AJP - Heart BIOPAC complete lab solutions
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Heart Circ Physiol 295: H1044-H1055, 2008. First published July 3, 2008; doi:10.1152/ajpheart.00516.2008
0363-6135/08 $8.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
295/3/H1044    most recent
00516.2008v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Web of Science (1)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Bourgoin, F.
Right arrow Articles by Nadeau, A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Bourgoin, F.
Right arrow Articles by Nadeau, A.

Endothelial and vascular dysfunctions and insulin resistance in rats fed a high-fat, high-sucrose diet

Frédéric Bourgoin,1 Hélène Bachelard,1 Mylène Badeau,1 Sébastien Mélançon,1 Maryse Pitre,1 Richard Larivière,2 and André Nadeau3

1Department of Medicine and Lipid Research Unit from Centre Hospitalier Université Laval (CHUL) Research Center, 2Research Center of L'Hôtel Dieu de Québec, 3Diabetes Research, Centre Hospitalier Universitaire de Québec (CHUQ), Laval University, Quebec, Canada

Submitted 15 May 2008 ; accepted in final form 27 June 2008


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This study was designed to examine the effects of a high-fat, high-sucrose (HFHS) diet on vascular and metabolic actions of insulin. Male rats were randomized to receive an HFHS or regular chow diet for 4 wk. In a first series of experiments, the rats had pulsed Doppler flow probes and intravascular catheters implanted to measure blood pressure, heart rate, and regional blood flows. Insulin sensitivity and vascular responses to insulin were assessed during a euglycemic hyperinsulinemic clamp performed in conscious rats. In a second series of experiments, new groups of rats were used to examine skeletal muscle glucose transport activity and to determine in vitro vascular reactivity, endothelial nitric oxide synthase (eNOS) protein expression in muscle and vascular tissues and endothelin content, nitrotyrosine formation, and NAD(P)H oxidase protein expression in vascular tissues. The HFHS-fed rats displayed insulin resistance, hyperinsulinemia, hypertriglyceridemia, hyperlipidemia, elevated blood pressure, and impaired insulin-mediated renal and skeletal muscle vasodilator responses. A reduction in endothelium-dependent vasorelaxation, accompanied by a decreased eNOS protein expression in muscles and blood vessel endothelium, and increased vascular endothelin-1 protein content were also noted in HFHS-fed rats compared with control rats. Furthermore, the HFHS diet induced a reduced insulin-stimulated glucose transport activity in muscles and increased levels of NAD(P)H oxidase protein and nitrotyrosine formation in vascular tissues. These findings support the importance of eNOS protein in linking metabolic and vascular disease and indicate the ability of a Westernized diet to induce endothelial dysfunction and to alter metabolic and vascular homeostasis.

vascular endothelial function; oxidative stress

THE METABOLIC SYNDROME is associated with a clustering of cardiovascular risk factors presently encountered in the affected individual, including hypertension, dyslipidemia, type 2 diabetes and obesity, one of the most common nutritional disorders in modern civilization. The fundamental feature in the pathogenesis of the metabolic syndrome is considered insulin resistance (8, 41). Obesity, in particular abdominal obesity, was pointed out as a primary contributor to acquired insulin resistance, as increasing adiposity is correlated with impaired insulin action (13, 23). Dietary factors are likely to be of importance in the etiology of the metabolic syndrome in humans. Studies in rodents have shown that chronic consumption of diets high in fat and sugar (HFHS) induces several of the abnormalities characterizing the metabolic syndrome (1, 7, 45, 53), while most of these abnormalities are completely reversed by chronic withdrawal of the obesity-inducing HFHS diet (36). In humans, the metabolic syndrome can be controlled by feeding a low-fat, complex-carbohydrate diet combined with aerobic exercise (2). Endothelial dysfunction, an independent predictor of cardiovascular events (21, 31), has been consistently associated with the metabolic syndrome (18) in a complex interplay with insulin resistance (8). Deficiency of endothelial-derived nitric oxide (NO) is believed to be the primary defect that links insulin resistance and endothelial dysfunction (8, 45). A decreased synthesis and/or release, in combination with exaggerated consumption of NO in tissues by high levels of reactive oxygen (ROS) and nitrogen (RNS) species, which are produced by cellular disturbances in glucose and lipid metabolism, could be important limiting factors contributing to impair endothelial vasomotor function by reducing NO bioavailability (8, 17, 19, 43, 44).

In addition to its classical glucoregulatory actions, insulin has important nonmetabolic hemodynamic actions. Previous studies carried out in humans, dogs, and rats have shown that insulin causes peripheral vasodilation, promotes capillary recruitment, and increases regional blood flow (5, 33, 38, 39). These NO-mediated effects (48, 49) are considered an important physiological determinant of insulin action on glucose metabolism, by enhancing glucose delivery and exposing previous underperfused tissues, i.e., skeletal muscle, to insulin (5, 56). In conditions of insulin resistance, such as in hypertensive, diabetic, or obese subjects, the endothelial-mediated vasodilator responses to insulin are impaired (3, 6, 55). Given the central role of the endothelium and its vasorelaxing factor NO in maintaining vascular tone, regulating blood flow to organs and tissues, and mediating the blood flow regulatory effects of insulin, we hypothesize that a reduction in NO availability, induced by the consumption of an obesity-inducing HFHS diet in rats, could be critically instrumental in the development and/or the progression of insulin resistance by reducing glucose and insulin delivery to insulin-sensitive tissues. Vascular endothelium could be an important physiological target of insulin that couples regulation of insulin sensitivity and glucose metabolism with hemodynamics (60, 61). The purpose of the present study was to characterize, in a rat model of diet-induced insulin resistance and obesity, the vascular endothelium function in relation with the glucoregulatory and hemodynamic actions of insulin. Insulin resistance was induced in rats by feeding them an HFHS diet, which mimics to some extent the high-energy diets frequently consumed by many individuals. This diet maintains a high lipid flux and leads to increased fat accretion and alterations in glucose and lipid metabolism (34) that are similar to those observed in the human metabolic syndrome (16). In the present study, the rats were randomized to either HFHS or a normal chow diet for 4–5 wk, as we were interested in evaluating the effect of early experimental obesity on endothelial function and the metabolic and vascular actions of insulin. Although some other groups have reported insulin resistance and endothelial dysfunction in similar dietary models (1, 36, 43), none of these has conducted experiments looking in parallel at metabolic and regional hemodynamic actions of insulin in conscious HFHS-fed rats.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animals and feeding protocol. This study was approved by the Animal Care and Handling Committee of Laval University; animals were cared and handled in accordance with the Canadian Guide for the Care and Use of Laboratory Animals. Male Sprague-Dawley rats (n = 103) aged 6 wk and initially weighing 175–230 g were purchased from Charles River Laboratories (St-Constant, QC, Canada). The rats were individually housed in a light-controlled (12-h light cycle starting at 6 AM) and temperature-regulated (20–22°C) space. They were randomly divided into two dietary groups and for 4 wk had free access to water and one of two diets. The control group (n = 52) was fed standard rat chow (Teklad Global 18% Protein Rodent Diet no. 2018), while the test group (n = 51) was fed a purified HFHS diet. The diet components are listed in Table 1. The fat content of the HFHS diet was predominantly saturated (49%).


