Effect of Creosote Bush-Derived NDGA on Expression of Genes Involved in Lipid Metabolism in Liver of High-Fructose Fed Rats: Relevance to NDGA Amelioration of Hypertriglyceridemia and Hepatic Steatosis

The following reagents were supplied by Sigma-Aldrich Corp. (St Louis, MO): ATP, dithiothreitol (DTT), palmitic acid, fatty acid free bovine serum albumin, deoxycholate, Tris-HCl, SDS, Triton X-100, Oil Red O, and EDTA. Akt (pan) (67E7) rabbit monoclonal antibody (mAb) (#4691), PhosphoPlus^®Akt (Ser473) (D9E)XP™ rabbit mAb (#4060), phospho-Akt (Thr308) (D25E6) XP® rabbit mAb (#13038), acetyl CoA carboxylase (ACC) rabbit polyclonal antibody (#3662), phospho-acetyl-CoA carboxylase (Ser79) rabbit polyclonal antibody (#3661), AMPKα (23A3) rabbit mAb (#2603), phospho-AMPKα (Thr172) rabbit polyclonal antibody (#2531), AMPKβ1 antibody (#4182), phospho-AMPKβ1 (Ser108) rabbit polyclonal antibody (#4181), phospho-GSK-3α/β (Ser21/9) (37F11) rabbit mAb, and GSK-3β (27C10) rabbit mAb (#9315) were purchased from Cell Signaling Technology (Danvers, MA). MLYCD polyclonal antibody (15265-1-AP) was obtained from Proteintech (Chicago, IL). PPARα (H-98) rabbit polyclonal antibody and SCD1 (E-15) rabbit polyclonal antibody (sc-14720) were obtained from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA). IRDye 800CW-conjugated goat anti-rabbit IgG (H + L) and IRDye 680LT-conjugated goat anti-mouse antibodies were the products of LI-COR Biosciences (Lincoln, NE). Glucose, TG and total cholesterol measurement kits were obtained from Stanbio Laboratory (Boerne, TX), and FFA measurement kits were obtained from Wako Diagnostics. Rat insulin RIA Kit was obtained from EMD Millipore (Billerica, CA). [1-¹⁴C]-Palmitic acid (40–60 mCi [1.48–2.22 GBq/mmol was purchased from PerkinElmer (Waltham, MA). Humulin^®R (recombinant human insulin) was the product of Eli Lilly and Company (Indianapolis, IN). NDGA purified from Creosote bush was obtained from Insmed Inc., Richmond, VA as reported earlier [17–19].

Ethics Statement: All animal experiments were performed according to the procedures approved by the VA Palo Alto Health Care System Institutional Animal Care and Use Committee (IACUC). The specific procedures for this study were approved by the IACUC of VA Palo Alto Health Care System.

Male Sprague-Dawley (SD) rats, obtained from Harlan Laboratory (Indianapolis, IN), were used in these studies. NDGA treatment was delivered to rats weighing 175-200g by oral gavage. Rats were first maintained on a chow diet (TD 2018; Harlan Teklad, Madison, WI) for approximately 1 week to allow them to acclimatize to a controlled new environment (25 ± 2^°C, 55 ± 5% relative humidity with a 12 h-light-dark cycle), and then divided into three groups, two of which were switched to a high-fructose diet (TD.89247; Harlan Teklad, Madison, WI) that provided 60% of total calories as fructose (HFrD group), while the third group was maintained on a chow diet. Eight weeks (56 days) after feeding, on day 57, the rats were fasted for 4h and tail vein blood was collected for baseline measurement of serum triglyceride (TG), glucose, total cholesterol (TC), insulin and FFA. The two HFrD groups of rats were then treated with vehicle (0.5% carboxymethyl cellulose; CMC) or NDGA (125 mg/kg BW) twice a day for 5 days, delivered by oral gavage. The chow group (diet control) was treated with vehicle. During the treatment regimen, the animals were maintained on the high-fructose diet. After 5 days of treatment, blood was collected from the tail vein 3 h after the last dose of vehicle or NDGA, and serum samples were analyzed for TG, glucose, insulin, TC and FC. Subsequently, rats were euthanized with CO₂ and liver tissues collected, weighed and quickly frozen in liquid nitrogen. The samples were stored at -80°C for further analysis including Oil Red O staining of frozen sections for steatosis evaluation, RNA extraction for qRT-PCR, lipid extraction for measurement of hepatic triglyceride and cholesterol and protein extraction for Western blot analyses.

