mRNA copy number and normalized to PolR2b copies with TaqMan gene expression assays (Thermo Fisher Scientific) as previously described (28).

Subcutaneous white adipose tissue (sWAT) was homogenized in 50 mmol/L Tris (pH 7.4), 150 mmol/L NaCl, 2 mmol/L EDTA, 50 mmol/L NaF, 1% Triton X-100, 1% sodium deoxycholate, 0.1% SDS, and 5 mmol/L sodium vanadate supplemented with Halt Protease and Phosphatase Inhibitor cocktail (#1861281; Thermo Fisher Scientific). Total protein concentration was determined by bicinchoninic acid analysis (Pierce BCA Protein Assay; Thermo Fisher Scientific, Rockford, IL). Immunoblots were run on the Western capillary electrophoresis system, and data were analyzed by Compass software (ProteinSimple, San Jose, CA) as in Checkley et al. (29). Primary antibodies included PPARγ (#2435S; Cell Signaling, Danvers, CT), fatty acid synthase (FASN) (#3180S; Cell Signaling), and perilipin 1 (PLIN1) (#70R-1297; Fitzgerald Industries International, Acton, MA) and were normalized to cyclophilin A (CypA) (#2175; Cell Signaling).

Total DNA was isolated by using the AllPrep DNA/RNA Mini Kit (QIAGEN, Germantown, MD) according to the manufacturer’s protocol, and DNA was quantified by using the Qubit Fluorometric Quantitation assay (Thermo Fisher Scientific). Bisulfite-converted DNA was PCR amplified by using primers designed in the PyroMark Assay Design Software (QIAGEN) as in Yang et al. (30). Pyrosequencing was performed on the PyroMark Q96 MD (QIAGEN), with reagents and protocols supplied by the manufacturer. Each plate contained 0%, 50%, and 100% methylated controls. Percent methylation was calculated from the peak heights of C and T by Pyro Q-CpG software (QIAGEN) in duplicate for each sample, and averaged values were used in the statistical analysis (30).

Whole-body composition was quantified by magnetic resonance (EchoMRI; Echo Medical Systems, Houston, TX) on PND14 for dams and litters or for adult offspring before, during, and after calorimetry. Energy balance of adults was conducted at 17 weeks old as previously described (24,31). Mice were housed individually at 27°C in an eight-chamber system (Oxymax Comprehensive Lab Animal Monitoring System; Columbus Instruments, Columbus, OH) with a 14-h light, 10-h dark cycle. Mice were acclimated for 1 week on chow and then challenged with an HF/HS diet for 5 days. Food and water intake were measured during the entire time while in calorimetry. VO₂, VCO₂, respiratory quotient, activity measurements, and energy expenditure were calculated as previously described (31). After calorimetry, mice were returned to individual housing for an additional 2 weeks and then sacrificed, and samples were collected.

PND14 WT offspring from litters of four to five independent dams per genotype were used in analyses. One-way or two-way ANOVA followed by Bonferroni multiple comparisons test was performed in GraphPad Prism version 7.0a for Macintosh software (GraphPad Software, La Jolla, CA). For microarray analysis, one-way ANOVA with a Bonferroni false discovery rate of 0.1 was performed by using Partek Genomics Suite version 6.6 software (Partek, St. Louis, MO) followed by Bonferroni multiple comparisons test.

When provided a low-fat diet, maternal fat-1 transgene expression did not influence body composition of either dams or their biological litters, and differences were not observed in the macronutrient composition of the milk (data not shown). Maternal fat-1 expression reduced the n-6/n-3 long-chain (LC) PUFA and AA/DHA + EPA ratios in milk (Fig. 2A) without affecting quantitative amounts (in micromoles per milliliter) of saturated fatty acids (SFAs), monounsaturated fatty acids (MUFAs), PUFAs, or the ratio of essential dietary fatty acids (LA [18:2n-6]/LNA [18:3n-3]). The plasma of PND14 pups qualitatively mimicked the composition of the milk with decreased LC-PUFA and AA/DHA + EPA ratios (Fig. 2B). Despite maternal provision of the same diet, WT offspring exposed to fat-1 mother’s milk had significantly reduced plasma amounts (in micromoles per milligrams of protein) of 20:2n-6 and 20:4n-6 (P ≤ 0.01), with enriched amounts of 22:6n-3 (P = 0.038) (Fig. 2B).

