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4 Center for Human Nutrition, University of Texas Southwestern Medical Center, Dallas, TX 75390-9052; 5 Department of Biomedical Engineering, University of Texas at Arlington, Arlington, TX 76019-0018; and 6 Department of Food Science and Nutrition, University of Minnesota, St. Paul, MN 55108
* To whom correspondence should be addressed. E-mail: elizabeth.parks{at}utsouthwestern.edu.
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ABSTRACT |
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 TOP ABSTRACT IntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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Introduction |
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TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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Materials and Methods |
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TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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Study design. Three different CHO solutions were tested to determine the effect of sequential replacement of glucose by fructose on the stimulation of hepatic lipogenesis. These solutions included a control bolus of 100% glucose and 0% fructose (hereafter denoted 100:0), a weight-for-weight mixture of 50% glucose:50% fructose (50:50), and one of 25% glucose:75% fructose (25:75). Information regarding solution preparation and quantity of CHO fed in the tests is described in detail below. For 3 d prior to each metabolic test, subjects were instructed to consume a weight-maintaining diet of constant energy based on the Harris-Benedict equation (18) with comparison to 3-d food records of usual intake recorded during screening. Following directions from a registered dietitian, pretest meals were prepared by the subjects themselves and consisted of whole foods. Because the recent energy balance of the subject can significantly affect fatty acid metabolism (19), the goal of the pretest diet was to have it be of known composition and yet match the subject's background diet, providing 53% of total energy from CHO, 32% from fat, and 15% from protein. The actual mean profile of the 3-d prestudy diet was 8277 ± 819 kJ/d with 52 ± 3% of total energy from CHO (23 ± 6% of total energy from sugars), 31 ± 1% from fat, and 17 ± 3% from protein, and included 19 ± 5 g of fiber and 1.1 ± 0.5 g/d of (n-3) fatty acids. Except for the different CHO solutions, the protocol for each metabolic test was the same (Fig. 1).
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51% of their total daily energy needs.
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To prepare for the protocol shown in Figure 1, on the morning of each test, the CHO and water were combined with unsweetened Kool-Aid lemonade mix and the solution was served ice cold to increase palatability. Subjects were given 15 min to consume the 8 ounces of liquid. Indirect calorimetry was performed for 30 min between 0630 to 0730 (fasting state) and 0900 to 0930 (1 h post-CHO bolus) using a Deltatrac II Metabolic Cart (Sensor Medix) in the hooded mode. Subjects rested, watched TV, or read during the study. Nonenergy containing, noncaffeinated drinks were available upon request. Starting at 0740 and continuing until 1800, blood samples were drawn and serum separated immediately by centrifugation (3000 x g; 10 min, 20°C). Serum samples were kept on ice, a preservative cocktail was added (22), and samples were divided into aliquots for glucose, insulin, NEFA, serum TG analysis, and lipoprotein isolation.
Lipoprotein isolation. The CHO bolus was consumed at time 0 and the lunch was consumed at time 4.33 h. Total TG-rich lipoproteins (tTRL) were isolated from serum (23) at 12 time points throughout d 2 including: –0.08, 0.92, 2.25, 2.75, 3.25, 4.25, 4.58, 5.75, 7.25, 8.25, 9.42, and 10.25 h. In the morning, this lipoprotein fraction contained particles predominantly of hepatic origin (VLDL), because only sugars had been fed in the morning meal. By contrast, the lunch contained fat and, therefore, the postlunch tTRL fraction included both chylomicrons and VLDL. For 6 of the time points after lunch (4.25, 4.58, 5.75, 7.25, 8.25, and 9.42 h), an additional procedure (24) was used to isolate lipoproteins with a Svedberg flotation rate (Sf) of >400, which includes primarily chylomicrons with small quantities of very large VLDL and Sf 60–400, which includes large VLDL with a very short half-life (24). After lunch, the apoB100 concentrations in the fraction Sf 60–400 were determined by SDS-PAGE (22). The mol:mol ratio of TG:apoB100 was calculated using molecular weights of TG and apoB100 of 850 and 530,000, respectively.