View this table:
[in this window]
[in a new window]

  		Table 1. Chow and HFHS diet composition

 
Surgical preparation. In vivo experiments were performed in HFHS- (n = 17) and chow-fed rats (n = 19). Three weeks after the start of the diets, the rats were anesthetized with a mixture of ketamine-xylazine (100 and 10 mg/kg, respectively, ip) and had pulsed Doppler flow probes implanted to monitor changes in renal, mesenteric, and hindquarter blood flows, as previously described (20, 38). After surgery, the rats were given subcutaneous injections of buprenorphine (0.05 mg/kg) and returned to their cages and allowed to recover for at least 7 days. The chow or HFHS diet continued during postsurgical recovery, and the latter was deemed satisfactory by the resumption of growth and normalization of 24-h food intake. After the recovery period, the rats were again anesthetized (as above), and two separate catheters were implanted in the right jugular vein (for iv infusions) and one catheter in the left carotid artery (for blood pressure and heart rate measurements) (35). The rats were given subcutaneous injections of buprenorphine (0.05 mg/kg) and returned to their cages. A euglycemic hyperinsulinemic clamp was then performed 72 h after this last surgical step. The rats were 10–11 wk old.

Euglycemic hyperinsulinemic clamp studies. Before each experiment, blood glucose and plasma insulin were determined, and resting heart rate, blood pressure, and regional blood flows were recorded over 30 min in quiet, unrestrained, and overnight-fasted rats. Following this, a euglycemic hyperinsulinemic clamp was carried out over 2 h in the designated groups as previously described (38), while heart rate, blood pressure, and regional blood flows were continuously measured. Insulin (Humulin R; Eli Lilly, Toronto, ON, Canada) was infused at a rate of 4 mU·kg–1·min–1 in a first group of chow-fed (n = 9) and HFHS-fed rats (n = 10). During this time, a dextrose solution (50% wt/vol) was infused at variable rates to maintain glycemia near the fasting level; the amount of dextrose infused to maintain euglycemia was plotted as whole body glucose infusion rate (GIR; expressed in mg·kg–1·min–1). In control experiments, a second group of chow-fed (n = 10) and HFHS-fed rats (n = 7) was infused with saline-0.2% BSA instead of insulin and dextrose to match approximately the saline load delivered during the clamp studies. At the end of the clamp, the rats were killed by an overdose of anesthetic (pentobarbital sodium, 75 mg/kg, iv) followed by decapitation.

Blood and tissue collection for in vitro studies and determinations. A second cohort of chow- and HFHS-fed rats was used to determine the effect of the diets on plasma glucose, insulin, triglycerides and free fatty acids, endothelial NO synthase (eNOS) protein expression, nitrotyrosine formation, NAD(P)H oxidase protein expression, and endothelin-1 (ET-1) concentration in isolated tissues, and glucose transport activity in skeletal muscles. Because of low tissue concentrations and different extraction procedures, ET-1 (mesenteric arterial bed) and eNOS, nitrotyrosine and NAD(P)H oxidase protein expression (thoracic aorta) could not be measured in the same blood vessel sample. The rats used in these different studies were 10 to 11 wk old. Rats from both dietary groups were anesthetized with pentobarbital sodium (75 mg/kg, ip) and had their soleus and extensor digitorum longus (EDL) muscles rapidly dissected out, weighed, and immediately proceeded for glucose transport activity measurement (see below) The thoracic cage of the animals was opened, and blood was collected by cardiac puncture and centrifuged (1,500 g, 15 min at 4°C). Plasma was stored at –20°C until later biochemical determinations. The thoracic aorta, from the diaphragm to the aortic arch, and the complete mesenteric vascular bed, from the aorta to the intestinal border, were removed and cleaned of blood and surrounding adipose tissue. The vessels were either snap frozen and stored at –80°C, immersed in Optimal Cutting Temperature (OCT) Embedding Medium (Tissue-Tek, Sakura Finetek, Torrance, CA), and quickly frozen for later immunofluorescence analysis or immediately processed for in vitro vascular reactivity studies. Epididymal white adipose tissues (WAT), gastrocnemius muscles, and liver were excised, weighed, snap frozen, and stored at –80°C.

Triacylglycerol, nonesterified fatty acids, glucose, and insulin measurements. Plasma glucose concentrations were measured by the glucose oxidase method (42), using a Beckman glucose analyzer (Beckman Instruments, Palo Alto, CA). Insulin levels were measured by radioimmunoassay (RIA), using porcine insulin standards and polyethylene glycol for separation (15). Plasma triglycerides were assayed by an enzymatic method (30), using a reagent kit from Roche Diagnostics (Basel, Switzerland) that allowed correction for free glycerol. Plasma nonesterified fatty acids (NEFA) were also determined enzymatically using commercially available kits (Wako Chemicals, Dallas, TX).

Western blot analysis. Immunoblotting was performed in thoracic aorta and gastrocnemius muscle of both diet groups to detect eNOS protein levels as previously described (35). Nitrotyrosine formation and NAD(P)H oxidase gp91phox protein levels were also determined in thoracic aorta. The aorta and gastrocnemius muscle of each rat were individually homogenized in 5 vol of homogenization buffer containing 50 mM Tris·HCl (pH 7.5), 1 mM EDTA, 1 mM EGTA, 10% glycerol, 1 mM DTT, 50 mM NaF, 5 mM Na pyrophosphate, 1% Triton X-100, 0.1 mg/ml phenylmethylsulfonyl fluoride (PMSF), and protease inhibitors cocktail (P-8340; Sigma-Aldrich, Oakville, ON, Canada). Homogenates were centrifuged (10,000 g for 10 min, 4°C), and protein concentrations of the supernatant were determined by the bicinchoninic acid method (Pierce), using BSA as the standard. Protein samples (45 µg) were separated by SDS-PAGE and electrophoretically transferred (100 V, 2 h) to polyvinylidene difluoride (PVDF) membranes (35, 47). PVDF membranes were incubated at 4°C overnight with monoclonal antibodies against eNOS (1:500; Transduction Labs, Lexington, KY), nitrotyrosine (1:500; Calbiochem, San Diego, CA) (35), or polyclonal anti-gp91phox (1:500; Transduction Labs). Monoclonal {alpha}-tubulin antibodies (1:5,000; Sigma-Aldrich) were used to standardize the results. Immunoreactive bands were detected by the enhanced chemiluminescence method.

Immunofluorescence analysis. Three 7-µm transversal sections of each frozen thoracic aorta of chow- (n = 7) and HFHS-fed rats (n = 7) were mounted on glass slides, fixed in acetone at –20°C for 10 min, and washed in phosphate-buffered saline (PBS). They were incubated in the blocking solution containing 10% BSA diluted in PBS at room temperature for 45 min and then incubated overnight at 4°C with 1:200 diluted primary antibody for eNOS (Transduction Laboratories) and the Alexa Fluor-488 phalloidin (Molecular Probes, Burlington, ON, Canada) diluted 1:150 in the blocking solution. Following that, the sections were washed three times with PBS for 10 min and incubated for 1 h with the Alexa Fluor-594 rabbit anti-mouse IgG, diluted 1:1,000 (Molecular Probes) in the blocking solution. After being rinsed in PBS, tissue slices were mounted on coverslips. Immunofluorescence was observed on a confocal microscope (MRC-1024 Confocal System, Bio-Rad) combined with the Laser Sharp image acquisition software v3.2. Comparison of fluorescence levels between groups was performed, under the same conditions of laser exposure, using the Metamorph software (Molecular Devices, Downington, PA).

