Effects of Dietary Monosaccharides on Sympathetic Nervous System Activity in Adipose Tissues of Male Rats Free

Male Sprague Dawley rats were obtained from Harlan (Indianapolis, IN) at 7 weeks of age and housed three per cage before study at a room temperature of 21 ± 2°C, with a light:dark cycle of 14:10 h. Animals used in this study were maintained in accordance with the guidelines and approval of the Animal Care and Use Committee of the Feinberg School of Medicine of Northwestern University.

In all experiments, animals were fed measured quantities of food for each cage of three rats for 6–7 days before study. All groups received the same quantity of standard lab food (Harlan Diet 7012; Harlan Teklab, Madison, WI), and in addition, animals fed the monosaccharide-supplemented diets received an amount of monosaccharide equal in energy to the food ration. Thus, monosaccharide-fed groups received twice the dietary energy of animals fed food alone. Monosaccharide supplements consisted of dextrose monohydrate, d-fructose, and d-galactose (all from Harlan Teklab). The various diets were continued throughout the 24 h of the [³H]NE turnover experiments. Animals had free access to water throughout these experiments.

6-Hydroxydopamine (6-OHDA; Sigma-Aldrich, St. Louis, MO) was diluted in isotonic saline containing 5 μl 2N HCl and 1 mg/ml ascorbic acid and injected intravenously. Each animal in the treatment group received two 6-OHDA injections (60 and 120 mg/kg) on successive days, the latter being 4–5 days before the start of a feeding protocol, whereas animals in the control group received injections of diluent. Tissues were removed for analysis of NE content and gene expression 10–11 days following 6-OHDA treatment.

l-[Ring-2,5,6-³H]NE (40–60 Ci/mmol specific activity; DuPont NEN Research Products, Boston, MA) was diluted with 0.9% NaCl and injected intravenously into the tail veins of unanesthetized animals in a total volume of 1.0 ml, beginning at ∼9:00 a.m. The dose of [³H]NE used in the current studies was 57–67 μCi/kg (∼0.21–0.25 μg NE/kg). The rats were killed 2, 6, 12, and 24 h following tracer injection by CO₂ inhalation. For each time point in the NE turnover studies, 4–6 animals were killed from each experimental group. The tissues were rapidly removed, frozen on dry ice, and stored at −20°C for later processing (usually within 2 weeks).

For NE analysis, the organs were weighed and homogenized in iced 0.2 N perchloric acid containing 1% Na₂S₂O₅ (by weight) and 1 mmol/l EDTA in a Polytron homogenizer (Brinkmann Instruments, Westbury, NY) to extract the catecholamines. After addition of the internal standard, 3,4-dihydroxybenzylamine (DHBA; Sigma, St. Louis, MO), catecholamines were isolated from the perchloric acid extract by adsorption onto alumina (Woelm neutral; ICN Nutritional Biochemicals, Cleveland, OH) in the presence of 2 mol/l Tris(hydroxymethyl)-aminomethane buffer (pH 8.7; Sigma) containing 2% EDTA. Catecholamines were eluted from the alumina with 0.2 N perchloric acid. Aliquots of the alumina eluate were injected onto a liquid chromatographic system for catecholamine analysis following the method of Eriksson and Persson (15) with slight modification. Unless otherwise specified, all chemicals were obtained from Fisher Scientific (Fair Lawn, NJ). Aliquots of the alumina eluates were counted for [³H]NE by scintillation spectrometry in a Packard Tri-Carb 2100TR liquid scintillation analyzer (Packard Instrument, Meriden, CT). Efficiency for ³H is ≥58% in this system.

