Biochimica et Biophysica Acta 1841 (2014) 1525–1537

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Cardiac diastolic dysfunction in high-fat diet fed mice is associated with lipotoxicity without impairment of cardiac energetics in vivo Desiree Abdurrachim a,1, Jolita Ciapaite a,b,1, Bart Wessels a, Miranda Nabben a, Joost J.F.P. Luiken c, Klaas Nicolay a, Jeanine J. Prompers a,⁎ a b c

Biomedical NMR, Department of Biomedical Engineering, Eindhoven University of Technology, Den Dolech 2, 5612 AZ Eindhoven, The Netherlands Department of Pediatrics, Center for Liver, Digestive and Metabolic Diseases, University of Groningen, University Medical Center Groningen, Hanzeplein 1, 9713 GZ Groningen, The Netherlands Department of Molecular Genetics, Cardiovascular Research Institute Maastricht (CARIM), Maastricht University, P.O. Box 616, 6200 MD Maastricht, The Netherlands

a r t i c l e

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Article history: Received 9 April 2014 Received in revised form 4 July 2014 Accepted 23 July 2014 Available online 30 July 2014 Keywords: Diet-induced obesity Cardiomyopathy Cardiac energetics Lipotoxicity Calcium homeostasis

a b s t r a c t Obesity is often associated with abnormalities in cardiac morphology and function. This study tested the hypothesis that obesity-related cardiomyopathy is caused by impaired cardiac energetics. In a mouse model of high-fat diet (HFD)-induced obesity, we applied in vivo cardiac 31P magnetic resonance spectroscopy (MRS) and magnetic resonance imaging (MRI) to investigate cardiac energy status and function, respectively. The measurements were complemented by ex vivo determination of oxygen consumption in isolated cardiac mitochondria, the expression of proteins involved in energy metabolism, and markers of oxidative stress and calcium homeostasis. We also assessed whether HFD induced myocardial lipid accumulation using in vivo 1H MRS, and if this was associated with apoptosis and fibrosis. Twenty weeks of HFD feeding resulted in early stage cardiomyopathy, as indicated by diastolic dysfunction and increased left ventricular mass, without any effects on systolic function. In vivo cardiac phosphocreatine-to-ATP ratio and ex vivo oxygen consumption in isolated cardiac mitochondria were not reduced after HFD feeding, suggesting that the diastolic dysfunction was not caused by impaired cardiac energetics. HFD feeding promoted mitochondrial adaptations for increased utilization of fatty acids, which was however not sufficient to prevent the accumulation of myocardial lipids and lipid intermediates. Myocardial lipid accumulation was associated with oxidative stress and fibrosis, but not apoptosis. Furthermore, HFD feeding strongly reduced the phosphorylation of phospholamban, a prominent regulator of cardiac calcium homeostasis and contractility. In conclusion, HFD-induced early stage cardiomyopathy in mice is associated with lipotoxicityassociated oxidative stress, fibrosis, and disturbed calcium homeostasis, rather than impaired cardiac energetics. © 2014 Elsevier B.V. All rights reserved.

1. Introduction Over-nutrition and consumption of high-fat diets are associated with obesity-related cardiomyopathy and increased risk for heart failure [1,2]. In obesity, the elevated plasma free fatty acid (FFA) concentration induces an increase in myocardial fatty acid (FA) uptake and oxidation [3]. Despite the increase in FA oxidation, the excessive availability of FA may lead to an imbalance between FA uptake and oxidation, resulting in an increased deposition of potentially toxic lipids in the heart [4,5]. Myocardial lipid accumulation has been implicated in the development of cardiomyopathy through a number of pathways, including lipid-induced apoptosis [6] and fibrosis [7,8]. ⁎ Corresponding author at: Department of Biomedical Engineering, Eindhoven University of Technology, PO Box 513, 5600 MB Eindhoven, The Netherlands. Tel.: +31 40 247 31 28. E-mail address: [email protected] (J.J. Prompers). 1 Authors contributed equally.

http://dx.doi.org/10.1016/j.bbalip.2014.07.016 1388-1981/© 2014 Elsevier B.V. All rights reserved.