In PND14 pups exposed to low n-6/n-3 PUFA ratios, adipocyte cellularity from digested sWAT revealed small-sized adipocytes (10–20 μmol/L) had a significantly greater percentage of total adipocytes, whereas the percentage of medium-sized adipocytes (30–60 μmol/L) was significantly lower (Fig. 3A). Exposure to low n-6/n-3 PUFA ratios yielded 1.48-fold more total adipocytes per gram of sWAT (P = 0.01), and sWAT depots were 1.33-fold heavier (Fig. 3B). By PND14, total adiponectin was significantly greater in the low n-6/n-3 PUFA ratio pups (P < 0.001) (Fig. 3C), driven by the low- (trimer) and medium- (hexamer) molecular-weight forms (total/high-molecular-weight adiponectin ratio P < 0.001). Other systemic factors, including blood glucose, insulin, and leptin, were unchanged (Fig. 3C). Together, these data indicate that low n-6/n-3 PUFA ratio perinatal exposures establish an early adipose cellularity associated with increased insulin sensitivity, producing more adiponectin.

Microarray gene expression identified multiple differentially expressed pathways in sWAT in PND14 pups with low plasma n-6/n-3 PUFA ratios (Fig. 4A). Several lipid metabolism genes were downregulated in pups with low plasma n-6/n-3 ratios, including actyl-CoA carboxylase, ATP citrate lyase, fatty acid transporter (CD36), elongase 5, fatty acid desaturase 1, FASN, lipoprotein lipase (LPL), malic enzyme, prostaglandin E synthase, prostaglandin reductase, stearoyl-CoA desaturase 2, and SREBP1. Conversely, acyl-CoA thioesterase 12, acyl-CoA synthetase medium-chain family member 4, fatty acid desaturase 6, lipoyltransferase 1, phospholipase A2, and retinol dehydrogenase 7 were upregulated in the low plasma n-6/n-3 ratio offspring. Signal transduction genes were downregulated in the low plasma n-6/n-3 ratio pups, including the adiponectin receptor 2, frizzled family members (Fzd-1, -5, -7), leptin receptor, LDL receptor–related protein 8 and 10, lysophosphatidic acid receptor 4, prostaglandin F2-α receptor, tumor necrosis factor (TNF) receptor superfamily member 11b, TNF receptor–associated factor 7, and TNF receptor–associated protein 1, as well as secreted ligands angiopoietin-related protein 1, C-C motif chemokines (2, 27a, 27b, and 7), and Wnt10a. Conversely, IGF-I and Wnt7b were both induced in low plasma n-6/n-3 PUFA pups (Fig. 4A). Regulatory and adipogenic markers decreased by low n-6/n-3 PUFA ratios were PPARγ, PPARγ coactivator 1 and estrogen-related receptor–induced regulator in muscle protein 1 (Perm1), Berardinelli-Seip congenital lipodystrophy gene (Bscl2, Seipin), fatty acid binding protein 4 (Fabp4), and Plin1. Significantly regulated genes are presented in Supplementary Table 1. Together, expression data suggest that low plasma n-6/n-3 ratio conditions during offspring adipogenesis exerts broad control over lipid uptake and metabolism, surface receptors, chemokine levels, protein hormones, and adipocyte regulatory factors.

