Heart Care Info - Heart Disease Prevention & Treatment

Metabolic stress–induced activation of FoxO1 triggers diabetic cardiomyopathy in mice

PK Battiprolu, B Hojayev, N Jiang... - The Journal of ..., 2012 - Am Soc Clin Investig Pavan K. Battiprolu 1, Berdymammet Hojayev 1, Nan Jiang 1, Zhao V. Wang 1, Xiang Luo 1, Myriam Iglewski 1, John M. Shelton 1, Robert D. Gerard 1, Beverly A. Rothermel 1, Thomas G. Gillette 1, Sergio Lavandero 1, 2 and Joseph A. Hill 1, 3 [CITATION] Cardiac-Targeted TRPM7 Deletion Induces Heart Block and Cardiomyopathy via Disrupted Embryonic Ventricular Development R Sah, C Bates-Withers, X Mason... - Biophysical Journal, 2012 - Elsevier

Proteolytic degradation of IRS1 results in impaired insulin signaling in a number of cell types (26). Additionally, low levels of IRS1 are correlated with the subsequent development of T2D (23). Indeed, decreased IRS1 activity is central to insulin resistance in a number of diet-induced and genetic models of T2D (27). Earlier studies from our lab showed that ca Fox O1 expression in cultured rat cardiomyocytes decreases insulin-induced Glut4 membrane localization and glucose uptake (7). Consistent with this, we found that HFD-exposed cardiomyocytes with selective Fox O1 inactivation manifested enhanced responsiveness to insulin-induced Akt activation (Figure 6, A and B, and ref. 7). Together, these data point to Fox O1-dependent increases in lipid uptake, imbalances in energy metabolism, inactivation of IRS1, and subsequent decreases in insulin responsiveness as a mechanism contributing to insulin resistance–induced diabetic cardiomyopathy (Figure 7C).

Diabetic cardiomyopathy — heart disease independent of hypertension or coronary atherosclerosis — is a significant contributor to the morbidity and mortality associated with diabetes and metabolic syndrome (2, 3). A large body of evidence implicates insulin resistance in the pathogenesis of these disorders. As Fox O transcription factors are inhibited by insulin/PI3K/Akt signaling and yet also signal upstream to govern the cascade, we set out to define a possible role for Fox O-dependent transcriptional activity in diabetic heart disease. Here, we report that Fox O factors were activated in cardiac tissue from 2 models of T2D. Moreover, we uncovered a signaling axis linking Fox O activation and shifts in metabolic substrate use, cardiomyocyte insulin resistance, myocyte steatosis, and the cardiomyopathic phenotype. Finally, we identified metabolic stress–induced downregulation of IRS1 as the molecular site of Fox O1-elicited insulin resistance. Together, these data uncover what we believe to be a previously unrecognized Fox O1-dependent negative feedback mechanism in the regulation of insulin signaling and implicate Fox O1 activation in the pathogenesis of diabetic cardiomyopathy.

Diabetes and cardiac metabolism. Diabetes mellitus is a complex disease characterized by hyperglycemia stemming from absolute or relative insulin deficiency. In the majority of instances, it is associated with insulin resistance. The hormone insulin is central to the control of intermediary metabolism, orchestrating substrate utilization for storage or oxidation in all cells (28). As a result, insulin has profound effects on both carbohydrate and lipid metabolism throughout the body as well as substantial influence on protein metabolism. Consequently, derangements in insulin signaling have widespread and devastating effects in numerous tissues, including the cardiovascular system.

