The objective of this study was to determine whether a grape seed procyanidin extract (GSPE) exerts a triglyceride-lowering effect in a hyperlipidemic state using the fructose-fed rat model and to elucidate the underlying molecular mechanisms. Rats were fed either a starch control diet or a diet containing 65% fructose for 8 weeks to induce hypertriglyceridemia. During the 9th week of the study, rats were maintained on their respective diet and administered vehicle or GSPE via oral gavage for 7 days. Fructose increased serum triglyceride levels by 171% after 9 weeks, compared to control, while GSPE administration attenuated this effect, resulting in a 41% decrease. GSPE inhibited hepatic lipogenesis via down-regulation of sterol regulatory element binding protein 1c and stearoyl-CoA desaturase 1 in the fructose-fed animals. GSPE increased fecal bile acid and total lipid excretion, decreased serum bile acid levels and increased the expression of genes involved in cholesterol synthesis. However, bile acid biosynthetic gene expression was not increased in the presence of GSPE and fructose. Serum cholesterol levels remained constant, while hepatic cholesterol levels decreased. GSPE did not modulate expression of genes responsible for esterification or biliary export of the newly synthesized cholesterol, but did increase fecal cholesterol excretion, suggesting that in the presence of GSPE and fructose, the liver may secrete more free cholesterol into the plasma which may then be shunted to the proximal small intestine for direct basolateral to apical secretion and subsequent fecal excretion. Our results demonstrate that GSPE effectively lowers serum triglyceride levels in fructose-fed rats after one week administration. This study provides novel insight into the mechanistic actions of GSPE in treating hypertriglyceridemia and demonstrates that it targets hepatic de novo lipogenesis, bile acid homeostasis and non-biliary cholesterol excretion as important mechanisms for reducing hypertriglyceridemia and hepatic lipid accumulation in the presence of fructose.
Citation: Downing LE, Heidker RM, Caiozzi GC, Wong BS, Rodriguez K, Del Rey F, et al. (2015) A Grape Seed Procyanidin Extract Ameliorates Fructose-Induced Hypertriglyceridemia in Rats via Enhanced Fecal Bile Acid and Cholesterol Excretion and Inhibition of Hepatic Lipogenesis. PLoS ONE 10(10): e0140267. https://doi.org/10.1371/journal.pone.0140267
Editor: Marcia B. Aguila, State University of Rio de Janeiro, Biomedical Center, Institute of Biology, BRAZIL
Received: June 9, 2015; Accepted: September 22, 2015; Published: October 12, 2015
Copyright: © 2015 Downing et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Data Availability: All relevant data are within the paper and its Supporting Information files.
Funding: This work was supported by start-up funding from the University of Nevada Reno and USDA Hatch (NEV 0738)/Multi-State project W-3122: Beneficial and Adverse Effects of Natural Chemicals on Human Health and Food Safety (http://nimss.umd.edu/lgu_v2/homepages/home.cfm?trackID=14436) to MLR. GCC was the recipient of a CONICYT Bicentennial Becas-Chile Scholarship. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Abbreviations: BA, Bile acid; GSPE, Grape seed procyanidin extract; TG, Triglyceride
Rates of obesity, type 2 diabetes, non-alcoholic steatohepatitis (NASH) and non-alcoholic fatty liver disease (NAFLD) have increased significantly in recent years, both in children and adults [1–4]. This surge has correlated with a significant increase in dietary fructose intake in the United States, due in large part to the rise in consumption of sugar-sweetened beverages . Fructose is a highly lipogenic dietary factor  and increasing evidence points to an obesogenic role for fructose via the generation of substrates for de novo lipogenesis, resulting from rapid hepatic metabolism . Since fructose metabolism is insulin-independent , there is less uptake and catabolism of triglyceride (TG)-rich lipoproteins by tissues, ultimately resulting in increased postprandial plasma TG levels . Increased hepatic lipogenesis, combined with decreased uptake of TGs in peripheral tissues, is an important mechanism by which fructose induces steatosis and elevates serum TG levels .
In contrast to fructose-induced metabolic dysregulation, evidence indicates that diets rich in fruits and vegetables, e.g. the Mediterranean diet, exert protective effects against the development of the metabolic syndrome . Such diets tend to be high in flavonoids, which exhibit cardioprotective effects in humans [11, 12]. Dietary procyanidins, a class of flavonoids commonly found in grapes, apples and red wine, have been shown to ameliorate risk factors associated with hypertriglyceridemia and steatosis [13–17]. We previously reported that a grape seed procyanidin extract (GSPE) exerts hypotriglyceridemic effects in vivo in a normolipidemic state [18–22]. We identified GSPE as a co-agonist ligand for the farnesoid x receptor (nuclear receptor subfamily 1, group H, member 4; FXR) , a transcription factor that regulates bile acid (BA), TG, cholesterol and glucose homeostasis [23–27]. Mechanistically, GSPE functions in conjunction with BAs, the endogenous ligands of FXR, to upregulate the expression of small heterodimer partner (nuclear receptor subfamily 0, group B, member 2; Shp), which represses sterol regulatory element binding protein 1c (Srebp-1c) expression, a key regulator of lipogenesis, resulting in the concomitant decrease in downstream lipogenic gene expression [19, 22]. Evidence also shows that GSPE administration increases fatty acid β-oxidation  and reduces VLDL-TG secretion .
Previous studies utilized a high-fat diet to examine the effects of GSPE [20, 28], however, since sugar intake is also a critical factor that can modulate metabolic homeostasis, particularly TG levels [29, 30], evaluation of the potential therapeutic impact of GSPE on animal models subjected to other dietary regimens, such as carbohydrate-induced hypertriglyceridemia, is warranted. Consequently, the aim of the current study was to more closely mimic a real-world scenario, by assessing whether GSPE can mitigate the effects of existing fructose-induced hypertriglyceridemia in vivo, and to determine the underlying molecular mechanisms. This investigation has important implications for further investigations in human subjects using GSPE as a potential natural therapy to counteract increased incidences of hypertriglyceridemia and steatosis.
