The effect of chronic exercise on reducing body fat and obesity has been well established (2). Previous work has primarily focused on the benefit of exercise in promoting fat oxidation during aerobic exercise (23). However, there is evidence that exercise may also down-regulate hepatic lipogenic enzymes, thereby reducing the availability of long-chain fatty acids required for synthesis of triglyceride (9,12,29). This anti-lipogenic effect of training is especially important when the diet is low in fat and high in simple sugars such as glucose and fructose, which have been shown to promote lipogenic enzyme induction and obesity (14).
In mammalian species, de novo lipogenesis in the liver and adipose tissue is controlled by lipogenic enzymes, especially fatty acid synthase (FAS), the rate-limiting enzyme for hepatic fatty acid synthesis (14). Insulin is the major hormone promoting hepatic lipogenesis via the stimulation of FAS and other lipogenic enzymes (14). The primary mechanism for FAS induction is transcriptional activation of the FAS gene in the liver (20). Furthermore, this transcriptional activation by insulin has been shown to result from the binding of transcription factors to the gene regulatory elements on the FAS promoter region, such as insulin response sequence (IRS/A), carbohydrate (CHO) response elements (ChoRE), and inverted CCAAT-box (ICE) (11,18,28).
A commonly used animal model to study obesity and insulin independent diabetes mellitus (NIDDM) is the obese Zucker rat (OZR) (3). The affected rats were both hyperinsulinemic and hyperlipidemic, inherited as an autosomal recessive trait located within the rat ob-receptor (3). One reason for the abnormal lipid profiles in OZR is the excessive rate of lipogenesis (1,26). It is estimated that the rate of fatty acid synthesis in adipose tissue and liver was 10-fold greater in 5-wk-old OZR than their lean litter mates. The ratio of total fatty acid synthesized in liver versus adipose tissue was 10.8 and 5.9 in 13-wk-old OZR and lean rats, respectively, indicating a predominant role for hepatic lipogenesis. Furthermore, lipogenic enzyme activities are dramatically elevated in OZR, which also appeared to be more responsive to high-CHO diet feeding (3). Interestingly, the inhibitory effects of food restriction on hepatic FAS activity were not found in OZR (26). These results indicate that OZR have an abnormal lipogenic enzyme regulation that might contribute to the excessive fat accumulation.
Previous work examining the effect of exercise training on OZR has focused primarily on changes in serum lipids and tissue insulin sensitivity. Durstine et al. (5) showed that serum triglyceride content in OZR was decreased after 9 wk of treadmill training. In addition, low-density lipoprotein in trained rats contained less triglyceride and cholesterol, whereas high-density lipoprotein was elevated. Insulin resistance observed in OZR has been shown to diminish with exercise training due to mobilization of the muscle glucose transporter (GLUT-4) (6). It is possible that depressed serum lipids was secondary to altered plasma insulin status and tissue insulin sensitivity with exercise training. However, whether or not a greater hepatic lipogenic rate observed in OZR can be modulated with training has never been studied.
The purposes of conducting the present study were threefold: first, to investigate whether chronic exercise training could down-regulated hepatic FAS gene expression, which constitutes an important phenotypic trait in OZR; second, to study the hormonal and molecular mechanisms underlying training down-regulation of FAS, especially the role of the nuclear protein binding to several well-established gene regulatory sequences located in the FAS promoter; and third, to evaluated whether a reduced FAS activity could lead to alterations in liver fatty acid composition as well as whole body fat deposition.
Female obese Zucker rats (O, N = 16) and their lean litter mates (L, N = 16) were purchased from Harlan Sprague Dawley Co. (Indianapolis, IN). Female rats were used as in our earlier experiments because male rats reduce food intake and body weight in response to endurance training, whereas female rats increase food intake with training without losing body weight (9,24). Rats were individually housed in steel cages within temperature-controlled rooms (22°C) with a reversed dark:light cycle (0700–1900, dark; 1900–0700, light). Animal care and use protocols were approved by the Research Animals Resource Center Review Board of the University of Wisconsin-Madison.