Vascular reactivity. The effects of diets on vascular reactivity were evaluated in vitro using aortic rings prepared as previously described (9) which were allowed to equilibrate for 1 h in the incubation medium. The incubation medium was Krebs-Henseleit physiological solution, containing the following (in mM): 118 NaCl, 4.7 KCl, 25 NaHCO3, 1.18 MgSO4, 1.18 KH2PO4, 2.5 CaCl2, and 5.5 glucose, pH 7.4, maintained at 37°C and gassed with 95% O2-5% CO2. In some experiments, the NOS inhibitor L-NAME (100 µM) was added at the start of the equilibration period and was present for the duration of the experiments. After equilibration, the arteries were constricted using a high-potassium Krebs solution (KCl, 18 mM) and were allowed to reequilibrate. The baths were then emptied and rinsed three times with Krebs solution and again allowed to equilibrate. The arteries were then constricted with 10–6 M phenylephrine (Sigma, St. Louis, MO). While the rings were constricted, the endothelium-dependent dilator responses to 10–5 M carbachol (Sigma) were obtained to ensure that the endothelium of each vascular ring was functioning. After washout of the phenylephrine and carbachol, a cumulative dose-response curve to phenylephrine (10–9–10–5M) was obtained. After a washout period, rings were constricted with phenylephrine (10–6 M), and cumulative relaxation curves to carbachol (10–8–10–4 M) and then to sodium nitroprusside (SNP; 10–10–10–7 M, Sigma) were quantified to assess the endothelium-dependent and endothelium-independent ability of the smooth muscle to relax. Each curve was separated by a 30-min washout period. The contracting curves to phenylephrine were expressed in gram of tension, and the relaxation curves were expressed as percent changes from the contraction induced by phenylephrine.

Measurement of endothelin level in mesenteric arterial bed. Immunoreactive ET-1 concentrations were measured in the complete mesenteric arterial beds from HFHS- (n = 8) and chow-fed rats (n = 8) using a specific RIA in C18 Sep-Pak-extracted samples as described in detail elsewhere (35, 47). Briefly, the mesenteric vascular bed of each rat was individually homogenized twice with a Tissue-Tearor for 15 s in 2 ml of ice-cold extraction solution (1 M HCl, 1% acetic acid, 1% TFA, and 1% NaCl). The homogenate was centrifuged at 3,000 g for 30 min at 4°C. The supernatant was then collected, and 100 µl of [125I]ET-1 (~1,000 counts/min; cpm) were added before extraction on a C18 Sep-Pak column. The Sep-Pak column was activated with 4 ml of 60% acetonitrile and 0.1% trifluoroacetic acid (TFA) and then rinsed twice with 10 ml of 0.1% TFA. After sample loading, the column was washed twice with 10 ml of 0.1% TFA, and the immunoreactive ET-1 (ir-ET-1) fraction was eluted with 3 ml of 60% acetonitrile and 0.1% TFA and then counted in a gamma counter (recovery is 90–95%). The sample extracts were dried overnight in a Speed-Vac and reconstituted in 500 µl of RIA buffer. Aliquots of 100 and 200 µl of extracted samples or 200 µl of standards (ET-1, Peninsula Laboratories) were added to 100 µl of anti-ET-1 antibody, and the final reaction volume was adjusted to 300 µl with RIA buffer. After a 24-h incubation period at 4°C, 100 µl of [125I]ET-1 (15,000 cpm) in RIA buffer were added, and the tubes were incubated for an additional 24 h at 4°C. Bound and free radioactivity was separated by the second antibody method. After a 2-h incubation period at room temperature, 0.5 ml of RIA buffer was added, and the tubes were centrifuged at 2,500 g for 20 min at 4°C. The supernatant was then discarded, and the pellet was counted in a gamma counter. ir-ET-1 concentrations were corrected for losses in extraction.

Glucose transport activity in isolated rat skeletal muscles. Basal and insulin-stimulated glucose utilization was examined in isolated soleus and EDL skeletal muscles from overnight-fasted chow- (n = 9) and HFHS-fed rats (n = 11). Glucose transport was measured by use of the glucose analog [3H]-2-deoxy-D-glucose, according to the method developed by Hansen et al. (24) and as previously described (47). [3H]-2-deoxy-D-glucose uptake rates were corrected for extracellular trapping using [14C]mannitol (24).

Data analysis. The results are expressed as means ± SE; n is the number of rats. Western blot and immunofluorescence analysis data are expressed relative to control, assigning a value of 1 to the chow-fed group baseline mean. Data describing the biological characteristics of the rats as well as those resulting from Western blot, RIA, and immunofluorescence analysis were evaluated using Student's t-test for unpaired data. Vascular ring experiments and glucose transport activity measurements as well as results obtained over time, such as those from cardiovascular responses to insulin in HFHS- and chow-fed rats, were analyzed for statistical significance using a two-way repeated-measures ANOVA. Post hoc analyses were performed when significant differences were noted, using the Fishers least significance difference test. A P value <0.05 was taken to indicate a significant difference.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Body weight and metabolic changes. Compared with chow, chronic ingestion for 4 wk of the HFHS diet resulted in larger increases in final body weight and epididymal adipose depot, whereas gastrocnemius muscle and liver weights were comparable (Table 2). These significant differences in final body weight and fat accretion were associated with a larger energy intake in the HFHS-fed group than in the chow-fed group. Furthermore, plasma insulin and NEFA levels were significantly higher in overnight-fasted HFHS-fed rats than in chow-fed rats, whereas no differences in plasma glucose or triglycerides levels were noted between the dietary cohorts (Table 3). The demonstration here of a higher plasma insulin level in HFHS-fed rats compared with their chow-fed counterparts is indicative of insulin resistance. In unfasted rats, we found significantly higher plasma insulin, glucose, triglycerides, and NEFA levels in HFHS-fed than in chow-fed rats.


View this table:
[in this window]
[in a new window]

  		Table 2. Body, WAT, and muscle weights and average daily food and energy intakes of rats chronically fed either rodent chow or an HFHS diet for 4 wk

 

View this table:
[in this window]
[in a new window]

  		Table 3. Fasting and unfasting plasma levels of glucose, insulin, triglycerides, and nonesterified fatty acids of chow-fed and HFHS-fed rats

 
Hemodynamic responses to insulin infusion during the euglycemic hyperinsulinemic clamp. Baseline values (before any iv infusion) for cardiovascular variables are listed in Table 4 for both groups of rats. Basal heart rate and mean blood pressure were higher in HFHS-fed rats than in the chow-fed group. This was accompanied by smaller basal superior mesenteric flow and vascular conductance in HFHS-fed rats than in chow-fed rats, while there was no significant difference in basal renal or hindquarter flows or vascular conductances between the two dietary groups.


View this table:
[in this window]
[in a new window]

  		Table 4. Baseline values of HR, MAP, and regional Doppler shift and vascular conductance in chow-fed and HFHS-fed rats

 
Figure 1 shows that insulin infusion at a rate of 4 mU·kg–1·min–1 in chow-fed rats caused significant increases in renal and hindquarter blood flows, but it had no effect on heart rate, mean arterial blood pressure, or superior mesenteric flow compared with measurements following a control infusion of vehicle (saline-0.2% BSA). These responses were associated with significant increases in renal and hindquarter vascular conductances, whereas no change was noted in superior mesenteric vascular conductance (Fig. 2). In HFHS-fed rats, the same infusion of insulin had no effect on heart rate, mean arterial blood pressure, or renal or hindquarter flows, while a significant decrease in superior mesenteric flow was observed when compared with the effects of control infusion of vehicle (Fig. 1). The renal, superior mesenteric, and hindquarter blood flow responses to insulin differed significantly from those seen in chow-fed rats. Furthermore, in HFHS-fed rats the euglycemic infusion of insulin elicited a significant fall in superior mesenteric vascular conductance but had no consistent effect on renal or hindquarter vascular conductances when compared with the effects of control infusion of vehicle (Fig. 2). These responses differed significantly from those seen in chow-fed rats, in which insulin produced marked vasodilation in renal and hindquarter vascular beds but had no effect in the superior mesenteric vascular bed.