In the present study, we show that a high-fructose diet for 8 weeks significantly increases serum TG levels in rats, while also markedly inducing hepatic lipid accumulation (steatosis). Co-administration of GSPE with the high-fructose diet for one week only, during the 9th week of the study, effectively ameliorated the adverse consequential effects on serum TGs resulting from the high-fructose diet. This attenuation was achieved via enhanced fecal BA, total lipid, cholesterol and non-esterified fatty acid excretion, inhibition of hepatic de novo lipogenesis and increased TG catabolism. We evaluated the gene regulatory effects exerted by GSPE in the liver in the presence of fructose to gain a better insight and understanding regarding the molecular effects leading to the observed hypotriglyceridemic effect.
Materials and Methods
Grape Seed Procyanidin Extract (GSPE) was obtained from Les Dérives Résiniques et Terpéniques (Dax, France). The extract was analyzed in-house using normal phase high performance liquid chromatography (HPLC), as previously described  to determine procyanidin composition based on the degree of polymerization. As shown in S1 Fig, GSPE is comprised of procyanidin monomers (68.68 ± 0.02%), dimers (26.16 ± 0.01%) and trimers (5.16 ± 0.02%).
Animal feeding studies and diets
Rats were housed under standard conditions and all experimental procedures were approved by the local Institutional Committee for Care and Use of Laboratory Animals (IACUC) at the University of Nevada, Reno (Protocol # 00502). Male Wistar rats, 7 weeks of age, were purchased from Charles River Laboratories. After one week of acclimation, rats were randomly assigned to either a control diet (n = 5) or fructose diet (n = 8) for 8 weeks (Harlan Teklad). As shown in Table 1, the starch control diet was a modification of AIN-93G (TD.94045) replacing all sucrose with starch, and was comprised of (% by weight) 17.7% protein, 58.9% carbohydrate and 7.2% fat, providing 3.7 Kcal/g (TD.110787). The fructose diet was a modification of AIN-93G replacing all sucrose and starch with fructose, and was comprised of 17.7% protein, 64.7% carbohydrate and 7.2% fat, providing 3.9 Kcal/g (TD.110786). The animals were maintained on the AIN-93G formulated diets throughout the 9-week study, consistent with previous reports [32, 33]. Food was replenished 3 times per week and food intake was estimated by subtracting the total amount of feed and the amount remaining in the box. Rats were weighed weekly.
Blood samples were collected at 0, 4 and 8 weeks to measure serum triglyceride levels. After 8 weeks on the diets, the rats were randomly assigned to receive either vehicle (water) or GSPE (250 mg/kg) via oral gavage for 7 days, while still consuming their assigned diets. The dose of GSPE used in this study is one-fifth of the no-observed-adverse-effect level (NOAEL) described for GSPE in male rats . This dose is effective in reducing serum TG levels in normolipidemic C57BL/6 mice [19, 22] and rats , and was chosen to aid in the identification of the primary, short-term effects of grape seed procyanidins on lipid metabolism in the presence of fructose, in order to gain insight into the mechanisms that underlie their potential longer-term effects. Using a translation of animal to human doses based on metabolic body size  and estimating the food intake for a 60-kg human, the dose of GSPE used herein corresponds to ~1.8 g, which is less than a 2 g/day dose previously tested in human subjects . On day 7, the rats were gavaged at 9am, food was then removed and they were sacrificed 5 hours later. Rats were anesthetized using isoflurane, blood was collected from the saphenous vein, after which the animals were euthanized using carbon dioxide, livers were excised and weighed, snap frozen in liquid nitrogen and stored at –80°C. For collection of the feces, rats were placed in clean cages 3 days prior to the end of the experiment and feces were manually collected at the end of the study, air-dried and weighed.
Gene expression analysis
Total RNA was extracted from liver using TRIzol (Life Technologies) according to the manufacturer’s instructions. Complimentary DNA (cDNA) was reverse transcribed using superscript III reverse transcriptase (Life Technologies), and real-time quantitative polymerase chain reaction (qPCR) was used to determine gene expression changes. qPCR was performed using a CFX96 Real-Time System (BioRad). Forward and reverse primers and probes were designed using Primer3Plus software  and purchased from Sigma-Aldrich or Integrated DNA Technologies. Expression of β-actin and TATA box binding protein (Tbp) were used as endogenous controls. All gene names, abbreviations and accession numbers can be found in S1 Table. Primer and probe sequences can be provided upon request.
Serum triglyceride and total cholesterol were measured enzymatically using InfinityTM kits (Thermo Scientific) according to the manufacturers’ instructions. Serum bile acid levels were measured using the Total Bile Acids Assay kit from Diazyme Laboratories, and serum free fatty acids were measured using a non-esterified fatty acid enzymatic colorimetric HR series NEFA-HR (2) assay from WAKO Chemicals USA Inc., performed according to the manufacturer’s instructions. Lipoprotein Lipase (LPL) activity was measured in serum using an ELISA kit (Cell Biolabs, Inc, San Diego, CA), and serum alanine aminotransferase (ALT) and aspartate aminotransferase (AST) were measured using Teco Diagnostic Kits (A526 and A561 respectively), according to the manufacturer’s instructions.
After removal from the rat, sections of excised liver were immediately immersed in 10% buffered formalin, and processed for hematoxylin and eosin (H&E) staining, which was performed by the Pathology Laboratory at University of Nevada, Reno.
Evaluation of Steatosis by Image Analysis
At least three non-consecutive microscopic fields per animal were randomly analyzed, by blindly moving the field of view, using an Olympus DP71 camera attached to a BX41 microscope, equipped with a UPlanFL 40X objective. Each image field was analyzed using a 94x70 point grid (PT). The volume density of hepatic steatosis (VV [steatosis, liver]) was then estimated as the ratio of the points marking the vesicles of fat (PP) compared to the number of test points using the following equation: VV [steatosis, liver] = PP [steatosis]/PT, as previously described .
Hepatic cholesterol measurement
Hepatic cholesterol was extracted using the Folch extraction method and levels were measured as previously described [39, 40]. Briefly, lipids were extracted from 100 mg of liver using a chloroform/methanol mixture. Cholesterol was then determined using a colorimetric Infinity cholesterol assay kit, performed according to the manufacturers’ instructions. Extracted cholesterol contents were normalized to wet liver weight.