Analysis of metabolic variables and GC-MS. Serum glucose concentrations were measured on a Vitros Analyzer 950 (Ortho-Clinical Diagnostics), whereas insulin was determined via chemiluminescent immunoassay (Diagnostic Products). Concentrations of NEFA (kit no. 994 79405E, Wako Chemicals) and serum TG in tTRL and VLDL fractions were determined via enzymatic assays (994–40491 Wako Chemicals) on a microtiter spectrophotometer (Model EL 340 Microplate, Bio-Tek Instruments). Serum leptin and adiponectin were measured using radioimmunoassay and gastric inhibitory polypeptide was measured using ELISA (Linco kits number no. HL-81HK, HADP-61HK, and EZHGIP-54K, respectively, Linco). To prepare for GC-MS, TG in tTRL and VLDL were separated and fatty acids derivatized to FAME as described previously (24). GC was performed to determine fatty acid composition on an Agilent Technologies 6890 GC fitted with a flame ionization detector with a split injection of 20:1 (25). tTRL- and VLDL-TG FAME were separated on a SP-2560 column (Sigma-Aldrich), 100-m x 0.25-mm i.d. x 0.20-µm film thickness with helium as a carrier gas. GC-MS was performed using a DB-225, 30-m column (250-mm i.d. and 0.25-µm film thickness, J&W, Chromtech) in an Agilent 6890 GC with helium as the carrier gas. Selected ion monitoring was used for ions with mass:charge ratios of 270, 271, 272, which were analyzed with an Agilent 5975 mass spectrometer. Newly made fatty acids from de novo lipogenesis were calculated by mass isotopomer distribution analysis (MIDA) (26).
Calculations and statistical analysis. The 16-carbon fatty acid, palmitate, was used as the indicator of changes in lipogenesis because it represents the primary product of the fatty acid synthesis pathway (27). Lipogenesis data are presented as the fractional lipogenic rate, which is derived from the MIDA calculation, and also as the absolute lipogenic rate, which was calculated as follows. At each time point, the concentration of lipoprotein-TG fatty acids (in µmol/L) derived from an enzymatic TG assay was multiplied by the molar percentage of fatty acids that were palmitate in that TG, measured by GC analysis of TG-FAME. This quantity was then multiplied by the percentage of TG-FAME palmitate derived from de novo lipogenesis, as measured from GC-MS using the MIDA calculation (26).
The Sf > 400 lipogenesis data were analyzed as the absolute area under the curve (AUC), because these values were decaying throughout the postlunch phase. All other metabolic data (glucose, NEFA, insulin, serum TG, tTRL-TG, and VLDL-TG) were analyzed by incremental AUC (iAUC). Data are presented relative to the fasting value for all areas during the morning post-CHO ingestion, postlunch, and for the entire day (total iAUC = post-CHO + postlunch areas). In addition, to assess the isolated effect of lunch on changes in metabolite concentrations, the iAUC after lunch was also calculated relative to the lowest value occurring at the start of the lunch. In the tables, this iAUC is denoted as the value "reset at lunch" and serves to limit the effect of any carryover of elevated concentration as a result of the morning CHO feeding. Statistical analyses were performed using Statview for Windows (version 5.0.1, SAS Institute). Unless otherwise noted, values in the text are means ± SEM. A P-value of < 0.05 was considered significant. P < 0.07 are also noted, given the sample size of 6. In addition to AUC analysis, differences between treatments were analyzed using repeated-measures ANOVA. The Bonferroni/Dunn test was used for post hoc analysis.