During the first 2 wk after arrival, all rats were accustomed to a meal-feeding regimen using a powdered chow diet. The rats were given access to the food for 4 h·d−1 starting at the beginning of the dark (active) cycle. At the end of the 4 h, the food was removed from the cages until the next meal. Two weeks later, the chow diet was replaced by a cornstarch diet, in which 50% of the weight consisted of a basal mix including the required amounts of amino acids, fats, carbohydrates, vitamins, and minerals, whereas the other 50% was cornstarch [cf. Table 1, ref. 9]. Body weights of all rats were measured weekly throughout the study, before the daily feeding.
Starting at the second week, all rats were acclimated to running on a Quinton rodent treadmill at 15.5 m·min−1 and 0% grade for 10 min·d−1, 5 d·wk−1. Treadmill speed and running time were increased daily, so that by the end of wk 4, O and L Zucker rats were able to run at 18 and 20 m·min−1, respectively, at 0% grade for 100 min. During the following weeks the treadmill speed and grade were gradually increased differently in order to provide similar workloads for O and L rats (Table 1), although relative work intensity (%V̇O2max) for each group was unknown. Training sessions were scheduled during 4–6 h after the beginning of the meal timed to coincide with peak dietary induction of FAS gene expression (9,12,29). Untrained O and L rats were run on the treadmill at 15 m·min−1 for 10 min·d−1. This allowed the control animals to experience the sounds and stress of training without gaining training effect.
Tissue collection and preparation.
At the end of the exercise-training program, all rats were killed by decapitation in the resting state 27 h after the end of their last training/handling session. Rats were equally distributed across four consecutive killing days. On the day of killing both the normal 4-h meals and killing times were staggered such that each rat was killed between 8 and 9 h after the start of last meal. Previous work indicated that the activities of hepatic lipogenic enzymes do not vary significantly over the course of a day during meal-feeding regimens (9).
After decapitation, the mixed arteriovenous blood was collected in a conical tube containing 100-μL heparin (6 g·L−1) and chilled on ice. The blood sample was centrifuged at 500 g for 15 min and blood plasma was stored at −80°C. The abdominal cavity was opened and the liver quickly excised, weighed, and frozen in liquid N2. The deep portion of the vastus lateralis muscle (DVL) from one hindleg was quickly removed. Liver and muscle samples were stored at −80°C until processing and assay.
Plasma glucose and hormonal analyses.
Plasma glucose concentration was measured using a glucose oxidase assay (Sigma Diagnostics Glucose Kit 510-DA, Sigma Chemical Co., St. Louis, MO). Plasma glucagon and insulin were determined using RIA kits obtained from Diagnostic Products (Los Angeles, CA) and Linco Research (St. Louis, MO), respectively.
Electromobility shift assays.
A portion of the liver samples was used for nuclear extract preparation as previously described (8). The following single-stranded oligonucleotides were purchased from Gibco Life Sciences (Gaithersburg, MD):
FAS-IRS-A (−71/-50): 5′-TCAGCCCATGTGGCGTGGCCGC-3′
FAS-ChoRE (+283/+303): 5′-GGCCGCTGTCACGTGGGCGCC-3′
FAS-ICE (−109/-85): 5′-CGACGCTCATTGGCCTGGGCGG-3′
NFκβ (consensus sequence): 5′-AGTTGAGGGGACTTTCCCAGGC-3′
The oligonucleotides were incubated with γ-P32-ATP and T4 polynucleotide kinase. The labeling reaction (25 μL) was allowed to proceed for 30 min at 37°C and stopped by adding 0.5 M EDTA and bringing to a volume of 100 μL with Tris-EDTA (TE) buffer. After chloroform extraction, free [γ-P32]-ATP was removed by spin chromatography and ethanol precipitation. Complementary labeled strands were annealed by combining equal amounts of each oligonucleotide in TE buffer (pH 8.0), heating to 90°C and allowing to cool slowly to room temperature.