Measurement of fecal bile acid, total lipid, cholesterol and free fatty acid excretion
A modified version of the method reported by Modica et al., was used to measure fecal BA content . Briefly, 0.2 g of dried feces was mixed with 2 ml of 2 mg/ml sodium borohydrate in ethanol and incubated at room temperature for 1 hour. Hydrochloric acid and sodium hydroxide were added and samples were vortexed and allowed to digest for 12 hours under reflux. The samples were then filtered and dried under nitrogen. Samples were re-suspended in milli-Q water and filtered through Sep-Pak C18 cartridges (Thermo Scientific), washed and eluted with methanol and dried under nitrogen. Samples were re-dissolved in 1 ml methanol and BA concentrations were measured enzymatically, using the Total Bile Acids Assay kit from Diazyme Laboratories. Fecal cholesterol and non-esterified fatty acids were extracted as previously described , and cholesterol levels were measured using a colorimetric Infinity cholesterol assay kit and non-esterified fatty acids were quantified using a Wako diagnostics HR Series NEFA-HR (2) assay. Total fecal lipids were assessed via gravimetric analysis following Folch extraction. Briefly, 0.5 ml aliquots of the chloroform layer were placed into pre-weighed glass tubes (three tubes per sample) and allowed to evaporate in a fume hood overnight. The next day, the weights were recorded and converted to percent lipids (mg lipid/mg dry fecal weight).
Data represents the mean ± SEM for the fold change relative to control (or endogenous gene expression) (n = 4 or 5 per treatment, per group, analyzed in triplicate). One-way analysis of variance (ANOVA) followed by Holm-Sidak post-hoc tests was employed to detect significant differences between groups. Treatment differences were considered statistically significant at p<0.05. All statistical analyses were performed using GraphPad Prism version 6.05 for Windows, GraphPad Software (San Diego, CA).
Effect of fructose feeding for 8 weeks and co-administration with GSPE for the 9th week of the study on body weight, liver weight and serum biochemical analysis
Fructose feeding for 8 weeks did not significantly increase body weight compared to the rats fed the control diet (Table 2). As shown in S2A Fig, the fructose fed rats tended to have a lower body weight compared to the control animals after the first week on the diets, a trend which continued through to week 8. Food intake was not statistically different between the groups.
GSPE administration during the 9th week of the study did not have any significant effect on the body weight of the rats in the fructose-GSPE group, compared to the fructose-vehicle group. As shown in S2B Fig, the group of animals assigned to receive GSPE in week 9 demonstrated no significant differences in body weight during the course of the study. In contrast, the fructose-fed rats had a significantly higher liver to body weight ratio compared to the control group at the end of the study, as shown in Table 2, with no significant differences observed following GSPE administration.
Consistent with the increased liver weight, fructose-fed animals had a significantly higher grade of microvesicular steatosis and increased hepatic lipid accumulation volume density, compared to the control group (Fig 1). The fructose-fed rats also showed evidence of mild inflammatory infiltration around the portal triad (Fig 1C). GSPE administration for one week resulted in a significant decrease in hepatic lipid volume density and therefore steatosis (Fig 1E, 1F and 1G). The fructose diet increased serum TG levels by 171% compared to the control group after 9 weeks (Fig 2A), which was ameliorated by treatment with GSPE resulting in a 41% reduction. No significant changes were observed in the fructose-fed rats with respect to serum cholesterol (Fig 2B) or NEFA levels (Fig 2C). Serum BA levels were not altered by the fructose diet alone, however, co-administration with GSPE for one week significantly reduced serum BA levels (Fig 2D). Neither fructose feeding nor GSPE administration caused liver damage, as evidenced by the fact that the values for both ALT and AST remained within normal limits (Fig 2E and 2F). Lipoprotein lipase activity did not change upon fructose consumption, nor following GSPE administration (Fig 2G). Hepatic cholesterol levels were significantly reduced following GSPE administration (Fig 2H). Fecal BA levels were significantly decreased by the fructose diet, and increased following GSPE administration (Fig 3A). Total fecal lipid excretion was significantly increased by GSPE in the fructose-fed animals (Fig 3B). Fecal cholesterol (Fig 3C) and NEFA (Fig 3D) were both increased by the fructose diet, while GSPE further enhanced their excretion.
Representative liver histology sections stained with hematoxylin and eosin from (A and B) Control, (C and D) Fructose-vehicle and (E and F) Fructose-GSPE treated rats. Lipid droplet infiltration is evident in the fructose-fed rats as indicated by the black arrows (BD: Bile duct; CV: Central vein; HA: Hepatic artery; PT: Portal triad); (G) Lipid accumulation induced by fructose consumption was evaluated using image analysis of the histological sections from each animal. The volume of lipid droplets was assessed as a percentage of the total hepatic volume. *p<0.05 and **p<0.01.
Serum was analyzed for (A) triglycerides (TG) (B) cholesterol (CHOL), (C) non-esterified fatty acids (NEFA), (D) bile acids (BA), (E) Alanine aminotransferase (ALT), (F) aspartate aminotransferase (AST); the dotted lines represent the normal upper and lower limits respectively. (G) lipoprotein lipase activity (LPL), and (H) hepatic cholesterol content (CHOL). *p<0.05, ***p<0.001 and ****p<0.0001.
Expression of genes involved in hepatic lipogenesis, cholesterol & bile acid synthesis and transport
To explore the molecular mechanism underlying the hypotriglyceridemic effect of GSPE observed in the fructose-fed animals, we next examined the expression of hepatic genes that regulate lipogenesis, cholesterol synthesis and transport, and BA synthesis and transport. Dietary fructose did not alter sterol regulatory element binding protein 1c (Srebp-1c) expression in this study; however, GSPE significantly reduced expression (Fig 4A). Peroxisome proliferator-activated receptor gamma, coactivator 1 beta (Pgc-1β activates the expression of genes involved in lipogenesis and TG secretion via direct co-activation of Srebp . Consequently, Pgc-1β, a regulator of both carbohydrate and lipid metabolism, has been proposed to play a pivotal role in fructose-induced lipogenesis . However, fructose feeding reduced Pgc-1β expression, compared to the control diet, in this study (Fig 4B). Fructose feeding, either with or without GSPE, had no effect on fatty acid synthase (Fasn) expression, a Srebp-1c lipogenic target gene (Fig 4C). Stearoyl-CoA desaturase (delta-9-desaturase) 1 (Scd1) another SREBP-1c target gene, although not affected by fructose, was markedly repressed by GSPE (Fig 4D).
