Single-Dose Dexamethasone Induces Whole-Body Insulin Resistance and Alters Both Cardiac Fatty Acid and Carbohydrate Metabolism

The investigation conforms to the guide for the care and use of laboratory animals published by the National Institutes of Health and the University of British Columbia. Adult male Wistar rats (270–290 g) were obtained from the University of British Columbia Animal Care Unit and fed a standard laboratory diet (PMI Feeds, Richmond, VA) and water ad libitum. The synthetic glucocorticoid hormone DEX (1 mg/kg) or an equivalent volume of ethanol was administered by intraperitoneal injection and the animals killed 1–4 h later (plasma half-life of DEX is ∼279 min). Previous studies have determined that this dose of DEX inhibits insulin-stimulated skeletal muscle glucose transport (17).

Whole-animal insulin resistance was assessed using a euglycemic-hyperinsulinemic clamp, as described previously (18,19). This procedure involves the simultaneous intravenous infusion of insulin (to inhibit endogenous hepatic glucose production) and glucose; the quantity of exogenous glucose required to maintain euglycemia is a reflection of the net sensitivity of target tissues (mainly skeletal muscle) to insulin. Briefly, after injection of vehicle or DEX for 4 h, animals were anesthetized with sodium pentobarbital (Somnotol; 65 mg/kg) and a cannula inserted into the left jugular vein. Surgical insertion of the cannula was rapid (∼10 min for each animal). Insulin (Humulin R; 3 mU · min⁻¹ · kg⁻¹) and d-glucose (50%) were continuously delivered for 3 h, with the glucose infusion started 4 min after commencement of insulin infusion. Insulin and glucose were dissolved in 0.9% saline before infusion. At regular intervals, a small amount of blood taken from the tail vein was analyzed for glucose (using a glucometer: AccuSoft Advantage) and fatty acid. The glucose infusion rate was adjusted accordingly to maintain euglycemia.

When metabolized, glucose passes through the glycolytic pathway to be oxidized to CO₂ or nonoxidatively catabolized to lactate and alanine. To measure glucose oxidation, isolated hearts were perfused for 30 min with Krebs-Henseleit buffer in the working mode at a preload of 11.5 mmHg and an afterload of 80 mmHg, as previously described (20). The buffer solution contained 0.4 mmol/l palmitate, 2.0 mmol/l CaCl₂, 5.5 mmol/l glucose, 0.5 mmol/l lactate, 100 mU/l insulin, and 3% BSA (pH 7.4). Rates of glucose oxidation (quantitative collection of ¹⁴CO₂ liberated from [U-¹⁴C]glucose at the pyruvate dehydrogenase reaction and in the citric acid cycle) and glycolysis (¹⁴C-labeled products, [¹⁴C]lactate and [¹⁴C]pyruvate) were determined as previously described (20). On transport into cardiac cells, glucose disposal can also occur via its conversion into glycogen, which serves as the primary storage form of glucose. Cardiac glycogen was determined as glucose residues by a glucose kinase method after acid hydrolysis (21).

The rate of glucose oxidation is dependent on mitochondrial PDC. Phosphorylation via PDK inhibits PDC. PDK gene expression was measured in the indicated groups using RT-PCR. Briefly, total RNA from hearts (100 mg tissue) was extracted using Trizol (Invitrogen). After spectrophotometric quantification and resolving of RNA integrity using a formaldehyde agarose gel, reverse transcription was carried out using an oligo (dT) primer and superscript II RT (Invitrogen). First-strand cDNA was amplified using PDK2- and PKD4-specific primers. PDK2: 5′-TCTACCTCAGCCGCATCTCT-3′ (left) and 5′-GTTGGTGGCATTGACTTCCT-3′ (right); PDK4: 5′-CCTTTGGCTGGTTTTGGTTA-3′ (left) and 5′-CACCAGTCATCAGCCTCAGA-3′ (right); the β-actin gene was amplified as an internal control using 5′-TTGTAACCAACTGGGACGATATGG-3′ (left) and 5′-GATCTTGATCTTCATGGTGCTAGG-3′ (right). The linear range was determined to be between 15 and 40 cycles. The amplification parameters were set at 94°C for 1 min, 58°C for 1 min, and 72°C for 1 min (40 cycles total). The PCR products were electrophoresed on a 1.7% agarose gel containing ethidium bromide. Expression levels were represented as the ratio of signal intensity for PDK mRNA relative to β-actin mRNA.