Principal adipogenic transcription factors and downstream target levels were verified by quantitative RT-PCR by using an independent set of PND14 offspring (Fig. 4B). Pparγ2 was significantly decreased (P = 0.024) in sWAT of low n-6/n-3 PUFA ratio offspring, but mRNA level of C/ebpα was unchanged. In addition, Pparγ2 transcriptional targets were either significantly decreased (Fabp4, Fasn, Plin1, and AdipoQ) (P ≤ 0.04) or tended to be decreased (Cidea, Lpl, and leptin) (P ≤ 0.075) (Fig. 4B). The mRNA level for the Bscl2 gene, which regulates lipid droplet formation and size in adipocytes (32,33), was significantly decreased by low n-6/n-3 PUFA ratio exposure (Fig. 4B). Reduction of PPARγ2 mRNA was consistent with its decreased protein levels (P = 0.007) and for PPARγ2 targets FASN and PLIN1 (P = 0.04 and 0.01, respectively) (Fig. 4C). Consistent with the decreased mRNA and protein levels, DNA methylation of the PPARγ2 proximal promoter at −3,195 and −322 upstream of the transcriptional start site was increased by 6.4% and 4.5%, respectively (P < 0.01) (Fig. 4D).

The long-term effects of early-life PUFA exposures were tested when adult offspring were challenged with an HF/HS diet. Between perinatal exposure groups, body fat percentage, lean mass, and body weight of adult offspring at 17, 18, and 19 weeks were not significantly different (Fig. 5A); however, the low n-6/n-3 PUFA ratio group had one-half the body fat relative to the high n-6/n-3 PUFA ratio group by week 22. Of note, the lean body mass was not affected after 4 weeks of HF/HS diet, so the significantly increased body weight (P ≤ 0.001) was due to greater adiposity. Metabolic phenotyping revealed a lower energy balance by −1.04 kcal/day during the lead-in period (P = 0.001) in adults from the low plasma n-6/n-3 PUFA perinatal exposure group, and these mice exhibited an attenuation of the positive energy imbalance in response to the HF/HS diet (Fig. 5B). Energy intake was significantly decreased in the low perinatal n-6/n-3 ratio group during the lead-in period (P = 0.046) as well as at days 2 (P = 0.003) and 5 (P = 0.04) of the HF/HS diet. Although the total energy expenditure was significantly greater in the lead-in period (P = 0.006), it did not differ significantly during the HF/HS diet challenge. The non–resting energy expenditure was significantly increased during the lead-in period and on day 3 of the HF/HS diet relative to the high perinatal n-6/n-3 ratio group (P < 0.05). The respiratory exchange ratio was significantly lower in the low n-6/n-3 ratio group during the lead-in period (P = 0.002) as well as at days 1 (P = 0.05) and 3 (P = 0.02). After 4 weeks of HF/HS diet feeding, adult offspring in the low perinatal n-6/n-3 ratio group had significantly lower fasting glucose levels and reduced response during an oral glucose tolerance test (area under the curve P = 0.003) (Fig. 5C) indicative of a healthier metabolic phenotype.

The novel observation of this study is that enriching n-3 and simultaneously reducing n-6 LC-PUFA exposure during perinatal development establishes epigenetic, gene expression, and morphological changes in adipose tissue as well as increases circulating adiponectin in 14-day-old offspring. Of note, these early-life responses to the low n-6/n-3 PUFA ratio were independent of confounding factors potentially introduced by maternal obesity or the mother’s high-fat feeding. Consequently, the low plasma n-6/n-3 ratio and high circulating adiponectin levels early in life, without significant differences in blood insulin, glucose, or leptin levels, set up a persistent metabolic phenotype that ameliorates the predisposition to develop obesity later in life. Early-life exposure to a low perinatal n-6/n-3 PUFA ratio resulted in an adipose phenotype characterized by a greater number of total adipocytes that were smaller in size, a broad-based reduction in adipogenic gene expression, and hypermethylation of two regions in the proximal promoter of PPARγ2. Cumulatively, the current observations indicate that perinatal n-6/n-3 exposure affects fundamental molecular and cellular characteristics of sWAT in mice, and these effects on adipose tissue development may have functional consequences for metabolic disease predisposition later in life.