Messenger RNA levels were quantitated using the Applied Biosystems TaqMan system (PE, Norwalk, CT). This method uses a double-labeled oligonucleotide probe (fluorescent dye/quenching molecule) that anneals between the PCR primers. Fluorescence occurs only after digestion by the polymerase during the extension phase of each PCR cycle. Emitted fluorescence is detected using the Sequence Detector 7700 instrument. Analysis is performed using the Sequence Detector software. The manufacturer’s protocol was followed, with the exception that the concentration of certain reagents was adjusted as follows: each reaction tube contained 0.25 μg total RNA, 2.5 μl TaqMan buffer A (10×), 2 μl dNTP mix (2.5 mmol/l dATP, 2.5 mmol/l dCTP, 2.5 mmol/l dGTP, and 5 mmol/l dUTP), 0.375 μl each of forward and reverse primers (20 μmol/l), 4 μl MgCl₂ (25 mmol/l), 0.025 μl probe (100 μmol/l), 0.25 μl MuLV reverse transcriptase (50 units/μl), and 0.25 μl Amplitaq Gold (5 units/μl) in a total volume of 25 μl. Values in parentheses indicate stock, not final, concentrations. RNA samples were prepared by tissue extraction with guanidine thiocyanate, purified using CsCl₂ centrifugation, and treated with DNase before assay to destroy any contaminating genomic DNA. Each RNA sample was assayed in triplicate; a fourth tube per sample containing no reverse transcriptase was included as a control for contaminating genomic DNA. One tube per assay contained water in place of RNA as a control for RNA or DNA contamination of the solutions. The temperature profile was 30 min at 42°C, 10 min at 95°C, 15 s at 95°C/1 min at 60°C for 40 cycles, and 5 min at 23°C. All equipment and reagents were from PE. Primers and probes used in the RT-PCR assays are presented in Table 1 (16–22). In this assay, the number of cycles to reach a given threshold value is inversely related to the initial abundance of a specific mRNA at the start of the reaction. Tissues for analysis of mRNA were stored at −80°C before assay.

Data are displayed as means ± SE. Statistical ANOVA and ANCOVA were performed using Data Desk 6.1 statistical software (Data Description, Ithaca, NY) (23). Post hoc pairwise comparisons following ANOVA utilized Scheffé’s test. Analyses employed P < 0.05 as the criterion for statistical significance; P values between 0.05 and 0.1 are provided in text and tables, whereas P values ≥0.1 are indicated by “NS”.

In studies of NE turnover, the method of least squares was used to calculate the slope (k) of decline in NE specific activity over time after tracer injection (24). In all measurements of [³H]NE turnover, no significant variation in endogenous NE was observed over the 24 h of the experiment. The statistical significance of each computed regression line was assessed by ANOVA, and ANCOVA was used in comparison of fractional turnover rates. Goodness of fit for each regression line was evaluated by examination of externally studentized residuals. NE turnover rates were calculated as the product of the fractional turnover rate, and the endogenous NE concentration (25) and CIs were computed as previously described (26).

In an initial study, [³H]NE turnover was measured in interscapular brown adipose tissue (IBAT) of rats fed diet alone or diet supplemented with glucose or fructose for 6 days before turnover measurement. Results from this experiment are presented in Table 2. Rats fed either of the monosaccharide supplements were heavier than diet-fed controls. Fractional rates of [³H]NE turnover in IBAT were greater in both monosaccharide-fed groups than in controls (P < 0.0001 for pairwise comparisons of both glucose- and fructose-fed rats versus diet-fed controls). In IBAT, the increases in total NE turnover over control were 77% in glucose-fed rats and 99% in fructose-fed animals. In retroperitoneal fat, [³H]NE turnover was 57% higher in the animals given glucose than in diet-fed controls (P < 0.03) and 73% greater in the fructose-fed rats than in controls (P < 0.001). NE turnover rates in retroperitoneal fat did not differ between glucose- and fructose-fed animals. By contrast, [³H]NE turnover rates in epididymal fat were only slightly (22–23%), but not significantly, higher in rats fed glucose or fructose than in those fed diet alone. Thus, oral intake of glucose and fructose over a 6-day period activates sympathetic nerves in IBAT and retroperitoneal fat, but not in epididymal fat.

In a second experiment of similar design, [³H]NE turnover rates were compared in animals fed glucose- and galactose-enriched diets for 6 days. Despite consuming a diet containing twice as much energy as that given to diet-fed controls, galactose-fed animals weighed the same as animals fed diet alone, and both groups were significantly lighter than the glucose-fed rats (P < 0.0001 for both pairwise comparisons). Total [³H]NE turnover rates were likewise greater in glucose- than in diet- or galactose-fed rats. In IBAT, glucose-fed rats showed an increase in [³H]NE turnover (124%, P < 0.05), in contrast to a slight, but not statistically significant, reduction (−21%) seen in galactose-fed animals relative to controls.

