| | PPAR-γ response element activity in intact primary human adipocytes: effects of fatty acidsReceived 10 January 2005; accepted 26 April 2005. published online 13 October 2005. Abstract ObjectiveWe studied the activity and regulation of the peroxisome proliferator-activated receptor-γ response element (PPRE) in primary human adipocytes. MethodsWe transfected primary human adipocytes with a plasmid-encoding firefly luciferase cDNA under control of a PPRE from the acyl-coenzyme A oxidase gene by using our newly developed electroporation-based method. Several fatty acids were added to the fat cells to study potential activation of peroxisome proliferator-activated receptor-γ. ResultsCells responded maximally to 5 μM of rosiglitazone at a 5.1 ± 1.4-fold over basal increase in luciferase activity. There was a positive correlation between body mass index and the response to 5 μM of rosiglitazone (r = 0.36, P = 0.03). Patients with type 2 diabetes had similar basal PPRE activity but responded more strongly to 5 μM of rosiglitazone than did non-diabetic subjects (10.2 ± 5-fold and 5.4 ± 1-fold over basal increase, respectively, P < 0.0001). Among saturated fatty acids, lauric acid was without effect, but 10 μM of palmitic or stearic acid increased PPRE activity 20% to 35% above basal levels. Monounsaturated palmitoleic acid at 1 μM induced a PPRE transcriptional activity that corresponded to half the therapeutic levels of rosiglitazone. ConclusionAdipocytes from obese subjects and patients with type 2 diabetes responded particularly strongly to the effect of rosiglitazone on PPRE. Because fatty acids in the diet can affect the transcriptional activity of peroxisome proliferator-activated receptor-γ over decades, the stimulation induced by stearic and palmitoleic acids can affect insulin sensitivity and, hence, cardiovascular morbidity and mortality in humans.
Introduction  Transcription factor peroxisome proliferator-activated receptor-γ (PPAR-γ) is strongly expressed in adipocytes and has been shown to affect several genes of importance for differentiation of fat cells and for insulin sensitivity [1], [2]. PPAR-γ heterodimerizes with retinoid-X receptor-α (RXR-α) after ligand binding [3]. The synthetic PPAR-γ agonist rosiglitazone increases mRNA levels of the insulin-inducible glucose transporter-4 [4]. Treatment with PPAR-γ activators such as rosiglitazone and pioglitazone is currently a treatment option that lowers blood glucose levels and improves other components of the metabolic syndrome in patients with type 2 diabetes [1]. Dominant negative mutations of PPAR-γ have been found in subjects who are severely insulin resistant and develop type 2 diabetes at a young age [5], [6]. This highlights the importance of a fully functioning PPAR-γ system for normal insulin signaling in humans. The physiologic ligand for PPAR-γ activation in humans is not known. However, according to in vitro studies, several fatty acids can bind to PPAR-γ [7], [8]. This is of particular interest because ingestion of diets rich in oils containing unsaturated fatty acids have been linked to a more insulin-sensitive phenotype and even to decreased cardiovascular morbidity and mortality according to epidemiologic surveys [9], [10], [11], [12]. In contrast, ingestion of other types of fatty acids, in particular unsaturated trans-fatty acids, has been linked to an unfavorable phenotype in humans [12], [13], [14], [15]. Treatment of insulin-resistant subjects with conjugated trans-10 cis-12 linoleic acid (CLA), marketed as a weight-reduction agent, has been shown to increase insulin resistance and markers of inflammation in humans [14], [16]. Interestingly, in rodents CLA increases insulin sensitivity [17], which highlights the importance of using relevant models when searching for mechanisms behind type 2 diabetes and the metabolic syndrome in humans. Dietary fatty acids might affect adiposity and insulin sensitivity by binding to PPAR-γ in adipocytes [12]. However, strong synthetic ligands of PPAR-γ, such as rosiglitazone and pioglitazone, often induce side effects including peripheral edema, dilutional anemia, and even overt heart failure. This is generally believed to be a drug class side effect, i.e., a direct consequence of potent PPAR-γ activation [18]. Because there are no similar reports of adverse effects caused by ingestion of fatty acids from food, it is likely that most fatty acids at best are weak activators of PPAR-γ. In