To determine whether changes in PDK4 mRNA are translated to an increase in protein expression, Western blot analysis for PDK4 was carried out in mitochondria isolated by a previously described method (10). Mitochondrial proteins (5 μg) were fractionated by 11% SDS-PAGE and transferred onto nitrocellulose membranes. Blots were probed with polyclonal rabbit anti-PDK4 antisera (1:500) (generously provided by Dr. K.M. Popov), followed by incubation with goat anti-rabbit horseradish peroxidase linked secondary antibody (1:3,000). Reaction products were visualized using chemiluminescence and quantified by densitometry.

Isolated heart function (rate pressure product − heart rate [in bpm] × peak systolic pressure [in mmHg]) was recorded using a Direcwin physiograph (Raytech) (20). Mean arterial pressure was measured in vivo using a cannula inserted into the carotid artery.

At varying times (1–4 h) following DEX, hearts were perfused retrogradely through the aorta by the nonrecirculating Langendorff technique (16). To measure endothelium-bound LPL, perfusion solution was changed to buffer containing 1% fatty acid-free BSA and heparin (5 units/ml). Coronary effluent was collected in timed fractions over 10 min and assayed for LPL activity by measuring the hydrolysis of a sonicated [³H]triolein substrate emulsion (16).

To compare DEX effects on LPL with those previously observed following hyoinsulinemia, β-cell death was induced with a single dose of STZ (55 mg/kg i.v.) (15). Diabetic rats were kept for 4 days after STZ injection, at which time they were killed and hearts removed. To determine whether DEX- and STZ-treated animals can maintain their augmented LPL activity in vitro, hearts from control and STZ- and DEX-treated animals were either perfused with heparin, immediately upon removal (0 min), or perfused with heparin-free buffer for 60 min. During this 60-min perfusion with heparin-free buffer, LPL activity in the buffer reservoir (total volume 30 ml) was determined at various intervals. Finally, a 10-min perfusion with heparin was carried out to determine the extent of residual LPL at the coronary lumen.

Control and DEX (4 h) hearts were fixed in 10% formalin overnight. Blocks were embedded in Paraplast and sectioned at 5 μm. For immunostaining, sections were incubated with affinity-purified chicken anti-bovine LPL polyclonal antibody (1:400). Samples were then incubated with the secondary biotinylated rabbit anti-chicken IgG (1:150; Chemicon), followed by incubation for 1 h with streptavidin-conjugated Cy3 fluorescent probe (1:1,000). Slides were visualized using a Biorad 600 Confocal Microscope at 630× magnification.

LPL gene expression was measured using RT-PCR as described previously (22).

Acute (90 min) treatment of STZ-induced diabetic rats with rapid-acting insulin reduces heparin-releasable LPL activity to normal levels (15). To determine whether insulin could also influence LPL activity following DEX, insulin was infused over 3 h via the euglycemic-hyperinsulinemic clamp and cardiac LPL activity subsequently measured. For comparison, some STZ rats were treated intravenously with a rapid-acting insulin (8 units/animal); rats were killed after 180 min, and LPL activity was determined. Some hearts from DEX-treated animals were also perfused for 60 min with dichloroacetate (DCA; 1 mmol/l) for quantification of LPL. DCA, by inhibiting PDK, stimulates pyruvate dehydrogenase and promotes glucose utilization (23).

Control rats were injected with DEX at 10:00 a.m. (fed state). Following DEX, blood samples from the tail vein were collected at varying intervals in heparinized glass capillary tubes. Blood samples were immediately centrifuged, and plasma was collected and assayed. Diagnostic kits were used to measure glucose, triglyceride (Sigma), nonesterified fatty acid (Wako), and insulin (Linco).

To assess whether DEX causes accumulation of lipid droplets, morphological evaluation of hearts was carried out using transmission electron microscopy. Briefly, left ventricular tissue was fixed in 1.5% glutaraldehyde and paraformaldehyde, cut into small blocks (∼1 × 0.5 × 0.2 mm), and fixed for 8 h at 4°C. After washing, tissue was postfixed with 1% osmium tetroxide and further treated with 1% uranyl acetate and dehydrated using increasing concentrations of ethanol (50–100%). Blocks were embedded in epoxy resin and sectioned at ∼90 nm. Sections were stained with 1% uranyl acetate and Reynolds lead citrate. Images of the longitudinal sections were obtained with a Hitachi H7600 electron microscope.