Gene expression changes were analyzed for (A) Srebp-1c, (B) Pgc-1β, (C) Fasn, (D) Scd1, (E) Pparα, (F) Fgf21 and (G) Mlxipl. *p<0.05, **p<0.01 and ****p<0.0001.
An alternative mechanism by which fructose can increase lipogenesis is via repression of peroxisome proliferator-activated receptor alpha (Pparα) . Consequently, we also assessed the effects of fructose feeding on the expression of Pparα and fibroblast growth factor 21 (Fgf21), a down-stream target of Pparα [43–45]. As shown in Fig 4E, fructose feeding did not alter the expression of Pparα, however, Fgf21 expression was significantly repressed (Fig 4F). Pparα expression was significantly repressed by GSPE, compared to the control, while no difference was seen with respect to Fgf21 expression in the presence of GSPE, compared to fructose only. No significant changes in the expression of MLX interacting protein-like (Mlxipl), also referred to as carbohydrate response element binding protein, were observed following fructose ingestion or GSPE administration in this study (Fig 4G).
Based on the combined observations of reduced serum TG and BA levels and increased fecal BA excretion, we postulated that serum TG levels were reduced due to the need to synthesize cholesterol and then BAs. Therefore, we next examined potential regulatory effects on genes involved in both cholesterol and BA synthesis. Dietary fructose had no effect on the expression of 3-hydroxy-3-methylglutaryl-CoA synthase 1 (Hmgcs1), while GSPE significantly increased expression (Fig 5A). In addition, no changes in expression were seen in the fructose-fed animals with respect to 3-hydroxy-3-methylglutaryl-CoA reductase (Hmgcr) (Fig 5B). However, GSPE increased expression of Hmgcr compared to both the control and fructose-vehicle-treated animals. In addition, several genes important in cholesterol synthesis were significantly upregulated in the presence of GSPE, including farnesyl-diphosphate farnesyltransferase 1 (Fdft1) (Fig 5C), squalene epoxidase (Sqle) (Fig 5D), lanosterol synthase (2,3-oxidosqualene-lanosterol cyclase) (Lss) (Fig 5E) and 7-dehydrocholesterol reductase (Dhcr7) (Fig 5F), indicating that there was increased cholesterol synthesis in the presence of GSPE. Consequently, we next evaluated the effects of GSPE on BA biosynthesis. No significant effects were seen with respect to cytochrome P450, family 7, subfamily A, polypeptide 1, (cholesterol 7 alpha-monooxygenase; Cyp7a1) expression, which initiates the classical (neutral) BA biosynthetic pathway, following GSPE administration (Fig 6A), while cytochrome P450, family 8, subfamily B, polypeptide 1, (sterol 12-alpha-hydroxylase; Cyp8b1) levels were significantly reduced (Fig 6B). We then assessed the effects of GSPE on genes involved in the alternative (acidic) BA biosynthesis pathway. No significant effects were observed with respect to cytochrome P450, family 27, subfamily A, polypeptide 1 (steroid 27-hydroxylase; Cyp27a1) (Fig 6C) or cytochrome P450, family 7, subfamily B, polypeptide 1 (oxysterol 7-alpha-hydroxylase; Cyp7b1) (Fig 6D). In addition, no changes were observed in the expression of ATP-binding cassette, sub-family B (MDR/TAP), member 11 (bile salt export pump; Abcb11) (Fig 6E).
Gene expression changes were analyzed for (A) Hmgcs1, (B) Hmgcr, (C) Fdft1, (D) Sqle, (E) Lss, and (F) Dhcr7. *p<0.05, **p<0.01, ***p<0.001 and ****p<0.0001.
Gene expression changes were analyzed for (A) Cyp7a1, (B) Cyp8b1, (C) Cyp27a1, (D) Cyp7b1, and (E) Abcb11. ** p<0.01.
Since we observed an increase in endogenous cholesterol synthesis but no changes in the expression of genes regulating BA biosynthesis, despite reduced hepatic cholesterol levels, we next analyzed expression levels of acetyl-CoA acetyltransferase 1 (Acat1) and acetyl-CoA acetyltransferase 2 (Acat2) to determine whether the newly synthesized cholesterol was being esterified and exported, for assembly into lipoproteins. No changes in Acat1 (Fig 7A) or Acat2 expression (Fig 7B) were observed. ATP-binding cassette, subfamily (ABC1), member 1 (Abca1) either directly or indirectly mediates the transport of cholesterol and phospholipids across cell membranes, where they are removed from cells by apolipoproteins . We therefore measured Abca1 and found no changes in expression (Fig 7C). Our previous studies showed that GSPE reduced levels of apolipoprotein B (ApoB) in HepG2 cells , and measurement of microsomal triglyceride transfer protein (Mttp) expression, which helps to deliver TG to ApoB was not affected by either fructose feeding or GSPE administration in this study (Fig 7D). We next analyzed the expression of ATP-binding cassette, subfamily G (WHITE), member 5 (Abcg5) and ATP-binding cassette, subfamily G (WHITE), member 8 (Abcg8) to determine whether the unesterified cholesterol underwent biliary export. No significant changes in expression were observed for either Abcg5 or Abcg8 (Fig 7E and 7F), and fructose feeding did not alter low density lipoprotein receptor (Ldlr) expression (Fig 7G).
The study presented herein provides new evidence establishing that one week administration with GSPE effectively lowers serum TG levels in a model of fructose-induced hypertriglyceridemia. Previous reports showed that GSPE decreases serum TGs in a normolipidemic state in several animal models, including rats [18, 19, 22]. We now show that, under conditions of severe hypertriglyceridemia, GSPE causes a significant 41% reduction in serum TG levels. This is comparable to that seen with fenofibrate, one of the most commonly prescribed lipid-lowering agents in the world , which reduces serum TGs by 36% . Cholesterol synthesis gene expression was increased in the fructose-GSPE treated animals, and was accompanied by a concomitant increase in fecal excretion of cholesterol, which could occur via transintestinal cholesterol efflux (TICE) . Additionally, compared to the control diet, fructose caused a marked increase in hepatic lipid droplet accumulation, indicative of steatosis (>5% lipid volume), which was significantly attenuated following co-administration with GSPE.
