Palmitic acid, regardless of obesity, impairs leptin and insulin’s ability to regulate food intake and body weight and decreases activation of PI3K. In agreement with previous reports (10, 29, 30), we first confirmed that maintenance on a high-fat–saturated (HFS) diet reduces hypothalamic insulin and leptin sensitivity. Rats fed a diet high in saturated fat (40% of calories from HFS diet) or a diet high in oleic acid (40% of calories from oleic acid diet) for 3 months weighed more and had more body fat (Table 1) than rats fed a matched low-fat control diet (for nutrient information, see ref. 31 and Table 2). The fatty acid composition of the HFS diet was 10% myristate/27% palmitate/12% stearate/25% oleate. The composition of the oleic acid diet was 11% palmitate/ 70% oleate/13% linoleate. It is important to note the HFS contains a high amount of saturated fatty acids, but it also contains unsaturated fatty acids, including oleic acid. The critical variable we are testing is the role of palmitate, and the HFS diet has substantially more palmitate than the oleic acid–enriched diet. The HFS diet–fed rats have an attenuated anorexic response to third-ventricular (i.c.v.) insulin (Figure 1A) compared with the animals maintained on the low-fat or the high-fat oleic acid diets. To assess whether the failure of insulin to reduce body weight in HFS diet–fed rats was associated with reduced insulin-induced phosphorylation of AKT (p-AKT), insulin was infused i.c.v. and rats were sacrificed 10 minutes later. Insulin-induced p-AKT was assessed in medial basal hypothalamic tissue (the brain was rapidly removed, snap frozen, and a ~70-mg wedge of hypothalamus was excised caudal to the mammillary bodies, rostral to the optic chiasm, lateral to the optic tract, and superior to the apex of the hypothalamic third ventricle). Animals maintained on the HFS diet have reduced insulin-induced p-AKT relative to those on the low-fat and the oleic acid diets (Figure 1, B and C), implying that maintenance on a saturated fatty acid–diet, which is high in palmitic acid, but not a diet high in oleic acid, attenuates insulin activity and induces insulin resistance.

Figure 1 Diets high in saturated fats impair insulin- and leptin-induced anorexia and hypothalamic insulin-signaling. Rats (n = 8–10/group) were maintained on a low-fat diet (low fat), a HFS diet, or diet high in oleic acid (Oleic; see Table 2 for nutrient composition). Rats received i.c.v. infusions of 8 mU insulin or saline and food intake was measured at 24 hours. (A) Food intake after drug and saline (vehicle) control injections (*P < 0.05 compared with saline intakes). Rats maintained on low-fat, oleic acid, or HFS diet received an i.c.v. infusion of 8 mU insulin or saline and were sacrificed after 10 minutes. (B) Representative Western blots for p-AKT and GAPDH. (C) Densitometry analysis for all rats. ANOVA and post-hoc tests confirmed that rats maintained on HFS diet had lower p-AKT (P < 0.05 compared with other groups) and that only rats consuming low-fat and oleic acid diets exhibited insulin-induced p-AKT (*P < 0.05 compared with saline). (D) A separate group of rats were maintained on the same diets and received 10 μg i.c.v. leptin or saline (*P < 0.05 compared with saline). (E) Animals on the HFS-R diet (Table 1) were insulin resistant relative to the low-fat–fed animals, implying that the HFS diet, rather than increased body fat per se, is sufficient to reduce central insulin sensitivity. ANOVA and post-hoc tests confirmed only rats maintained on low-fat diet exhibited insulin-induced anorexia (data are mean ± SEM; *P < 0.05 compared with saline injection).

Table 1 Characteristics of rats

Table 2 Composition (g/kg of diet) of the low-fat, HFS, and oleic acid diets

Leptin (3) and insulin (4) exert their catabolic action through PI3K, and we found reductions in leptin’s ability to reduce food intake in HFS diet–fed versus low-fat– or the oleic acid–fed rats (Figure 1D). Additionally, we found that consumption of the HFS diet reduced leptin-induced p-STAT3 immunoreactivity, a marker of leptin receptor activation (32, 33) (data not shown). These data imply that the HFS diet (with elevated palmitic levels), impairs both signaling pathways.

