Saturated and monounsaturated fatty acids in membranes are determined by the gene expression of their metabolizing enzymes SCD1 and ELOVL6 regulated by the intake of dietary fat

Six-to-eight-week-old female standard C57BL6 (Strain Code 027) mice, purchased from Charles River (Budapest, H), were first fed ad libitum for 2 weeks with regular chow (VRF1, Altromin GmbH, Lage, D).

Total RNA was isolated from liver and quantified by QRT-PCR (quantitative real-time PCR). Samples of liver tissue (50 mg) were homogenized in Trizol (10 mg tissue/100 µl Trizol, Sigma-Aldrich, Budapest, H) and extracted with chloroform (20 µl/100 µl Trizol). The aqueous phase was mixed with 700 µl of ethanol (70% v/v) and loaded on the RNA isolation column (GenElute Mammalian Total RNA Miniprep Kit, Sigma-Aldrich, Budapest, Hungary). RNA was isolated from tissue according to the protocol of Sigma-Aldrich and eluted in nuclease free water. Concentration and purity of RNA were measured by Nanodrop (Thermo, Budapest, Hungary). cDNA was obtained by reverse transcription (10 min 25 °C, 120 min 42 °C, 5 min 72 °C) and amplified via QRT-PCR (40 cycles: 12 s 94 °C, 45 s 60 °C, 60 s 94 °C). The primer and probe for the expression analysis (Taq-Man-Gene Expression Assay) as well as the quantitative real-time PCR detection system (ABI-PRISM, 7900HT Sequence Detection System) were purchased from Applied Biosystems (Budapest, Hungary). The expression of genes was normalized to cyclophilin A (house-keeping gene): primer 77“+”5´-CGATGACGAGCCCTTGG-3´, primer 142”-“5´-TCTGCTGTCTTTGGAACTTTGTC-3´, probe (69+ , 96 +): FAM-CGCGTCTCCTTCGAGCTGTTTGCA, quencher tetramethylrhodamine (TAMRA). The amplification signal was detected and analyzed by the SDS2.1 program from Applied Biosystems, Budapest, Hungary. The expression of the house-keeping gene was stable and was not influenced by dietary treatment. The hepatic gene expression of target genes SCD1, ELOVL6, and CYP26A1 was normalized to coconut fat supplementation diet (= 1).

Food and plasma samples were analyzed by GC. Frozen samples were thawed and the pentadecanoylphosphatidylcholine (L-α phosphatidylcholine, dipentadecanoyl, Sigma-Aldrich, Budapest, Hungary) internal standard was added. Lipids were extracted by the addition of 3 ml chloroform and 1 ml methanol according to the method of Folch [39]. The mixture was vortexed at 3000 rpm for 15 min. The lower layer was then aspirated into vials and evaporated under an N₂ stream. Lipid extracts were reconstituted in 70 µl chloroform and lipid classes were separated by thin-layer chromatography (TLC). The solvent mix for TLC of plasma lipids was as follows: hexane: diethyl ether: chloroform: acetic acid (21:6:3:1, v/v). The bands were stained with dichlorofluorescein and removed by scraping and transesterified in 1 ml of 3 N HCl in methanol solution (3 N methanolic HCl, Supelco, Budapest, H) at 84 °C for 45 min [40]. For plasma samples, the phospholipid fraction was analyzed after TLC separation. Fatty acids were analyzed by high-resolution capillary GC using a Finnigan 9001 gas chromatograph (Finnigan/Tremetrics Inc., Austin, TX, USA) with split injection (ratio 1:25), an automatic sampler (A200SE; CTC Analytic, Zwingen, CH, USA), and a flame ionization detector with a DB-23 cyanopropyl column of 40 m length (J&W Scientific, Folsom, CA, USA). The temperature program was set to the following parameters: temperature of the injector at 80 °C/min up to 280 °C, temperature of the detector at 280 °C, temperature of the column area at 60 °C for 0.2 min, temperature increase by 40 °C/min up to 180 °C, 5 min isothermal period, temperature increase by 1.5 °C/min up to 200 °C, 8.5 min isothermal period, temperature increase by 40 °C/min up to 240 °C, and 13 min isothermal period. The constant linear velocity was 0.3 m/s (referred to 100 °C). Peak identification was confirmed by comparison with authentic mixtures of weighed fatty acid methylesters (GLC-463: Nu-Chek Prep, Elysian, MN, USA; and Supelco 37 FAME Mix: Supelco, Bellefonte, PA, USA). Individual fatty acid responses determined from weighed standards were used to calculate the percentage by weight for each fatty acid (between 12 and 24 carbon atoms) based on the area under the curve. Values were normalized to coconut fat (= 1) and compared with the other diet groups. The normalized values of enzyme-specific fatty acids for SCD1 and ELOVL6 analyzed by GC are displayed for diet and phospholipids. Normalization of values of the enzyme-specific fatty acid profile in diet and phospholipids enabled a direct comparison.