Figure 1
View larger version (21K):
[in this window]
[in a new window]

  		Fig. 1. Effects of a high-fat, high-sucrose (HFHS) diet on the cardiovascular responses to euglycemic infusion of insulin in conscious rats. Changes in mean arterial blood pressure (MAP), heart rate (HR), and regional Doppler shift were measured following control iv infusion of saline-0.2% BSA in chow-fed (n = 10) or HFHS-fed (n = 7) rats or euglycemic infusion of insulin at a rate of 4 mU·kg–1·min–1 in chow-fed (n = 9) or HFHS-fed (n = 10) rats. Values are means ± SE. Data were analyzed for statistical significance using a 2-way repeated-measures ANOVA followed by Fishers test. *P < 0.05, chow vs. vehicle; §P < 0.05, HFHS vs. chow; and {dagger}P < 0.05, HFHS vs. vehicle.

 

Figure 2
View larger version (25K):
[in this window]
[in a new window]

  		Fig. 2. Effects of HFHS diet on the changes in regional vascular conductances elicited by euglycemic infusion of insulin in conscious rats. Changes in regional vascular conductances were measured following control iv infusion of saline-0.2% BSA in chow-fed (n = 10) or HFHS-fed (n = 7) rats or euglycemic infusion of insulin at a rate of 4 mU·kg–1·min–1 in chow-fed (n = 9) or HFHS-fed (n = 10) rats. These data were derived from the data shown in Fig. 1. Values are means ± SE. Data were analyzed for statistical significance using a 2-way repeated-measures ANOVA followed by Fishers test. *P < 0.05, chow vs. vehicle; §P < 0.05, HFHS vs. chow; and {dagger}P < 0.05, HFHS vs. vehicle.

 
Responses during euglycemic hyperinsulinemic clamp. Figure 3 shows that, in the animals subjected to the euglycemic hyperinsulinemic clamp, the HFHS-fed rats had slightly but not significantly higher preinfusion insulin levels compared with their chow-fed counterparts (137 ± 14 vs. 96 ± 18 pmol/l). During the euglycemic infusion of insulin, we found that plasma insulin levels in both dietary cohorts rose acutely and achieved similar plateaus (770 ± 84 vs. 735 ± 75 pmol/l in HFHS- and chow-fed rats, respectively). Preinfusion glucose levels were similar in both dietary groups (4.3 ± 0.1 vs. 4.3 ± 0.1 mmol/l in HFHS- and chow-fed rats, respectively), and glycemia achieved during insulin infusion was comparable in the chow- and HFHS-fed rats (average levels between 60 and 120 min of infusion: 4.1 ± 0.1 mmol/l and 4.3 ± 0.1 mmol/l, respectively). The variation in glycemia during the infusions never exceeded 10%. The control infusion of saline-0.2% BSA did not alter plasma glycemia or insulin (data not shown). The steady-state glucose infusion rate needed to maintain euglycemia during the last hour of the clamp was found to be significantly lower in the HFHS-fed rats than in the chow-fed group, indicating frank whole body insulin resistance in the former.


Figure 3
View larger version (21K):
[in this window]
[in a new window]

  		Fig. 3. Summary of steady-state blood glucose and plasma insulin concentrations and glucose infusion rate during a euglycemic hyperinsulinemic clamp. The experiments were performed in conscious, unrestrained chow-fed (n = 9) or HFHS-fed (n = 10) rats. Insulin was infused at a rate of 4 mU·kg–1·min–1. Glucose infusion rates necessary to maintain euglycemia during steady-state hyperinsulinemia were significantly lower in HFHS-fed rats than in chow-fed rats. Values are means ± SE. Data were analyzed for statistical significance using a 2-way repeated-measures ANOVA followed by Fishers test. *P < 0.05, HFHS-fed rats vs. chow-fed rats.

 
Effect of HFHS diet on eNOS protein expression in thoracic aorta and skeletal muscle. The Western blot analysis revealed that eNOS proteins were detectable in vascular and muscle tissues from both dietary groups. The proteins migrated as a single band of ~140,000 M (Fig. 4A). Immunoreactivity of eNOS was quantified by scanning densitometry, and the mean data are presented. The results indicate that eNOS protein expression was significantly lower in thoracic aorta and gastrocnemius muscle from HFHS-fed rats than from chow-fed rats (Figs. 4B). Using confocal microscopy, we found a threefold reduction in eNOS immunofluorescence level in the endothelium of thoracic aorta from HFHS-fed rats compared with the chow-fed group (Fig. 5).


Figure 4
View larger version (18K):
[in this window]
[in a new window]

  		Fig. 4. Effects of HFHS diet on endothelial nitric oxide synthase (eNOS) protein expression in thoracic aorta and skeletal muscle. eNOS protein content was measured in thoracic aortas and gastrocnemius muscles isolated from overnight-fasted HFHS-fed (n = 8) and chow-fed rats (n = 8). A: representative immunoblots showing eNOS immunoreactivity in aorta and gastrocnemius muscle of one animal in each dietary group are shown. B: densitometric band intensities of eNOS are summarized. The mean level of eNOS in chow-fed rats was assigned a value of 1, and the relative levels of eNOS in each group were adjusted with {alpha}-tubulin. Data are expressed as means ± SE and were analyzed using unpaired Student's t-tests. *P < 0.05, significant difference between HFHS-fed rats and chow-fed rats.

 

Figure 5
View larger version (50K):
[in this window]
[in a new window]

  		Fig. 5. Effects of HFHS diet on eNOS immunofluorescence level in the endothelium of thoracic aortas from chow- and HFHS-fed rats. A: confocal microscopy photographs of the thoracic aortas from a chow-fed rat (left) and an HFHS-fed rat (right). Endothelium-specific eNOS protein expression was revealed using an Alexa Fluor-594-conjugated rabbit anti-green fluorescent protein (GFP) (red). The aortas were counterstained with Alexa Fluor-488-conjugated phalloidin (green), which has a high affinity for F-actin. B: densitometric band intensities of eNOS are summarized. The mean level of eNOS in chow-fed rats (n = 7) was assigned a value of 1, and the relative level of eNOS in HFHS-fed rats (n = 7) was calculated. Data are means ± SE and were analyzed using unpaired Student's t-tests. *P < 0.05, significant difference between HFHS-fed rats and chow-fed rats.

 
Vascular reactivity. Contractions induced by increasing concentrations of phenylephrine were not significantly altered by the HFHS diet compared with the chow diet (Fig. 6). The addition of L-NAME in the incubation medium was found to significantly enhance to the same extent the contracting responses to phenylephrine in the two dietary cohorts. Figure 7 shows that the endothelium-dependent relaxations induced by increasing concentrations of carbachol were slightly but significantly lower in the aortic rings from the HFHS-fed rats than in those from the chow-fed group (significant at 3 x 10–7–10–4 M). The addition of L-NAME in the incubation medium completely abrogated the vasorelaxing effects of carbachol in the two experimental groups. The evaluation of the endothelium-independent vasorelaxation by use of SNP in cumulative concentrations revealed that HFHS-fed rat vascular smooth muscles had a similar ability to relax as those from chow-fed rats (Fig. 7). The addition of L-NAME slightly but significantly reduced the vasorelaxations elicited by low doses of SNP (10–10–10–9 M) in both groups. However, because we were using a consistent dose of phenylephrine to create the constriction, it is likely that the reduction observed here was due to changes in preconstriction, as L-NAME was found to enhance the contracting responses to phenylephrine.


Figure 6
View larger version (20K):
[in this window]
[in a new window]

  		Fig. 6. Cumulative dose-response curves to phenylephrine in rat isolated aortic rings. The aortic rings were incubated in the absence (HFHS, n = 12; chow, n = 11) or presence of L-NAME (100 µM) (HFHS, n = 8; chow, n = 8). Developed tensions are expressed in grams, and values are means ± SE. Data were analyzed using a 2-way repeated-measures ANOVA, and post hoc analyses were performed (Fishers test) when appropriate. §P < 0.05, HFHS + L-NAME vs. HFHS; {dagger}P < 0.05, chow + L-NAME vs. chow.