To confirm that the observed insulin resistance is the result of dietary fatty acids, rather than obesity per se, we performed an additional study in which 1 group of rats had restricted access to the HFS diet and were not allowed to become obese, relative to controls. These animals were given the same number of calories consumed by the low-fat diet–fed control animals (HFS-restricted diet [HFS-R diet]). HFS-R diet–fed rats developed insulin resistance relative to the low-fat diet–fed controls (Figure 1E), implying that maintenance on the high-saturated-fat diet, rather than increased body fat per se (Table 1), is sufficient to reduce central insulin sensitivity and confirming our recent results.

Palmitic acid inhibits CNS insulin regulation of hepatic glucose production. Insulin regulates glucose use throughout the body, including the brain (1, 34). To test the hypothesis that a component of saturated fatty acids diets, specifically palmitic acid, results in impaired insulin regulation of hepatic glucose production, we administered low doses of palmitic acid, oleic acid, or vehicle directly into the brain, followed by an intracarotid infusion of insulin. Intracarotid insulin infusions mimic the route by which endogenous insulin physiologically accesses the brain and act at central sites without altering plasma insulin, glucose, or fatty acid levels (35). i.c.v. infusions of palmitic acid, but not oleic acid or vehicle alone, attenuate insulin-induced suppression of hepatic glucose production, as demonstrated by a euglycemic clamp in awake, unrestrained rats maintained on the low-fat diet (Figure 2). Importantly, the central infusions of fatty acids did not significantly change peripheral FFA concentrations (vehicle, 384 ± 30 μmol/l; palmitic acid, 419 ± 39 μmol/l; oleic acid, 392 ± 51 μmol/l), implying that changes in palmitic acid concentrations in the CNS have profound effects on CNS actions and do not increase circulating FFA.

Figure 2 i.c.v. palmitic acid (but not oleic acid) attenuates insulin-induced suppression of hepatic glucose production. Rats (6–8/group) received a 3-day i.c.v. infusion of palmitic acid, oleic acid, or control (cont) (PBS), before a carotid-artery insulin infusion under euglycemic clamp conditions. Comparable glucose use rates (~100 mg/dl) were achieved in all groups, beginning 90 minutes after the start of the experiment. All groups had plasma insulin values of approximately 900 pmol/l (data not shown). Carotid insulin infusions reduced hepatic glucose production in control and oleic acid–infused rats but not in rats infused with palmitic acid (data are mean ± SEM; ***P < 0.001 compared with control infusions).

Palmitic acid inhibits CNS insulin signaling in vitro. To further test the specific effects of palmitic acid on insulin signaling, we exposed hypothalamic neuronal cells in culture to palmitic acid or oleic acid, using a cell line that expresses both leptin and IRs (36) and is responsive to both leptin and insulin (data not shown). Similar to our in vivo findings, palmitic acid, but not oleic acid, attenuated insulin-induced p-AKT (Figure 3, A and B) in vitro. Because fatty acids can be toxic to cells in culture, we also assessed the toxicity of the concentration of palmitic acid used in our experiments, with the Trypan blue exclusion test of cell viability (Figure 3, C and D). Following exposure to 100 μmol/l palmitic acid, cells were normal and viable (Figure 3C; as used in Figure 3, A and B), whereas toxicity and cell death was demonstrated following exposure to a 300 μmol/l dose (Figure 3D). Additionally, we found, following exposure to 100 μmol/l palmitic acid, levels of pJNK or other inflammatory intermediates did not change. These data are consistent with our hypothesis that palmitic acid directly impairs insulin-induced p-AKT, rather than exerting nonspecific toxic effects.

Figure 3 Palmitic acid attenuates insulin signaling in vitro. Hypothalamic IVB cells (36) were exposed to 100 μmol/l palmitic acid, 100 μmol/l oleic acid, or vehicle (10% fatty acid–free BSA in PBS) for 4 hours and then tested with insulin (50 ng/ml in PBS) or vehicle (PBS) for 10 minutes. (A) Quantified data (mean ± SEM) from 2 separate Western blot analyses for p-AKT, using GAPDH as a control (n = 4–6/treatment). Insulin increased p-AKT relative to vehicle (*P < 0.05 compared with vehicle), though this increase was significantly reduced by palmitic acid infusion (P < 0.05). (B) A representative Western blot is shown (the top bands depict p-AKT, vehicle, vehicle/insulin, palmitic acid, and palmitic acid/insulin lanes were run on the same gel but were noncontiguous, and oleic acid and oleic acid/insulin were run on a separate gel but under the same conditions; and the bottom band depicts GAPDH, lanes were run on a different gel). To test for the viability of the cells following exposure to 100 μmol/l palmitic acid, we used the Trypan blue exclusion test of cell viability. (C) Cells exposed to 100 μmol/l palmitic acid. (D) Cells exposed to 300 μmol/l palmitic acid. Viable cells have a clear cytoplasm, whereas nonviable cells have a blue cytoplasm. Original magnification, ×10 (C and D).

