Department of Medicine III, Division of Endocrinology & Metabolism, University of Vienna, Austria
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ABSTRACT |
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INTRODUCTION |
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TZDs are agonistic ligands of peroxisome proliferatoractivated receptor- (PPAR-
), which belongs to the nuclear hormone receptor superfamily of transcription factors (5,6,10,11,12). Upon stimulation, PPAR-
binds to responsive elements located in the promotor regions of many genes and modulates their transcriptive activities (2,10). Convincing evidence for an important role of PPAR-
in TZD-induced insulin sensitization includes insulin sensitization by PPAR-
agonists that do not belong to the TZD class (13,14,15,16) and by LG-100.268, an agonist of retinoid X receptor (RXR), which is the heterodimeric partner of PPAR-
(17). Furthermore, a strong correlation of antidiabetic efficacies of PPAR-
agonists in vivo with their respective potentials to bind and activate PPAR-
in vitro has been demonstrated (6,12,16).
Although there is evidence to support PPAR- having an important role in TZD-induced insulin sensitization, TZDs also seem to address other mechanisms that do not involve PPAR-
. That metabolic responses to TZDs in several experimental setups are independent of PPAR-
induced gene transcription is indicated by their rapid occurrence (5,7,18,19,20,21,22,23,24) and by a failure of the amplitudes of such responses to reflect the PPAR-
activating efficacies of different TZDs (5,20). The relevance of PPAR-
is specifically unclear in skeletal muscle, which quantitatively is the most important target tissue for insulin and plays a predominant role in TZD-induced improvement of glucose homeostasis (25,26). Although PPAR-
mRNA is abundant in fat and hardly found in skeletal muscle (10,27,28), the abundance of PPAR-
protein in muscle was recently reported to be 67% of that in fat (29). However, clear experimental evidence for any functional PPAR-
signaling in skeletal muscle has never been provided. Because relevance of direct TZD effects on skeletal muscle for antidiabetic action is still being debated, we examined the interaction of TZDs with isolated skeletal muscle fuel metabolism.
We recently reported that troglitazone had rapid and direct action on fuel metabolism of freshly isolated rat skeletal muscle in vitro, action that was characterized by distinct inhibition of CO2 production from palmitate and glucose (19). Troglitazone shifted glycolytic flux from the aerobic toward the anaerobic pathway simultaneous with glycogen depletion, which was marked after exposure of muscle specimens to troglitazone for 25 h (19). In the present study, we aimed to investigate the mechanisms responsible for direct interaction of TZDs with in vitro fuel handling in skeletal muscle using specimens of isolated rat soleus muscle. In particular, the studies were designed to provide evidence for or against the hypothesis that TZDs directly affects muscle fuel metabolism independent of PPAR--induced modulation of gene expression.
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RESEARCH DESIGN AND METHODS |
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Compounds.
The PPAR- agonistic TZDs troglitazone, pioglitazone, and rosiglitazone, as well as the RXR agonist LG-100.268 (17), were generously provided by Sankyo (Tokyo, Japan); the TZDs BM13.1258 and BM15.2054 (5) were generously provided by Boehringer-Mannheim/LaRoche (Mannheim, Germany); the TZD T-174 (8) was generously provided by Tanabe (Saitama, Japan); the non-TZD PPAR-
plus PPAR-
agonist JTT-501 (13,14) was generously provided by Tobacco (Osaka, Japan); and the natural PPAR-
ligand 15-deoxy-
12,14-prostaglandin J2 (30) and vitamin E succinate were purchased from Calbiochem (La Jolla, CA).
Long-term muscle incubation (5 or 25 h).
Immediately after rats were killed, two (SD rats) or three (Zucker rats) longitudinal soleus muscle strips per leg (i.e., four or six strips per rat) were prepared, weighed (25 mg/strip), and tied under tension on stainless steel clips, as previously described (31). According to procedures used earlier (19,32), muscles were immediately put into 50-ml Erlenmeyer flasks coated with BlueSlick solution (Serva, Heidelberg, Germany) and placed into a shaking waterbath (four or six strips per flask, 37°C, 130 cycles/min). Each flask contained 20 ml Cell Culture Medium 199 (5.5 mmol/l glucose [pH 7.35]; cat. no. M-4530; Sigma, St. Louis, MO) supplemented with 0.3% (wt/vol) fatty acid-free bovine serum albumin (BSA), 5 mmol/l HEPES, 25,000 units/l penicillin G, and 25 mg/l streptomycin. Palmitate was dissolved in ethanol and added to the medium to give final concentrations of 300 µmol/l palmitate and 0.5% ethanol (vol/vol). An atmosphere of 95% O2/5% CO2 was continuously provided within the flasks.