[³H]triolein was purchased from Amersham Canada. Heparin sodium injection (Hapalean; 1,000 USP units/ml) was obtained from Organon Teknika. All other chemicals were obtained from Sigma Chemical.

Values are means ± SE. LPL activity in response to heparin perfusion over time was analyzed by multivariate (two-way) ANOVA using the Number Cruncher Statistical System. Wherever appropriate, one-way ANOVA followed by the Tukey or Bonferroni tests or the unpaired and paired Student’s t test were used to determine differences between group mean values. The level of statistical significance was set at P < 0.05.

Injection of DEX for 4 h was not associated with either hyperinsulinemia or hyperglycemia (Table 1). However, by using the euglycemic-hyperinsulinemic clamp, a direct measure of insulin sensitivity, the unfavorable effect of DEX on glucose metabolism was established. Accordingly, our clamp data revealed that the glucose infusion rate necessary to maintain euglycemia was lower after DEX administration (Fig. 1). Development of insulin resistance was not linked to any change in blood pressure (Table 1). Additionally, mechanical function of hearts isolated from DEX-treated animals remained unchanged throughout the perfusion period (Table 1).

Insulin resistance involves multiple organs and various mechanisms. Thus, although DEX is known to inhibit insulin-stimulated skeletal muscle glucose transport, its influence on cardiac metabolism is unknown. Upon transport into cardiac cells, glucose disposal occurs via oxidative metabolism to CO₂, nonoxidative metabolism to lactate and alanine, or conversion into glycogen. Myocardial glucose metabolism in control and DEX-treated hearts is summarized in Fig. 2. Mean steady-state rates of glycolysis and glucose oxidation were determined from data obtained during the initial portion of the heart perfusion. Compared with control, although rates of glycolysis were unaffected in DEX-treated hearts (Fig. 2A), the rate of cardiac glucose oxidation following DEX treatment was significantly decreased (Fig. 2B). In a separate experiment, cardiac glycogen was also evaluated from whole hearts after injection of DEX for 4h. Myocardial glycogen content in DEX hearts increased approximately twofold compared with control (Fig. 2C).

Inhibition of cardiac glucose oxidation by DEX could occur through inactivation of PDC by PDK. As PDK2 and -4 are the main isoforms present in the heart, their expressions were determined following acute DEX administration. DEX increased PDK4 (Fig. 3B) but not PDK2 mRNA (Fig. 3A). More importantly, the increased PDK4 mRNA was associated with augmented PDK4 protein, as measured by Western blot analysis (Fig. 3C).

LPL-mediated hydrolysis of circulating triglyceride-rich lipoproteins at the coronary lumen provides the heart with fatty acid. To measure this LPL activity, retrograde perfusion of hearts with heparin results in the release of LPL into the coronary perfusate (Fig. 4A). In a preliminary study, to determine the kinetics of LPL regulation at the vascular lumen, hearts from some fed control animals treated with a single injection of DEX were isolated at 1–4 h and LPL activity measured. Interestingly, increase in LPL activity became apparent as early as 1 h after injection of DEX (Fig. 4A, inset) and was maintained for an additional 4 h (Fig. 4A). The DEX-induced increase in heparin-releasable LPL activity at the vascular lumen after 4 h was substantial compared with control (approximately threefold) (Fig. 4A). Additionally, immunofluorescence microscopy of mycocardial sections from DEX hearts confirmed a more intense LPL immunofluorescence in blood vessels (Fig. 4B).

We have previously demonstrated that hypoinsulinemia also causes a substantial increase in coronary LPL activity and immunofluroscence at the vascular lumen (22). To determine whether this high LPL activity can be maintained in vitro, hearts from STZ and DEX animals were perfused with normal Krebs buffer for 1 h. Four days of STZ-induced diabetes caused a decline in plasma insulin (control 2.3 ± 0.4, diabetic 0.9 ± 0.1 ng/ml; P < 0.05) with ensuing hyperglycemia (control 8.7 ± 0.2, diabetic 14 ± 1.1 mmol/l; P < 0.05). Interestingly, STZ but not DEX hearts demonstrated a decline in heparin-releasable LPL activity to control values, presumably through dissociation of the enzyme from HSPG binding sites into the perfusate (Fig. 5). The mechanism by which DEX hearts are able to maintain high LPL in vitro suggests either an increased recruitment from myocytes or decreased displacement from the coronary lumen. To further investigate this process, basal LPL activity (in the absence of heparin) was determined in the buffer reservoir over time. DEX hearts showed greater basal release of LPL throughout the perfusion, suggesting that these hearts were able to maintain coronary lumen LPL through accelerated transfer from the myocytes (Fig. 5, inset).