Upregulation of FA oxidation may also result in a disturbance in cardiac energetics. Compared to glucose oxidation, stoichiometrically, FA oxidation produces 10% less ATP per oxygen consumed (i.e. 10% decrease in cardiac efficiency). However, up to 30% lower cardiac efficiency has been observed as a consequence of increased FA oxidation, suggesting that also other mechanisms than the inefficiency of FA oxidation affect the cardiac efficiency [9]. The increase in the supply of reducing equivalents (i.e. NADH and FADH2) to the electron transport chain (ETC), without a parallel increase in the oxidative phosphorylation (OXPHOS) capacity, might result in the loss of electrons from the ETC and subsequently, the generation of reactive oxygen species (ROS) [10]. In turn, increased ROS may promote mitochondrial uncoupling [11–15], as a mechanism to reduce the electrochemical proton gradient required for ROS formation, which will however result in lower ATP generation. ROS may induce mitochondrial defects, such as shown by lower expression of OXPHOS proteins in obese and diabetic mice [16,17]. Increased ROS production might also stress sarco/endoplasmic reticulum, which could lead to disturbed calcium homeostasis

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and thereby impair cardiac muscle contraction/relaxation [18]. Although in vitro data suggest that cardiac energy metabolism is disturbed in obesity, in vivo evaluation of cardiac energetics is currently lacking. In animal models, cardiac energetics is usually assessed ex vivo in isolated perfused heart setups by measuring cardiac power and oxygen consumption to calculate cardiac energy efficiency [19,20]. Reduced cardiac efficiency has been observed in rodents with diet-induced obesity [15,21], obese ob/ob mice [16,20,22], and diabetic db/db mice [19,20, 23]. Unfortunately, the ex vivo setup does not entirely mimic the complexity of the in vivo situation as the ex vivo heart is usually perfused with glucose and FA at a constant concentration, while in obesity the in vivo heart is exposed to a hyperlipidemic and, possibly, a hyperinsulinemic environment, which affects the choice of substrates. In vivo cardiac energetics can be assessed using 31P magnetic resonance spectroscopy (MRS), in which the PCr/ATP ratio is used as a measure of cardiac energy status [24,25]. The implementation of 31P MRS in vivo in the mouse heart has proven very challenging, because of its small size (~ 2000 times smaller than the human heart) and high heart rates (~10 times faster than the human heart). In a number of recent studies, in vivo cardiac 31P MRS has been applied in mouse models of heart failure, demonstrating a robust association between heart failure and cardiac energy deficiency [26,27]. However, in vivo cardiac PCr/ATP data in obesity-related cardiomyopathy is currently not available. In this study, we applied cardiac 31P MRS in a mouse model of dietinduced obesity to investigate whether obesity-related cardiomyopathy is associated with an impairment in cardiac energetics in vivo. For this purpose, C57BL/6J mice were fed a high-fat diet (HFD) for 20 weeks. Cardiac systolic and diastolic function were examined using cinematic magnetic resonance imaging (MRI). The 31P MRS measurements of in vivo cardiac energy status were complemented by ex vivo determinations of oxygen consumption in isolated cardiac mitochondria, the expression or activity of various proteins involved in energy metabolism, as well as markers of oxidative stress and calcium homeostasis. Additionally, we assessed whether HFD feeding induced myocardial lipid accumulation using in vivo localized 1H MRS and if this was associated with apoptosis and fibrosis.

2. Materials and methods 2.1. Animals Male C57BL/6J mice were purchased from Charles River (Elsene, Belgium). The animals were housed under controlled temperature (23 °C) and humidity (50%) with a 12:12-h dark–light cycle and were given ad libitum access to food and water. Starting from the age of 12 weeks, the animals were divided into two groups (n = 8 per group) and fed either low fat diet (LFD; 10 kcal% (4.3 g%) palm oil-based fat, 70 kcal% (67.3 g%) carbohydrate, 20 kcal% (19.2 g%) protein; based on OpenSource Diets No. D12450B, Research Diet Services, Wijk bij Duurstede, the Netherlands) or HFD (45 kcal% (24 g%) palm oil-based fat, 35 kcal% (41 g%) carbohydrate, 20 kcal% (24 g%) protein; based on OpenSource Diets No. D12451, Research Diet Services, Wijk bij Duurstede, The Netherlands). The animals received the diet for 20 weeks. The animals then underwent MR measurements in the fed state, after which they were sacrificed within one week, during which they were maintained on the same diet. The heart was excised and part of it was used immediately for the isolation of mitochondria. The other part was either snap-frozen in liquid nitrogen and stored at − 80 °C until further biochemical analyses, or immersed in 4% paraformaldehyde (PFA) in PBS at 4 °C overnight and embedded in paraffin the following day. Animal handling procedures and experimental protocols conformed to and were approved by the Animal Experimental Committee of Maastricht University (The Netherlands).