contrast, fatty acids in the diet have the potential to affect transcription through PPAR-γ over decades; thus, even a relatively weak agonist activity could be of clinical interest. The ability or potency of different fatty acids to activate PPAR-γ and the PPAR-γ response element (PPRE) has not been examined in human primary fat cells because no method to transfect these cells has hitherto been available. We recently developed an electroporation-based transfection method that can be used for molecular biological studies of signal transduction in primary human adipocytes [19]. In the present study, we examined the effects of fatty acids on activation of PPRE in the physiologically most relevant cells, primary human adipocytes. Materials and methods Materials The expression plasmid pAOX-luc, the GAL4-PPAR-γ construct and 5XGAL4-TK-LUC have been described previously [20]. Plasmid pRluc was from BioSignal (Packard, CT, USA). The pCDNA3-hSRC-1 was kindly provided by Dr. R. Kurokawa (University of California, San Diego, La Jolla CA, USA), and pCDNA3-mTIF2 and pCDNA3-mpCIP were kindly provided by Dr. J Torchia (London Regional Cancer Center, London, ON, Canada). The PPAR-γ antibody sc-6285 was from Santa Cruz Biotechnology (Santa Cruz, CA, USA) and fatty acids were from Larodan (Malmö, Sweden). All fatty acids except CLA were of greater than 99% purity as determined by high-performance liquid chromatography. CLA was 95% pure. Fatty acids were stored at −20°C and used well within the time frames of stability. Ethics The study was approved by the ethics committee of Linköping University (Linköping, Sweden). Informed consent was obtained from all participating patients. Statistics Statistical calculations were made using StatView 4.5 (Abacus Concepts Inc., Berkeley, CA, USA). Comparisons within and between groups were made with Student’s paired and unpaired two-tailed t tests and linear correlations with Pearson’s test unless stated otherwise. Data are presented as mean ± standard deviation. Statistical significance was set at the 5% level (P ≤ 0.05). Subjects and isolation of adipocytes Subcutaneous abdominal human adipose tissue was removed during surgery. The patients were women who underwent surgery that usually required hysterectomy for various gynecologic diseases. The 43 non-diabetic patients were 61 ± 17 y old with a body mass index (BMI) of 26.5 ± 4 kg/m2 (mean ± standard deviation). Patients were not investigated with regard to insulin sensitivity. Three subjects with diagnosed type 2 diabetes mellitus were analyzed separately and received treatment with insulin plus sulfonylurea and/or metformin. Patients with diabetes were 58 ± 9 y old and had a BMI of 31 ± 4 kg/m2 and hemoglobin A1c level of 5.8 ± 0.4% (range 5.5–6.3%, reference range 4.5–5.0%). Adipose tissue was cleared from vascular and fibrous structures and cut with scissors into millimeter-size pieces. Fat tissue weighting 5 to 10 g was rinsed in 0.9% (w/v) NaCl and digested in 5 mL of Krebs-Ringer solution (0.12 M of NaCl, 4.7 mM of KCl, 2.5 mM of CaCl2, 1.2 mM of MgSO4, and 1.2 mM of KH2PO4) containing 20 mM of Hepes, pH 7.4, 3.5% (w/v) fatty acid-free bovine serum albumin, 200 nM of adenosine, 2 mM of glucose, and 260 U/mL of collagenase (type 1; Worthington Biochemical Corporation, Lakewood, NJ, USA Worthington, NJ, USA) for 1.5 h at 37°C in a shaking water bath. After collagenase digestion, adipocytes were separated from tissue debris by filtering. Cells were then washed at 40% cells by volume in Krebs-Ringer solution containing 20 mM of Hepes, pH 7.4, 1% (w/v) fatty acid-free bovine serum albumin, 200 nM of adenosine, and 2 mM of glucose and kept in a shaking water bath at 37°C [21] for a maximum of 30 min until electroporation. Electroporation and luciferase assay Each 0.4-cm gap of the electroporation cuvette was filled with 200 μL of adipocyte solution. An additional amount of 200 μL of phosphate buffered saline (137 mM of NaCl, 2.7 mM of KCl, 10 mM of Na2HPO4, and 1.8 mM of KH2PO4, pH 7.5) containing 2 μg of pAOX-luc and 0.1 μg of pRluc plasmid DNA was then mixed with the cells. In some experiments plasmid DNA coding for the coactivators SRC-1, pCIP, and TIF2 (0.2 μg of each) was added. Each cuvette was electroporated with a 400-V 4-ms square-wave pulse using a Bio-Rad GenePulser II (Bio-Rad Laboratories Inc., Hercules, CA, USA). Cells from three cuvettes were pooled, transferred to Petri dishes, and kept at 37°C in 10% CO2. One