TZDs, JTT-501, prostaglandin J2, LG-100.268, and vitamin E succinate were dissolved in DMSO (Sigma) and added to the medium to give the final indicated drug concentrations. In some experiments, the medium also contained 1 mg/l of the protein synthesis inhibitor cycloheximide or 1 mg/l of the nucleic acid synthesis inhibitor (i.e., gene transcription inhibitor) actinomycin D. The final DMSO concentration in incubation media, including controls, was 0.1% (vol/vol) in all experiments. In experiments designed for direct comparison of the various agents, the first of four muscle strips from the same SD rat was incubated in the absence of any drug (control), the second was exposed to troglitazone, and the third and fourth were exposed to other compounds. Hence an intraindividual control was available, and troglitazone action was confirmed for each rat.
After pre-exposure periods of 4 or 24 h, muscles were immediately transferred into 25-ml flasks and incubated in 3 ml of identical buffer solution (one strip per flask). Muscles were then incubated for another 60 min, during which metabolic rates were determined (measurement period). During the measurement period, the media alternatively contained trace amounts of D-[U-14C]glucose, [U-14C]palmitic acid, or 2-deoxy-D-[2,6-3H]glucose ([3H]2DG) plus [U14C]sucrose (all from Amersham, Amersham, U.K.), and if not stated otherwise, 100 nmol/l insulin. At the end of the measurement period, muscles were quickly removed, blotted, and frozen in liquid nitrogen. Later, muscle strips were lysed in 1 mol/l KOH at 70°C; the lysate was then used for further analytical procedures, as described below.
The method used for determining CO2 production was based on the addition of radiolabeled palmitate or glucose to the incubation medium. Hence the data reflect CO2 production from the extracellular substrate only, but neglect CO2 production from intracellular lipid or glycogen pools. Some experiments were designed to determine CO2 production from intracellular lipid and glycogen stores prelabeled with [U-14C]palmitic acid or D-[U-14C]glucose, respectively. In these experiments, trace amounts of the radioactive compounds were added to the medium during a 24-h pretreatment period. Subsequently, CO2 production was measured without radioactive tracers in the medium.
Short-term muscle incubation (30 or 60 min).
Muscles from SD rats were put into coated 25-ml Erlenmeyer flasks that were placed into the waterbath immediately after preparation (one strip per flask). Each flask contained a continuous atmosphere of 95% O2/5% CO2 and 3 ml Krebs-Ringer buffer solution (pH 7.35) supplemented with 5.5 mmol/l glucose, 0.3% (wt/vol) BSA, 300 µmol/l palmitate, 0.5% (vol/vol) ethanol, and 0.1% (vol/vol) DMSO.
In the short-term experiments, the pretreatment period lasted for 30 min, after which muscles were immediately transferred into another set of flasks. Then muscles were incubated for another 30 min in 3 ml of identical buffer solution, which was supplemented with the above-described radioactive tracers and 30 nmol/l insulin (measurement period). A total of 20 µmol/l troglitazone was added to the incubation medium during the measurement period only or during both pretreatment and measurement periods, resulting in troglitazone exposure periods of 30 and 60 min, respectively. Finally, muscles were quickly removed, blotted, frozen, and lysed in KOH for further analytical procedures.
Analytical procedures.
Net uptake of [3H]2DG was determined using [14C]sucrose as a marker of extracellular space by previously described methods (19). Under the applied experimental conditions, insulin-stimulated [3H]2DG uptake did not reach saturation within the measurement period of 60 min (data not shown). The net rate of glucose incorporation into glycogen, referred to as glycogen synthesis, was determined by measuring the conversion of [14C]glucose to [14C]glycogen, as previously described (31). Rates of CO2 production were calculated from the conversion of [14C]glucose or [14C]palmitate into 14CO2, which was trapped with a solution containing methanol and phenethylamine (1:1) (33). Rates of lactate release were calculated from the amount of lactate accumulated in the incubation medium during the measurement period; this concentration was determined enzymatically using the lactate dehydrogenase method (34). For the determination of muscle glycogen content prevailing at the end of the experiment, glycogen in the muscle lysate was completely degraded to glucose with amyloglucosidase (33). Glucose was then measured enzymatically by a commercial kit (Human, Taunusstein, Germany).
Positive control experiments on cycloheximide and actinomycin D.