In patients with T2D, endogenous glucose production is accelerated (29). Resulting hyperglycemia can trigger glucotoxicity, which contributes to cardiac injury through multiple mechanisms, including direct and indirect effects of glucose on cardiomyocytes, cardiac fibroblasts, and endothelial cells. Chronic hyperglycemia promotes the overproduction of ROS through the electron transport chain, which can induce apoptosis (30) and activate poly (ADP-ribose) polymerase-1 (PARP). This enzyme mediates the direct ribosylation and consequent inhibition of GAPDH, diverting glucose from the glycolytic pathway toward alternative biochemical cascades that participate in hyperglycemia-induced cellular injury. These include increases in advanced glycation end products (AG Es) and activation of the hexosamine pathway, the polyol pathway, and protein kinase C (30, 31). Hyperglycemia-induced apoptosis is stimulated by ROS (32), PARP (33), AG Es (34), and aldose reductase (35). Hyperglycemia also contributes to altered cardiac structure and function through posttranslational modification of extracellular matrix components (e.g., collagens) and altered expression and function of both the ryanodine receptor (Ry R) and sarco(endo)plasmic reticulum Ca2+-AT Pase (SERCA), which in aggregate contribute to decreased systolic and diastolic function (30).

Enhanced lipid synthesis in hepatocytes and increased lipolysis in adipocytes together lead to increases in circulating F As and T Gs in patients with diabetes. Also, insulin stimulates FA transport into cardiomyocytes (36). Thus, elevated circulating lipids and hyperinsulinemia together increase FA delivery to cardiac cells, which rapidly adapt by promoting FA utilization. However, if FA delivery exceeds the oxidative capacity of the cell, F As accumulate and lipotoxicity ensues (37). Several mechanisms contribute to cardiac lipotoxicity, including ROS accumulation, ceramide production, insulin resistance, and impaired contractility due to altered electromechanical coupling. Thus, high FA uptake and metabolism not only stimulate accumulation of FA intermediates, but also increase oxygen demand, provoke mitochondrial uncoupling and ROS generation, decrease ATP synthesis, induce mitochondrial dysfunction, and trigger apoptosis. Together, these events play an important role in the pathogenesis of diabetes-associated heart disease.

Diabetic cardiomyopathy and myocardial insulin resistance. Diabetes and insulin resistance are powerful predictors of cardiovascular morbidity and mortality and are independent risk factors for death in patients with established heart failure (38). Patients with diabetes often develop atherosclerosis and hypertension, both of which are major contributors to the development of heart disease. However, cardiomyopathy can also develop in the absence of established risk factors (39). Indeed, Rubler et al. coined the term diabetic cardiomyopathy 4 decades ago to describe this form of disease (40). Currently, it is clear that this process, which sometimes emerges in isolation, is widespread and synergizes with the numerous diabetic comorbidities.

In a normal heart, approximately two-thirds of the energy required for cardiac contractility derives from FA oxidation, with the remainder deriving from glucose and lactate metabolism. In conditions of insulin resistance or diabetes, myocardial glucose utilization is significantly reduced, and a greater proportion of substrate utilization shifts to β-oxidation of FA (41). Associated with the reduced glucose utilization by diabetic myocardium is depletion of the glucose transporter proteins GLUT-1 and GLUT-4. Indeed, altered myocardial substrate metabolism favoring F As over glucose as energy source has been identified as a metabolic target of relevance. The diabetic heart relies on FA oxidation and is unable to switch to glucose, despite the lower oxygen consumption requirement of that substrate. As a consequence, cardiac efficiency (the ratio of cardiac work to myocardial oxygen consumption) decreases; diminished cardiac efficiency has been reported in humans and experimental animals with diabetes (42–45).

Insulin resistance is defined as diminished insulin-dependent stimulation of myocardial glucose uptake (42–45). Underlying mechanisms include FA accumulation that impairs insulin-mediated glucose uptake through inhibition of IRS and Akt. The serine protein kinases PKC-θ and IKK, which elicit serine phosphorylation of IRS, are activated (23). Phosphorylation and activation of PI3K and Akt are reduced, with substantial consequences for insulin actions in the heart (46).

Fox O transcription factors. Fox O proteins are emerging as important targets of insulin and other growth factor action in the myocardium (4, 6). Originally identified by their involvement in chromosomal translocations associated with leukemias and rhabdomyosarcomas (4, 6), abundant evidence now suggests that 3 members of the Fox O subfamily, Fox O1, Fox O3, and Fox O4, are critical for maintenance of cardiac function and cardiac stress responsiveness. The direct metabolic effects of Fox O signaling are not yet entirely clear, and actions of Fox O in nonmyocyte cellular elements of the heart are largely unknown.