Electromobility shift assays (EMSA) were performed with liver nuclear extracts at room temperature in 30 μL, as previously described (8). For FAS-IRS/A, the assay contained 10 mM Tris HCl (pH8.0), 50 mM NaCl, 1 mM EDTA, 1 mM DTT, 10% glycerol, and 0.5-μg poly(dI-dC). For FAS-ChoRE, the assay included 10 mM HEPES, pH 8.0, 50 mM NaCl, 50 mM KCl, 5 mM MgCl2, 2 mM DTT, 17.5% glycerol, and 1-μg poly (dI-dC). For FAS-ICE: 20 mM HEPES, pH 7.9, 1 mM DTT, 60 mM KCl, 0.5 mM EDTA, 5% glycerol, and 0.5-μg poly [dI-dC]. Each reaction contained 40,000 cpm (0.1–0.5 ng) of oligonucleotides and the indicated amounts of nuclear extracts. To ensure equal loading of nuclear protein, we initially measured protein concentration by the Bradford method and later confirmed by staining the SDS-PAGE gels. For competition experiments, unlabeled competitor oligonucleotides were added to the mixture before addition of labeled probe. After 20 min at room temperature, the samples were subjected to 4% nonreducing PAGE in 1X Tris-glycine buffer. The dried gels were exposed to x-ray film at −70°C with an intensifying screen.
Northern blot analysis.
Total RNA was isolated from frozen livers with Trizol reagent (Gibco Life Technologies) as previously described (12). The cDNA probes for FAS and 18s were labeled using random primer extension (7) with a labeling kit using [α-32P]dCTP (Megaprime, Amersham, Arlington Heights, IL). The hybridization solution consisted of dextran sulfate (10%) and radiolabeled probes (106 cpm·mL−1) and were allowed to hybridize overnight. The stringency washes consisted of two 20-min washes with 1x SSC; 0.5% SDS at 45°C and two 20-min washes with 0.5x SSC; 0.5% SDS at 50°C, and one 20-min wash with 0.1x SSC; 0.5% SDS at 60°C. After autoradiography, the probe was removed from filters with a solution of 50% formamide, 2x SSPE at 65°C for 60 min. Quantification of the FAS signals was achieved by use of a scanning densitometer (BioRad model GS-670, BioRad, Richmond, CA). FAS mRNA abundance was expressed relative to the density of the respective 18s values.
Maximal activity of FAS (EC 126.96.36.199) was measured in liver cytosol by following the rate of NADPH utilization in palmitate synthesis at 30°C, as previously described (12). Citrate synthase (CS, EC 188.8.131.52) activity was measured in DVL as previously cited (9). Cytosolic protein content was determined by the Bradford method with bovine serum albumin as the standard.
Hepatic fatty acid composition.
Hepatic fatty acid composition was analyzed by gas chromatography, as previously described (9). Briefly, approximately 1 g of frozen liver tissue was homogenized in chloroform:methanol (2:1, vol:vol) using a Polytron homogenizer (setting #6) for 40 s. Fatty acid methyl esters (FAME) were analyzed with a Varian 3400 gas chromatography equipped with a flame-ionization detector, a Varian 1093 SPI temperature programmable injector (Palo Alto, CA), a Supelcowax 10 fused silica column (30 m × 0.32 mm i.d. 0.25-μm film, Supelco), and an In-Board Data handling System (IBDHTM, Sugarland, TX). The detector temperature was 300°C. Fatty acids were identified by comparing the retention times with those of known standards (Nu-Chek-Prep, Elysian, MN) and expressed as the weight percent distribution of FAME.
Body composition analysis.