Palmitic acid increases CNS diacylglycerol levels. Our working hypothesis is that fatty acids increase intracellular diacylglycerol (DAG), which facilitates the translocation of PKC-θ and reduces in insulin activity. We assessed CNS DAG levels following palmitic and oleic acid exposure, both in vitro and in vivo. Here, we report animals exposed to the HFS diet have significantly (P < 0.05) elevated DAG levels relative to the oleic acid– and low-fat–fed animals (low fat, 549.2 ± 22.5 pmol/mg; HFS, 863.7 ± 121.1 pmol/mg; oleic acid, 599.1 ± 45.5 pmol/mg). Additionally, following the gavage of the fatty acids, we again found significantly (P < 0.05) elevated DAG levels in palmitate acid–gavaged animals relative to the oleic acid–gavaged or control gavaged animals (control, 320 ± 25.6 pmol/mg; palmitate, 449 ± 28.9 pmol/mg; oleic acid, 338 ± 49.7 pmol/mg). In vitro, we observed that cells exposed to palmitic acid exhibited a significant (P < 0.05) increase in DAG levels, which did not occur in the cells exposed to oleic acid (control, 1.0 ± 0.04 pmol/mg; palmitate, 3.2 ± 0.9 pmol/mg; oleic acid, 1.1 ± 0.05 pmol/mg). Therefore, exposure to diets elevated in palmitic acid results in increased CNS DAG levels, providing a mechanism by which PKC-θ translocation is activated.

PKC-θ is expressed in hypothalamic nuclei critical for hepatic glucose production and body weight regulation. In order to determine the mechanism by which fatty acids were inhibiting insulin signaling, we next sought to determine whether PKC-θ is expressed in the brain and whether it is a critical mediator in reducing hypothalamic insulin sensitivity. We designed primers against rat Prkcq mRNA (GenBank accession number AB020614) and performed RT-PCR on muscle and hypothalamic tissue from untreated control rats. The resulting product sequence was identical to both rat and human PKC-θ with only 10% (nonprimer) homology to the reported PKC-δ sequences (see Methods). Consistent with a recent report by Dewing et al. (37), PKC-θ was expressed in the medial basal hypothalamus and muscle of rats (Figure 4A). To determine which regions within the hypothalamus express PKC-θ, we performed immunocytochemistry for PKC-θ and found PKC-θ–like immunoreactivity in discrete neuronal populations in the arcuate nucleus (Figure 4, B and C). We confirmed the specificity of our antibody by comparing PKC-θ labeling in WT mice and mice with a targeted deletion of the Prkcq gene (Prkcq–/–; Figure 4, D and E). These results demonstrate that the PKC-θ labeling in Figure 4, B and C, is highly specific. To assess the phenotype of hypothalamic cells expressing PKC-θ, we used 2 different transgenic mouse lines that express GFP either in neuropeptide Y (NPY) or pro-opiomelanocortin (POMC) neurons. We performed dual immunocytochemistry to colocalize PKC-θ expression with POMC (Figure 4, F–H) and NPY (Figure 4, I–K). There was 20% colocalization of PKC-θ with NPY (Figure 4K), and no colocalization with POMC (Figure 4H). Additionally, as demonstrated in Figure 4, L and M, PKC-θ immunoreactivity colocalized with NeuN, a marker of neuronal cells, but not the glial-cell marker GFAP. To assess whether PKC-θ is expressed in neurons that respond to leptin, we used a transgenic line that expresses GFP only in cells that also express the leptin receptor (38). We identified cells in the arcuate nucleus that had 22% colocalization for leptin receptor and PKC-θ (Figure 4, N and O). Together, our data demonstrate for the first time to our knowledge that PKC-θ is located in arcuate NPY and leptin-responsive neurons, providing evidence that PKC-θ is critically positioned to be responsive to fatty acids and to modulate hepatic glucose production and body weight regulation.