Emerging evidence links perinatal fatty acid exposure with DNA methylation changes in liver, skeletal muscle, and adipose tissue (11,34–38). Consistent with these reports, the current findings of increased PPARγ2 proximal promoter DNA methylation, in combination with suppressed lipogenic and adipocyte markers, support the hypothesis that low n-6/n-3 PUFA ratios modulate perinatal adipogenesis epigenetically independent of maternal obesity. Others found that MUFA (trans 18:1n-9) provided in the maternal diet is associated with DNA hypermethylation in offspring adipose tissue, although the exact genes were not identified (39), and that in vitro treatment of cells with n-6 PUFA causes dose-dependent DNA methylation patterns consistent with those observed in obese individuals and those with diabetes (40). In an elegant maternal DIO study in rats, Borengasser et al. (5) identified comprehensive DNA methylation modifications in adipogenic genes present in the white adipose tissue of cross-fostered offspring, which enhanced in vitro differentiation of AP cells present in the stromal vascular fraction. Together, enriching n-3 while decreasing n-6 LC-PUFA throughout gestation and lactation could regulate a common genetic program in AP cells to establish important epigenetic modifications. Future investigations should determine how DNA methylation, particularly in AP cells, is changed during the reduction of n-6 PUFAs (20:2 and 20:4) or enrichment of n-3 PUFAs (22:6) or whether the balance of n-6/n-3 PUFAs is most critical in epigenetic control of ATE potential.

That ATE occurs both by increasing total adipocyte number (proliferation) and increasing mature adipocyte size (hypertrophy) is well established (15,41). In Zucker rats, LC-MUFA and LC-PUFA favor adipose hyperplasia, whereas LC-SFA favors adipose hypertrophy, indicating different saturation affects of adipocyte cellularity in vivo (42). In PND14 pup plasma, we observed no significant difference in absolute amounts of SFAs, MUFAs, or PUFAs (data not shown); however, offspring with a low plasma n-6/n-3 PUFA ratio had more small adipocytes in their sWAT, a morphology consistent with the regulation of AP cell proliferation and/or differentiation (43). We identified significantly increased 20:2n-6 and 20:4n-6 in PND14 high n-6/n-3 PUFA ratio plasma. Arachidonic acid (20:4n-6) and its downstream biosynthetic product prostacyclin promote preadipocyte proliferation by stimulating G-coupled protein receptor PTGIR signaling in a paracrine loop and by driving differentiation by activating PPARs (9,12,16). Conversely, decreasing endogenous n-6/n-3 PUFA ratios by transient fat-1 expression in 3T3-L1 cells inhibits preadipocyte proliferation, suppresses PPARγ and FABP4, and inhibits lipid accumulation (44) consistent with what we observed in vivo. In the mouse, HFD initiates extensive in vivo proliferation of CD24⁺ adipocyte progenitors, which then lose CD24 expression and are rapidly committed to differentiate into lipid-laden mature adipocytes (15,43). With respect to early-life adipogenesis, the current findings support that low perinatal n-6/n-3 PUFA ratios might act 1) on the CD24⁺ AP cells to regulate proliferation and adipocyte number or 2) on the CD24⁻ preadipocytes to regulate differentiation genes. Consequently, the early-life n-6/n-3 ratio may regulate AP cells, establishing a sWAT morphology capable of synthesizing greater adiponectin and ultimately influencing adult appetite regulation and metabolic fuel preference during an obesogenic diet.