 

Figure 7
View larger version (12K):
[in this window]
[in a new window]

  		Fig. 7. Cumulative dose-response curves to carbachol or sodium nitroprusside (SNP) in rat isolated aortic rings. The aortic rings were precontracted with phenylephrine and incubated in the absence (HFHS, n = 12; chow, n = 11) or presence of L-NAME (100 µM) (HFHS, n = 8; chow, n = 8). Relaxations are expressed as percent changes from the initial precontraction induced by phenylephrine, and values are means ± SE. Data were analyzed using a 2-way repeated-measures ANOVA, and post hoc analyses were performed (Fishers test) when appropriate. *P < 0.05, HFHS-fed rats vs. chow-fed rats; §P < 0.05, HFHS + L-NAME vs. HFHS; and {dagger}P < 0.05, chow + L-NAME vs. chow.

 
Effect of HFHS diet on nitrotyrosine expression in vascular tissue. Western blot analysis of the thoracic aortas isolated from both experimental groups revealed higher levels of nitrosylated proteins in vascular tissues isolated from HFHS-fed rats than from chow-fed rats (Fig. 8). This was associated with higher NAD(P)H oxidase protein expression in vascular tissues isolated from HFHS-fed rats than from chow-fed rats (Fig. 9).


Figure 8
View larger version (19K):
[in this window]
[in a new window]

  		Fig. 8. Effects of HFHS diet on nitrotyrosine expression in vascular tissue. Representative Western blots (A) and group data (B) depicting nitrotyrosine expression in the thoracic aorta isolated from overnight-fasted HFHS-fed (n = 8) and chow-fed (n = 8) rats. The mean level of nitrotyrosine in chow-fed rats was assigned a value of 1, and the relative levels of nitrotyrosine in each group were adjusted with {alpha}-tubulin. Data are expressed as means ± SE and were analyzed using unpaired Student's t-tests. *P < 0.05, significant difference between HFHS-fed rats and chow-fed rats.

 

Figure 9
View larger version (26K):
[in this window]
[in a new window]

  		Fig. 9. Effects of HFHS diet on NAD(P)H oxidase expression in vascular tissue. Representative Western blots (A) and group data (B) depicting protein abundance of gp91phox subunit of NAD(P)H oxidase in the thoracic aorta isolated from overnight-fasted HFHS-fed (n = 8) and chow-fed (n = 8) rats. The mean level of gp91phox in chow-fed rats was assigned a value of 1, and the relative levels of NAD(P)H oxidase in each group were adjusted with {alpha}-tubulin. Data are expressed as means ± SE and were analyzed using unpaired Student's t-tests. *P < 0.05, significant difference between HFHS-fed rats and chow-fed rats.

 
ET-1 protein content in vascular tissues. Higher concentrations of ir-ET-1 were noted in the mesenteric arterial beds isolated from the HFHS-fed rats than from the chow-fed rats (Fig. 10).


Figure 10
View larger version (12K):
[in this window]
[in a new window]

  		Fig. 10. Effects of HFHS diet on immunoreactive endothelin-1 (ir-ET-1) concentration in vascular tissue. ir-ET-1 concentration was measured in mesenteric arterial bed isolated from overnight-fasted HFHS-fed (n = 8) and chow-fed (n = 8) rats. Values are means ± SE. *P < 0.05, HFHS vs. chow, Student's t-test for unpaired data.

 
Effect of HFHS diet on [3H]-2-deoxy-D-glucose uptake in isolated skeletal muscles. The effect of the diets on basal and insulin-stimulated glucose transport activities in isolated soleus and EDL muscles is shown in Fig. 11. In both skeletal muscles, we found that HFHS feeding induced a significant decrease in basal glucose transport activity compared with chow feeding. Furthermore, in the presence of insulin, we found a significantly smaller glucose transport activity in soleus (at doses of 0.002, 0.02, and 0.2 mU/ml insulin) and EDL muscles (at doses of 0.02, 0.2 and 2 mU/ml insulin) isolated from HFHS-fed rats than in those isolated from chow-fed rats.


Figure 11
View larger version (14K):
[in this window]
[in a new window]

  		Fig. 11. Insulin dose-response curve for stimulation of glucose uptake in soleus and extensor digitorum longus (EDL) muscles. Muscles were dissected out from chow-fed rats (n = 9) and HFHS-fed rats (n = 11). Values are means ± SE. Comparisons were made between insulin-evoked responses in chow-fed rats and those in HFHS-fed rats, and data were analyzed using a 2-way repeated-measures ANOVA followed by Fishers test. *P < 0.05, HFHS-fed rats vs. chow-fed rats.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The rats fed an HFHS diet for 4 wk were found to be significantly heavier than the chow-fed rats. They had increased fat accretion and developed insulin resistance that was associated with vascular endothelium dysfunction and significant increases in blood pressure and unfasting plasma levels of insulin, glucose, triglyceride, and free fatty acid. Increased plasma levels of insulin and free fatty acid were also noted in overnight-fasted HFHS-fed rats. Therefore, the HFHS-fed rat represents a good model of diet-induced metabolic perturbations that are quite similar to those that define the metabolic syndrome in obese subjects (40). They also represent a suitable model to further explore vascular endothelium function in relation to the glucoregulatory and hemodynamic actions of insulin. Although some others have shown insulin resistance, hyperinsulinemia, hyperlipidemia, hypertension, and endothelial dysfunction in similar dietary models (1, 36, 37, 43, 45, 58), none of them has conducted experiments looking in parallel at vascular and metabolic actions of insulin in conscious, unrestrained HFHS-fed rats. An obvious strength of this study is the present status of the preparation in which the metabolic, vascular, and NOS responses can be evaluated in the same animal. These in vivo experiments were completed with in vitro studies to assess skeletal muscle glucose transport activity, vascular reactivity, eNOS and ET-1 protein expression and localization, and parameters of oxidative stress and metabolic homeostasis.

The major findings that emerged from this study are that a 4-wk HFHS diet in rats markedly impaired the vascular responses to insulin and induced insulin resistance, as concomitantly determined during a euglycemic hyperinsulinemic clamp. Thus, in chow-fed rats, the euglycemic infusion of insulin was found to induce vasodilations in renal and hindquarter vascular beds, whereas in HFHS-fed rats, the same euglycemic infusion of insulin had no vasodilatory effects but caused a vasoconstriction in the superior mesenteric vascular bed. The mechanism underlying the impaired vascular responses to insulin in the HFHS-fed group remains unclear. However, since a role for endothelium-derived NO has been shown in the mediation of the insulin vasodilator effect (48, 49), it is likely that a reduced production of NO, consecutive to a downregulation of eNOS protein in the vasculature of HFHS-fed rats, could have contributed, at least in part, to the impaired renal and hindquarter vasodilator responses. Indeed, our results clearly indicate a decreased abundance of eNOS protein expression in skeletal muscles and thoracic aortas from HFHS-fed rats compared with their chow-fed counterparts. This contention is further supported by the demonstration here of a threefold reduction in eNOS immunofluorescence level in the endothelium of thoracic aorta, using confocal microscopy, and by the depressed endothelium-dependent vasorelaxation response to carbachol noted in aortas from the HFHS-fed rats. The latter finding cannot be explained by smooth muscle dysfunction, because the vascular smooth muscle contracting response to phenylephrine, and the response to the endothelium-independent vasodilator SNP, did not differ between control and insulin-resistant rats. These findings indicate that the ability of vascular smooth muscle to contract or to relax in response to exogenous NO was not impaired in the HFHS-fed rats, and that diet selectively impaired endothelium-dependent vasodilation. These findings agree with previous data from this laboratory and others showing impaired endothelium-mediated relaxation and depressed eNOS protein expression in obese subjects, and in animal models with insulin resistance (36, 37, 43, 45, 47, 50).