hour after electroporation, an equal volume of Dulbecco’s Modified Eagle’s Medium, pH 7.5, containing 25 mM of glucose, 50 UI/mL of penicillin, 50 μg/mL of streptomycin, 200 nM of phenyl-isopropyl-adenosine, 7% (w/v) fatty acid-free bovine serum albumin, and 25 mM of Hepes was added. Rosiglitazone or fatty acids were added in an ethanol solution 1 h after electroporation. Final ethanol concentration was 0.5% (v/v) and final albumin concentration was 3.75% (w/v). Luciferase activity was analyzed after incubation for 18 h at 37°C in 10% CO2 after electroporation. Cells were homogenized and assayed for firefly and renilla luciferase using the Dual-Luciferase Reporter Assay Systems (Promega, Madison, WI, USA). Cell lysates were prepared with 200 μL of supplied buffer by passing twice through a 25-gauge needle. The firefly luciferase assay was initiated by adding 100 μL of Luciferase Assay Reagent II to each 50-μL aliquot of cell lysate. After quantifying the firefly luminescence, the reaction was quenched and the renilla luciferase reaction was activated by adding 100 μL of Stop & Glo Reagent (Promega). The firefly and renilla luciferase activities were measured using a Victor 1420 multilabel counter (Wallac, Turku, Finland). Immunoblotting Whole cell lysates were subjected to sodium dodecylsulfate polyacrylamide gel electrophoresis and transferred to polyvinylidene diflouride membrane for immunoblotting with antibodies against human PPAR-γ (diluted 1:250). Bound antibodies were detected with horseradish peroxidase–conjugated anti-immunoglobulin G according to the ECL+ protocol (Amersham Biosciences, Uppsala, Sweden). Results Response to rosiglitazone Primary human adipocytes were transfected with a plasmid that encodes firefly luciferase cDNA under the control of a PPRE from the acyl-coenzyme A oxidase gene (pAOX). The cells were also transfected with a plasmid encoding renilla luciferase controlled by a constitutively active promoter (pRluc). PPRE-inducible firefly luciferase activity was normalized according to renilla luciferase, thus correcting for differences in the amount of transfected cells. Cells treated with 0.01 to 5 μM of the synthetic PPAR-γ agonist rosiglitazone responded in a dose-dependent manner, with a maximal 5.1 ± 1.4-fold increase in PPRE (Fig. 1). All potential ligands to PPAR-γ examined in this study were added to the cells in ethanol solution. There was no effect of ethanol on PPRE activity (no vehicle: 0.0035 ± 0.003 firefly luciferase/renilla ratio, ethanol: 0.0033 ± 0.001 firefly luciferase/renilla ratio, n = 6, P = 0.15 by paired t test). In some cells and species, addition of RXR-α ligands have been found to increase PPRE activity [22], [23]. However, addition of the RXR-α agonist LG1069 yielded variable and statistically non-significant effects on PPRE activity (P = 0.17 by paired t test; Fig. 2). Thus, in human adipocytes, ligand binding to PPAR-γ rather than RXR-α increases PPRE activity. Levels of coactivators are crucial for the transcriptional activity of PPAR-γ. We thus tested the three coactivators SRC-1, TIF2, and pCIP. Overexpression of pCIP or TIF2, but not of SRC-1, slightly increased the response to rosiglitazone (Fig. 3). Because coactivators generally are believed to mainly affect the activated PPAR-γ receptor, combinations of SRC-1, TIF2, and pCIP and 10 μM of palmitoleic acid were added to the cells. However, these combinations did not increase the effect when compared with addition of only the coactivator (all three corresponding P values were greater than 0.2 by paired t tests calculated from five individual experiments, not shown). Relations of PPRE to BMI and diabetes There was no correlation between the age of the non-diabetic subjects and basal PPRE activity in the adipocytes (Fig. 4A) or the maximal response to rosiglitazone (P = 0.7, not shown). However, when analyzed in relation to BMI, there was a weak trend toward a negative correlation to basal PPRE activity in non-diabetic subjects (r = −0.31, P = 0.058, Spearman’s rank test P = 0.07; Fig. 4B; corresponding figures when the three patients with type 2 diabetes were included: r = −0.29, P = 0.054, Spearman’s rank test P = 0.047). There was a positive correlation between BMI and the response to 5 μM of rosiglitazone (r = 0.36, P = 0.03, Spearman’s rank test P = 0.03; Fig. 4C; corresponding figures when the three patients with type 2 diabetes were included: r = 0.45, P = 0.002, Spearman’s