To confirm that cycloheximide efficiently blocked protein synthesis in our experimental setup, muscle strips were incubated in the absence or presence of 1 mg/l cycloheximide under the described conditions, except that culture medium devoid of methionine was used (modified Eagles medium; cat. no. 31900-020, Life Technologies, Paisley, U.K.), which was supplemented with all ingredients as listed above for Medium 199 plus 0.25 µCi/ml [35S]methionine. The effects of troglitazone and cycloheximide on soleus muscle fuel handling were not influenced by the medium used (Medium 199 versus modified Eagles medium; data not shown). After 4 or 24 h, muscles were quickly removed, blotted, and frozen for the determination of net [35S]methionine incorporated into protein. Later, muscle specimens were thawed and immediately lysed in 0.5 ml NaOH (1 mol/l), and the protein was precipitated with 0.6 ml perchloric acid (1 mol/l). After centrifugation, the supernatant was discarded and the pellet was redissolved in 0.5 ml NaOH. After this procedure was repeated twice, 0.6 ml perchloric acid were added and the sample was counted for 35S radioactivity.
To provide evidence that 1 mg/l actinomycin D inhibited transcription in our experimental setup, muscle strips were homogenized immediately after incubation for 24 h in Medium 199, as described above. Total RNA was isolated from muscle homogenates with RNAzol B following the instructions of the manufacturer (Tel-Test, Friendswood, TX), and the RNA content of the extracts was determined photometrically (ratio 260:280 nm >1.8).
Statistics.
All results are given as means ± SE. P values were calculated by two-tailed paired Students t test, and P < 0.05 was considered significant.
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RESULTS |
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Time dependence of troglitazone action.
Exposure of SD rat muscle to 20 µmol/l troglitazone for 30 min significantly increased the rate of insulin-stimulated lactate release (+ 27%; P < 0.01). After 60 min, the troglitazone-induced increase in lactate release was accompanied by significant reductions in mitochondrial fuel oxidation and glycogen storage (Table 2).
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Marked effects of both cycloheximide and actinomycin D on fuel handling of isolated rat muscle were observed after 25 h, whereas only minor effects of actinomycin D but no effects of cycloheximide were observed after 5 h. All results were consistent, in that they clearly demonstrated the failure of cycloheximide and actinomycin D to block troglitazones inhibitory action on fuel oxidation and glycogen synthesis. Thus, the troglitazone-induced reductions in glycogenesis or CO2 production from palmitate and glucose were not affected by inhibition of gene expression (Fig. 1).
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Regarding the non-TZD PPAR- agonists prostaglandin J2 and JTT-501, 5 µmol/l of these substances did not modulate fuel metabolism in the manner observed in response to TZDs. JTT-501 failed to affect any parameter measured, and prostaglandin J2 moderately stimulated glycogenesis and palmitate oxidation to CO2 (P < 0.05 each).
Parallel experiments were performed using LG-100.268 and vitamin E succinate, which are not ligands for PPAR-. Both compounds failed to influence glucose transport or glycogen metabolism. LG-100.268 inhibited CO2 production from palmitate (P < 0.0001) and marginally increased lactate release (P < 0.05), and vitamin E moderately decreased CO2 production from both palmitate and glucose (P < 0.05 each).
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DISCUSSION |
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First, the experiments on time dependence and interaction with actinomycin D and cycloheximide argue against any role for PPAR--mediated transcription or translation in troglitazones direct effects on muscle fuel metabolism in vitro. Thus distinct responses to troglitazone occurred within 30 min and became marked within 60 min, a finding that would argue against the established view that triggering metabolic effects via the modulation of gene expression requires a more prolonged time period. In addition, actinomycin D and cycloheximide, currently regarded as potent blockers of gene transcription and protein synthesis, respectively, clearly failed to counteract troglitazone action in our experimental setup.
Second, our experiments revealed that direct troglitazone action on isolated skeletal muscle is independent of concomitant insulin stimulation (see also 19), and that muscles isolated from obese Zucker rats, which exhibit lower rates of glucose metabolism than muscles from lean littermates (31), had similar dose-dependent responses to troglitazone as did muscle strips from normally insulin-sensitive SD rats (detailed dose-response curve for SD rat muscle in 19). These findings indicate major differences between TZD in vivo versus in vitro action, because the metabolic effects of TZDs in vivo are seen only in the presence of insulin, and TZD action is hardly detectable in healthy rodents but is very distinct in insulin-resistant animals (7,9). Furthermore, the inhibition of fuel oxidation observed in obese rat muscle exposed to TZDs in vitro contrasts with the increased glucose oxidation and insulin sensitization of the glycogenic pathway prevailing in skeletal muscle isolated from insulin-resistant rodents orally treated with TZDs in vivo (5,36,37). Given the widely accepted assumption that TZD-induced insulin sensitization in vivo relies on PPAR- activation (2,6,10,11,12,16), such divergences in TZD in vivo versus in vitro action support our conclusion that a differenti.e., a PPAR-
-independentmechanism must underlie modulation of fuel metabolism in isolated muscle.