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Results |
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TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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For tTRL-TG, the post-CHO concentration tended to be higher after 50:50 compared with 100:0 (Fig. 3A; Table 3). Similar to the serum TG concentration, tTRL-TG after 100:0 was associated with a negative iAUC. The postlunch tTRL-TG iAUC for 25:75 was >50:50 and both of these were greater than after 100:0. Resetting the baseline at lunch demonstrated that 25:75 remained significantly >50:50. For the total day, 25:75 had a significantly greater iAUC for tTRL-TG compared with 100:0. The percentage of tTRL-TG fatty acids that were palmitic acid (16:0, our marker of a lipogenic fatty acid) remained constant throughout the day (Supplemental Fig. 1). Palmitoleic acid (16:1) may be a marker of lipogenesis with prolonged feeding of high CHO diets, because it has been shown to be higher in serum TG during eucaloric CHO feeding for 25 d (28), during 4 d of overfeeding (27), and lower in blood TG following 12 wk of dietary CHO restriction (29). In this study, the percentage of palmitoleic did not change in the morning and decreased after consumption of the meal (Supplemental Fig. 1).
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Because the meal fed at noon contained 32% of energy from fat, chylomicrons would be found in the tTRL fraction after lunch and dilute the apparent lipogenic rate. To determine the de novo production of fatty acids from the liver, lipoproteins in the Sf > 400 were separated from Sf 60–400 in the tTRL fraction at 6 time points after lunch. The percentage of de novo fatty acids evident in the Sf > 400 fraction (Fig. 4B) at 4 h averaged 3–8% and decreased throughout the afternoon, resulting in a small, absolute quantity of de novo fatty acids in Sf > 400 lipoproteins. Early after lunch for all of the treatments, the fractional hepatic lipogenic rate (Fig. 4E) decreased after the morning stimulation and increased again postlunch after a short delay, resulting in similar postlunch absolute lipogenesis rates in VLDL between the treatments (Fig. 4F; Table 3). After lunch, the apoB100 content in the Sf 60–400 lipoprotein fraction did not differ between the tests (21 ± 11, 22 ± 7, and 56 ± 35 nmol/L for 100:0, 50:50, and 25:75, respectively; P < 0.161), nor did the amount of TG per particle in these fractions differ (60,516 ± 26,245, 33,831 ± 6340, and 33,986 ± 165,61 mol TG/mol apoB, respectively; P < 0.225).
Fractional lipogenesis values from each individual's 3 tests were analyzed to assess the repeatability of this measurement within a single subject (Supplemental Table 1). Among the subjects, the fasting lipogenesis varied from 0.0 to 15.3%; however, the within-subject SD was low (0.9%). The results of respiratory gas measurements revealed that for all treatments, compared with fasting, post-CHO respiratory quotients were elevated, rising from 0.77 ± 0.02 to 0.88 ± 0.03 1 h after the CHO bolus (P < 0.03; data not shown). CHO oxidation increased (P < 0.02) and fat oxidation decreased (P < 0.04) to the same extent after all treatments. Serum concentrations of the hormones, gastric-inhibitory peptide, leptin, and adiponectin did not differ between the treatments (Supplemental Fig. 2), likely as a result of the limited number of subjects.
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Discussion |
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TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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In this study, after the 100:0 glucose bolus, fractional lipogenesis rose to only 8%, which is quite low compared with the 23% observed previously (32). After consumption of the 50:50 and 25:75 solutions used here, the peak in lipogenesis was 17%. These "meals" were liquid in nature and the timing of the serum glucose excursion curves in the present research suggests that the solutions were all readily absorbed. The lower values of de novo lipogenesis observed here with 85 g glucose compared with our previous study's 112-g dose resulted from the lower amount of CHO fed and from the absence of fat and protein in the solutions used herein (i.e. less total energy in the meal). Thus, one conclusion from these studies is that lipogenesis is only slightly increased after a glucose bolus similar to that routinely used for a medical oral glucose tolerance test. Second, the timing of the morning peak in fractional lipogenesis (Fig. 2, 3–4 h across all treatments) was consistent with past literature (32,34) and shows that this delay occurs whether fat is fed in the meal or not, as in the present study. Third, fructose significantly stimulated lipogenesis. By contrast, after lunch the quantities of newly made fatty acids were similar between the treatments, consistent with a set amount of fatty acids being made from the constant amount of CHO fed in this meal.