Body composition of the OZR and LZR was analyzed by dual-energy x-ray absorptiometry (DXA) using a Hologic DXA unit (Hologic QDR 2000, Waltham, MA) equipped with small animal software developed by Hologic. To validate that body composition could be evaluated using the rat carcass, pilot experiments were performed. Two rats were immobilized with pentobarbitol injection (40 mg·kg, i.p.) and subjected to DXA measurements. Thereafter, the same rats were decapitated and a portion of the liver (∼2 g) was removed. Body composition of the rat carcass was analyzed with DXA immediately, and then frozen for 24 h at −20°C before being thawed and immediately subjected to DXA measurement for a third time. There was no difference in body fat, lean body mass (LBM), or mineral content among the three measurements, whereas the loss of body water (reflected by the difference between live and carcass weights) was less than 5%. Therefore, carcass body composition without the sampled liver was estimated on frozen-thawed carcasses.
Two-way ANOVA was used to determine significant differences (P < 0.05) in the means of the various treatment groups. The main effects were obesity (L vs O) and exercise training (T vs U). After an overall F was found to be significant, a post hoc least significant difference (LSD) test (SYSTAT, Evanston, IL) was used to evaluate the significance of differences between treatment groups.
Body and liver weights and food consumption.
Both body weight and food consumption were monitored weekly. Obese rats were heavier than L rats throughout the experimental period, as expected. Body weights increased similarly in OT and OU rats during the first 6 wk. Starting at the seventh wk, OT rats gained significantly less weight than OU rats (P < 0.05, Fig. 1). No significant difference in body weight gain was observed between LT and LU rats
There was no difference in absolute food consumption between O versus L rats (data not shown). When expressed relative to body weight (i.e. g·100 g−1 body weight), O rats consumed significantly less food than L rats (P < 0.05, Table 2). Exercise training had no effect on food consumption in either O or L rats. Obese rats had higher absolute liver weights than L rats (P < 0.05, Table 2), but liver weights per unit body weight were similar between the two groups (data not shown).
Citrate synthase activity.
CS activity was measured in the DVL muscle and showed no difference between UO and UL rats (Table 2). Training increased CS activity by 31% in L rats and by 136% in O rats (P < 0.05), confirming a metabolic effect of training regimen. CS activity in TO rats was significantly higher than that in TL rats (67%, P < 0.05;Table 2).
Plasma glucose, insulin, and glucagon.
Plasma glucose concentrations were not significantly different between untrained O and L rats (Table 2). Furthermore, plasma glucose was not altered with T in L rats. However, T significantly lowered glucose levels in O rats (P < 0.05).
Obese rats showed a severe hyperinsulinemia, with fourfold higher plasma insulin levels than their lean litter mates. This elevated insulin level was not significantly affected with T. In contrast to the hyperinsulinemia seen in O rats, plasma glucagon was normal in O rats and was unaltered by T.
Fatty acid synthase activity and mRNA abundance.
Obese rats had 2.5-fold higher FAS activity than L rats (P < 0.01, Fig. 2). Training resulted in a significant 18% reduction of FAS activity in O rats (P < 0.05) but had no significant effect on FAS activity in L rats. Northern blot analysis revealed a double band with sizes of 8.6 and 9.0 kb for FAS mRNA signals (Fig. 3A). O rats showed a greater mRNA abundance than L rats, whereas the abundance of 18s rRNA, used to normalize FAS mRNA values, showed no change. Data analysis of individual samples obtained by slot blot analysis revealed that O rats had sixfold higher levels of FAS mRNA relative to 18s RNA (P < 0.05, Fig. 3B). However, T had no significant effect on FAS mRNA abundance in L or O rats.
Transcription factor binding.