On the other hand, enhanced ROS-mediated inactivation and sequestration of NO, leading to depressed NO bioavailability (43, 45) and then limiting the NO-dependent vasodilator capacity at the level of the vascular smooth muscle (36, 37, 43), might have contributed to the altered vascular responses noted in our group of HFHS-fed rats. In agreement with that, we found an increased level of NAD(P)H oxidase protein expression, a major vascular source of ROS, as well as an increased formation of nitrotyrosine, which is the stable "footprint" of NO oxidation by ROS, in vascular tissues of rats fed an HFHS diet. Therefore, these findings indicate that increased vascular oxidative stress induced by the HFHS diet, leading to NO breakdown and endothelial dysfunction, might be responsible, at least in part, for the impaired insulin-mediated vasodilator responses in renal and skeletal muscle vascular beds. This is supported by two recent studies carried out in female Fischer rats and demonstrating that long-term consumption of an HFHS diet (for 2 mo and longer) induces endothelial dysfunction and oxidant/antioxidant imbalance, resulting in upregulation of the ROS-generating enzyme NAD(P)H oxidase and downregulation of several key antioxidant enzymes in the kidney and vascular tissues (43, 44). Added to that, accelerated ROS production and diminished bioavailable NO caused by eNOS uncoupling in vascular tissues of our rat model might be an interesting hypothesis that remains to be verified.

Another potential mechanism that could contribute to impaired vascular responses to insulin in HFHS-fed rats is an exaggerated production of endothelium-derived contracting factors such as ET-1, which is a powerful vasoconstrictor that has been previously reported to modulate endothelium-dependent vasoconstriction in rats and mice fed a high-fat diet (12, 54). Consistent with that is the demonstration here that the mesenteric arteries from the HFHS-fed rats contained a greater amount of ET-1 protein than those from the chow-fed rats. This could have contributed to the insulin-mediated vasoconstrictor effect noted in this vascular bed. Insulin is known to stimulate gene expression of vascular ET-1 in vitro and to modulate ET-1 production and release both in vivo and in vitro (26, 57). Moreover, elevated vascular expression of ET-1 and ETA receptor mRNA and increased plasma levels of ET-1 have been shown in fructose-fed rats (27, 28), and we recently reported increased ET-1 concentration in mesenteric arteries isolated from sucrose-fed rats (47). Therefore, diet-induced insulin resistance/hyperinsulinemia may have created an environment resulting in activation of the ET system, thus causing impaired endothelium-dependent vasodilation.

On a quantitative basis, skeletal muscle is the predominant site of insulin-stimulated glucose disposal and the major tissue responsible for postprandial hyperglycemia in insulin-resistant states (4, 14). Therefore, we believe that the impaired hindquarter vasodilator response to insulin could have contributed to the insulin resistance noted in our HFHS-fed rats by reducing delivery of glucose and insulin to muscle beds, as previously suggested in humans and rats (5, 10, 11). However, to distinguish the direct effects of insulin on glucose uptake from its effects on blood flow, we carried out experiments in isolated skeletal muscles to examine the effect of HFHS feeding on glucose transport activity, thus in the absence of blood flow influence. We found that, compared with the chow diet, the HFHS diet caused resistance of skeletal muscle glucose transport to stimulation by insulin, suggesting that there is an intrinsic defect in the tissue itself. Thus impairment of insulin's hemodynamic actions in vivo would contribute to insulin resistance by reducing access for hormone and nutrient but would not be the sole cause of insulin resistance in these animals. These findings are consistent with our previous data and with those of other laboratories indicating an impaired ability of insulin to stimulate glucose uptake in muscles isolated from rats fed a high-fat and/or high-sucrose diet (1, 25, 47). The extent to which a reduction in the total cellular content of GLUT4 proteins or an impairment in the translocation process of the GLUT4 protein to the cell surface and/or a change in its intrinsic activity contributes to the reduced insulin-stimulated glucose transport activity remains to be verified.

In the present study, the HFHS diet-induced insulin resistance was accompanied with a significant rise in blood pressure. The reduction in baseline value of the superior mesenteric vascular conductance noted in the HFHS-fed group, together with the altered endothelial function and the insulin resistance/hyperinsulinemia induced by the diet, could be the trigger for the development of high blood pressure, as previous data indicated that insulin resistance and endothelial dysfunction precede hypertension in comparable animal models (1, 29). Another factor that could potentially contribute to the rise in blood pressure is the presence of increased sympathetic drive and hyperlipidemia in the HFHS-fed rats. Fat feeding in rats was shown to stimulate sympathetic nervous system activity (59), and increased sympathetic activity is known to stimulate free fatty acid release from fat depots (46). Previous studies have indicated an association between increased circulating levels of triglycerides or free fatty acids and endothelial dysfunction and hypertension (22, 32, 51, 52). The sucrose component of the diet could have enhanced the deleterious effect of the saturated fats on blood pressure and endothelial function, as we previously reported that feeding a high-sucrose diet (for 4 wk) to rats leads to insulin resistance, impaired insulin-mediated skeletal muscle vasodilation, and reduced eNOS protein content in muscle, but it has no effect on resting blood pressure (47). In the latter study, neither effect of the sucrose diet was observed on body weight or plasma levels of triglycerides or free fatty acids. Therefore, the hypertensive effect of the HFHS diet could be a facet of fat deposition and/or the fat content of the diet, as saturated fat feeding without sucrose was shown to cause endothelial dysfunction and hypertension (58, 59). Further studies are required to clarify this point.

In conclusion, metabolic insulin resistance is a problem of utmost clinical importance and a major risk factor for cardiovascular morbidity and mortality. Loss of the modulatory role of the endothelium may be a critical and initiating factor in the development of diabetic vascular disease. Our findings in HFHS-fed rats provide further evidence of the importance of eNOS protein in linking metabolic and vascular disease and indicate the ability of a relatively short-term HFHS diet to induce insulin resistance and endothelial dysfunction and to affect regulation of vascular NO synthesis and glucose and vascular homeostasis.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by grants to H. Bachelard from the Canadian Institutes of Health Research and the Heart and Stroke Foundation of Quebec. F. Bourgoin was supported by a studentship from the Association Diabète Québec.


   ACKNOWLEDGMENTS
 
We thank Marie Tremblay for technical assistance. We also thank Geneviève Pilon and Patrice Dallaire from Dr. André Marette's laboratory for technical assistance and judicious advice on Western blot analysis. We are grateful to Nadia Chbinou from Dr. Richard Larivière's laboratory for expert assistance with confocal microscopy.