A 4th observation made here was constant serum and tTRL-TG concentrations during the first 4 h after fructose at either dose, compared with a slight reduction in TG following 100:0 (Figs. 2D and 3A). Whether this difference was due to increased clearance of blood TG following 100:0 and/or maintenance of TG secretion with fructose cannot be definitively evaluated by the present study design. However, the greater fractional lipogenic rates in the morning after fructose suggest that elevated secretion of TG was the mechanism. Focusing on the postlunch metabolism, when a meal was fed after the 100:0, the fractional lipogenic rate rose (Fig. 3B) to yield equivalent absolute de novo fatty acids postlunch compared with the other treatments (Fig. 3C). Yet, this stimulation was not sufficient to bring the postlunch serum TG and tTRL-TG concentrations up to the level of the 25:75 test (Figs. 2D and 3A). The data show that fructose acutely and significantly increased lipogenesis, which served to maintain TG concentrations in the morning and also led to greater TG concentrations after lunch. Further, the effect after lunch was greater than could be accounted for by the slight elevation in the morning. This suggests that the morning rise in de novo lipogenesis led to increased use of other fatty acid sources (plasma NEFA, dietary or stored TG) for TG synthesis after lunch. In rodent liver perfusate experiments, fructose increased fatty acid esterification (33) and a recent study by Chong et al. (16) provided direct support of this hypothesis in humans. In this latter report, the authors did not show a significant increase in absolute fatty acid production after the bolus of fructose. However, the lipogenic pathway was labeled only during breakfast consumption via the addition of labeled fructose added to a sweet drink. This bolus-labeling scheme would not have allowed enough time for isotopic labeling of fatty acids used for subsequent VLDL-TG synthesis. It can take anywhere from 12 to 36 h of labeling to be able to accurately identify newly made fatty acids in VLDL, because de novo fatty acids enter a delay pool before being used for VLDL-TG synthesis (34). Furthermore, it appears that the consumption of a meal facilitates the movement of these fatty acids through this cytosolic delay pool (Supplemental Fig. 3).
The observed significant increase in fractional lipogenesis during the morning after fructose is not likely a mathematical result of a reduction in the liver's use of fatty acids derived from the plasma NEFA pool. This is because the concentration of tTRL-TG remained unchanged in the morning (iAUC were close to 0) and NEFA concentrations decreased after all 3 treatments, whereas the pattern of change in lipogenesis varied among the tests. In previous studies, we demonstrated that the reduction in use of plasma NEFA for VLDL-TG synthesis occurs extremely fast and on a different timeline than the rise in de novo lipogenesis (Supplemental Fig. 1). Furthermore, late in the postprandial period (4 h), the reappearance of plasma NEFA in VLDL-TG takes place at the same time as the rise in lipogenesis (i.e. the 2 are not reciprocal). The early postprandial reduction in the use of plasma NEFA for VLDL-TG synthesis coincides with an appearance of unlabeled fatty acids in the particle (20,32), which could originate from a liver TG storage pool. We speculate that the liver calls upon stored TG to support and maintain VLDL-TG secretion postprandially when a significant decrease in the flow of plasma NEFA to liver has occurred.
It appears that de novo fatty acid synthesis does not result in the addition of excess TG fatty acid that are added to VLDL. Rather, their presence may prevent apoB degradation, resulting in more particles being secreted that are smaller in size, as suggested by the apoB data. Given the constant tTRL-TG concentrations across treatments, it is possible that a fructose-induced increase in lipogenesis displaces the use of stored TG for VLDL synthesis, or, more likely, that the stimulation of lipogenesis represents an intracellular signal for liver to esterify fatty acids from any source into TG. This effect may also depend on the sensitivity of the liver to insulin. If fructose does increase hepatic TG storage, fructose feeding could lead to fatty liver. Using magnetic resonance spectrometry, Le et al. (10) found no increase in hepatic TG stores in healthy men eucalorically consuming a high-fructose diet for 4 wk. Interestingly, in this article, the greater the serum TG concentration, the lower the level of hepatic TG post-treatment and the authors hypothesized that "exportation of newly formed TG as VLDL-TG is a key element to prevent liver-TG stores from expanding" (10). Our data support this concept.