Specificity of nuclear protein binding to IRS/A, ChoRE, and ICE was determined using the competition gel mobility shift assay. A single major DNA-protein complex was detected with labeled FAS-IRS/A (Fig. 4a, panel A, lane 1). This complex was successfully competed away by increasing concentrations (1- to 100-fold) of unlabeled IRS/A oligonucleotide (panel A, lanes 2–5). However, adding increasing concentrations of unlabeled ICE (panel A, lanes 6–10) or a consensus NFκB oligonucleotide probe (data not shown) did not affect binding to the labeled IRS/A, indicating that the band detected was specific. To determine whether the binding of transcription factors was affected by nutritional and metabolic factors, liver nuclear extracts were prepared from pooled nuclear extracts from each treatment group, as shown in Figure 4a, panel B. Results of ANOVA on the means of various treatment groups derived from gel shift experiments for each individual rat are depicted in Fig. 4b. Transcription factor binding to FAS-IRS/A was lower in O versus L rats (P < 0.05). The binding was unaffected by T in either group.
Competition experiments with FAS-ChoRE were performed to determine the specificity of transcription factor binding (Fig. 5a, panel A). A single band representing DNA-protein complex was detected (lane 1). This complex was successfully competed away with increasing concentrations of unlabeled ChoRE oligonucleotide (lanes 2–6), whereas adding increasing concentrations of unlabeled consensus NFkB oligonucleotide probe did not affect ChoRE binding (lanes 7–12), indicating that the band was specific. ChoRE binding patterns in the various treatment groups are shown in Fig. 5a, panel B. Binding of FAS-ChoRE was attenuated in O rats versus L rats, with no apparent T effect observed (Fig. 5b).
Incubation of nuclear proteins with FAS-ICE produced two major complexes, both of which were competed away with excess unlabeled ICE at concentrations greater than 10-fold molar excess (Fig. 6a, panel A, lanes 1–5). Addition of excess NFκB oligonucleotide probe did not show competition with ICE (panel A, lanes 6–10). FAS-ICE binding was increased in O versus L rats (P < 0.05, Fig. 6a, panel B;Fig. 6b). As seen for FAS-ChoRE, FAS-ICE binding was not affected by T in either group.
Hepatic fatty acid profiles.
Obese rats had significantly higher levels of 16:0, 16:1, and 18:1 fatty acids (P < 0.05) in the liver than L rats (Table 3). Conversely, liver 18:0, 18:2, and 20:4 fatty acid levels were lower (P < 0.05) in O versus L rats, whereas levels of 18:3 fatty acids were unchanged. Training had no effect on levels of most of liver fatty acids, except 18:3, which was decreased (P < 0.05). The levels of 20:4 tended to increase with T (P < 0.08).
Body composition data of O and L rats are shown in Table 4. As expected, carcass weights of O rats were heavier than those of L rats (P < 0.01). Carcass weights of L rats were no significantly altered by T, whereas T decreased carcass weight in O rats (P < 0.01). Gut weights were not different between groups. Body fat, expressed as fat weight per carcass and percentage body weight, was more than four- and two-fold greater (P < 0.01), respectively, in O rats compared to L rats. Training decreased fat weight and percentage by 24 and 21% in L rats (P < 0.01), and by 34 and 20% (P < 0.01), respectively, in O rats. Absolute and percentage LBM were lower in O versus L rats (P < 0.01), whereas T significantly increased %LBM in both O and L rats (P < 0.01).
Hepatic FAS activity was 2.5-fold higher in O rats compared with their lean litter mates. Training significantly decreased hepatic FAS activity in O rats. In contrast, training only caused a small and nonsignificant effect in L rats. This discrepancy is likely due to a higher relative exercise intensity imposed on O versus L rats because of the large body weight difference. We attempted to adjust workload by altering speed and grade to elicit similar metabolic stress, but the actual exercise intensity might be greater for O rats. We previously showed a significant training down-regulation of FAS activity in Sprague-Dawley rats meal-fed an identical diet with similar exercise intensity and duration (9). Other than genetic differences was a markedly (34%) lower plasma insulin concentration in the lean Zucker rats (148 and 220 μIU·mL−1 for untrained and trained, respectively) compared to Sprague-Dawley rats (224 and 283 μIU·mL−1). Resting FAS activity in the untrained lean Zucker rats (5 mU·mg−1) was ∼30% lower than the Sprague-Dawley rats (7.2 mU·mg−1). Perhaps the lower baselines make it more difficult to suppress insulin and FAS activity and subsequently require a more rigorous exercise protocol.