   FOOTNOTES
 

Address for reprint requests and other correspondence: H. Bachelard, Lipid Research Unit, CHUL Research Center, CHUQ, 2705 blvd. Laurier, Ste-Foy, Quebec, Canada G1V 4G2 (e-mail: helene.bachelard{at}crchul.ulaval.ca)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Barnard RJ, Roberts CK, Varon SM, Berger JJ. Diet-induced insulin resistance precedes other aspects of the metabolic syndrome. J Appl Physiol 84: 1311–1315, 1998.[Abstract/Free Full Text]
  2. Barnard RJ, Wen SJ. Exercise and diet in the prevention and control of the metabolic syndrome. Sports Med 18: 218–228, 1994.[Web of Science][Medline]
  3. Baron AD. Insulin and the vasculature. Old actors, new roles. J Investig Med 44: 406–412, 1996.[Web of Science][Medline]
  4. Baron AD, Brechtel G, Wallace P, Edelman SV. Rates and tissue sites of noninsulin- and insulin-mediated glucose uptake in humans. Am J Physiol Endocrinol Metab 255: E769–E774, 1988.[Abstract/Free Full Text]
  5. Baron AD, Clark MG. Role of blood flow in the regulation of muscle glucose uptake. Annu Rev Nutr 17: 487–499, 1997.[CrossRef][Web of Science][Medline]
  6. Baron AD, Quon MJ. Insulin action and endothelial function. In: Insulin Resistance, edited by Reaven G and Laws A. Totowa, NJ: Humana, 1999, p. 247–263.
  7. Brindley DN. Role of glucocorticoids and fatty acids in the impairment of lipid metabolism observed in the metabolic syndrome. Int J Obes 19: S69–S75, 1995.
  8. Cersosimo E, DeFronzo RA. Insulin resistance and endothelial dysfunction: the road map to CV diseases. Diabetes Metab Res Rev 22: 423–436, 2006.[CrossRef][Web of Science][Medline]
  9. Christon R, Marette A, Badeau M, Bourgoin F, Mélançon S, Bachelard H. Fatty acid-induced changes in vascular reactivity in healthy adult rats. Metabolism 54: 1600–1609, 2005.[CrossRef][Web of Science][Medline]
  10. Clark ADH, Barrett EJ, Rattigan S, Wallis MG, Clark MG. Insulin stimulated laser Doppler signal by rat muscle in vivo, consistent with nutritive flow recruitment. Clin Sci 100: 283–290, 2001.[CrossRef][Web of Science][Medline]
  11. Cleland SJ, Petrie JR, Ueda S, Elliott HL, Connell JMC. Insulin-mediated vasodilation and glucose uptake are functionally linked in humans. Hypertension 33: 554–558, 1999.[Abstract/Free Full Text]
  12. daSilva AA, Kuo JJ, Tallam LS, Hall JE. Role of endothelin-1 in blood pressure regulation in a rat model of visceral obesity and hypertension. Hypertension 43: 383–387, 2004.[Abstract/Free Full Text]
  13. DeFronzo RA, Ferrannini E. Insulin resistance: a multifaceted syndrome responsible for NIDDM, obesity, hypertension, dyslipidemia, and atherosclerotic cardiovascular disease. Diabetes Care 14: 173–194, 1991.[Abstract]
  14. DeFronzo RA, Gunnarson R, Bjorkman O, Olson M, Wahren J. Effect of insulin on peripheral and splanchnic glucose metabolism in non-insulin-dependent (type II) diabetes mellitus. J Clin Invest 76: 149–155, 1985.[Web of Science][Medline]
  15. Desbuquois B, Aurbach GD. Use of polyethylene glycol to separate free and antibody-bound peptide hormones in radioimmunoassays. J Clin Endocrinol Metab 37: 732–738, 1971.
  16. Despres JP. Dyslipidaemia and obesity. Baillieres Clin Endocrinol Metab 8: 629–660, 1994.[CrossRef][Web of Science][Medline]
  17. Erdei N, Toth A, Pasztor E, Papp Z, Edes I, Koller A, Bagi Z. High-fat diet-induced reduction in nitric oxide-dependent arteriolar dilation in rats: role of xanthine oxidase-derived superoxide anion. Am J Physiol Heart Circ Physiol 291: H2107–H2115, 2006.[Abstract/Free Full Text]
  18. Fornoni A, Raij L. Metabolic syndrome and endothelial dysfunction. Curr Hypertens Rep 7: 88–95, 2005.[Web of Science][Medline]
  19. Galili O, Versari D, Sattler KJ, Olson ML, Mannheim D, McConnell JP, Chade AR, Lerman LO, Lerman A. Early experimental obesity is associated with coronary endothelial dysfunction and oxidative stress. Am J Physiol Heart Circ Physiol 292: H904–H911, 2007.[Abstract/Free Full Text]
  20. Gardiner SM, Bennett T. Regional haemodynamic responses to adrenoceptor antagonism in conscious rats. Am J Physiol Heart Circ Physiol 255: H813–H824, 1988.[Abstract/Free Full Text]
  21. Gokce N, Keaney JF, Hunter LM, Watkins MT, Nedeljkovic ZS, Menzoian JO, Vita JA. Predictive value of noninvasively determined endothelial dysfunction for long-term cardiovascular events in patients with peripheral vascular disease. J Am Coll Cardiol 41: 1769–1775, 2003.[Abstract/Free Full Text]
  22. Goodfriend TL, Egan BM. Nonesterified fatty acids in the pathogenesis of hypertension: theory and evidence. Prostaglandis Leukot Essent Fatty Acids 57: 57–63, 1997.[CrossRef]
  23. Haffner SM, Fong D, Zuda HP. Hyperinsulinemia, upper body obesity and cardiovascular risk factors in non-diabetics. Metabolism 37: 336–345, 1988.
  24. Hansen PA, Gulve EA, Holloszy JO. Suitability of 2-deoxyglucose for in vitro measurement of glucose transport activity in skeletal muscle. J Appl Physiol 76: 979–985, 1994.[Abstract/Free Full Text]
  25. Hansen PA, Han DH, Marshall BA, Nolte LA, Chen MM, Mueckler M, Holloszy JO. A high-fat diet impairs insulin stimulation of glucose transport in muscle. J Biol Chem 273: 26157–26163, 1998.[Abstract/Free Full Text]
  26. Hu RM, Levin ER, Pedram A, Frank HJL. Insulin stimulates production and secretion of endothelin from bovine endothelial cells. Diabetes 42: 351–358, 1993.[Abstract]
  27. Juan CC, Fang VS, Hsu YP, Huang YJ, Hsia DB, Yu PC, Kwok CF, Ho LT. Overexpression of vascular endothelin-1 and endothelin-A receptors in a fructose-induced hypertensive rat model. J Hypertens 16: 1775–1782, 1998.[CrossRef][Web of Science][Medline]
  28. Katakam PV, Pollock JS, Pollock DM, Ujhelyi MR, Miller AW. Enhanced endothelin-1 response and receptor expression in small mesenteric arteries of insulin-resistant rats. Am J Physiol Heart Circ Physiol 280: H522–H527, 2001.[Abstract/Free Full Text]
  29. Katakam PVG, Ujhelyi MR, Hoenig ME, Winecoff-Miller A. Endothelial dysfunction precedes hypertension in diet-induced insulin resistance. Am J Physiol Regul Integr Comp Physiol 275: R788–R792, 1998.[Abstract/Free Full Text]
  30. Kohlmeier M. Direct enzymatic measurement of glycerides in serum and lipoprotein fractions. Clin Chem 32: 63–66, 1986.[Abstract/Free Full Text]
  31. Lerman A, Zeiher AM. Endothelial function: cardiac events. Circulation 111: 363–368, 2005.[Free Full Text]
  32. Lewis TV, Dart AM, Chin-Dusting JP. Endothelium-dependent relaxation by acetylcholine is impaired in hypertriglyceridemic human with normal levels of plasma LDL cholesterol. J Am Coll Cardiol 33: 805–812, 1999.[Abstract/Free Full Text]
  33. Liang CS, Doherty JU, Faillace R, Maekawa K, Arnold S, Gavras H, Hood WB. Insulin infusion in conscious dogs: effects on systemic and coronary hemodynamics, regional blood flows, and plasma catecholamines. J Clin Invest 69: 1321–1336, 1982.[Web of Science][Medline]
  34. Mantha L, Palacios E, Deshaies Y. Modulation of triglyceride metabolism by glucocorticoids in diet-induced obesity. Am J Physiol Regul Integr Comp Physiol 277: R455–R464, 1999.[Abstract/Free Full Text]