Figure 4 PKC-θ is expressed in the hypothalamus. (A) PKC-θ expression, using RT-PCR in the hypothalamus and muscle of rats (con, water control). To determine regions within in the hypothalamus that express PKC-θ, we assessed PKC-θ–like immunoreactivity in cells by DAB and fluorescent immunocytochemistry. (B and C) Hypothalamic PKC-θ–like immunoreactivity in the arcuate assessed by fluorescent and DAB immunocytochemistry, respectively. (D and E) The specificity of the PKC-θ antibody is confirmed by comparing PKC-θ labeling in the hypothalamus of WT mice and PKC-θ–knockout mice (Prkcq–/–) respectively. We used 2 different transgenic mouse lines that express GFP either in NPY or POMC neurons. In the arcuate, we did not observe colocalization of PKC-θ with POMC neurons, (F) in POMC neurons (green), (G) with PKC-θ (red), or (H) PKC-θ/POMC colocalization. We did observe 20% colocalization of (I) NPY neurons (green) (arrow identifies an NPY neuron), (J) with PKC-θ (red) (arrow identifies a PKC neuron), and (K) NPY/PKC-θ colocalization (arrow identifies a colocalized NPY/PKC neuron). The mice express humanized renilla GFP (hrGFP) on the NPY promoter. To determine whether PKC-θ is expressed in neurons or on glia, (L) we colocalized PKC-θ (green) with NeuN (red) but not with (M) GFAP (red), a glial-cell marker. To assess whether PKC-θ is expressed in neurons that respond to leptin, we used a transgenic line that expresses GFP in only cells that also express the leptin receptor (LepR-GFP). We identified 22% colocalization of cells in the arcuate nucleus that were positive for (N) leptin receptor (green) or (O) both leptin receptor and PKC-θ (yellow) (arrow identifies a PKC/LepR-GFP colocalized neuron). Original magnification, ×10 (B, D, and E); ×20 (C and N); ×40 (L and M); ×120 (O). Scale bar: 120 μm (F–H); 100 μm (I–K); 300 μm (N); 50 μm (O). ARH, arcuate; ME, median eminence; PKC-ir, PKC immunoreactivity; 3v, third ventricle; VMH, ventral medial hypothalamic nucleus.

Palmitic acid induces translocation and activation of CNS PKC-θ. Saturated fat diets, specifically those with elevated levels of palmitic acid, induce translocation of PKC-θ to the membrane in peripheral tissues (5–7, 14, 28). We therefore assessed whether palmitic acid induces translocation of PKC-θ in brain. To this end, we used ultracentrifugation to accurately assess membrane and cytosolic protein content to determine translocation of PKC-θ. Validation of this technique is demonstrated in Figure 5A, showing immunolabeling of membrane-specific fragment markers, SNAP25 and IR, and labeling for a nuclear protein, Histone 3.

Figure 5 Saturated fatty acids increased PKC-θ translocation to the membrane. To confirm our protocol to isolate membrane from cytosolic fractions, medial basal hypothalamic cell lysate was ultracentrifuged. (A) The membrane fraction was stained with membrane-associated proteins, IR, SNAP 25, and Histone 3, a nuclear protein. Rats (n = 8–10/group), gavaged 3 times per day for 3 days with palmitic acid, oleic acid, or vehicle had no change in body weight (data not shown). Western blots were performed in 3 replications. There was a significant (P < 0.05) increase in medial basal hypothalamic membrane translocation of PKC-θ in the palmitic acid–gavaged animals but not the oleic acid– or vehicle-gavaged animals. Cyt, cytosolic; NUC, nuclear; TOT, total. (B) The GAPDH control was run on the same blot as AKT. Quantification (mean ± SEM) of 3 representative blots (n = 3/group) in C (vehicle and palmitic acid lanes were run on the same gel but were noncontiguous; oleic acid was run on a separate gel but under the same conditions; GAPDH lanes were run on separate gels). There was a significant decrease in medial basal hypothalamic cytosolic location of PKC-θ. (D) Quantified data (mean ± SEM) from 2 separate Western blots (n = 4/group), of which E is a representative blot. Cell membrane content of medial basal hypothalamic PKC-θ was also increased by i.c.v. osmotic minipump infusion of palmitic acid relative to oleic acid or vehicle. (F) Quantification (mean ± SEM) from 2 separate Western blots (n = 4–5/group), with a representative of the blot shown in G (vehicle, palmitic acid, and oleic acid lanes were run on the same gel but were noncontiguous). No significant change in cytosolic PKC-θ was found. (H) Quantification (mean ± SEM) from 2 separate Western blots (n = 4–5/group), representative of the blot shown in I (vehicle, palmitic acid, and oleic acid lanes were run on the same gel but were noncontiguous). (J) Represents increased serine phosphorylation of IRS-1, which is associated with reductions in insulin signaling in the animals i.c.v. infused the palmitic acid but not the oleic acid or vehicle infusion. (*P < 0.05 compared with vehicle treatment; quantification [mean ± SEM] from 3 separate Western blots [n = 5–6/group]).