2.2. MR measurements All MR measurements were performed on a 9.4 T horizontal bore MR scanner (Bruker, Germany). MRI and 1H MRS were performed using a 35-mm quadrature birdcage coil (Bruker, Germany) for both signal reception and transmission. For 31P MRS, a 54-mm double tuned quadrature 1H and linear 31P birdcage coil (Rapid Biomedical, Germany) was used for signal transmission, while a 15-mm diameter, home-built, actively decoupled, two-turn 31P surface coil was used for signal reception. Before the experiments, the animals were sedated in a chamber with 3% isoflurane in medical air at a flow rate of 0.4 L/min. During the measurements, the anesthesia was maintained at 1–2% isoflurane through a customized anesthesia mask. Temperature was maintained at 36–37 °C with a heating pad. Rectal temperature, ECG signal, and breathing rate were monitored throughout the measurements. All measurements were performed with respiratory gating and cardiac triggering. 2.2.1. Cardiac function measurement using cardiac cine MRI Cardiac systolic and diastolic function were measured as described previously [28]. For systolic function measurement, cine movies from the beating heart (15–18 frames/cardiac cycle) were acquired using prospectively cardiac-triggered gradient echo imaging of 5–6 contiguous short axis and 2 long axis slices (slice thickness: 1 mm). The imaging parameters were as follows: repetition time: 7 ms, echo time: 1.8 ms, flip angle: 15°, matrix: 192 × 192, and field of view: 30 × 30 mm2. For diastolic function, cardiac movies of only the mid-ventricular slice were acquired using retrospectively-triggered gradient echo imaging, allowing data reconstruction with a much higher temporal resolution (50–60 frames/cardiac cycle). The parameters used were as follows: repetition time: 4.7 ms, echo time: 2.35 ms, flip angle: 15°, matrix: 128 × 128, field of view: 30 × 30 mm2, and effective time resolution: 2 ms. Image segmentation and data analysis were performed using CAAS MRV 2.0 (Pie Medical, Maastricht, the Netherlands) or Segment (version 1.8 R1145, http://segment.heiberg.se). 2.2.2. Cardiac energy status measurements using 31P MRS Cardiac 31P MR spectra were acquired using the image selected in vivo spectroscopy (ISIS) sequence, on a voxel of typically ~ 6 × 6 × 6 mm 3 covering the left ventricle, at the end of the diastolic phase, as described previously [29]. The parameters were as follows: repetition time: ~ 2 s, 1.2 ms 90° sinc-shaped excitation pulse (bandwidth: 32.0 ppm), 6.25 ms 180° adiabatic hyperbolic secant inversion pulses (bandwidth: 37.5 ppm), 96 ISIS cycles (768 scans), and γ-ATP on resonance. The 90° sinc-shaped excitation pulse was calibrated during the in vivo scan, by performing a series of single pulse measurements with varying pulse power on a 5-mm diameter glass phantom containing 15 M phosphoric acid, which was positioned underneath the surface coil. Data fitting and analysis were performed using AMARES in jMRUI [30], as described as in [29]. The γ-ATP line widths (LWγ-ATP) were constrained relative to the PCr line width (LWPCr) according to an empirically determined relation: LWγ-ATP = LWPCr + 14.8 Hz (n = 63 data sets, R = 0.78, P b 0.001). As a measure of cardiac energy status, the ratio of PCr to γ-ATP was determined. The PCr/ATP ratio was corrected for partial T1 saturation using correction factors of 1.75 and 1.31 for PCr and γ-ATP, respectively, which were determined by analyzing ISIS spectra acquired at repetition times of 2 s and 15 s (n = 19 data sets from the current and previous studies). The contamination of the ATP from the blood to the spectra was shown to be less than 4%, and was considered negligible [31]. 2.2.3. Myocardial lipid measurements using 1H MRS 1 H MR spectra were acquired from a 1 × 2 × 2 mm3 voxel positioned in the interventricular septum during the diastolic phase of the cardiac cycle, using point resolved spectroscopy (PRESS) with chemical shift selective (CHESS) water suppression, as described in [32]. The parameters were as follows: repetition time: ~ 2 s, echo time: 9.1 ms,

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0.41 ms 90° Hermite-shaped excitation pulse, 0.9 ms 180° Mao-type refocusing pulses, and 256 averages. The spectra were processed and analyzed using AMARES in the jMRUI software package [30]. All metabolites (taurine, choline/carnitine, creatine, and seven peaks from lipids) were fitted to Gaussian line shapes. Myocardial metabolite levels were then calculated from the metabolite signal relative to the unsuppressed water signal measured from the same voxel.