Acute treatment of STZ rats with a rapid-acting insulin reduced hyperglycemia (STZ 18.7 ± 1.7, STZ + insulin 6.4 ± 0.8 mmol/l; P < 0.05). Acute insulin treatment also lowered peak heparin-releasable LPL activity to control levels (Fig. 6, inset). Unlike insulin reversal of LPL in STZ rats, LPL activity remained high even after 3 h of insulin perfusion during the clamp, suggesting ongoing resistance to the action of insulin. DCA, a stimulator of glucose oxidation, was also unable to reverse the DEX-induced increase in cardiac LPL (data not shown). This was not surprising given the evidence that PDK2 is more sensitive to inhibition by DCA, whereas PDK4 is insensitive (9).

Previously, we have reported that STZ-induced diabetes does not influence LPL mRNA and that increases in cardiac heparin-releasable LPL activity are likely due to a posttranslational mechanism (22). To determine whether the change in LPL activity following DEX is related to augmented synthesis, we measured LPL mRNA in heart homogenates. Four hours of DEX increased LPL mRNA levels (Fig. 4C).

Cardiac-specific overexpression of LPL has been suggested to be an important determinant of plasma triglyceride levels in mice (24). Given the increase of LPL at the coronary lumen, we evaluated both plasma triglyceride and fatty acid at varying times (1–4 h) following DEX. Interestingly, although plasma triglyceride declined progressively, there was no increase on plasma fatty acid (Fig. 7A).

Electron micrographs illustrate increases lipid-like vacuoles in myocytes following DEX treatment (Fig. 7B, arrows). These spherical lipid droplets were closely associated with the mitochondria. No other ultrastructural difference was detected between control and DEX hearts.

Chronic DEX treatment induces insulin resistance, hyperinsulinemia, and hyperglycemia (25,26). In our study, a single bolus dose of DEX had no influence on either plasma insulin or glucose after 4 h. Nevertheless, exploiting the euglycemic-hyperinsulinemic clamp, we demonstrate that DEX induced whole-body insulin resistance at this time point. Although this insulin resistance could embrace metabolic abnormalities in multiple organs (27), the effects of DEX on insulin-sensitive cardiac tissue have not been entirely elucidated. Assessment of cardiac glycolytic rates revealed no change following DEX, whereas glucose oxidation decreased significantly. Moreover, cardiac glycogen content increased almost twofold after DEX. These data suggest that in the short term, DEX is capable of inducing insulin resistance and switching cardiac glucose disposal from oxidation to storage, likely compromising energy production in the heart.

Glucose utilization provides the heart with ∼30% of its energy requirements. After glucose uptake and conversion to pyruvate, PDC facilitates subsequent entry and oxidation of pyruvate in the mitochondria. By phosphorylating PDC, PDK can decrease the rate of glucose oxidation. In Morris hepatoma cells, DEX is known to stimulate PDK4 but not PDK2 expression (12). Because these two PDKs are the major isoforms in the heart, we examined their cardiac expression. DEX increased the expression of PDK4 but was without effect on the high basal level of PDK2. Our data suggest that acute DEX could lower cardiac glucose oxidation through augmentation of PDK4 gene and protein expression. Whether pyruvate dehydrogenase phosphatase also plays a role in explaining the effect of DEX on cardiac glucose metabolism, as suggested in STZ-induced diabetes (28), is currently unknown.

Typically, fatty acids provide most of the energy required by the normal heart, with the balance coming from oxidation of other substrates like glucose. Since cardiac glucose oxidation is impaired following DEX, we predicted that fatty acid consumption would make up the energy deficit. Based on reports that 1) LPL is the major fatty acid supplier to the heart and 2) glucocorticoids are known to enhance LPL activity in postheparin serum (29), heart homogenates (30), and isolated myocytes (31), we examined LPL regulation following DEX. The present study is the first to report a rapid increase in LPL activity and protein at its functionally relevant location, the coronary lumen. Given the observation that cardiac LPL is a major determinant of plasma triglyceride (24), the increase in cardiac luminal LPL could be associated with the decline in circulating triglyceride. However, as no apparent change was noted in plasma fatty acid levels, our data suggest that following LPL-mediated triglyceride hydrolysis, fatty acid can be taken up rapidly and directly into tissues. In support of this suggestion, cardiac- and skeletal muscle-specific overexpression of LPL decreased plasma triglyceride and elevated triglyceride storage in muscle tissue but was without effect when plasma fatty acid was measured (32). In this study, visualization using electron microscopy also revealed high triglyceride storage in DEX-treated hearts.