With respect to cardiac function, Fox O factors participate in remodeling (47, 48), autophagy (49), apoptosis (50), responses to oxidative stress (51), regulation of metabolism (52), and cell cycle control (53). Through a variety of transcriptional targets, Fox O factors facilitate the response to changes in the environment via regulation of metabolic enzymes and energy-dependent proteins. Work in Caenorhabditis elegans has demonstrated a link among hormonal inputs, Akt signaling, and Fox O (51, 54). Whereas the traditional notion is that Fox O-dependent transcriptional activity is inhibited by PI3K-Akt signaling, a more complex feedback regulatory network has been reported by our group (7), positioning Fox O proteins as central elements in the control of insulin signaling. Forced expression of Fox O in primary cardiomyocytes triggers Akt phosphorylation via a calcineurin/PP2A-dependent mechanism, culminating in reduced insulin sensitivity and impaired glucose metabolism. Our present results extended these findings by uncovering a role for Fox O1 in the pathogenesis of this clinically prevalent disease process, demonstrating that Fox O1 activation, observed in 2 models of T2D, provoked insulin resistance, altered substrate metabolism, myocyte steatosis, and cardiomyopathy. Our findings further pinpointed the molecular defect at IRS1 within the insulin signaling cascade.

Potential therapeutic implications. The pathogenesis of diabetic cardiomyopathy is at once intricate, multifactorial, and clinically significant. The multiple, interlacing events occurring in patients with diabetes culminate in an environment which, coupled with insulin resistance, leads to diabetic cardiomyopathy. However, the central role of myocyte insulin resistance in the pathogenesis of cardiomyopathy suggests that this signaling cascade is a logical starting point for targeted treatment. The well-established myocardial dysfunction in diabetic individuals independent of hypertension and coronary artery disease suggests that the altered metabolic state of cardiac myocytes brought about by T2D is, in itself, sufficient to provoke cardiomyopathy (55). However, treatment of hypertension or atherosclerosis alone may not be enough to alter the development of heart disease in T2D patients (55).

Over time, constant and unremitting metabolic stress on the heart leads to progressive deterioration of myocardial structure and function. This suggests that therapeutic interventions early in the disease, targeting specific metabolic and structural derangements, may be required. This is especially relevant as strict control of hyperglycemia, however central to treatment, has not fulfilled hopes of meaningful morbidity and mortality benefit.

Perspective. Our present findings suggest that metabolic stress–induced activation of Fox O1 is central to the development of diabetic cardiomyopathy. Indeed, our data lend strong credence to a model in which persistent activation of Fox O1 under conditions of T2D provokes alteration in cardiac metabolism, which may be driven, at least in part, by a PDK4-induced shift in metabolic substrate utilization and an increase in lipid uptake. This change is exacerbated by the attenuation in insulin signaling through Fox O1-dependent inhibition of protein phosphatases (7) and by inactivation of IRS1, ultimately culminating in cardiomyopathy, heart failure, and death. Data reported here suggest that normalization of myocardial substrate metabolism by targeting Fox O1 activity may diminish the prevalence of heart failure and improve long-term survival of patients with T2D.

Animals. 8-week-old male C57BL/6 and db/db (leptin receptor mutant) mice were purchased from Jackson Labs. Starting at 8 weeks of age, C57BL/6 mice were maintained on a HFD (60% fat) for 25 weeks or longer, as indicated in Results. Controls were fed either 10%-fat diet (initial studies) or standard rodent food (chow) for the same duration as the respective HFD group. HFD is calorie rich (5.24 kcal/g, compared with 3.1 kcal/g in controls) as a result of higher fat composition. 10- to 12-week-old db/db mice were included in the study.