As [³H]NE turnover rates in white adipose tissue (WAT) were not measured in the preceding study, the experiment was repeated in diet- and galactose-fed rats, and the results are presented in Table 3. Although body weights did not differ between diet- and galactose-fed groups, weights for IBAT and the two WAT pads were significantly lower in the animals fed galactose (P < 0.0001 for all comparisons). [³H]NE turnover rates in IBAT were slightly, but significantly, lower in galactose-fed rats (−33%, P < 0.05). In retroperitoneal fat, [³H]NE turnover was slightly, but not significantly, higher (14%) in the galactose-fed group. In contrast to findings in IBAT or retroperitoneal fat, however, [³H]NE turnover in epididymal fat was markedly elevated in the galactose-fed animals (138%, P < 0.05). Thus, oral intake of galactose for 6 days suppresses SNS activity in IBAT, exerts minimal effect in retroperitoneal fat, but stimulates sympathetic nerves in epididymal fat.

In the preceding studies, [³H]NE turnover in monosaccharide-fed rats was compared with animals fed a limited diet ration. To determine whether the differences observed were due to alterations in caloric intake or to ingestion of the monosaccharides themselves, [³H]NE turnover in glucose-fed rats was compared with that in rats fed either of two diet rations, one equivalent to the diet content of the diet:glucose diet (diet50) and the other equivalent to the combined energy intake of diet plus glucose (diet100). The results of these comparisons are presented in Table 4. Animals fed the higher diet ration (diet100) were significantly heavier after 7 days than either of the other two groups; glucose-fed animals were again heavier than animals given the reduced diet (diet50).

The effects of extra diet or glucose intake on [³H]NE turnover rates differed among adipose tissues. In IBAT, animals fed the full diet ration exhibited an intermediate rate for total NE turnover that was statistically distinguishable from the lower rate seen in the restricted diet group and from the higher rate observed in the glucose-fed rats. In retroperitoneal fat, the stimulatory effect of the higher diet ration was fully equivalent to that observed in glucose-fed animals. In epididymal fat, the increases in [³H]NE turnover were small and not statistically significant for either increased diet or glucose feeding. Consequently, the stimulatory effect of dietary monosaccharides on [³H]NE turnover in IBAT would appear to be influenced by both the level of energy intake and the specific nutrient ingested, whereas the response in retroperitoneal fat would appear to be related principally to energy intake. Since [³H]NE turnover rates in epididymal fat did not correspond with energy intake, the sympathetic response to galactose ingestion noted above (Table 3) must reflect intake of galactose per se.

To determine how differences in [³H]NE turnover might affect adipose tissue function, gene expression for several sympathetically related proteins was examined in IBAT and WAT obtained from rats fed diet alone or diet supplemented with each of the three monosaccharides. The results, presented as “cycles to threshold” in Table 5, are inversely related to the amount of a particular transcript present in 0.25 μg of total RNA extracted from each tissue. In IBAT, the data show three patterns of response: no effect of diet on β₃-adrenergic receptor (AR) expression, suppression of leptin expression in galactose-fed rats (P < 0.05), and increased expression of uncoupling protein (UCP)1, GLUT4, angiotensinogen (AGT), and ribosomal protein L19 (RPL19) in glucose- and fructose-fed animals compared with diet- or galactose-fed animals. The changes in gene expression in IBAT for UCP1, GLUT4, AGT, and RPL19 are complementary to the elevations in [³H]NE turnover noted in Tables 2 and 3.

In retroperitoneal fat, gene expression for GLUT4, β₃-AR, leptin, and AGT follow a pattern in which expression is greater (fewer cycles to threshold) in glucose- and fructose-fed rats than in galactose-fed animals or diet-fed controls. The pattern for RPL19 is similar, though the differences are not statistically significant. By contrast, in epididymal fat only gene expression for leptin followed the same pattern noted in retroperitoneal fat (increased expression in glucose- and fructose-fed animals); expression of GLUT4, β₃-AR, AGT, and RPL19 did not vary significantly among diet groups. Thus, expression of GLUT4, β₃-AR, AGT, and RPL19 in retroperitoneal fat show changes in response to monosaccharide intake that correspond to the differences noted previously with [³H]NE turnover, raising the possibility that increased expression of these four genes may occur in response to nutrient-induced changes in SNS activity. In epididymal fat, none of the genes examined showed increased expression with galactose intake.