Whereas the elevation of FAS activity was reflected in a huge increase in FAS mRNA levels, training was without effect. The finding that training decreased FAS enzyme activity without affecting FAS mRNA abundance is consistent with our previous study reporting training effects on diet-induced FAS in meal-fed rats (9). Obese Zucker rats demonstrated sixfold higher FAS mRNA levels than their lean litter mates, indicating an elevated gene expression of lipogenic enzymes. Our data suggest that exercise training, at least at the current intensity and frequency, is not sufficient to suppress the strong genetic tendency of high lipogenic enzyme expression in this line of animals. However, training may elicit translational and posttranslational modulation on FAS resulting in the down-regulation of FAS activity. It cannot be entirely ruled out that FAS mRNA levels were decreased transiently after each training session but returned to the resting levels after 27 h when the rats were killed. In a previous study, we showed that an acute bout of exercise in fasted-refed rats inhibited both FAS activity and mRNA abundance up to 12 h postexercise, whereas at 24 h we observed only decreased FAS activity but not mRNA levels (12).
Obese rats demonstrated a dramatic hyperinsulinemia, which is considered one of the main reasons for the elevated lipogenic status and hyperlipidemia observed in these animals (1,3,26). Meal-feeding might also have played a role but was not likely the main mechanism for the high plasma insulin concentration (13,22). We demonstrated previously that decreased hepatic FAS activity and mRNA abundance after an acute exercise bout were accompanied by a lower plasma insulin and a higher plasma glucagon concentration, as well as a lowered liver pyruvate content (12). These hormonal and metabolic profiles inhibitory to lipogenic enzyme induction likely have taken place during each training session and for a short period thereafter (9). A decreased resting plasma glucose concentration in O rats exemplified the training adaptations that could affect hepatic lipogenic enzyme regulation.
Altered nuclear protein binding to putative gene regulatory sequences in the FAS promoter in response to hormonal and dietary stimulations has been established as the most important molecular mechanism governing the gene expression of lipogenic enzyme (27). FAS-IRS/A and FAS–ChoRE binding by upper-stream stimulatory factors (USF) have been shown to enhance FAS gene transcription and the binding has been shown to increase with hyperinsulinemia and fructose feeding (11,18,20,28). In contrast, binding of FAS-ICE by NF-Y is associated with a down-regulation of FAS gene transcript under the influence of cAMP and glucagon (21,25). We reported in a recent study that liver nuclear protein binding to FAS-IRS/A and ChoRE were attenuated in favor of a decreased FAS gene transcription and hence enzyme protein synthesis after an acute bout of exercise in fasted rats refed a high-fructose diet (8). We therefore hypothesized that obese Zucker rats had an elevated steady-state binding of FAS-IRS/A and ChoRE, and that training could exert an inhibitory influence on the binding. The results from the EMSA, however, argued against our hypotheses. Despite the dramatically elevated plasma insulin levels and FAS mRNA levels, IRS/A and ChoRE binding was decreased in O versus L rats, whereas FAS-ICE binding was increased in O rats without altered glucagon status. Furthermore, these bindings were not influenced with training. These findings suggest that the regulation of FAS in O rats may be controlled either by mechanisms other than the transcriptional step, or by transcription factors other than IRS/A, ChoRE, and ICE. For example, the FAS promoter region contains an E-box motif (−64/−59) that is part of the well-defined IRS/A (−71/−50) and can bind with other basic helix-loop-helix transcription factors, such as adipocyte determination differentiation dependent factor (ADD) 1 and sterol regulatory element binding protein (SREBP) 1 to elicit transactivation of FAS (17). Furthermore, transcription of ADD1/SREBP1 in hepatocytes was found to be controlled positively by insulin and negatively by glucagon and cAMP (10). In addition, SREBPs have been found to regulate lipogenic enzyme gene expression, including FAS (4). It cannot be ruled out that each training session might have caused a transient alteration of IRS/A, ChoRE and ICE binding profiles, resulting in a down-regulation of FAS transcription and protein synthesis. These changes could have been reversed postexercise due to the prevailing plasma insulin levels in O rats.