Rats received intragastric gavage of nutritionally matched, fatty acid–enriched emulsions containing palmitic or oleic acid for 3 days. To our knowledge, this is a novel method, which tests the role of fatty acids independent of differences in palatability (hedonics), caloric consumption, or mixtures of fatty acids. The 3-day gavage of fatty acids elicited significant (P < 0.05) differences in CNS fatty acid content. The brain uptake index (BUI) (39) for animals gavaged with palmitic acid was 25.45% of that for the total fatty acids (oleic acid content in these animals was 15.98%) and the oleic acid–gavaged animals had a BUI of 23.19% of that for oleic acid (palmitic acid content in these animals was 15.56%). Additionally, we recently published elevations in hypothalamic long-chain fatty acyl-CoAs, following exposure to a high-fat diet or direct infusion of palmitate (10). Here, we extend those findings and demonstrate significantly (P < 0.05) elevated levels of palmitoyl (16:0) long-chain fatty acyl-CoA levels following the palmitate acid gavage relative to the control or oleic acid gavage (control = 20 ± 3.4 pmol/g tissue; palmitate, 35 ± 4.9 pmol/g tissue; oleic acid, 23 ± 3.2 pmol/g tissue). Palmitic acid–gavaged animals have increased hypothalamic membrane-bound PKC-θ relative to control or oleic acid–gavaged animals (Figure 5, B and C), and this is associated with concomitant reductions in cytosolic content (Figure 5, D and E). To test the ability of fatty acids, locally within the hypothalamus, to translocate PKC-θ to the membrane, independent of peripheral effects, we infused palmitic acid, oleic acid, or vehicle i.c.v. Similar to gavage, i.c.v. palmitic acid increased the membrane content of PKC-θ (Figure 5, F and G), with consequent reductions in cytosolic content (Figure 5, H and I), while oleic acid infusion had no effect. Consistent with the hypothesis that increased membrane translocation of PKC-θ induces insulin resistance, only the palmitic acid–infused animals exhibited an increase in hypothalamic serine 307 phosphorylation of IRS-1 (Figure 5J). To confirm that these effects were unique for PKC-θ, we found that neither PKC-δ nor PKC-ε were translocated following fatty acid exposure (data not shown).

PKCs also engage a number of downstream signaling systems that can be measured as surrogate indicators of PKC activity. Cytosolic myristoylated alanine-rich C kinase substrate (MARCKS) is phosphorylated by activation and translocation of many members of the PKC family, including PKC-θ (40). Therefore, to specifically test whether the fatty acids are impacting downstream targets of PKC-θ, we determined that i.c.v. palmitic acid but not oleic acid increased p-MARCKs in the cytosol (Figure 6, A and B). These data together demonstrate that exposure to palmitic but not oleic acid results in membrane localization and activation of PKC-θ in the hypothalamus, resulting in attenuation of insulin signaling.

Figure 6 Translocation of PKC-θ is associated with increased cytosolic p-MARCKS. A 3-day i.c.v. infusion of palmitic acid but not oleic acid or vehicle (n = 8–10/group) increased cytosolic p-MARCKS. (A) Quantification (mean ± SEM) from 3 separate Western blots (n = 8–10/ group) representative of the blot shown in B (vehicle and palmitic acid lanes were run on the same gel but were noncontiguous; oleic acid was run on a separate gel but under the same conditions; the bottom band depicts GAPDH from a separate blot). *P < 0.05.