2.3. Isolation of mitochondria Following excision, the heart was rinsed in ice cold 0.9% KCl, quickly minced with scissors in 0.5 ml of medium containing 160 mM KCl, 10 mM NaCl, 20 mM Tris, 5 mM EGTA and 0.05 mg/ml bacterial proteinase type XXIV (pH 7.7), and incubated for ~1 min on ice. The mixture was then homogenized using a Potter–Elvehjem homogenizer. Mitochondria were isolated through a differential centrifugation procedure as described in [33] and resuspended in buffer containing 180 mM KCl, 20 mM Tris, 3 mM EGTA and 1 mg/ml bovine serum albumin (pH 7.4). Protein content was determined using a BCA protein assay kit (Pierce, Thermo Fisher Scientific Inc., Rockfort, IL, USA).

2.4. High-resolution respirometry A 2-channel high-resolution Oroboros oxygraph-2k (Oroboros, Innsbruck, Austria) was used to measure oxygen consumption rates at 37 °C. Isolated mitochondria (0.15 mg/ml) were incubated in 1 ml of assay medium (110 mM KCl, 20 mM Tris, 2.3 mM MgCl2, 5 mM KH2PO4 and 1 mg/ml bovine serum albumin, pH 7.2) with i) 5 mM pyruvate plus 5 mM malate, or ii) 25 μM palmitoyl-L-carnitine plus 2 mM malate as the oxidizable substrates. The maximal O2 consumption rate in a coupled state of oxidative phosphorylation (state 3) was measured after the addition of 0.1 mg/ml hexokinase, 12.5 mM glucose and 1 mM ATP. The basal O2 consumption rate (state 4) was measured after blocking ATP synthesis with 1.25 μM carboxyatractyloside (CAT). The uncoupled O2 consumption rate (state U) was determined after the addition of 1 μM carbonyl cyanide 3-chlorophenyl hydrazone (CCCP). Data acquisition and analysis were performed using Oxygraph-2 k-Datlab 4.3.1.15 software (Oroboros, Innsbruck, Austria).

2.5. Citrate synthase activity Tissues were weighed and 20% (w/v) homogenates were prepared in ice cold PBS, pH 7.4. Homogenates were sonicated for 30 s in the pulse mode (pulse duration 1 s, interval between the pulses 1 s, power input 10 W) on ice. Half of each sample was used for the determination of acylcarnitine concentrations (see below). The remainder of the sample was centrifuged at 1000 g for 10 min at 4 °C. Protein content in the supernatant was measured using BCA protein assay kit (Pierce, Thermo Fisher Scientific Inc., Rockford, IL, USA). Citrate synthase activity was determined spectrophotometrically as described in [34].

2.6. Determination of ATP, ADP, and AMP concentration Tissues were weighed, grinded in liquid nitrogen and homogenized in 0.5 M perchloric acid at 4 °C, followed by centrifugation for 10 min at 14,000 g, 4 °C. The supernatant was neutralized with 2 M KOH to ~pH 7 and centrifuged for 10 min at 14,000 g, 4 °C. Nucleotides in 100 μl of supernatant were separated with HPLC as described in [35]. The concentrations in the eluate were calculated from ATP, ADP and AMP (SigmaAldrich, Zwijndrecht, The Netherlands) standard curves and expressed as nmol/mg wet tissue.

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2.7. Quantification of acylcarnitines Sonicated 20% homogenates were centrifuged for 10 min at 14,000 g, 4 °C. Next, 10 μl of the supernatant was mixed with 100 μl acetonitrile and 100 μl of methanol–water (80:20 v/v) containing internal standards [1,1,1-N-methyl-2H3]-L-carnitine, [2H3]acetyl-L-carnitine, [3,3,3-2H3] propionyl-L-carnitine, [8,8,8-2H3]octanoyl-L-carnitine, [10,10,10-2H3] decanoyl-L-carnitine and [16,16,16-2H3]hexadecanoyl-L-carnitine (VU Medical Center, Amsterdam, The Netherlands), vortexed and centrifuged for 10 min at 14,000 g at 4 °C. Concentrations of acylcarnitines were measured in the supernatant with an API 3000 LC–MS/MS equipped with a Turbo ion spray source (Applied Biosystems/MDS Sciex, Ontario, Canada) as described in [36].