Serum glucose, triglyceride, and total cholesterol levels were determined with commercial assay kits (Stanbio Laboratory, Boerne, TX). Serum free-fatty acids levels were determined with kits from Wako Diagnostic (Richmond, VA). Serum insulin levels were quantified using a rat specific insulin RIA Kit (EMD Millipore, Billerica, CA).

Suitable aliquots of liver tissue homogenates were extracted with chloroform-methanol according to the procedure of Folch et al [22], and extracted lipid samples were analyzed for their TG content with an enzymatic assay kit as noted above.

Liver fatty acid (palmitate) oxidation rate was determined in fresh homogenates by a modification of the method of Mannarets et al [23] as described previously from this laboratory [19] using [1-¹⁴C]palmitate. Briefly, approximately 50 to 100 mg of liver samples were homogenized on ice in 20 volumes of SET buffer (250 mM sucrose, 1 mM EDTA, 10 mM Tris-HCl, and 2 mM ATP; pH 7.4) in a Potter-Elvehjem homogenizer with a tight-fitting Teflon pestle. The supernatant fractions in each case were employed for the measurement of total and peroxisomal β-oxidation activities. Briefly, for the determination of total β-oxidation activity, the reactions were carried out in a final volume of 0.4 ml in a 20 ml glass scintillation vial containing 0.2 mM palmitate + 0.2 μCi/ml [1-¹⁴C]palmitate (complexed to BSA), 100 mM sucrose, 10 mM Tris-HCl, pH 7.4, 10 mM potassium phosphate, 100 mM KCl, 1 mM MgCl₂, 1 mM L-carnitine, 0.1 mM malic acid, 2 mM ATP, 50 μM Coenzyme A (CoA) and 1 mM DTT. The vial also contained 400 μl of 1N NaOH with filter paper in a well suspended from the top of the vial to trap the ¹⁴CO₂ produced during the reaction. Before initiation of the reaction, the vial was sealed with a rubber cap and the reaction was initiated by the addition of 80 μl of the liver homogenate via syringe through the rubber cap. The vial was incubated for 60 min at 30°C before termination with 200 μl 6M H₂SO₄. After termination, the flasks were shaken continuously for 2h to ensure quantitative collection of CO₂ [24]. The paper insert containing the NaOH and trapped ¹⁴CO₂ was then removed to a scintillation vial and the radioactivity counted. Peroxisomal β-oxidation assays were carried out under identical conditions as described above except in each case the incubation mixture also contained 2 mM KCN, but without 0.5% BSA. The rate of mitochondrial fatty acid β-oxidation was calculated as the difference of total fatty acid β-oxidation minus peroxisomal fatty acid β-oxidation.

RNA isolation from the liver tissue samples (~20 mg) was carried out using the RNeasy Plus mini kit (Qiagen, Valencia, CA) as per the manufacturer’s instruction and was reverse-transcribed (1 μg total RNA) using a High Capacity RNA-to-cDNA^™ Kit (Life Technologies, Grand Island, NY). Real-time PCR was performed in a final volume of 10 μl containing 50 ng of cDNA template and specific sets of forward and reverse primers using a FastStart Universal SYBR Green Master PCR Kit (Roche Applied Science, Indianapolis, IN) and an ABI Prism 7700 system (Applied Biosystems^® Life Technologies, Grand Island, NY) according to the manufacturer’s protocols. The oligonucleotide primers and their sequences used in RT-PCR are shown in Table 1. The amplification process time was as follows: 95°C for 2 min; 40 cycles at 95°C for 20 s, 60°C for 20 s, 72°C for 30 s, and a final 5-min extension at 72°C. Results were normalized to the housekeeping gene 36B4, and the values of the control (chow-fed) group were set to 1.