In hypoinsulinemic and hyperglycemic STZ rats, we reported elevated cardiac luminal LPL activity (15,16) that was independent of shifts in mRNA levels, suggesting a posttranslational increase of LPL at this location (22). With insulin resistance induced by DEX, although comparable results were observed when luminal LPL activity was measured, in this instance, changes in LPL activity are coordinated to an increase in LPL mRNA. Another interesting dissimilarity between STZ-mediated hypoinsulinemia and DEX-induced insulin resistance is when hearts are perfused in vitro with heparin-free buffer. STZ hearts lose the augmented pool of heparin-releasable LPL at the coronary lumen, implicating hypoinsulinemia as an important reason for this effect in vivo, with continuous displacement of the enzyme in its absence in vitro. It should be noted that we are unable to detect any alteration in basal enzyme displacement between control and STZ hearts due to the large volume (30 ml) into which LPL was being shifted. In contrast, hearts from DEX animals continued to demonstrate augmented basal and heparin-releasable LPL activity in vitro, implying that the rate of displacement is overcome by intrinsic and enduring changes, likely an increased synthesis and transfer of enzyme, that act to keep LPL high at this location.

Regulation of cardiac luminal LPL may be an important means for maintaining cardiac function during metabolic stress by providing excess fatty acid to the heart. Acute treatment (180 min) of 4-day STZ-induced diabetic rats with a rapid-acting insulin reduced peak heparin-releasable LPL activity to control levels. It is likely that in vivo, insulin, by facilitating glucose entry and utilization, overcomes any energy deficit, eventually decreasing the requirement for LPL. DEX is considered a long-acting steroid, with a single dose lasting ∼2–2.5 days, and an “every other day” schedule is recommended for treatment (33). Interestingly, estimation of cardiac LPL from DEX hearts revealed an increase in enzyme, both immediately and following 3 h of insulin infusion (during clamp). These data suggest that unlike hypoinsulinemia, provision of exogenous insulin during DEX-induced insulin resistance is unable to normalize LPL. It should be noted that in humans, insulin secretegogues like sulfonylureas are also ineffective in overcoming glucocorticoid-induced insulin resistance (34).

In summary, our studies suggest that under circumstances where glucose utilization is compromised due to a glucocorticoid-induced increase of PDK4, augmented LPL will amplify triglyceride hydrolysis and the fatty acids supplied to the heart are used as additional sources of substrate to maintain continuous energy production. On the other hand, intracellular availability of fatty acid could regulate glucose metabolism, as suggested by the “glucose–fatty acid cycle” (12,35,36). Accordingly, DEX effects on glucose oxidation are prevented by Acipimox, which reduces plasma fatty acid (37), and in Morris hepatoma cells, fatty acids directly increase PDK4 expression (12). Irrespective of the mechanisms that increase LPL or decrease glucose oxidation, a role for LPL in cardiac pathology has been demonstrated in transgenic mouse lines overexpressing human LPL in skeletal and cardiac muscle (24,38). These animals exhibited insulin resistance, severe myopathy characterized by muscle fiber degeneration, and extensive proliferation of mitochondria and peroxisomes. In a more recent study using genetically engineered mice that specifically overexpressed cardiomyocyte surface-bound LPL, lipid oversupply and impaired contractile function (cardiomyopathy) were observed (39). It has not been determined whether these effects of DEX on cardiac metabolism can be translated into increased cardiovascular risk (40).

The studies described in this article were supported by operating grants from the Canadian Diabetes Association (in honor of Lillian I. Dale) (to B.R.) and the CIHR and Heart and Stroke Foundation of B.C. and Yukon (both an operating [to B.R. and M.A.] and a program [to B.R. and R.B.] grant). The financial support of the Health Research Foundation/Canadian Institutes of Health Research (to S.G., T.P., and R.B.), the Heart and Stroke Foundation of Canada (to T.P.), and the University of BC Graduate Fellowship (to S.G.) is gratefully acknowledged.

Dr. K.M. Popov generously provided the PDK4 polyclonal antibody.