2.8. Biochemical determination of intramyocelullar lipids Intramyocellular lipids were determined in cardiac muscle homogenates as described previously [37]. In short, samples containing 400 μg of protein were used for intracellular lipid extraction in methanol/chloroform, and an internal standard and water were added. Afterwards thin-layer chromatography was used to separate lipids. Bands were resolved with a hexane/diethylether/propanol (87:10:3) resolving solution. Triacylglycerol (TAG), diacylglycerol (DAG), monoacylglycerol (MAG), and cholesterol bands were detected with a Molecular Imager (ChemiDoc XRS, BioRad) and analyzed with Quantity One® (BioRad).

2.9. Immunoblotting Total cardiac protein extracts were prepared, resolved with SDSPAGE and transferred to polyvinylidene difluoride membranes (Millipore, Bedford, MA, USA) as described in [38]. After blocking with TBS containing 0.1% Tween 20 and 5% skim milk powder for 1 h at room temperature, the membranes were incubated overnight at 4 °C with one of the following antibodies: rabbit polyclonal antiperoxisome proliferator-activated receptor γ coactivator 1α (PGC1α) (1:500), rabbit anti-caspase 3 (1:500), rabbit polyclonal anti-collagen α1 Type I (1:1000), rabbit anti-cardiac troponin T (1:500), goat antisarcoplasmic reticulum calcium ATPase2 (SERCA2) (1:200), rabbit anti-phospholamban (1:200), rabbit anti-phospho Thr17 phospholamban (1:200) (all antibodies were from Santa Cruz Biotechnology, Santa Cruz, CA, USA), rabbit anti-uncoupling protein 3 (UCP3; 1:500; SigmaAldrich), rabbit polyclonal anti-caspase 9 (1:1000; Stressgen Biotechnologies Corporation, San Diego, CA, USA), rabbit polyclonal anti-Cu/ Zn superoxide dismutase (SOD; 1:1000; Stressgen Biotechnologies Corporation), rabbit polyclonal anti-Mn superoxide dismutase (1:1000; Stressgen Biotechnologies Corporation), or MitoProfile® Total OXPHOS Rodent WB Antibody Cocktail (1:2000, MitoSciences, Eugene, OR, USA) containing mouse monoclonal antibodies against Complex I, II, III, IV and V subunits. Membranes were washed 3 × for 5 min with TBS containing 0.1% Tween 20 and incubated with a corresponding horse-radish peroxidase-conjugated secondary antibody for 1 h at room temperature. After the final wash of 3× for 5 min with TBS containing 0.1% Tween 20 and 1× 5 min with TBS, the immunocomplexes were detected using SuperSignal West Dura Extended Duration Substrate (Pierce, Thermo Fisher Scientific Inc., Rockford, IL, USA), visualized using ChemiDoc™ XRS + imaging system and quantified using Image Lab™ analysis software version 3.0 (Bio-Rad Laboratories Inc., Hercules, CA, USA). All data were normalized to glyceraldehyde 3phosphate dehydrogenase (GAPDH) expression level and expressed relative to the LFD-fed controls. Immunoblot detection of protein carbonylation in the total cardiac protein extracts was done using the OxyBlot™ Protein Oxidation Detection Kit (Chemicon/Millipore, Billerica, MA, USA) following recommendations of the manufacturer.

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2.10. Histology

compared with LFD mice, indicating hyperlipidemia in HFD mice (Table 1).

The paraffin-embedded hearts were sliced in 5 μm-thick sections. The collagen was stained with Picrosirius red (Direct Red 80, SigmaAldrich, Zwijndrecht, The Netherlands) according to standard histological procedures. To detect apoptotic cells, terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) staining was performed using Apoptag peroxidase in situ apoptosis detection kit (Millipore Corporation, Billerica, MA, USA) according to manufacturer's protocol. The stained sections were digitalized with a Pannoramic MIDI scanner (3DHISTECH, Budapest, Hungary). Fibrotic area, represented by red area in Sirius Red staining, was quantified from 15 random locations in 2–4 heart slices (magnification 40×), using a macro in ImageJ 1.47v (NIH, USA). TUNEL-positive cells were counted from 5 random locations (40 × magnification, about 200–300 cells per location) in a slice, and quantified as a percentage of the total number of cells detected using ‘analyze particles’ in ImageJ 1.47v (NIH, USA). 2.11. Plasma FFA, triglyceride, insulin, and glucose determination Blood samples were collected in EDTA coated tubes, at 20 weeks of feeding after 4 h fasting and at sacrifice (fed conditions). Plasma was obtained by centrifuging the blood at 1000 g for 10 min and stored at −80 °C until plasma insulin, FFA, and triglyceride (TG) determination. Fasting plasma glucose levels were measured immediately after blood collection in one drop of blood using a Glucose-201 glucose meter (HemoCue, Ängelholm, Sweden). Plasma insulin levels were determined using a mouse insulin determination kit (Mercodia, Uppsala, Sweden). Plasma TG levels were determined using a TG determination kit (Sigma-Aldrich, Zwijndrecht, The Netherlands). Plasma FFA levels were determined using a NEFA-HR kit (Wako Chemicals, Neuss, Germany). 2.12. Statistical analysis All data are presented as means ± standard deviation. Statistical analysis was performed with SPSS 17.0 (SPSS Inc.) using independentsamples T-test. Multivariate ANOVA was applied to analyze different lipid peaks in the cardiac 1H MR spectra. The level of significance was set at P b 0.05.