To examine the contribution of SNS activity in WAT to these diet-induced elevations in gene expression, the effects of 6 days of fructose feeding on gene expression were compared in intact and chemically sympathectomized rats (Table 6). Fructose, rather than glucose, was used in this comparison to minimize any potential confounding effects from direct insulin action on the WAT pad under study. Although body weights were reduced by 6-OHDA treatment, the sympathectomized rats consumed equivalent amounts of diet and fructose compared with intact controls during the 6 days of pair feeding. Although WAT weight was not affected by sympathectomy, tissue NE contents in 6-OHDA-treated WAT were 73 and 84% lower in retroperitoneal and epididymal fat, respectively (P < 0.0001 for both). Of note, sympathectomy reduced expression of all genes examined in retroperitoneal fat. Moreover, fructose increased UCP1 and GLUT4 expression in intact animals (P < 0.03 for both), but not in sympathectomized rats (P = NS for both). On the other hand, despite an overall reduction in leptin and AGT gene expression in sympathectomized rats, the response to fructose was similar in both treatment groups.

Effects of sympathectomy were less evident in epididymal fat. Treatment with 6-OHDA exerted no discernible effects on expression of GLUT4, β₃-AR, or leptin compared with the reductions observed in retroperitoneal fat. Moreover, while expression of AGT and AGTR_1A was slightly reduced in sympathectomized rats, fructose intake was without effect on expression of either gene. Thus, although chemical sympathectomy abolished the fructose-induced increase in UCP1 and GLUT4 expression in retroperitoneal fat, sympathetic denervation exerted lesser effects on gene expression in epididymal fat.

The results reported here provide further evidence that SNS activity in peripheral tissues is responsive to carbohydrate content of the diet. As noted previously in heart (6), the addition of glucose to the basic diet ration consistently increased SNS activity in IBAT and retroperitoneal fat, though not in epididymal fat. Sympathetic responses in IBAT and retroperitoneal fat to fructose intake were qualitatively similar to those observed with glucose. Although galactose feeding, by contrast, did not stimulate [³H]NE turnover in either IBAT or retroperitoneal fat, it markedly raised [³H]NE turnover in epididymal fat despite the minimal increases noted in this tissue in response to glucose, fructose, or added diet. These data, thus, indicate that SNS activity, while responsive to dietary monosaccharides in both tissues, is regulated differently in retroperitoneal and epididymal fat.

In addition to assessment of SNS activity via measurement of NE kinetics in peripheral tissues, the current studies also compared effects of dietary monosaccharides on gene expression in WAT. The genes monitored included ones like UCP1, GLUT4, leptin, β₃-AR, and RPL19, which have been shown to be responsive to sympathetic stimulation in IBAT (27) and AGT, which is reduced by fasting (28). Dietary effects on gene expression were similar in IBAT and retroperitoneal fat and paralleled alterations in [³H]NE turnover rates. Moreover, the increase in gene expression of UCP1 and GLUT4 induced by fructose feeding was abolished by sympathetic denervation. This correspondence both supports the contention that [³H]NE turnover rates reflect tissue-specific differences in SNS activity and indicates that changes in SNS activity have consequences for gene expression in these two tissues. Expression of the index genes in epididymal fat was less responsive to diet-induced changes in SNS activity or to sympathetic denervation than that in retroperitoneal fat, a further indication that sympathetic nerves serve different functions in these two adipose tissues.

The present studies were a continuation of work begun to examine sympathetic responses in WAT to fasting. In contrast to expectations, SNS activity in retroperitoneal fat was more often noted to decrease than increase with fasting (29). The current experiments were designed, therefore, to determine whether carbohydrate feeding would elicit an opposite response. The findings confirm such a hypothesis, but also show that SNS activity in retroperitoneal fat is related to energy intake (Table 4), which may also explain a suppressive effect of fasting (29). Although the effects of dietary monosaccharides noted here are, in the authors’ view, more likely due to differences in monosaccharide metabolism, it is possible that the variation among diet groups in body weight or body fat may have played a contributory role. Of the three adipose tissues examined, SNS activity in retroperitoneal fat most closely paralleled diet-induced differences in body weight (Tables 2–4). Further studies will be required to clarify the mechanisms underlying these sympathetic responses.