It is well established that high-fructose feeding causes diet-induced alterations in lipid metabolism . In the present study, we observed a significant 171% increase in serum TGs in the rats after 9 weeks on the fructose diet. Increased expression of Srebp-1c was previously shown to be one mechanism underlying fructose-induced hypertriglyceridemia, however, it is not the only mechanism involved . Pparα serves as an essential regulator of lipid metabolism, with gene ablation disrupting normal lipid homeostasis . Hepatic suppression of Pparα was also identified as a mechanism contributing to serum hypertriglyceridemia induced by a high-fructose diet . Additionally, reduced Fgf21 levels have been shown to decrease lipolysis .
In the current study, GSPE administration in the fructose-fed rats significantly decreased the expression of both Srebp1c and Scd1, indicating decreased TG synthesis, thereby contributing to reduced serum TG levels. We did not observe an increase in Srebp-1c following fructose consumption, likely due to the fact that the rats were fasted for 5 hours prior to sacrifice. It is well known that Srebp-1c is sensitively suppressed by fasting or nutritional deprivation [52, 53]. Therefore, fasting could have resulted in the lack of induction in the expression of Srebp-1c and its downstream targets, including Pgc-1β and Fasn, in addition to Mlxipl, in the fructose-fed animals. Despite no effect on Srebp1c, Fgf21 expression was reduced in the fructose group, indicating decreased lipolysis . Therefore, it is possible that repression of Fgf21 is the underlying mechanism by which fructose induced hypertriglyceridemia in these rats.
Reduced serum BA levels observed in the fructose-GSPE treated rats correlate with increased fecal BA output. Indeed, decreased intestinal BA absorption, combined with reduced hepatic lipogenesis, may be linked to the observed reduction in serum TG levels. Reduced BA absorption necessitates increased conversion of cholesterol into BAs. Since the rats consumed a cholesterol-free diet, increased endogenous cholesterol synthesis is necessary for the production of BAs. This is consistent with the observed increase in cholesterol synthesis gene expression observed in this study. Interestingly, there was no effect on Cyp7a1, which encodes cholesterol 7α-hydroxylase, the rate-limiting enzyme in BA biosynthesis. Cyp8b1, responsible for canonical BA synthesis, was decreased, with no changes in alternative BA biosynthetic gene expression, indicating that the newly synthesized cholesterol is not then shuttled into the pathway for BA production. In addition, the most readily available source of acetyl CoA for use in cholesterol synthesis would be from catabolized TGs, which provides an elegant explanation for the observed reduction in serum TGs, even in the presence of fructose.
GSPE administration profoundly increased total fecal lipid excretion, as well as BA excretion. It is known that hepatobiliary cholesterol excretion is not the only way to remove cholesterol from the body. The proximal part of the small intestine is now known to actively secrete cholesterol, via a pathway called transintestinal cholesterol efflux (TICE) . The rate of TICE strongly depends on the presence of a cholesterol acceptor, and increased levels of BAs in the intestinal lumen are known to increase TICE . When bile salts are combined with phospholipid, the TICE pathway is strongly stimulated . Therefore, increased levels of both BA and total lipid within the intestine could stimulate TICE and therefore contribute to enhanced fecal cholesterol excretion. Consequently, serum cholesterol levels remained the same due to an equilibrium being achieved between the rate of endogenous cholesterol synthesis and the amount secreted from the liver into the blood to be subsequently excreted via TICE. Importantly, and in agreement with this notion, the amount of cholesterol excreted in the feces exceeds the levels that could have originated from dietary intake.
In addition to decreased serum TG levels, GSPE treatment significantly decreased hepatic steatosis induced by fructose, as evidenced by the reduction in hepatic lipid droplet accumulation and cholesterol content. Steatosis by itself is considered to be a relatively benign and reversible condition . However, transition from steatosis into NASH represents a key step in pathogenesis, since it sets the stage for further liver damage, including development of fibrosis, cirrhosis and eventually hepatocellular carcinoma . Progression from simple steatosis to NASH usually involves a “second hit”, e.g. oxidative stress and inflammation, and previous studies have shown that accumulation of cholesterol, rather than fatty acids or triglycerides, is critical for this progression . Reduction in hepatic cholesterol content has been proposed as a fundamental treatment strategy for NAFLD . It may be speculated that one reason why the newly synthesized cholesterol is not converted to BAs, esterified or moved to the plasma membrane for export via the apolipoprotein/Abca1 pathway by GSPE in the presence of fructose, is because it is removed from the liver as a protective mechanism against further progression of steatosis.
In conclusion, this study provides valuable and innovative insight into the molecular regulatory actions of GSPE in treating hypertriglyceridemia in the fructose-fed rat model. This is achieved at the molecular level via GSPE-induced down-regulation in the hepatic expression of Srebp1c and Scd1, to decrease lipogenesis, combined with increased endogenous hepatic cholesterol synthesis, without any corresponding increase in de novo BA biosynthesis from the newly synthesized cholesterol in the fructose-fed rats. The novel observations resulting from this study demonstrate that, in the presence of fructose, GSPE alters the conversion of endogenously synthesized cholesterol, directing it through TICE for export via the feces. These results, combined with the decreased levels of steatosis, indicate that GSPE warrants further investigation as a treatment strategy against metabolic dysregulation and potential for amelioration of hepatic steatosis.
S1 Fig. HPLC analysis of the procyanidin composition in grape seed procyanidin extract (GSPE).
S2 Fig. Body Weight during the 8 week feeding study for (A) Control versus Fructose-fed rats, and (B) Fructose-Vehicle versus Fructose-GSPE treated rats.
The authors would like to thank Dr. Mike Teglas, DVM, Ph.D., Department of ANVS, for assistance with histological analysis of rat liver sections.