Liver samples (~200 mg) were homogenized using a Potter-Elvehjem homogenizer in three volumes of detergent containing lysis buffer (50 mM Tris.HCl, pH 7.4, 1% Triton X-100 (v/v), 0.1% SDS, 0.25% deoxycholate 150 mM NaCl and 1X Halt™ Protease and Phosphatase Inhibitor Cocktail (Thermo Scientific/Pierce Biotechnology, Rockford, IL) and incubated on ice for 10 min. Following centrifugation at 10,000 x g for 10 min, the supernatant fractions were analyzed for protein content by a BCA™ Protein Assay Kit (Thermo Scientific/Pierce Biotechnology). Nuclear fractions were prepared using NE-PER nuclear and cytoplasmic extraction reagents (Thermo Scientific/Pierce Biotechnology, Rockford, IL). Protein extracts(20 μg) of total or nuclear proteins were subjected to 10% SDS-PAGE under denaturing conditions and transferred to nitrocellulose membranes. Blotted membranes were blocked with Odyssey blocking buffer (LI-COR Biosciences, Lincoln, NE) for 1h and then incubated with anti-phospho Akt, anti-Akt, anti-phospho GSK-3 α/β, anti-GSK-3β, anti-SCD1, anti-PPARα, anti-phospho-AMPKα, anti-phospho-AMPKβ, anti-AMPKβ, anti-phospho ACC1, anti-ACC1, anti-ACC2, or β-actin mAb (Sigma-Aldrich) for 16 h at 4°C. After three washes with tris-buffered saline containing 0.1% Tween 20, the membranes were incubated with IRDye 800CW goat anti-rabbit and IRDye 680LT goat anti-mouse secondary antibodies (LI-COR Biosciences) for 1h. Proteins were detected with the Odyssey Infrared Imaging System (LI-COR Biosciences).

Statistical analysis was performed using the Prism software (Prism 6.0, GraphPad, San Diego, CA). All of the data are expressed as the means ± S.E. ANOVA analyses were performed with Bonferroni’s multiple comparisons test, P < 0.05 was considered statistically significant.

Table 2 compares the effects of feeding a high-fructose diet (HFrD) and oral gavaging of HFrD fed rats with NDGA on body weight, serum glucose, TG, FFA, total cholesterol and insulin concentrations. Body weights were similar among the chow-fed, HFrD and HFrD-NDGA groups, consistent with many previous studies demonstrating that chronic high fructose diets do not cause increases in weight [25]. In contrast, feeding rats the high-fructose diet caused a dramatic elevation in plasma TG levels and also significantly raised plasma FFA and insulin levels. After 4 days of NDGA treatment, NDGA reduced plasma TG levels to chow-fed control levels. NDGA significantly decreased fasting serum insulin, fasting blood glucose and FFA levels. These changes reflected a reversal of insulin resistance induced by fructose feeding by NDGA, as assessed by HOMA-IR. Interestingly, NDGA treatment reduced the FFA levels in HFrD rats to levels that were 40% lower than that of chow-fed control (Table 2).

The results presented in Fig 1A show that fructose feeding increased hepatic weights, which was unaffected by NDGA treatment. Hepatic TG content of HFrD animals was elevated compared with chow controls (P < 0.05). NDGA treatment showed a trend in reducing hepatic TG (Fig 1B and 1C), an effect consistent with the restoration of normal plasma TG levels (Table 2).

In an effort to investigate the underlying mechanisms involved in NDGA-mediated amelioration of hypertriglyceridemia, we first measured hepatic mitochondrial and peroxisomal fatty acid β-oxidation rates in liver samples from chow-fed, HFrD-fed and HFrD-NDGA-treated animals. The results presented in Fig 2 demonstrate that hepatic total and mitochondrial β-oxidation ¹⁴CO₂ production rates were similar between HFrD fed and chow-fed control groups. Feeding the high-fructose diet, however, significantly decreased the rate of hepatic peroxisomal dependent β-oxidation production of ¹⁴CO₂ as compared to data obtained with chow-fed control livers (P < 0.01). Treatment of animals with NDGA further decreased the rate of peroxisomal fatty acid oxidation (P <0.05 HFrD vs HFrD-NDGA; P <0.0001 Chow vs HFrD-NDGA). In contrast, total and mitochondrial fatty acid oxidation was 40–50% greater in HFrD-NDGA-treated animals compared to vehicle administered HFrD-fed rats (P <0.05 total HFrD vs total HFrD-NDGA; P <0.01 Mito HFrD vs Mito HFrD-NDGA).