Network graphical analysis was created using the mixOmics package in R (3.5 version) [41]. The resulting color codes for the mouse data as well as the food data were incorporated into a self-constructed graphic representation.

The hepatic gene expression of CYP26A1 was significantly higher in the fish oil diet compared with the margarine, olive oil, and sunflower oil diets (Fig. 3a). In addition, CYP26A1 was significantly increased exclusively after administration of a specific RAR ligand and not by any other specific ligand of the nuclear hormone receptor family such as PPARα, β/δ or γ, RXR and LXR (Fig. 3b).

Our results clearly showed that the content of MUFAs and SAFAs in phospholipids is highly dependent on the hepatic gene expression of the lipid-metabolizing enzymes SCD1 and ELOVL6 and to a lesser extent on the ingestion of MUFA or SAFA (summarized in Fig. 4a). We found that the ratio 18:1n9/18:0 highly reflects the hepatic expression of SCD1, however this was only partly observed in the ratio 16:1n7/16:0. Our observation confirms previous findings in human plasma [43, 44]. In our study, 16:1n7/16:0 was further directly affected by fatty acids highly present in the fish oil diet. This fish oil diet contained high amounts of 16:1n7 and consumption leads to a high amount of 16:1n7 in plasma phospholipids in mice which could be an explanation for the elevated 16:1n7/16:0 ratio (Fig. 1a, e). In addition to this, we observed that in animals fed the fish oil diet, the hepatic expression of SCD1 was significantly lower compared with other dietary fats which explains the low levels of 18:1n9 incorporated into plasma phospholipids and the reduced 18:1n9/18:0 ratio. On the other hand, higher 18:1n9/18:0 and 16:1n7/16:0 ratios were observed in phospholipids of animals fed the coconut fat and margarine diets. These results highly reflect the increased hepatic gene expression of SCD1 following the coconut and margarine diets in comparison with those animals fed fish oil diet.

We also investigated specific fatty acid ratios in plasma 18:0/16:0 and 18:n7/16:1n7, because they have the potential to reflect the ELOVL6 hepatic gene expression [21]. Knock-out models of ELOVL6 have shown the importance of this enzyme for the elongation of fatty acids derived from diet or de novo fatty acid synthesis to provide sufficient amounts of 18:0 and 18:1n9 in the liver of mice [21]. Thereby, specific fatty acid ratios of 18:0/16:0 and 18:n7/16:1n7 were suggested by Matsuzaka et al. to reflect the ELOVL6 enzymatic reaction [21]. In one of our previous studies, we induced an overexpression of ELOVL6 in the liver by administration of specific ligands to activate the NHRs, LXR, and RXR, which induced the ELOVL6 hepatic gene expression leading further to an increase in the specific fatty acid ratios of ELOVL6 in liver and plasma [24]. In this study, we showed that feeding mice with fish oil, margarine, olive oil, and sunflower oil diets which contained high levels of 18:1n7 (Fig. 2a) resulted in low plasma phospholipid levels of this specific fatty acid (Fig. 2e). This result was more pronounced in the fish oil diet group, as the hepatic expression of ELOVL6 was significantly lower, which could further explain the low fatty acid ratio of 18:n7/16:1n7 observed in plasma phospholipids of this diet group. The low ratio of 18:0/16:0 in phospholipids of the fish oil diet group which was observed seems to result from the low hepatic gene expression of ELOVL6. Other suppressing effects by diet on ELOVL6 gene expression have been previously reported by our group. High fat diets rich in n6-PUFAs also suppressed ELOVL6 hepatic gene expression in a dose-dependent manner [22]. This suppression led to reduced enzymatic products of ELOVL6 in phospholipids [22].