Conceived and designed the experiments: MLR. Performed the experiments: LED RMH GCC BSW KR FDR MLR. Analyzed the data: LED RMH GCC MLR. Wrote the paper: LED GCC MLR.
- 1. Ludwig DS, Peterson KE, Gortmaker SL. Relation between consumption of sugar-sweetened drinks and childhood obesity: a prospective, observational analysis. Lancet. 2001;357(9255):505–8. Epub 2001/03/07. pmid:11229668.
- 2. Montonen J, Jarvinen R, Knekt P, Heliovaara M, Reunanen A. Consumption of sweetened beverages and intakes of fructose and glucose predict type 2 diabetes occurrence. The Journal of nutrition. 2007;137(6):1447–54. Epub 2007/05/22. pmid:17513405.
- 3. Marchesini G, Brizi M, Bianchi G, Tomassetti S, Bugianesi E, Lenzi M, et al. Nonalcoholic fatty liver disease: a feature of the metabolic syndrome. Diabetes. 2001;50(8):1844–50. Epub 2001/07/27. pmid:11473047.
- 4. Marchesini G, Bugianesi E, Forlani G, Cerrelli F, Lenzi M, Manini R, et al. Nonalcoholic fatty liver, steatohepatitis, and the metabolic syndrome. Hepatology. 2003;37(4):917–23. Epub 2003/04/02. pmid:12668987.
- 5. Marriott BP, Cole N, Lee E. National Estimates of Dietary Fructose Intake Increased from 1977 to 2004 in the United States. Journal of Nutrition. 2009;139(6):S1228–S35. WOS:000266290500030.
- 6. Dekker MJ, Su Q, Baker C, Rutledge AC, Adeli K. Fructose: a highly lipogenic nutrient implicated in insulin resistance, hepatic steatosis, and the metabolic syndrome. American journal of physiology Endocrinology and metabolism. 2010;299(5):E685–94. Epub 2010/09/09. pmid:20823452.
- 7. Stanhope KL, Schwarz JM, Havel PJ. Adverse metabolic effects of dietary fructose: results from the recent epidemiological, clinical, and mechanistic studies. Current opinion in lipidology. 2013;24(3):198–206. WOS:000319446300003. pmid:23594708
- 8. Chong MFF, Fielding BA, Frayn KN. Mechanisms for the acute effect of fructose on postprandial lipemia. American Journal of Clinical Nutrition. 2007;85(6):1511–20. WOS:000247196300011. pmid:17556686
- 9. Samuel VT. Fructose induced lipogenesis: from sugar to fat to insulin resistance. Trends in endocrinology and metabolism: TEM. 2011;22(2):60–5. Epub 2010/11/12. pmid:21067942.
- 10. Kastorini CM, Milionis HJ, Esposito K, Giugliano D, Goudevenos JA, Panagiotakos DB. The effect of Mediterranean diet on metabolic syndrome and its components: a meta-analysis of 50 studies and 534,906 individuals. Journal of the American College of Cardiology. 2011;57(11):1299–313. Epub 2011/03/12. pmid:21392646.
- 11. Hertog MG, Feskens EJ, Hollman PC, Katan MB, Kromhout D. Dietary antioxidant flavonoids and risk of coronary heart disease: the Zutphen Elderly Study. Lancet. 1993;342(8878):1007–11. Epub 1993/10/23. pmid:8105262.
- 12. Hertog MG, Kromhout D, Aravanis C, Blackburn H, Buzina R, Fidanza F, et al. Flavonoid intake and long-term risk of coronary heart disease and cancer in the seven countries study. Archives of internal medicine. 1995;155(4):381–6. Epub 1995/02/27. pmid:7848021.
- 13. Rasmussen SE, Frederiksen H, Struntze Krogholm K, Poulsen L. Dietary proanthocyanidins: occurrence, dietary intake, bioavailability, and protection against cardiovascular disease. Molecular nutrition & food research. 2005;49(2):159–74. Epub 2005/01/07. pmid:15635686.
- 14. Caimari A, Del Bas JM, Crescenti A, Arola L. Low doses of grape seed procyanidins reduce adiposity and improve the plasma lipid profile in hamsters. International journal of obesity. 2012. Epub 2012/05/16. pmid:22584454.
- 15. Osakabe N. Flavan 3-ols improve metabolic syndrome risk factors: evidence and mechanisms. Journal of clinical biochemistry and nutrition. 2013;52(3):186–92. Epub 2013/05/25. pmid:23704807; PubMed Central PMCID: PMC3652297.
- 16. Auger C, Gerain P, Laurent-Bichon F, Portet K, Bornet A, Caporiccio B, et al. Phenolics from commercialized grape extracts prevent early atherosclerotic lesions in hamsters by mechanisms other than antioxidant effect. Journal of agricultural and food chemistry. 2004;52(16):5297–302. Epub 2004/08/05. pmid:15291511.
- 17. Zern TL, Wood RJ, Greene C, West KL, Liu Y, Aggarwal D, et al. Grape polyphenols exert a cardioprotective effect in pre- and postmenopausal women by lowering plasma lipids and reducing oxidative stress. The Journal of nutrition. 2005;135(8):1911–7. Epub 2005/07/28. pmid:16046716.
- 18. Del Bas JM, Fernandez-Larrea J, Blay M, Ardevol A, Salvado MJ, Arola L, et al. Grape seed procyanidins improve atherosclerotic risk index and induce liver CYP7A1 and SHP expression in healthy rats. FASEB journal: official publication of the Federation of American Societies for Experimental Biology. 2005;19(3):479–81. Epub 2005/01/08. pmid:15637110.
- 19. Del Bas JM, Ricketts ML, Baiges I, Quesada H, Ardevol A, Salvado MJ, et al. Dietary procyanidins lower triglyceride levels signaling through the nuclear receptor small heterodimer partner. Molecular nutrition & food research. 2008;52(10):1172–81. Epub 2008/08/23. pmid:18720348.
- 20. Quesada H, del Bas JM, Pajuelo D, Diaz S, Fernandez-Larrea J, Pinent M, et al. Grape seed proanthocyanidins correct dyslipidemia associated with a high-fat diet in rats and repress genes controlling lipogenesis and VLDL assembling in liver. International journal of obesity. 2009;33(9):1007–12. Epub 2009/07/08. pmid:19581912.