Although experiments described here and in a previous report (6) consistently show a stimulatory effect of fructose on SNS activity, the available literature is not uniform in agreement. Of all of the human and animal studies examining sympathetic responses to fructose based upon NE measurements or nerve impulse recordings (6–8,30–33), the only reports demonstrating fructose-induced SNS activation are those in which fructose was given orally, such as in the current studies (6–8). All of those (30–33) in which fructose was administered parenterally show no sympathetic response. Consequently, the afferent signal for SNS stimulation by fructose probably originates from peripheral chemoreceptors, possibly those located in the gastrointestinal tract and/or liver.

Activation of the SNS in IBAT and retroperitoneal fat by glucose and fructose, but not by galactose, indicates that the mechanisms regulating sympathetic responses to carbohydrate ingestion in these tissues are capable of distinguishing among the three principal monosaccharides in contemporary diets. Because fructose is less insulinogenic than glucose (9), SNS responses to fructose are likely to be less dependent on a primary role for insulin than those due to glucose. Following oral ingestion, fructose is largely taken up by liver and rapidly phosphorylated to fructose-1-phosphate via a relatively specific fructokinase (9,34). Further metabolism produces 3-carbon intermediates in the glycolytic pathway that are identical to those derived from glucose (34). If sensing mechanisms for glucose and fructose are dependent on products of intermediary metabolism rather than on the monosaccharides themselves, then a fructose-sensing apparatus would likely be responsive to glucose as well.

Differential sympathetic responses to carbohydrate in retroperitoneal and epididymal fat raise the possibility that SNS involvement in the regulation of lipid metabolism, likewise, may differ among WAT depots. In the case of retroperitoneal fat, if SNS outflow falls with fasting and rises with carbohydrate intake, it would appear unlikely for the principal role of sympathetic nerves to be stimulation of lipolysis. Rather, sympathetic nerves may promote α₂-adrenoceptor-mediated antilipolysis (35); alternatively, they may stimulate lipogenesis, as they do in IBAT (36). In such a model of fat cell metabolism, suppression of SNS activity by fasting, along with the reduction in insulin secretion, may allow lipolysis to proceed unimpeded, whereas circulating epinephrine from the adrenal medulla may provide the predominant adrenergic stimulus for lipolysis under physiological circumstances.

Finally, sympathetic nerves mediate the increase in GLUT4 expression seen in retroperitoneal fat of rats fed fructose-supplemented diets for 6 days. Support for this contention derives from the parallel changes (relative to diet-fed controls) in GLUT4 expression and in [³H]NE turnover in IBAT and retroperitoneal and epididymal fat of fructose-fed animals and from the lack of effect of fructose on GLUT4 expression in retroperitoneal fat of sympathectomized rats. Since glucose-fed rats exhibited qualitatively similar changes in GLUT4 expression and in [³H]NE turnover in adipose tissues, the SNS likely plays a contributory role in mediating these responses as well. The current findings thus provide preliminary evidence that oral intake of glucose and fructose activates a neural pathway, probably originating in gut or liver, which leads to enhanced expression of GLUT4 in specific adipose tissue depots.

While enhanced gene expression for GLUT4, the insulin-sensitive glucose transporter (37), does not necessarily signify an increase in glucose uptake or an increase in insulin sensitivity, a growing literature indicates that sympathetic nerves are capable of stimulating peripheral glucose uptake in other circumstances. During cold exposure, after electrical stimulation of the hypothalamus, or following central administration of leptin, glucose uptake into peripheral tissues increases via mechanisms that are sensitive to sympatholytic treatments and appear independent of and/or synergistic with insulin (38–41). Whether sympathetic activation by dietary carbohydrate also promotes glucose uptake into peripheral tissues, like IBAT and retroperitoneal fat, was not addressed in the current study. If stimulation of the SNS by diet does increase glucose uptake, however, such effects may relate to insulin action in two ways. First, the importance of dietary carbohydrate intake before glucose tolerance testing may be related to carbohydrate activation of sympathetic mechanisms assisting glucose uptake. Second, impairments in sympathetic responses to dietary carbohydrate may contribute to insulin resistance under some circumstances. (Although animals fed high-fructose diets for extended periods are frequently insulin resistant [12,42], the current studies were conducted within the first week of fructose exposure.) A role for sympathetic nerves to promote glucose uptake and enhance insulin action in peripheral tissues appears likely, though definitive characterization of such effects requires further study.

These studies were supported, in part, by U.S. Public Health Service (USPHS) Grant DK 20378.

The skillful technical assistance of Yiu-Kuen Chow and Siddique Mahmood is gratefully acknowledged.