Obese rats had significantly higher levels of saturated and monounsaturated fatty acids in the liver, i.e., palmitic acid (16:0), palmitoleic acid (16:1), and oleic acid (18:1). Although there was a decrease in stearic acid (18:0) in O rats, this was not unexpected because steroyl-CoA desaturase is regulated similarly to FAS and other lipogenic enzymes (19). An increase in this enzyme activity in O rats can lead to a decrease of 18:0. In contrast, the levels of polyunsaturated fatty acids (PUFA), especially linoleic (18:2n-6) and arachidonic acid (20:4n-6) were dramatically decreased in O versus L rats. These changes in hepatic PUFA composition may not only be a result of, but also a cause of, altered lipogenic enzyme activity. It has been known for some time that PUFA-feeding depresses hepatic lipogenesis (14). More recently, several lipogenic enzymes have been found to contain PUFA-responsive regions within their gene promoter (16). Although the exact mechanism remains unknown, the finding that the PUFA effect is only observed in tissues where peroxidation occurs indicates that specific eicosanoid products are involved (16). These fatty acid profiles indicate that the etiology of the drastic accumulation of body fat in these animals is uncontrolled de novo lipogenesis, which synthesizes primarily 16:0 but not 18:2 or 20:4 (15). It is interesting to note that the changes in liver fatty acid profile in O rats were similar to those found in normal rats meal-fed a high (50%)-fructose diet for 10 wk, although the magnitude of changes was much smaller (9). Based on our previous findings that training could decrease fructose-induced hepatic FAS activity in meal-fed normal rats, we hypothesized that down-regulation of FAS in trained O rats would reduce de novo lipogenesis and hence reduce body fat. Supporting evidence for this hypothesis came from the following: (a) training dramatically decreased the weight and percentage of body fat, which accounted solely for the decreased body weight after training in O rats; and (b) there was a strong tendency for 20:4 composition to be increased after training. The latter finding suggests that although the dramatic reduction in body fat in O rats may be explained primarily by increased fatty acid utilization during training, an attenuation of de novo fatty acid synthesis may also have played a role. However, our data did not allow us to assess the quantitative significance of the decreased de novo lipogenesis in the reduction of whole-body fat in O rats.
In conclusion, the current study revealed that endurance training performed after the daily high-carbohydrate meal markedly suppressed hepatic FAS enzyme activity in obese Zucker rats without affecting their resting plasma insulin level. The decreased FAS activity with training was not caused by a reduced steady-state binding of nuclear proteins to the FAS gene promoter. Training dramatically decreased the body fat in obese Zucker rats, which accounted solely for the observed loss in their body weight.
This study was supported in part by the Vilas Trust Fund of the University of Wisconsin-Madison. R. Fiebig was a recipient of the American Heart Association Predoctoral Fellowship. The cDNA for FAS was a kind gift from Stuart Smith at the Oakland Children’s Hospital, California. The 18s probe was obtained from Karyn Esser at the University of Illinois, Chicago. We would also like to thank Mike Grahn for his assistance in measuring fatty acid composition and Molly Brown for her assistance in measuring rat body composition.
Address correspondence to: Li Li Ji, Ph.D., Department of Kinesiology, 2000 Observatory Drive, Madison, WI 53706; E-mail: email@example.com.
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Keywords:©2002The American College of Sports Medicine
EXERCISE TRAINING; FATTY ACID SYNTHASE; LIPOGENESIS; NUCLEAR PROTEIN BINDING; OBESITY