The reduction of early-life n-6/n-3 PUFA ratio exposure through maternal fat-1 expression appears to have persistent, functional consequences because these adult offspring have proven to be less susceptible to the development of obesity in response to HF/HS feeding. The attenuated intake, enhanced metabolic requirements, and lower respiratory exchange ratio in the perinatal low n-6/n-3 ratio offspring generally is consistent with the diet-resistant phenotype, which remains lean in response to this obesogenic diet challenge (45). Diet-resistant rodents are able to sense nutrient excess, tend to burn more dietary fat, and respond by adjusting food intake and increasing energy expenditure accordingly. Although speculating that the altered predisposition to develop obesity in the face of this HF/HS challenge results from the molecular and cellular characteristics observed in adipose tissue is tempting, other key tissues involved in energy homeostasis (hypothalamus, muscle, liver, etc.) likely are affected in a concordant manner to affect appetite, satiety, nutrient trafficking, and/or metabolic regulation. The combined milieu of PUFA and high adiponectin circulation in the low n-6/n-3 ratio pups could mediate programming of peripheral tissues. For example, no changes in beige or brown adipocyte regulators (Ucp1, Prdm16, or Zfp423 [46]) were observed in the sWAT of pups, so skeletal muscle or alternative adipose depots could have increased lipid oxidation to account for the elevated energy expenditure in adults. In addition, reduction of the n-6/n-3 ratio has been shown to enhance beneficial gut bacteria and reduce low-grade inflammation and metabolic disease in adult mice (47), providing another possible long-term effector that could be manipulated early in life by low n-6/n-3 ratio exposure. Together, metabolic phenotyping data suggest that an early-life low n-6/n-3 ratio programs adult diet resistance by suppressing calorie intake and potentially diverting fat from storage to oxidative pathways, thereby preventing excessive adipose accumulation. In any case, more studies are needed to place the contribution of early-life adipose tissue programming into the context of the numerous other critical nodes of homeostatic regulation, which collectively explain the phenotype we observed in adult offspring.

This mouse model is potentially limited by the timing of adipogenesis in rodents versus humans. In human infants, adipose tissue develops progressively by increasing both adipocyte size and number until gestational week 23, when in the weeks following, adipose growth is almost exclusively accomplished through hypertrophy (48). Mice develop adipose tissue postnatally (49), but despite any temporal differences between infants and mouse pups, the mechanisms underlying adipogenesis are believed to be equivalent (15,49). Moreover, human offspring are likely not to be weaned onto a singular dietary source as in our controlled model where weanlings were maintained on chow until challenge with an HF/HS diet. Rather, human children likely have access to potentially obesogenic foods earlier than in adulthood. Evaluation of an obesogenic diet provided at weaning and during adolescence in the mouse is the focus of future studies.

Our observations implicate enriched n-3 and reduced n-6 PUFAs in the control of a genetic program that regulates neonatal ATE in mice. With regard to the human mother/infant dyad, these findings strengthen support for enhanced maternal dietary intake of n-3 PUFAs in combination with a more measured intake of n-6 PUFAs to lower the perinatal n-6/n-3 PUFA ratio. The possibility that maternal-supplied PUFAs conditions early-life adipogenic potential and appetite regulation in offspring, protecting against childhood obesity, opens an opportunity for early intervention to reduce the incidence of obesity in future generations.

Acknowledgments. Instruments were provided by the Mass Spectrometry Lipidomics Core Facility of the University of Colorado School of Medicine.

Funding. M.C.R. is supported by National Institute of Diabetes and Digestive and Kidney Diseases (NIDDK) grant K01-DK-109079, an Office of Research on Women’s Health Building Interdisciplinary Research Careers in Women’s Health Scholar Award (K12-HD-057022), and a Eunice Kennedy Shriver National Institute of Child Health and Human Development (NICHD) training grant (T32-HD-007186). M.C.R. and J.E.F. are supported by the Colorado NIDDK-Nutrition and Obesity Research Center (P30-DK-048520). J.E.F. is supported by NIDDK grant R24-DK-090964. P.S.M. is supported by NICHD grants P01-HD-038129 and R01-HD-075285.

Duality of Interest. No potential conflicts of interest relevant to this article were reported.

Author Contributions. M.C.R. designed the experiments, generated and analyzed the data, and wrote the manuscript. M.R.J., D.M.P., J.A.H., P.G.W., G.C.J., T.K.S., and B.A.d.l.H. generated the data and reviewed and edited the manuscript. I.V.Y. assisted with DNA methylation/gene expression studies and reviewed and edited the manuscript. J.E.F. and P.S.M. designed the experiments, contributed to the discussion, and reviewed and edited the manuscript. M.C.R., J.E.F., and P.S.M. are the guarantors of this work and, as such, had full access to all the data in the study and take responsibility for the integrity of the data and the accuracy of the data analysis.