To generate cardiomyocyte-specific Fox O KO mice, MCM mice were crossed with DKO and Fox O1 KO lines. Cre was activated by i.p. injection of tamoxifen (40 mg/kg) for 5 days. Mice were allowed to recover from the tamoxifen/Cre-induced toxicity for 4–5 weeks before starting them on HFD. Recovery was judged by periodic assessment of cardiac function by 2D echocardiography. DKO, Fox O1 KO, and corresponding control floxed or MCM mice were maintained on both FVB and mixed genetic backgrounds.

Whole cell extract and subcellular fractionation. Nuclear and cytosolic extracts were prepared using lysis buffer (4 m M HEPES, p H 7.4; 320 m M sucrose; 1 m M dithiothreitol; 10 m M Mg Cl2; 5 m M K Cl dithiothreitol; and 0.1% Triton X-100) containing protease inhibitors and phosphatase inhibitors. Rapidly flash-frozen tissues were homogenized with a Dounce homogenizer in ice-cold lysis buffer. An aliquot was collected and resuspended with 2% SDS sample buffer (65 m M Tris H Cl, p H 6.8; 4% SDS; 20% glycerol; 50 m M dithiothreitol; and bromophenol blue) to serve as whole cell extract. The remaining homogenate was centrifuged at low speed (2,000 g) for 3 minutes at 4°C. The supernatant was further recentrifuged at 2,000 g for 10 minutes at 4°C, mixed with an equal volume of 2× sample buffer, and used as the cytosolic fraction. The pellet (nuclear fraction) was washed with lysis buffer several times before resuspending with sample buffer at a volume equal to the final volume of the supernatant, to ensure that equal volumes of each fraction contained protein from the same number of cells. All extracts were passed through glass wool. Equal volumes (adjusted to protein concentration) of each fraction were used for Western blotting. Separation between the nuclear and cytosolic fractions was verified by blotting for the cytosolic protein GAPDH and the nuclear membrane protein lamin A/C.

Western blotting. Proteins were separated by SDS/PAGE, transferred to a supported nitrocellulose membrane, and immunoblotted. Antibodies were purchased from Cell Signaling Technology, excepting α-tubulin (Sigma-Aldrich) and GAPDH (Santa Cruz Biotechnology). Blots were scanned, and bands were quantified using Odyssey Licor (version 3.0) imaging system.

Echocardiography. Echocardiograms were performed on conscious, gently restrained mice using either a Sonos 5500 system with a 15-M Hz linear probe or a Vevo 2100 system with a MS400C scanhead. LVEDD and LVESD were measured from M-mode recordings. FS was calculated as (LVEDD — LVESD)/LVEDD and expressed as a percentage. Measurements of diastolic dysfunction, such as mitral valve E/A ratio, isovolumetric extraction and relaxation times, mitral valve deceleration time, and ejection time, were made from 2D parasternal short axis views in diastole. LV mass was calculated by the cubed method as 1.05 × ([IVS + LVID + LVPW]3 – LVID3), where IVS is interventricular septum thickness, LVID is LV internal diameter, and LVPW is LV posterior wall thickness, and expressed in milligrams (56). All measurements were made at the level of the papillary muscles.

CSA. Images of tissues stained with wheat germ agglutinin were paraffin fixed (Vector Laboratories), and images were acquired on a confocal microscope (TCS SP5; Leica) with Leica LAS AF software. The following lenses were used: HC PL APO 20×/0.70, HCX PL APO 40×/1.25-0.75 oil, and HCX PL APO 63×/1.40-0.60 oil. All images were taken at room temperature and processed in Image J for CSA analysis. Occasionally, images were linearly rescaled to optimize brightness and contrast uniformly without altering, masking, or eliminating data. CSA was calculated from at least 15 cells per condition and from representative triplicate experiments.