3.2. HFD feeding induced increased left ventricular mass and diastolic dysfunction Ejection fraction, end diastolic volume, end systolic volume, and stroke volume were all similar between LFD and HFD mice (P = 0.16, 0.56, 0.73, 0.23, respectively; Table 2). However, left ventricular (LV) mass and LV mass normalized to tibia length were 13–16% higher in HFD mice compared with LFD mice (P b 0.05; Table 2 and Fig. 1A). MRI with high temporal resolution (time resolution: 2 ms) was performed on the midventricular slice to determine cardiac diastolic function parameters. From the images, the LV cavity volume–time curve (Fig. 1B) and the time derivative of the LV cavity volume–time curve (Fig. 1C) were generated to analyze the early and late diastolic filling phases. Early peak filling rates (E) were 15% lower (P b 0.05) in HFD mice compared with LFD mice (Table 2 and Fig. 1C), which indicates an impairment in the active relaxation phase of the left ventricle. The late peak filling rate (A), i.e. the rate of diastolic filling due to atrial contraction, did not differ significantly between LFD and HFD mice (P = 0.11; Table 2 and Fig. 1C). In both LFD and HFD mice, the early filling phase contributed about 60% of the total diastolic filling (P = 0.85; Table 2). 3.3. HFD feeding did not alter cardiac energy status Cardiac energy status in vivo was determined using 31P MRS localized in the left ventricle (Fig. 2A). A typical 31P MR spectrum contains a phosphocreatine (PCr) peak originating from myocardium, a 2,3diphosphoglycerate (DPG) peak from blood in the LV lumen (obscuring the inorganic phosphate (Pi) peak from myocardium), and γ-, α-, and β-ATP peaks from mainly myocardium with a minor contamination from blood (Fig. 2B). PCr/γ-ATP ratios were similar for HFD and LFD mice (P = 0.73; Fig. 2C), indicating that 20 weeks of HFD feeding did not affect cardiac energy status in vivo. ATP, ADP, and AMP concentrations, determined biochemically, were similar between HFD and LFD mice (P = 0.27, P = 0.85, and P = 0.73, respectively; Fig. 2D). 3.4. HFD feeding increased mitochondrial FA oxidation capacity, without altering mitochondrial protein levels

3. Results 3.1. Mouse characteristics After 20 weeks of feeding, body weight of HFD mice was 50% higher compared with LFD mice (P b 0.001, Suppl Fig. 1A). The average caloric intake over the 20 weeks of feeding tended to be higher for HFD mice than for LFD mice (18%, P = 0.052, Suppl Fig. 1B). Fasting plasma glucose levels were higher in HFD compared with LFD mice (P = 0.018; Table 1). Fasting plasma insulin tended to be higher in HFD compared with LFD mice (P = 0.082; Table 1). Plasma TG and FFA levels, determined in the fed state, were about 1.5-fold higher (P b 0.05) in HFD Table 1 Plasma insulin, glucose, total TG, and FFA concentrations. LFD Insulin (fasted, ng/mL) Glucose (fasted, mM) Total TG (fasted, mM) Total TG (fed, mM) FFA (fed, mM)

0.87 9.03 0.63 0.59 0.39

HFD ± ± ± ± ±

0.50 0.99 0.10 0.20 0.09

2.25 11.23 0.62 0.92 0.63

± ± ± ± ±

1.02# 1.97⁎ 0.07 0.30⁎ 0.17⁎⁎

Data are means ± SD (n = 8 per diet group). TG: triglycerides, FFA: free fatty acids. # P = 0.08 vs. LFD. ⁎ P b 0.05 vs. LFD. ⁎⁎ P b 0.01 vs. LFD.