The large increase in de novo lipogenesis postlunch after 100:0, which had barely stimulated lipogenesis in the morning, resulted in postlunch lipogenic rates for all treatments being similar. This observation suggests that under healthy, well-regulated metabolism, there may be a maximum rate of lipogenesis that can be achieved given the quantity of the CHO fed or a maximum rate depending on the time of day. This robust increase in de novo lipogenesis postprandially in healthy subjects, in contrast to the lack of an observed increase in obese patients with fatty liver (35), suggests that a significant increase in fatty acid synthesis after CHO consumption can be used as one characteristic to test for metabolic flexibility (36,37).
In an overfeeding study, Lammert et al. (14) noted that within their group of research subjects, the same individuals repeatedly had the highest de novo lipogenesis values, no matter which diet was fed chronically (eucaloric, or excess energy from fat or CHO). Here, we were also able to demonstrate good reproducibility in the fasting lipogenic rate of individual subjects during the repeated tests and we think this was due to the relatively tight control of the subjects' food and alcohol intakes and activity levels before the tests. This fact will support future studies focusing on environmental and genetic factors that predict elevated fatty acid synthesis postprandially so as to investigate how lipogenesis may contribute to atherogenic risk during positive energy balance in obese and insulin-resistant subjects.
Strengths of this study include the blinded and random order of the treatments and adjustment for the quantity of CHO fed depending on the subjects' total daily energy needs. The key limitation of the study is a small sample size. However, the repeated measures design supports the notion that the differences observed among these 18 metabolic tests were real and would be reproducible. Another limitation may lie in the fact that the subjects consumed the CHO solutions in the fasted state, which would underestimate lipogenesis for 3 reasons. First, as described earlier, to achieve maximal labeling of lipogenesis a full day of acetate infusion may be needed (34,38); second, palmitate was used as the marker of lipogenesis when other fatty acids (16:1, 18:1) may have been made de novo but were not specifically tested; and third, at this time of day, much of the CHO consumed would be burned as energy and/or go to refill hepatic glycogen stores (6,33). Serving the beverages in the morning in the fasted state, did, however, isolate the effect of the CHO from the effects of excess energy intake, a confounder in overfeeding studies. Further, the consumption of a sweetened beverage in lieu of breakfast is not unheard of in real life.
In summary, these data show a significant immediate lipogenic effect of fructose in healthy, lean research subjects that was associated with greater serum TG concentrations in the morning and after a subsequent meal. The data suggest that an early stimulation of lipogenesis after fructose, consumed in the fasted state (even as a small amount in a mixture of sugars) augments subsequent postprandial lipemia. The additional postlunch elevation in TG was only partially due to a carry over from the morning, suggesting that fructose consumption creates a metabolic milieu that alters subsequent handling of nutrients flowing to the liver. This study, together with the strong, recent data presented by Chong et al. (16), provide compelling evidence that fructose causes hyperlipidemia postprandially both directly, through the synthesis of fatty acids, and indirectly, by increasing liver reesterification of fatty acids from all sources.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Author disclosures: E. J. Parks, L. E. Skokan, M. T. Timlin, and C. S. Dingfelder, no conflicts of interest.
3 Supplemental Table 1 and Supplemental Figures 1–3 are available with the online posting of this paper at jn.nutrition.org.
7 Abbreviations used: AUC, area under the curve; CHO, carbohydrate; iAUC, incremental area under the curve; GCRC, General Clinical Research Center; MIDA, mass isotopomer distribution analysis; Sf, Svedberg flotation rate; TG, triacylglycerol; tTRL, total triacylglycerol-rich lipoprotein; 100:0: a mixture of 100% glucose and 0% fructose wt:wt; 75:25: 75:25 glucose: fructose wt:wt; 50:50: 50:50 glucose:fructose wt:wt.
Manuscript received 9 October 2007. Initial review completed 9 November 2007. Revision accepted 28 February 2008.
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LITERATURE CITED |
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TOPABSTRACTIntroductionMaterials and MethodsResultsDiscussionLITERATURE CITED |
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