High-resolution PET. Myocardial glucose uptake was noninvasively assessed in vivo by monitoring FDG uptake in intact mice. Mice were fasted for 12 hours prior to imaging, then anesthetized with isoflurane (1.5%) and maintained at 37°C. Each mouse was injected with 10 M Bq FDG in 100 μl 0.9% saline i.v. (tail vein). After 60 minutes, mice were positioned on the bed of a submillimeter-resolution PET camera, and a 15-minute acquisition was initiated. Coronal images were then reconstructed, and myocardial FDG uptake was quantified as the ratio of radioactivity in a myocardial region of interest relative to the total injected dose (expressed as percent injected dose).

Adult mouse cardiomyocyte isolation. Adult mouse ventricular cardiomyocytes were isolated after enzymatic dissociation as previously described (58). Briefly, after retrograde perfusion with Krebs-Ringer solution (2 ml/min for 5 minutes), the heart was perfused with fresh solution containing 0.8 mg/ml collagenase (Worthington type II) for another 12–15 minutes. The LV was removed and cut into small pieces in KB solution (10 m M taurine; 70 m M glutamic acid; 25 m M K Cl; 10 m M KH2PO4; 22 m M glucose; and 0.5 m M EGTA, p H 7.2). After filtration, cells were kept in KB buffer and studied within 4–6 hours. All isolation steps were carried out at 36°C with continuous gassing with 95% O2, 5% CO2. Only Ca2+-tolerant, quiescent, rod-shaped cells showing clear cross striations were used.

Primary cardiac cell preparation and adenovirus infection. Neonatal rat ventricular myocytes were isolated from the ventricles of Sprague-Dawley rat pups on postnatal day 1–2. Cells were preplated (2 hours) to enrich for cardiac myocytes, plated at a density of 1,200 cells/mm2, and cultured for 24 hours in DMEM/M199 (3:1) containing 5% FBS, 10% horse serum, and 100 μmol/l Brd U (Sigma-Aldrich) (7). For adenovirus-mediated protein overexpression, cells were infected 48 hours after plating with GFP or GFP-tagged ca Fox O1 adenovirus at a multiplicity of infection of 15 plaque-forming units per cell. Cells were cultured for an additional 24 hours in fresh DMEM/M199 supplemented with 3% FBS, then harvested in M-PER buffer (Thermo Scientific) containing phosphatase and protease inhibitors (Roche).

RNA isolation, RT, and quantitative PCR analysis. Mouse tissues were harvested and frozen immediately in liquid nitrogen and stored at –80°C until use. Total RNA was isolated using the R Neasy Mini kit according to the recommendations of the manufacturer (Qiagen). A total of 1 μg RNA from each sample was used for RT using i Script c DNA synthesis kit (Bio-Rad). The c DNA was diluted by 10 using dd H2O and used for quantitative PCR analysis (Roche). Primer pairs used for RT-PCR are listed in Supplemental Table 2. A ΔCt method was used to calculate relative gene expression.

Statistics. Depending on the experimental design, averaged data (expressed as mean ± SD) were analyzed either by Student’s unpaired, 2-tailed t test or by 2-way ANOVA followed by Holm-Sidak post-hoc test. Comparison of FS over time between groups was performed using 2-way repeated-measures ANOVA. For statistical comparisons, a P value less than 0.05 was considered statistically significant. All statistical analyses were performed using Sigma Stat (version 3.1) software.

We thank Willie Young for assistance with the genotyping and Anwarul Ferdous and Craig Malloy for helpful discussions. We also thank Patrick Thomas and Xiankai Sun for guidance and assistance with the PET. This work was supported by grants from the NIH (HL-075173, HL-080144, and HL-090842 to J.A. Hill, and HL-072016 and HL 097768 to B.A. Rothermel), American Heart Association (0640084N to J.A. Hill and 0655202Y to B.A. Rothermel), American Diabetes Association Mentor-Based Postdoctoral Fellowship (7-08-MN-21-ADA to J.A. Hill and P.K. Battiprolu), the American Heart Association–Jon Holden De Haan Foundation (0970518N to J.A. Hill), and the Fondo Nacional de Desarrollo Cientifico y Tecnologico, Chile (FONDECYT 1120212, and FONDAP 1500006 to S. Lavandero).