The fish oil supplementation diet significantly reduced the hepatic gene expression of SCD1 and ELOVL6 in comparison with other sources of fat. This resulted in lower levels of MUFAs and higher levels of SAFAs in plasma phospholipids (summarized in Fig. 4a). The depletion and down regulation of SCD1 and ELOVL6 gene expression (indicating low levels of phospholipid MUFAs and high levels of SAFAs) might contribute to the protective mechanisms against diet-induced obesity, hepatic steatosis, and/or type 2 diabetes, as was shown in SCD1 and ELOVL6 knock-out models [45, 46]. This postulates different beneficial effects of fish oil/n3-PUFA diets besides functioning as precursors for anti-inflammatory and pro-resolving lipid mediators [21, 23, 47].

Although the hepatic gene expression of SCD1 and ELOVL6 by dietary supplemented fat and oils has been well investigated, it is still not clear why a fish oil diet rich in n3-PUFAs (20:5n3, 22:6n3) and in n6-PUFAs results in a comparable low hepatic expression of these enzymes [31, 48‐50]. For SCD1 and ELOVL6, we previously showed that the application of a synthetic LXR or RXR ligand can strongly induce hepatic SCD1 and ELOVL6 expression [24]. A retinoic acid is proposed as an endogenous RXR ligand [51], which our group found recently to be most likely 9-cis-13,14-dihydroretinoic acid (9CDHRA) [26, 52]. Afterwards, retinoic acid can be oxidized by the CYP26A1 and becomes inactive for receptor binding [29, 53]. Interestingly, 22:6n3 also has an inductive effect on the gene expression of CYP26A1 [30] via an unknown mechanism of action. We found that the fish oil-supplemented diet high in 22:6n3 strongly induced hepatic gene expression of CYP26A1 (Fig. 3a). The increased CYP26A1 expression might further catabolize 9-cis retinoic acid (9CRA) and 9CDHRA and reduce their availability as endogenous RXR ligand [26]. This might lead to reduced RXR-mediated signaling which may suppress hepatic SCD1 and ELOVL6 gene expression as observed in our experiment. The application of synthetic ligands to activate NHR showed that just the specific RAR ligand was able to strongly and significantly induce hepatic CYP26A1 expression (Fig. 3b). It seems likely that fish oil supplementation may mediate RAR-supported CYP26A1 expression via still unknown pathways (Fig. 4b). The activation of RAR-mediated signaling by a fish oil diet rich in n3-PUFAs may thereby be a major regulator of gene expression of the fat metabolizing target genes SCD1 and ELOVL6. Our findings suggest a potential new and important regulatory mechanism in which a fish oil diet rich in n3-PUFAs can alter lipid metabolism and might explain protective effects of PUFAs in obesity and diabetes [23] (summarized in Fig. 4b). We have used a mouse model with hepatic fatty acid-metabolizing enzymes that are also expressed in human liver [54‐57]; for example, PPARs hepatic expression is similar in mice and humans [58]. PPARγ expression and modulation is crucial in adipogenesis, lipid metabolism, and inflammation. PPARγ expression is modulated by dietary components belonging to the lipid family which undergoes extensive metabolism in the liver and acts as an inductor or suppressor of its expression [59]. The interaction between dietary fat type, circulating fatty acid patterns, and hepatic enzyme expression is likely to result in altered liver metabolism. This might have other major metabolic implications. For example, fish oil supplementation compared to olive oil increased beta-oxidation rate resisting the inhibitory effect of insulin which did not occur in olive oil supplementation. Fish oil also reduced fatty acid incorporation into triglycerides and secretion of VLDL triglycerides [60]. However, the applicability of our findings for humans remains speculative. We used different regimes of fat supplementation in controlled animal feeding studies which might not be relevant to human diets; thus, further studies using human cell lines, ex vivo human tissues, or human dose relevant clinical supplementation trials are needed for translational purposes.