To better understand the NDGA mediated changes in plasma and liver triglyceride levels and hepatic fatty acid β-oxidation rates, mRNA levels of key proteins and lipogenic transcription regulators involved in hepatic lipid metabolism were quantified by real-time reverse-transcriptase polymerase chain reaction (qRT-PCR). The results are summarized in Fig 3. The mRNA levels of two key fatty acid binding/transport proteins, Fabp (L-FABP) and Cd36 (CD36/FAT), were approximately 2.5-fold higher in liver samples from HFrD-NDGA treated rats as compared to rats fed HFrD alone (Fig 3A; P <0.01 HFrD vs HFrD-NDGA and P <0.0001 HFrD vs HFrD-NDGA, respectively). In contrast, no changes were noted in the expression levels of Fabp4 (FABP4) and Slc27a5 (FATP5) in the HFrD-NDGA group compared to the HFrD or chow control group. With regard to fatty acid oxidation, we observed a significant increase in Acox1 expression in HFrD-NDGA treated rats (Fig 3B; P< 0.05HFrD-NDGA vs HFrD); the protein encoded by this gene, acyl-CoA oxidase (ACO), is the first enzyme in the fatty acid β-oxidation pathway and catalyzes the desaturation of acyl-CoAs to 2-trans-enoyl-CoAs, and is the rate-limiting step in peroxisomal fatty acid β-oxidation [26]. Expression of Cpt2, which encodes carnitine palmitoyl-transferase II (CPT-II), another enzyme that participates in fatty acid oxidation, was also upregulated by HFrD-NDGA treatment (Fig 3B; P <0.0001 HFrD-NDGA vs HFrD). The expression of other mitochondrial β-oxidation enzymes, such as Cpt1 (CPT-1) and Acadl (acyl CoA dehydrogenase, long chain), was not impacted by either HFrD feeding or HFrD-NDGA treatment.

We measured the expression of three genes involved in fatty acid synthesis (Acac1, Fasn, and MLYCD). Acac1 encodes acetyl CoA carboxylase, which catalyzes irreversible carboxylation of acetyl CoA to produce malonyl CoA. The Fasn gene product, fatty acid synthase complex, catalyzes the synthesis of palmitate from acetyl CoA and malonyl CoA [26, 27], whereas MLYCD encodes malonyl CoA decarboxylase, which catalyzes malonyl CoA degradation [28]. The expression levels of Acac1 and Fasn mRNA were up-regulated in response to high-fructose feeding (P <0.05 Chow vs HFrD and P <0.0001 Chow vs HFrD, respectively), but down-regulated with NDGA treatment (Fig 3C; P <0.05 HFrD vs HFrD-NDGA and P <0.001 HFrD vs HFrD-NDGA, respectively). In contrast, the expression levels of MLYCD mRNA were unaffected by either the HFrD or NDGA treatment (Fig 4). Moreover, the fatty acid desaturation enzyme, Scd1 (SCD1), but not Scd2 (SCD2), which participates in TG synthesis showed similar expression patterns to the fatty acid synthesis enzymes except high-fructose feeding greatly upregulated (~9-fold) Scd1 mRNA levels (Fig 3D). Likewise, mRNA expression of other TG synthesizing enzymes, Gpat1 and Dgat2, was up-regulated in response to HFrD feeding, whereas oral administration of NDGA reduced their expression to chow fed control levels (Fig 3E). Similarly, the expression of Pklr (L-PK), which also contributes to lipogenesis, was up-regulated in HFrD animals (P <0.0001 Chow vs HFrD) and reduced to normal levels following NDGA treatment (Fig 3F, P <0.001, HFrD vs HFrD-NDGA). In contrast, the expression levels of key genes involved in gluconeogenesis (Gck, Pck1 and G6pc) were not altered in HFrD or HFrD-NDGA groups (Fig 3F). We examined the expression of genes that participate in VLDL-TG assembly/secretion (Mttp [MTP], Arf1 [ARF1], and Pld1 [PLD1]) or lipid hydrolyzing enzymes, Lipe [HSL], Abhd5 [CGI-58], Pnpla2 [ATGL] and Lpl [LPL]), all of which showed comparable mRNA expression levels among the three groups (Fig 3G and 3H).