Oxygen consumption rates when ETC is coupled with ATP synthase (coupled state, state 3) and in the uncoupled state (state U) were similar in isolated heart mitochondria from LFD and HFD mice, when pyruvate plus malate was used as the oxidizable substrate (P = 0.38 and P = 0.57, respectively; Fig. 3A). This indicates that glucose oxidation capacity did not change upon HFD feeding. When palmitoyl-Lcarnitine plus malate was used as the substrate, state 3 and state U Table 2 MRI parameters of cardiac function. LFD EF (%) EDV (μL) ESV (μL) SV (μL) PER (%EDV/ms) E (%EDV/ms) A (%EDV/ms) E contribution (%EDV) LV mass (mg) LV mass/tibia length (mg/mm)

72.3 71.6 20.2 51.4 2.3 2.5 1.6 60.4 101.8 5.6

HFD ± ± ± ± ± ± ± ± ± ±

3.6 14.5 6.4 8.2 0.2 0.2 0.2 5.3 13.5 0.7

74.7 75.4 19.2 56.2 2.1 2.1 1.3 61.0 114.9 6.5

± ± ± ± ± ± ± ± ± ±

2.7 11.0 4.5 7.0 0.3 0.3⁎ 0.4 5.4 7.8⁎ 0.5⁎

Data are means ± SD (n = 8 per diet group, except for PER, E, A, and E contribution n = 7). EF: ejection fraction, EDV: end diastolic volume, ESV: end systolic volume, SV: stroke volume, PER: peak ejection rate, E: early peak filling rate, A: late peak filling rate. ⁎ P b 0.05 vs. LFD.

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The activity of citrate synthase, a mitochondrial marker enzyme, was similar in cardiac muscle from both diet groups (LFD: 2.43 ± 0.16 vs HFD: 2.43 ± 0.19 μmol/min/mg protein, n= 8 per group, P = 0.92), suggesting that mitochondrial density was not affected by HFD feeding. In addition, the expression of peroxisome proliferator-activated receptor γ cofactor-1α (PGC1α), which regulates mitochondrial biogenesis, was not altered in HFD hearts (P = 0.75; Table 3). Furthermore, the expression of all mitochondrial oxidative phosphorylation complexes was similar between LFD and HFD hearts (P = 0.83, 0.65, 0.40, 0.80, 0.09 for Complexes I, II, III, IV, and V, respectively; Table 3 and Suppl. Fig. 2).

3.5. HFD feeding induced accumulation of myocardial lipids and lipid intermediates Myocardial lipid content was measured using 1H MRS from a single voxel located in the septum (Fig. 4A). Seven peaks associated with lipids are visible in the spectra (Fig. 4B and C). After 20 weeks of feeding, the peak area from the most prominent lipid signal from methylene protons (peak 9; CH2) was 1.6-fold higher (P b 0.001) in HFD compared with LFD mice (Fig. 4D). In addition, peak areas of the lipid signals from allylic methylene protons (peak 7; CH2\CH_CH\CH2), and β-methylene protons (peak 8; CβH2CH2COO) were significantly higher (P b 0.01) in HFD compared with LFD mice (Fig. 4D). The peak areas from other lipid resonances and those from creatine, choline/carnitine, and taurine were similar in both diet groups (Fig. 4D). In agreement with the higher myocardial lipid content in HFD mice in vivo, ex vivo biochemical analysis also showed 1.6-fold higher TAG levels in HFD mouse hearts compared with LFD mouse hearts (P b 0.001, Fig. 5A). The higher myocardial TAG content in HFD mice was accompanied by elevated levels of DAG (1.4-fold, P = 0.006), MAG (1.7fold, P b 0.001), and cholesterol (1.6-fold, P = 0.004), indicating increased lipotoxicity in the HFD heart (Fig. 5A). Furthermore, HFD feeding resulted in the accumulation of acylcarnitines in the heart (Fig. 5B). As a consequence, the content of free carnitine (C0) in the heart was 40% lower (P b 0.001) in HFD compared with LFD mice (Fig. 5B). This suggests stimulation of FA oxidation by HFD feeding. The observation that the content of acetylcarnitine (C2) was 23% higher (P b 0.001) in the hearts of HFD-fed mice may suggest that the increase in mitochondrial βoxidation capacity was not matched by the TCA cycle capacity to utilize acetyl-coenzyme A (CoA) produced by the former.