rank test P = 0.007). The maximal response to rosiglitazone was negatively correlated to basal PPRE activity in non-diabetic subjects (r = −0.38, P = 0.01, Spearman’s rank test P = 0.005; corresponding figures when the three patients with type 2 diabetes were included: r = −0.32, P = 0.03, Spearman’s rank test P = 0.02). The three patients with type 2 diabetes were also examined separately. These patients tended to be more obese than the non-diabetic subjects (BMI 31 ± 4 and 26 ± 4 kg/m2, respectively, P = 0.06) but basal levels of PPRE activity were similar (P = 0.5). Patients with type 2 diabetes responded more strongly to 5 μM of rosiglitazone than did non-diabetic subjects (Fig. 5) and this difference was independent of BMI in multivariate analysis (P = 0.005 after correction for BMI). Effects of fatty acids We next examined whether a range of fatty acids, which are normally present in different food sources, could activate PPRE. Table 1 lists the fatty acids examined and in which type of nutrients they can be found. Physiologic levels of the tested fatty acids in the cytosol of human fat cells are not known. We examined the fatty acids at concentrations of 1 and 10 μM because in vitro assessment of ligand binding of mono- and polyunsaturated fatty acids have demonstrated a half maximal effective concentration lower than 10 μM [7], [8]. Of the saturated fatty acids, the short 12-carbon lauric acid was without any statistically significant effect, but long-chain palmitic and stearic acids increased PPRE activity (Fig. 6A). The three examined long-chain monounsaturated fatty acids also increased PPRE activity, whereas a range of polyunsaturated ω-6 fatty acids, except for 1 μM of linoleic acid, had no statistically significant effect (Fig. 6B,C). The ω-3 fatty acids linolenic acid and eicosapentaenoic acids, but not docosahexaenoic acid, also stimulated PPRE activity (Fig. 6D). Because free fatty acids can reach levels up to 100 μM in human plasma [24], we examined the addition of 50 and 200 μM of oleic and palmitoleic acids and compared it with the effect seen for 10 μM. These concentrations did not further increase PPRE transcriptional activity (data not shown; n = 4, P = 0.7 for either paired comparison). | | |  | Fatty acid | Composition | Classification | Examples of dietary source | |
|---|
| Lauric | C12H24O2 | Saturated | Coconut oil | |
| Palmitic | C16H32O2 | Saturated | Most fats and oils | |
| Stearic | C18H36O2 | Saturated | Meat and chocolate | |
| Palmitoleic | C16H30O2 | Monounsaturated, ω-7 | Most fats and oils | |
| Oleic | C18H34O2 | Monounsaturated, ω-9 | Olive oil and generally in seed oils | |
| Petroselinic | C18H34O2 | Monounsaturated, ω-12 | Parsley and coriander seed oil | |
| Linoleic | C18H32O2 | PUFA, ω-6, 2 double bonds | Vegetable oils, particularly in corn oil | |
| γ-Linolenic | C18H30O2 | PUFA, ω-6, 3 double bonds | Vegetable oils and black currant | |
| CLA | C18H32O2 | PUFA, ω-6, trans-10, cis-12 CLA | Meat and diary products | |
| Linoelaidic | C18H32O2 | PUFA, ω-6, 9 trans, 12 trans-isomer of linoleic acid | Artificially hardened oils | |
| Linolenic | C18H30O2 | PUFA, ω-3, 3 double bonds | Soybean oils | |
| EPA | C20H30O2 | PUFA, ω-3, 5 double bonds | Fish | |
| DHA | C22H32O2 | PUFA, ω-3, 6 double bonds | Fish | | | | |
CLA, found in cows’ milk, is a PPAR-γ ligand in vitro and has been shown to increase insulin resistance when given to obese subjects [16]. Thus CLA could act to block PPAR-γ activity in humans. However, CLA was without effect on PPRE activity in human adipocytes (Fig. 6C). Exploring the possibility that CLA might interfere with other PPAR-γ ligands, we added 10 μM of CLA to increasing doses of rosiglitazone, but CLA did not interfere with the effects of rosiglitazone (Fig. 7). We also tested whether CLA decreases the amount of PPAR-γ protein in primary human adipocytes, as has been demonstrated in 3T3-L1 adipocytes [25]. There was no effect on the amount of PPAR-γ receptor by treatment with 1 or 10 μM of CLA for 18 h (Fig. 7, insert). The trans-fatty acid linoelaidic acid, a byproduct of artificial hardening of vegetable oils, was also without effect on PPRE activity (Fig. 6C). To ascertain that the increase in PPRE activity by fatty acids was mediated by PPAR-γ, we transfected the cells with a plasmid encoding GAL4-PPAR-γ fusion protein (amino acids 174 to 475 of PPAR-γ corresponding to the ligand binding domain, pGAL4-PPAR-γ) and a plasmid