  35. Mélançon S, Bachelard H, Badeau M, Bourgoin F, Pitre M, Larivière R, Nadeau A. Effects of high-sucrose feeding on insulin resistance and hemodynamic responses to insulin in spontaneously hypertensive rats. Am J Physiol Heart Circ Physiol 290: H2571–H2581, 2006.[Abstract/Free Full Text]
  36. Naderali EK, Fatani S, Williams G. Chronic withdrawal of a high-palatable obesity-inducing diet completely reverses metabolic and vascular abnormalities associated with dietary obesity in the rat. Atherosclerosis 172: 63–69, 2004.[Medline]
  37. Naderali EK, Williams G. Prolonged endothelial-dependent and independent arterial dysfunction in the rat by short-term feeding with a high-fat, high-sucrose diet. Atherosclerosis 166: 253–259, 2003.[Medline]
  38. Pitre M, Nadeau A, Bachelard H. Insulin sensitivity and hemodynamic responses to insulin in Wistar-Kyoto and spontaneously hypertensive rats. Am J Physiol Endocrinol Metab 271: E658–E668, 1996.[Abstract/Free Full Text]
  39. Rattigan S, Clark MG, Barrett EJ. Hemodynamic actions of insulin in rat skeletal muscle. Evidence for capillary recruitment. Diabetes 46: 1381–1388, 1997.[Abstract]
  40. Reaven GM. Insulin resistance, hyperinsulinemia, hypertriglyceridemia, and hypertension. Parallels between human disease and rodent models. Diabetes Care 14: 195–202, 1991.[Abstract]
  41. Reaven GM. Role of insulin resistance in human disease. Diabetes 37: 1595–1607, 1988.[Abstract]
  42. Richterich R, Dauwalder H. Zur bertimmung der plasmaglucokonzentration mit der hexokinase-glucose-6-phosphat-dehydrogenase methode. Schweiz Med Wochenschr 101: 615–618, 1971.[Web of Science][Medline]
  43. Roberts CK, Barnard RJ, Sindhu RK, Jurczak M, Ehdaie A, Vaziri ND. A high-fat, refined-carbohydrate diet induces endothelial dysfunction, oxydant/antioxidant imbalance and depresses NOS protein expression. J Appl Physiol 98: 203–210, 2005.[Abstract/Free Full Text]
  44. Roberts CK, Barnard RJ, Sindhu RK, Jurczak M, Ehdaie A, Vaziri ND. Oxidative stress and dysregulation of NAD(P)H oxidase and antioxidant enzymes in diet-induced metabolic syndrome. Metabolism 55: 928–934, 2006.[CrossRef][Web of Science][Medline]
  45. Roberts CK, Vaziri ND, Wang XQ, Barnard RJ. Enhanced NO inactivation and hypertension induced by a high-fat, refined-carbohydrate diet. Hypertension 36: 423–429, 2000.[Abstract/Free Full Text]
  46. Romijn JA, Fliers E. Sympathetic and parasympathetic innervation of adipose tissue: metabolic implications. Curr Opin Clin Nutr Metab Care 8: 440–444, 2005.[Web of Science][Medline]
  47. Santuré M, Pitre M, Marette A, Deshaies Y, Lemieux C, Larivière R, Nadeau A, Bachelard H. Induction of insulin resistance by high-sucrose feeding does not raise mean arterial blood pressure but impairs hemodynamic responses to insulin in rats. Br J Pharmacol 137: 185–196, 2002.[CrossRef][Web of Science][Medline]
  48. Scherrer U, Randin D, Vollenweider P, Vollenweider L, Nicod P. Nitric oxide release accounts for insulin's vascular effects in humans. J Clin Invest 94: 2511–2515, 1994.[Web of Science][Medline]
  49. Steinberg HO, Brechtel G, Johnson A, Fineberg N, Baron AD. Insulin-mediated skeletal muscle vasodilation is nitric oxide dependent. A novel action of insulin to increase nitric oxide release. J Clin Invest 94: 1172–1179, 1994.[Web of Science][Medline]
  50. Steinberg HO, Chaker H, Leaming R, Johnson A, Brechtel G, Baron A. Obesity/insulin resistance is associated with endothelial dysfunction. Implication for the syndrome of insulin resistance. J Clin Invest 97: 2601–2610, 1996.[Web of Science][Medline]
  51. Steinberg HO, Paradisi G, Hook G, Crowder K, Cronin J, Baron AD. Free fatty acid elevation impairs insulin-mediated vasodilation and nitric oxide production. Diabetes 49: 1231–1238, 2000.[Abstract]
  52. Stepniakowski KT, Goodfriend TL, Egan BM. Fatty acids enhance vascular alpha-adrenergic sensitivity. Hypertension 25: 774–778, 1995.[Abstract/Free Full Text]
  53. Storlien LH, James DE, Burleigh KM, Chisholm DJ, Kraegen EW. Fat feeding causes widespread in vivo insulin resistance, decreased energy expenditure, and obesity in rats. Am J Physiol Endocrinol Metab 251: E576–E583, 1986.[Abstract/Free Full Text]
  54. Traupe T, D'Uscio LV, Muenter K, Morawietz H, Vetter W, Barton M. Effects of obesity on endothelium-dependent reactivity during acute nitric oxide synthase inhibition: modulatory role of endothelin. Clin Sci 103: 13S–15S, 2002.[Web of Science][Medline]
  55. Vincent D, Ilany J, Kondo T, Naruse K, Fisher SJ, Kisanuki YY, Bursell S, Yanagisawa M, King GL, Kahn CR. The role of endothelial insulin signaling in the regulation of vascular tone and insulin resistance. J Clin Invest 111: 1373–1380, 2003.[CrossRef][Web of Science][Medline]
  56. Vincent MA, Clerk LH, Lindner JR, Klibanov AL, Clark MG, Rattigan S, Barrett EJ. Microvascular recruitment is an early insulin effect that regulates skeletal muscle glucose uptake in vivo. Diabetes 53: 1418–1423, 2004.[Abstract/Free Full Text]
  57. Wolpert HA, Steen SN, Istfan NW, Simonson DC. Insulin modulates circulating endothelin-1 levels in humans. Metab Clin Exp 42: 1027–1030, 1993.[Medline]
  58. Yoshioka S, Uemura K, Tamaya N, Tamagawa T, Miura H, Iguchi A, Nakamura J, Hotta N. Dietary fat-induced increase in blood pressure and insulin resistance in rats. J Hypertens 18: 1857–1864, 2000.[CrossRef][Web of Science][Medline]
  59. Young JB, Daly PA, Uemura K, Chaouloff F. Effects of chronic lard feeding on sympathetic nervous system activity in the rat. Am J Physiol Regul Integr Comp Physiol 267: R1320–R1328, 1994.[Abstract/Free Full Text]
  60. Zeng G, Nystrom FH, Ravichandran LV, Cong LN, Kirby M, Mostowski H, Quon MJ. Role for insulin receptor, PI3-kinase, and Akt in insulin-signaling pathways related to production of nitric oxide in human vascular endothelial cells. Circulation 101: 1539–1545, 2000.[Abstract/Free Full Text]
  61. Zeng G, Quon M. Insulin-stimulated production of nitric oxide is inhibited by wortmannin. Direct measurement in vascular endothelial cells. J Clin Invest 98: 894–898, 1996.[Web of Science][Medline]



This article has been cited by other articles:


Home page
 J. Pharmacol. Exp. Ther.Home page
K. Sachidanandam, J. R. Hutchinson, M. M. Elgebaly, E. M. Mezzetti, M.-H. Wang, and A. Ergul
Differential Effects of Diet-Induced Dyslipidemia and Hyperglycemia on Mesenteric Resistance Artery Structure and Function in Type 2 Diabetes
J. Pharmacol. Exp. Ther., January 1, 2009; 328(1): 123 - 130.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
295/3/H1044    most recent
00516.2008v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Web of Science (1)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Bourgoin, F.
Right arrow Articles by Nadeau, A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Bourgoin, F.
Right arrow Articles by Nadeau, A.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2008 by the American Physiological Society.