- 21. Quesada H, Diaz S, Pajuelo D, Fernandez-Iglesias A, Garcia-Vallve S, Pujadas G, et al. The lipid-lowering effect of dietary proanthocyanidins in rats involves both chylomicron-rich and VLDL-rich fractions. The British journal of nutrition. 2012;108(2):208–17. Epub 2011/10/21. pmid:22011563.
- 22. Del Bas JM, Ricketts ML, Vaque M, Sala E, Quesada H, Ardevol A, et al. Dietary procyanidins enhance transcriptional activity of bile acid-activated FXR in vitro and reduce triglyceridemia in vivo in a FXR-dependent manner. Molecular nutrition & food research. 2009;53(7):805–14. Epub 2009/06/06. pmid:19496086.
- 23. Watanabe M, Houten SM, Wang L, Moschetta A, Mangelsdorf DJ, Heyman RA, et al. Bile acids lower triglyceride levels via a pathway involving FXR, SHP, and SREBP-1c. The Journal of clinical investigation. 2004;113(10):1408–18. Epub 2004/05/18. pmid:15146238; PubMed Central PMCID: PMC406532.
- 24. Ma K, Saha PK, Chan L, Moore DD. Farnesoid X receptor is essential for normal glucose homeostasis. The Journal of clinical investigation. 2006;116(4):1102–9. Epub 2006/03/25. pmid:16557297; PubMed Central PMCID: PMC1409738.
- 25. Duran-Sandoval D, Mautino G, Martin G, Percevault F, Barbier O, Fruchart JC, et al. Glucose regulates the expression of the farnesoid X receptor in liver. Diabetes. 2004;53(4):890–8. Epub 2004/03/30. pmid:15047603.
- 26. Sinal CJ, Tohkin M, Miyata M, Ward JM, Lambert G, Gonzalez FJ. Targeted disruption of the nuclear receptor FXR/BAR impairs bile acid and lipid homeostasis. Cell. 2000;102(6):731–44. Epub 2000/10/13. pmid:11030617.
- 27. Houten SM, Volle DH, Cummins CL, Mangelsdorf DJ, Auwerx J. In vivo imaging of farnesoid X receptor activity reveals the ileum as the primary bile acid signaling tissue. Molecular endocrinology. 2007;21(6):1312–23. Epub 2007/04/12. pmid:17426284.
- 28. Terra X, Montagut G, Bustos M, Llopiz N, Ardevol A, Blade C, et al. Grape-seed procyanidins prevent low-grade inflammation by modulating cytokine expression in rats fed a high-fat diet. The Journal of nutritional biochemistry. 2009;20(3):210–8. Epub 2008/07/08. pmid:18602813.
- 29. Chan TF, Lin WT, Huang HL, Lee CY, Wu PW, Chiu YW, et al. Consumption of sugar-sweetened beverages is associated with components of the metabolic syndrome in adolescents. Nutrients. 2014;6(5):2088–103. Epub 2014/05/27. pmid:24858495; PubMed Central PMCID: PMC4042561.
- 30. Reaven GM, Ho H. Sugar-induced hypertension in Sprague-Dawley rats. American journal of hypertension. 1991;4(7 Pt 1):610–4. Epub 1991/07/01. pmid:1873016.
- 31. Kelm MA, Johnson JC, Robbins RJ, Hammerstone JF, Schmitz HH. High-performance liquid chromatography separation and purification of cacao (Theobroma cacao L.) procyanidins according to degree of polymerization using a diol stationary phase. Journal of agricultural and food chemistry. 2006;54(5):1571–6. Epub 2006/03/02. pmid:16506802.
- 32. Shrestha S, Ehlers SJ, Lee JY, Fernandez ML, Koo SI. Dietary green tea extract lowers plasma and hepatic triglycerides and decreases the expression of sterol regulatory element-binding protein-1c mRNA and its responsive genes in fructose-fed, ovariectomized rats. The Journal of nutrition. 2009;139(4):640–5. Epub 2009/02/06. pmid:19193814; PubMed Central PMCID: PMC2666357.
- 33. Masterjohn C, Park Y, Lee J, Noh SK, Koo SI, Bruno RS. Dietary fructose feeding increases adipose methylglyoxal accumulation in rats in association with low expression and activity of glyoxalase-2. Nutrients. 2013;5(8):3311–28. Epub 2013/08/24. pmid:23966111; PubMed Central PMCID: PMC3775256.
- 34. Yamakoshi J, Saito M, Kataoka S, Kikuchi M. Safety evaluation of proanthocyanidin-rich extract from grape seeds. Food and chemical toxicology: an international journal published for the British Industrial Biological Research Association. 2002;40(5):599–607. Epub 2002/04/17. pmid:11955665.
- 35. Rucker R, Storms D. Interspecies comparisons of micronutrient requirements: metabolic vs. absolute body size. The Journal of nutrition. 2002;132(10):2999–3000. Epub 2002/10/09. pmid:12368385.
- 36. Clifton PM. Effect of Grape Seed Extract and Quercetin on Cardiovascular and Endothelial Parameters in High-Risk Subjects. Journal of biomedicine & biotechnology. 2004;2004(5):272–8. Epub 2004/12/04. pmid:15577189; PubMed Central PMCID: PMC1082891.
- 37. Untergasser A, Nijveen H, Rao X, Bisseling T, Geurts R, Leunissen JA. Primer3Plus, an enhanced web interface to Primer3. Nucleic acids research. 2007;35(Web Server issue):W71–4. Epub 2007/05/09. pmid:17485472; PubMed Central PMCID: PMC1933133.
- 38. Aguila MB, Pinheiro Ada R, Parente LB, Mandarim-de-Lacerda CA. Dietary effect of different high-fat diet on rat liver stereology. Liver international: official journal of the International Association for the Study of the Liver. 2003;23(5):363–70. Epub 2004/01/08. pmid:14708898.
- 39. Folch J, Lees M, Sloane Stanley GH. A simple method for the isolation and purification of total lipides from animal tissues. The Journal of biological chemistry. 1957;226(1):497–509. Epub 1957/05/01. pmid:13428781.