encoding luciferase under control of five copies of a GAL4 response element (5xGAL4-TK-LUC). The cells responded to 5 μM of rosiglitazone, with a 6.1 ± 2.8-fold increase over basal increase in luciferase activity. Likewise, addition of 10 μM of stearic or palmitoleic acid increased the activity 54 ± 22% and 26 ± 17%, respectively, over basal levels (Fig. 8). Discussion In humans PPAR-γ is expressed at particularly high levels in fat cells. The transcriptional activity of PPAR-γ varies depending on species and cell type [26] and is regulated not only by ligand binding but also by coactivators and corepressors [20], the levels of which vary greatly between cell types and species. Thus, for potential clinical implications of in vitro studies, it is important to use cells of human origin. The thiazolidinedione rosiglitazone is a well-characterized synthetic ligand to PPAR-γ that is used for treatment of insulin resistance and type 2 diabetes in humans [1]. We found that treatment of primary human adipocytes with rosiglitazone for 18 h induced a maximal five-fold increase in PPRE activity. The subjects who donated the fat cells varied considerably with respect to age and BMI. Although insulin sensitivity decreases with age, we found no correlation between age and basal PPRE activity or maximal response to rosiglitazone. However, there was a positive correlation between BMI and maximal response to rosiglitazone. There was also a weak trend toward a linear negative relation between basal PPRE activity and BMI that reached statistical significance in the total material (including the three patients with type 2 diabetes) when calculated with Spearman’s rank test. This implies that obese subjects are particularly sensitive to the effects of rosiglitazone on PPRE transcriptional activity due to a low endogenous activity of PPRE. There was a negative correlation between basal PPRE transcriptional activity and maximal response to rosiglitazone. The patients with type 2 diabetes had similar basal PPRE activity levels but responded more strongly to rosiglitazone than did non-diabetic subjects and this was unrelated to degree of obesity. In some cell types, PPRE activity can be induced by addition of RXR-α ligands [22], [23]. However, the RXR-α ligand LG1069 did not affect PPRE activity in human adipocytes. Thus it is unlikely that treatment of patients with this kind of compound will affect insulin sensitivity, at least through an effect on PPRE in fat cells. When the effect of coactivators was investigated in the presence of rosiglitazone, pCIP and TIF2 were found to increase the response to the thiazolidinedione but not to the weaker activator palmitoleic acid. This may suggest that endogenous levels of the coactivators pCIP and TIF2 might be of particular interest with regard to insulin sensitivity in humans. Having demonstrated that our system can reproducibly measure the effect of rosiglitazone on PPRE in primary human adipocytes, we assessed the potential effect of a range of fatty acids. Fatty acids are normally taken up and converted to triacylglycerols in response to insulin by adipocytes, whereas stored triacylglycerols are hydrolyzed in response to lipolytic hormones. To limit interference from these processes, we incubated the cells without hormonal stimulation and with controlled levels of adenosine [27]. In the absence of adenosine, lipolysis is strongly stimulated in adipocytes. We therefore added the stable adenosine analogue phenyl-isopropyl-adenosine. Numerous trials of the effect of ingestion of different fatty acids on markers of insulin sensitivity have been reported [12]. Although there is evidence suggesting favorable effects of mono- and polyunsaturated fatty acids on markers for insulin sensitivity and components of the metabolic syndrome, the mechanisms behind such effects in humans are not known. Further, in many cases, the results have been contradictory and some studies have found negative effects of polyunsaturated or ω-3 acids [28], [29] and stimulation of PPAR-γ by saturated fatty acids [7], [8]. Many trials have focused on subjects being overweight or overtly insulin resistant, making it difficult to assess whether the results are applicable to the general population [12]. Our assessment of the specific activation of PPRE in intact primary human adipocytes demonstrated an increase in transcriptional activity when cells were treated with the saturated fatty acids palmitic and