Adeno-associated virus PKC-θ shRNA reduces PKC-θ expression and improves insulin sensitivity. In a final series of experiments, we assessed whether selective knockdown of hypothalamic PKC-θ by administration of shRNA-expressing adeno-associated virus (AAV) would improve glucose homeostasis, decrease body weight gain, and improve hypothalamic insulin signaling following exposure to the HFS diet elevated in palmitic acid. To accomplish this, we allowed ad libitum consumption of the diet, specifically to test whether knockdown of PKC-θ attenuated diet-induced obesity. Whereas the previous experiments attempted to avoid the confound of obesity, here we specifically sought to test whether reductions in PKC-θ in the arcuate had any influence on reducing body weight gain and improving CNS insulin sensitivity when animals are exposed to a HFS diet. We generated shRNA sequences, which are annealed into a hairpin AAV vector. Sequences of these oligos are identical to the publicly available mouse PKC-θ sequences: shRNA 1 (5′-AAACCACCGTGGAGCTCTACT-3′) and shRNA 2 (5′-AAGAGCCCGACCTTCTGTGAA-3′). To test the efficacy of this sequence, PKC-θ shRNA knocked down expression of PKC-θ in neuronal culture (Figure 7A). Next, we site specifically infused the PKC-θ shRNA bilaterally into the arcuate nucleus of mice (or performed control site–specific injections of scrambled oligo control AAV). Following the injections, mice were placed on HFS diet (with elevated levels of palmitic acid) or low-fat diet for 2.5 weeks. Mice that received the PKC-θ shRNA gained less weight on HFS diet than did mice with the control site–specific injections of scrambled oligo AAV. However, there were no differences in weight gain between PKC-θ shRNA and scrambled AAV–injected mice maintained on the low-fat diet (Figure 7B), providing further evidence that the PKC-θ–induced effects are directly downstream of elevated hypothalamic saturated fatty acids. Using an i.p. glucose tolerance test (0.75 g/kg body weight of 20% d-glucose [Phoenix Pharmaceutical St.]), we compared glucose sensitivity in mice maintained on the HFS diet that were either injected PKC-θ shRNA or scrambled oligo control AAV, and found a significant improvement in glucose tolerance in the mice that received the PKC-θ shRNA AAV injections (Figure 7C), demonstrating the peripheral effects of HFS diet–induced hypothalamic PKC-θ activation. Moreover, to further show that HFS diet–induced PKC-θ activation in the hypothalamus directly attenuates hypothalamic insulin signaling, we assayed p-AKT as a measure of insulin responsiveness. Two weeks following PKC-θ shRNA or scrambled oligo control AAV administration, 1 cohort of mice was peripherally injected with insulin (5 U/kg), sacrificed 30 minutes later, and hypothalamic tissue was processed for p-AKT immunoreactivity (Figure 7D). These results clearly demonstrate an improvement in insulin signaling (as measured by p-AKT) in those mice with PKC-θ knock down. In agreement with our overall hypothesis, we found that mice infused with PKC-θ shRNA and not the control scrambled oligo AAV, have significantly (P < 0.05) higher levels of insulin-induced arcuate p-AKT. To confirm the specificity of the injections, the PKC-θ shRNA and the scrambled oligo AAV control were tagged with GFP. As shown in Figure 7E, 2.5 weeks following i.c.v. administration of 0.5 μl of PKC-θ shRNA, GFP immunoreactivity was present in the arcuate nucleus, demonstrating successful transfection and expression of viral proteins. In a second cohort of mice, the arcuate was dissected and Western blot analysis demonstrated a significant reduction in PKC-θ protein levels only in the arcuate, following PKC-θ shRNA and not the control scrambled oligo AAV administration (Figure 7F), with no changes in PKC-θ in the rest of the hypothalamus (Figure 7G). Additionally, this reduction was specific for PKC-θ and not for PKC-δ, a PKC isoform with high sequence homology to PKC-θ (Figure 7, H and I). As an additional control, the PKC-θ shRNA and the control scrambled oligo AAV administration had no effect on muscle levels of PKC-θ (Figure 7J). These data confirm our hypothesis that arcuate-specific shRNA-induced knock down of PKC-θ “protects” against the deleterious effects of the HFS diet (elevated with palmitic acid) on CNS-mediated insulin resistance and the resultant obesity.