Third, we compared the efficacies of different PPAR- agonistic compounds and found further evidence against the involvement of PPAR-
. Our results clearly indicated that the observed shift in fuel utilization from aerobic toward anaerobic pathways is specific for drugs belonging to the TZD class. This conclusion is based on the finding that all six TZDs tested markedly stimulated lactate release and inhibited CO2 production, whereas the two non-TZD PPAR-
agonists, JTT-501 and prostaglandin J2, failed to trigger such responses. Considering the inability of non-TZD PPAR-
agonists to elicit responses as triggered by the same concentration of a TZD, it is of note that the RXR agonist LG-100.268 inhibited the conversion of palmitate to CO2. Although final conclusions cannot be drawn from the present study, it is possible that different mechanisms mediate the actions of TZDs and LG-100.268, as RXR is the heterodimeric partner for a number of other receptors beside PPAR-
(17).
Comparison of the relative potencies of different compounds belonging to the TZD class further corroborates our interpretation that the observed responses to TZDs in vitro were not related to PPAR- activation. Focusing on the three TZDs that have been used for regular patient treatment, the relative PPAR-
agonistic efficacy of rosiglitazone is greater than that of pioglitazone, which in turn is greater than that of troglitazone (5,6,12,16,20). This ranking, however, is reversed by these TZDs respective potentials to inhibit mitochondrial fuel oxidation in vitro, in which case troglitazone is ranked higher than pioglitazone, which in turn is ranked higher than rosiglitazone. The relative efficacies of TZDs to modulate fuel metabolism in isolated muscle therefore clearly fail to reflect their PPAR-
agonistic potentials. In this context, it should be emphasized that the strong correlation of PPAR-
agonism in vitro with insulin-sensitizing efficacy in vivo is often regarded as the most convincing evidence for PPAR-
activation being an early and essential step in TZD-induced insulin sensitization (6,12,16). In contrast, we conclude that the different rankings of efficacy in our experimental setting provide good evidence that PPAR-
activation is not relevant for direct interaction of TZDs with skeletal muscle fuel handling in vitro.
The fact that all TZDs share the ability to inhibit aerobic fuel metabolism in vitro suggests a role for the TZD ring structure common to this group of molecules (2,12). If so, differences in efficacy among the TZDs used are likely to rely on other structures attached to the TZD ring. In the case of troglitazone, the TZD ring is attached to a molecular structure also found in vitamin E, which we showed to be a moderate inhibitor of CO2 production in isolated muscle. Hence independent potentials to inhibit fuel oxidation to CO2 may be intrinsic to both parts of the troglitazone molecule, and may interact in a synergistic manner to render troglitazone with extraordinary potency to directly influence fuel metabolism of isolated muscle. The different pharmacokinetic properties of the specific TZD, however, may also be of relevance.
The marked reduction of CO2 production from palmitate observed in TZD-exposed isolated rat muscle is in line with findings from several other studies likewise reporting that TZD can inhibit oxidative steps of lipid metabolism in vitro. Thus TZD exposure has been demonstrated to distinctly reduce ketone body production from oleate in rat hepatocytes (21), progesterone production in porcine granulosa cells (22), and cholesterol synthesis in several cell lines related to adipose tissue, liver, and muscle (20). Notably, many parallels exist with our findings, including 1) superiority of troglitazone over pioglitazone and rosiglitazone (20,21,22), 2) the occurrence of responses within a short time range (20,21,22), 3) the lack of modulation by cycloheximide and actinomycin D (20), and 4) the independence of the concomitant presence of insulin (19,20,21,22).
In conclusion, our results demonstrated that TZDs directly and rapidly affect fuel metabolism of isolated native rat skeletal muscle independent of PPAR- activation and gene expression. The observed shift of fuel utilization from aerobic to anaerobic pathways obviously has no functional consequence for the ability of TZDs to act as insulin sensitizers in vivo, a finding that agrees with the suggested importance of PPAR-
for antidiabetic TZD action. Nevertheless, the obtained data suggest a TZD-specific potential to affect fuel metabolism via PPAR-
-independent mechanisms of action. The contribution of such PPAR-
-independent actions to the beneficial and/or unwanted actions of TZDs in vivo, however, is unclear and may differ considerably among the various antidiabetic agents belonging to the TZD class.
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ACKNOWLEDGMENTS |
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We thank Sankyo (Tokyo, Japan), Tobacco (Osaka, Japan), Boehringer-Mannheim/LaRoche (Mannheim, Germany), and Tanabe (Saitama, Japan) for generously providing the compounds. We also thank the staff at the Biomedical Research Center, University of Vienna, for taking care of the rats.
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FOOTNOTES |
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Received for publication 15 June 2000 and accepted in revised form 27 June 2001.
BSA, bovine serum albumin; [3H]2DG, 2-deoxy-D-[2,6-3H]glucose; PPAR, peroxisome proliferatoractivated receptor; RXR, retinoid X receptor; TZD, thiazolidinedione.
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REFERENCES |
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