stearic acids. In contrast, the polyunsaturated fatty acids linoleic acid, γ linolenic acid, and docosahexaenoic acid did not induce any statistically significant changes in PPRE activity. The monounsaturated palmitoleic acid increased basal PPRE activity by 35% at 1 μM, thus being more effective than the fish oil eicosapentaenoic acid in this regard. The concentration of rosiglitazone in plasma at therapeutic doses is 0.01 to 0.3 μM thus varying considerably depending on the interval between drug intake, or injection, and sampling plasma for analysis [30]. A physiologic level of rosiglitazone in the higher range of this interval corresponds to about a 2.5-fold increase in PPRE activity (Fig. 1). Hence, at 1 μM, palmitoleic acid induces a transcriptional activity level of PPRE in human fat cells that is 54% (1.35/2.5) of the effect of therapeutic plasma levels of rosiglitazone. Because fatty acids in the diet can affect transcriptional activity over decades compared with drugs that are used once disease has already developed, the activation of PPAR-γ induced by dietary fatty acids can have considerable effect on insulin sensitivity and, hence, cardiovascular morbidity and mortality. It is likely that the rather high final concentration of albumin, 3.75%, affected the response to rosiglitazone and to the tested fatty acids. However, this amount of albumin corresponds well to physiologic levels of albumin in human plasma, being in the range of 3.6% to 4.8% according to our clinical laboratory at the University Hospital of Linköping. Because stearic and palmitoleic acids activated the GAL4-PPAR-γ fusion protein in a manner similar to that of PPRE on the pAOX construct, which is dependent on endogenous PPAR-γ levels, saturated and unsaturated fatty acids can activate PPRE through the PPAR-γ receptor in intact human adipocytes. Our findings thus contradict the statement that ingestion of saturated fatty acids negatively affect cardiovascular risk and insulin sensitivity [12], [31], [32], [33]. This discrepancy between previous clinical trials and our present investigation may be due to an inability to isolate the effect of particular fatty acids when effects of diets are studied. It may also be due to a detrimental effect of the amounts of fat rather than of particular types of fatty acids [12]. Transcription factors other than PPAR-γ may also be involved when the effects on insulin sensitivity, or markers thereof, are tested with different diets. Because the trans-fatty acid CLA has been shown to induce insulin resistance and to worsen other markers of the metabolic syndrome, we paid particular attention to this fatty acid [16], [34]. The effect of CLA on PPRE activity was very variable, and although a stimulatory trend was seen, this was not statistically significant. We hypothesized that CLA might diminish the effect of coadministered rosiglitazone but found no such effect. In addition, CLA did not affect the amount of endogenous PPAR-γ protein in human adipocytes. Thus we found no effect of CLA on PPAR-γ activity in human adipocytes, which suggests that an increased insulin resistance due to CLA in humans may be mediated by other transcription factors, e.g., PPAR-α [35]. In this report we demonstrated differences in PPRE activity that were related to the degree of obesity in humans and that saturated and monounsaturated fatty acids induced activation of PPRE and PPAR-γ. 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MEDLINE a Department of Cell Biology, Faculty of Health Sciences, Linköping University, Linköping, Sweden b Diabetes Research Centre, Faculty of Health Sciences, Linköping University, Linköping, Sweden c Department of Molecular and Clinical Medicine, Faculty of Health Sciences, Linköping University, Linköping, Sweden d Department of Medicine and Care, Faculty of Health Sciences, Linköping University, Linköping, Sweden  Corresponding author. Tel.: +46-13-22-77-49; fax: +46-13-22-35-06.
This study was supported by the Swedish Society for Medical Research, the Swedish Research Council (grants 14254 and 12157), the Swedish National Board for Laboratory Animals, the Lions Foundation, the Östergötland County Council, the Swedish Diabetes Association, the Swedish Medical Society, and the University Hospital of Linköping Research Funds. PII: S0899-9007(05)00234-0 doi:10.1016/j.nut.2005.04.011 © 2006 Elsevier Inc. All rights reserved. | |
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