We next evaluated the effect of HFrD and HFrD-NDGA treatment on several key hepatic lipogenic transcription regulators. Among these, the carbohydrate response element-binding protein (ChREBP, also known as MLX-interacting protein-like) and sterol regulatory element binding protein-1c (SREBP-1c) are the two major transcription factors that induce key lipogenic enzymes to promote lipogenesis and steatosis [29–32]; ChREBP also increases the expression of a glycolytic gene, Pklr, and its protein product, L-PK, which in turn, stimulates both glycolysis and lipogenesis [30, 32]. SREBP-1c is activated by insulin via both transcriptional and posttranscriptional mechanisms [32, 33], whereas ChREBP is activated by high glucose independently of insulin [30, 32]. Feeding a high-fructose diet increased both Mixipl (ChREBP) and Srebf1 (SREBP-1c) mRNA levels by approximately 2-fold (P <0.001 and P <0.01 vs Chow, respectively) (Fig 3I). In HFrD-NDGA livers, however, mRNA levels of both Mixipl and Srebf1 were reduced (P <0.01 and P <0.01 vs HFrD, respectively). The mRNA levels of Ppara (PPARα), a nuclear receptor and master regulator of the fatty acid oxidation pathway in the liver [34, 35], was increased in HFrD-NDGA treated (P <0.01) rats as compared to animals maintained on a chow diet (Fig 3I), but high-fructose feeding had no effect. In contrast, the expression of Ppard (PPARδ), which also participates in the regulation of energy metabolism, was decreased by 70–80% in response to HFrD-NDGA treatment as compared to the chow-fed group. The expression of nuclear receptor Pparg (PPARγ), which promotes lipogenesis, as well as Cebpa (C/EBPα), Nr1h2 (LXRβ) and Nr1h3 (LXRα), was not affected by either of the treatments (Fig 3I). The expression levels of Foxo1a (FOXO1A) and Foxa2 (FOXA2) were down-regulated and up-regulated by HFrD-NDGA treatment (P <0.05) and HFrD feeding (P <0.05), respectively. Similarly, Xbp1 (XBP-1) mRNA levels were up-regulated both in high-fructose fed and HFrD-NDGA treated groups. PGC1α mRNA expression was down-regulated by fructose feeding and unaffected by NDGA, whereas levels of PGC1ß mRNA were elevated by fructose, but was not changed by NDGA treatment (Fig 4). The mRNA levels of malonyl-CoA decarboxylase (MLYCD) were not impacted by either fructose feeding or NDGA treatment (Fig 4).

The data presented above indicate that NDGA treatment stimulates and inhibits the gene expression of key enzymes involved in fatty acid oxidation and lipogenesis, respectively. Here, we measured the protein levels of two lipogenic enzymes, ACC and SCD1, and the PPARα, the master regulator of fatty acid oxidation together with malonyl-CoA decarboxylase. As shown in Fig 5, NDGA treatment suppressed the fructose-induced increases in SCD1 protein. Hepatic PPARα protein level was almost undetectable in HFrD fed mice, butwas increased significantly by NDGA treatment. At the same time, NDGA prevented the fructose suppression of MLYCD (Fig 5). In addition, we measured the total and phosphorylated levels of AMPK, ACC, AKT, and GSK3α/β. AMPK, a heterotrimeric protein complex consisting of a catalytic α subunit and regulatory β and γ subunits, plays a key role in the regulation of energy homeostasis [36, 37]. Phosphorylation of Thr172 within the α subunit and Ser108 of the β1 subunit by upstream kinases results in the activation of AMPK activity [36, 37]. AMPK phosphorylates ACC1 on Ser79 (the major phosphorylation site), Ser1200 and Ser1215 residues and deactivates its activity [38]. AKT, a core component of the phosphoinositol 3-kinase (PI3K)-AKT/protein kinase B (PKB), which is responsible for most of the metabolic actions of insulin [39], is activated by phosphorylation at Ser473 and Thr308. GSK3 is rapidly phosphorylated at Ser21 in GSK3α or Ser9 in GSK3β by AKT, resulting in inhibition of GSK3 kinase activity [40]. Results presented in Figs 5 and 6 demonstrate that protein levels of ACC1, ACC2 and SCD1, similar to their mRNA levels, were up-regulated in livers of high-fructose fed rats as compared to chow-fed control livers. Treatment of HFrD fed animals with NDGA decreased the protein levels by approximately 75–80%, further confirming that NDGA can inhibit hepatic lipogenesis. In contrast, the protein expression of PPARα was barely detectable in control or HFrD livers, but was markedly induced in response to HFrD-NDGA feeding (Fig 5).

Additionally, Western blotting analysis demonstrated that HFrD feeding decreased, whereas HFrD-NDGA treatment restored, the levels of the phosphorylated form of AMPKα (P-AMPKα) (Fig 6). Protein expression of hepatic total AMPKα, P-AMPKβ and total AMPKβ, however, was similar between chow-fed, HFrD and HFrD-NDGA animals (Fig 6). Interestingly, ACC1 phosphorylation was markedly increased by HFrD feeding, which was further increased in response to NDGA treatment (Fig 6; P <0.05). Total AKT levels were comparable among the three groups, but NDGA treatment significantly increased the basal Ser473 phosphorylation of AKT by approximately 2.5-fold (Fig 7). Although basal phosphorylation of both GSK3α (Ser21) and GSK3β (Ser9) tend to be increased, this difference was not statistically significant when data were normalized to total GSK3β (Fig 7). Moreover, total GSK3β levels showed a trend to be increased in response to both HFrD and HFrD-NDGA compared to chow fed animals; however, this did not achieve statistical significance.

All relevant data are within the paper and its Supporting Information files.