- 40. Modica S, Murzilli S, Moschett A. Characterizing Bile Acid and Lipid Metabolism in the Liver and Gastrointestinal Tract of Mice. Current Protocols in Mouse Biology. 2011;1:289–321. pmid:26069056
- 41. Nagai Y, Yonemitsu S, Erion DM, Iwasaki T, Stark R, Weismann D, et al. The role of peroxisome proliferator-activated receptor gamma coactivator-1 beta in the pathogenesis of fructose-induced insulin resistance. Cell metabolism. 2009;9(3):252–64. Epub 2009/03/04. pmid:19254570; PubMed Central PMCID: PMC3131094.
- 42. Roglans N, Sanguino E, Peris C, Alegret M, Vazquez M, Adzet T, et al. Atorvastatin treatment induced peroxisome proliferator-activated receptor alpha expression and decreased plasma nonesterified fatty acids and liver triglyceride in fructose-fed rats. The Journal of pharmacology and experimental therapeutics. 2002;302(1):232–9. Epub 2002/06/18. pmid:12065722.
- 43. Inagaki T, Dutchak P, Zhao G, Ding X, Gautron L, Parameswara V, et al. Endocrine regulation of the fasting response by PPARalpha-mediated induction of fibroblast growth factor 21. Cell metabolism. 2007;5(6):415–25. Epub 2007/06/07. pmid:17550777.
- 44. Lundasen T, Hunt MC, Nilsson LM, Sanyal S, Angelin B, Alexson SE, et al. PPARalpha is a key regulator of hepatic FGF21. Biochemical and biophysical research communications. 2007;360(2):437–40. Epub 2007/07/03. pmid:17601491.
- 45. Badman MK, Pissios P, Kennedy AR, Koukos G, Flier JS, Maratos-Flier E. Hepatic fibroblast growth factor 21 is regulated by PPARalpha and is a key mediator of hepatic lipid metabolism in ketotic states. Cell metabolism. 2007;5(6):426–37. Epub 2007/06/07. pmid:17550778.
- 46. Oram JF. HDL apolipoproteins and ABCA1: partners in the removal of excess cellular cholesterol. Arteriosclerosis, thrombosis, and vascular biology. 2003;23(5):720–7. Epub 2003/03/05. pmid:12615680.
- 47. Moutzouri E, Kei A, Elisaf MS, Milionis HJ. Management of dyslipidemias with fibrates, alone and in combination with statins: role of delayed-release fenofibric acid. Vascular health and risk management. 2010;6:525–39. Epub 2010/08/24. pmid:20730069; PubMed Central PMCID: PMC2922314.
- 48. Birjmohun RS, Hutten BA, Kastelein JJ, Stroes ES. Efficacy and safety of high-density lipoprotein cholesterol-increasing compounds: a meta-analysis of randomized controlled trials. Journal of the American College of Cardiology. 2005;45(2):185–97. Epub 2005/01/18. pmid:15653014.
- 49. van der Velde AE, Brufau G, Groen AK. Transintestinal cholesterol efflux. Current opinion in lipidology. 2010;21(3):167–71. Epub 2010/04/23. pmid:20410820.
- 50. Kelley GL, Azhar S. Reversal of High dietary fructose-induced PPARα suppression by oral administration of lipoxygenase/cyclooxygenase inhibitors. Nutr Metab (Lond). 2. London2005. p. 18.
- 51. Lee SS, Pineau T, Drago J, Lee EJ, Owens JW, Kroetz DL, et al. Targeted disruption of the alpha isoform of the peroxisome proliferator-activated receptor gene in mice results in abolishment of the pleiotropic effects of peroxisome proliferators. Molecular and cellular biology. 1995;15(6):3012–22. Epub 1995/06/01. pmid:7539101; PubMed Central PMCID: PMC230532.
- 52. Horton JD, Bashmakov Y, Shimomura I, Shimano H. Regulation of sterol regulatory element binding proteins in livers of fasted and refed mice. Proceedings of the National Academy of Sciences of the United States of America. 1998;95(11):5987–92. Epub 1998/05/30. pmid:9600904; PubMed Central PMCID: PMC27572.
- 53. Kim JB, Sarraf P, Wright M, Yao KM, Mueller E, Solanes G, et al. Nutritional and insulin regulation of fatty acid synthetase and leptin gene expression through ADD1/SREBP1. The Journal of clinical investigation. 1998;101(1):1–9. Epub 1998/02/14. pmid:9421459; PubMed Central PMCID: PMC508533.
- 54. van der Velde AE, Vrins CL, van den Oever K, Kunne C, Oude Elferink RP, Kuipers F, et al. Direct intestinal cholesterol secretion contributes significantly to total fecal neutral sterol excretion in mice. Gastroenterology. 2007;133(3):967–75. Epub 2007/09/15. pmid:17854600.
- 55. van der Velde AE, Vrins CL, van den Oever K, Seemann I, Oude Elferink RP, van Eck M, et al. Regulation of direct transintestinal cholesterol excretion in mice. American journal of physiology Gastrointestinal and liver physiology. 2008;295(1):G203–G8. Epub 2008/05/31. pmid:18511744.
- 56. Wouters K, van Gorp PJ, Bieghs V, Gijbels MJ, Duimel H, Lutjohann D, et al. Dietary cholesterol, rather than liver steatosis, leads to hepatic inflammation in hyperlipidemic mouse models of nonalcoholic steatohepatitis. Hepatology. 2008;48(2):474–86. Epub 2008/07/31. pmid:18666236.
- 57. Fernandez A, Colell A, Garcia-Ruiz C, Fernandez-Checa JC. Cholesterol and sphingolipids in alcohol-induced liver injury. Journal of gastroenterology and hepatology. 2008;23 Suppl 1:S9–15. Epub 2008/04/11. pmid:18336673.
- 58. Enjoji M, Nakamuta M. Is the control of dietary cholesterol intake sufficiently effective to ameliorate nonalcoholic fatty liver disease? World journal of gastroenterology: WJG. 2010;16(7):800–3. Epub 2010/02/10. pmid:20143458; PubMed Central PMCID: PMC2825326.