3.6. HFD feeding increased the production of reactive oxygen species

Fig. 1. In vivo cardiac function measured using MRI. (A) Two-chamber long axis, 4chamber long axis, and short axis cardiac MR images of a LFD and a HFD mouse. The larger heart size in the HFD compared with the LFD mouse can be appreciated. (B) The LV cavity volume-time curve and (C) the time derivative of LV cavity volume-time curve, showing the peak ejection rate (PER), early peak filling rate (E), and late peak filling rate (A). Open symbols: LFD, closed symbols: HFD.

oxygen consumption rates in isolated HFD heart mitochondria were 1.3- (P b 0.01) and 1.4-fold (P b 0.05) higher than those in isolated LFD heart mitochondria, respectively (Fig. 3B), indicating an increased FA oxidation capacity in the HFD group. The oxygen consumption rates in state 4 were similar in HFD and LFD groups, for both pyruvate plus malate and palmitoyl-L-carnitine plus malate as substrates (P = 0.07 and P = 0.85, respectively; Fig. 3), indicating that HFD feeding did not affect the leakiness of the inner mitochondrial membrane to protons. In agreement, the expression of mitochondrial uncoupling protein 3 (UCP3) was similar between LFD and HFD hearts (P = 0.90; Table 3).

To investigate the effect of HFD feeding on the production of reactive oxygen species (ROS), we measured the expression of cytosolic superoxide dismutase (Cu/ZnSOD) and mitochondrial superoxide dismutase (MnSOD), two enzymes involved in the scavenging of superoxide anion radicals produced mostly during mitochondrial respiration. The expression of these enzymes was higher (Cu/ZnSOD: 16%, P b 0.05; MnSOD: 20%, P b 0.001) in HFD compared with LFD mice (Fig. 6A–B). In agreement, we observed an increase in protein oxidation in HFD compared with LFD hearts (Fig. 6C–D).

3.7. HFD feeding and apoptosis The expression of caspases involved in the initiation of apoptosis, procaspase-9 and cleaved (mature) caspase-9, was 37% (P b 0.05) and 32% (P b 0.01) higher, respectively, in hearts of HFD mice compared with LFD mice (Fig. 7A–B). However, the expression level of executing caspase, procaspase-3, which is processed by caspase-9 into mature caspase-3, was similar between HFD and LFD groups (Fig. 7C). The mature caspase-3 was not detectable in both groups (data not shown). Accordingly, the number of TUNEL-positive cells was not different between TUNEL-stained LFD and HFD heart slices (P = 0.56; Fig. 7D–E).

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Fig. 2. Cardiac energy status measured using 31P MRS. (A) Two-chamber long axis, 4-chamber long axis, and short axis MR images from a LFD mouse showing the positioning of the 6 × 6 × 6 mm3 voxel used for 31P MRS in white. (B) Typical example of an in vivo cardiac 31P MR spectrum. (C) PCr/γ-ATP ratio. (D) ATP, ADP, and AMP concentrations, as determined biochemically. Data are means ± SD (n = 8 per diet group). White bar: LFD, black bar: HFD.

3.8. HFD feeding induced myocardial fibrosis The expression of procollagen type I was similar in both diet groups (Fig. 8A). The expression of mature collagen type I, the main collagen

type expressed in the heart, was 40% increased (P b 0.01) in response to HFD feeding (Fig. 8B). This increase in collagen content indicates excess formation of fibrous tissue, which was confirmed by a higher fraction of Sirius Red-enhanced areas in HFD compared with LFD heart

Fig. 3. Oxygen consumption rates in isolated cardiac mitochondria. For (A) pyruvate plus malate, and (B) palmitoyl-L-carnitine plus malate as the substrate. Data are means ± SD (n = 8 per diet group). White bars: LFD, black bars: HFD. *P b 0.05 and **P b 0.01 vs. LFD.

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4. Discussion

Table 3 Cardiac protein expression. LFD UCP3 PGC1α OXPHOS Complex I OXPHOS Complex II OXPHOS Complex III OXPHOS Complex IV OXPHOS Complex V

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1.00 1.00 1.00 1.00 1.00 1.00 1.00

HFD ± ± ± ± ± ± ±

0.10 0.08 0.11 0.11 0.09 0.08 0.09

1.01 0.98 1.01 1.04 1.05 0.99 1.15

± ± ± ± ± ± ±

0.06 0.05 0.09 0.07 0.02 0.03 0.07

Data are means ± SD (n = 3 per diet group). All data were normalized to glyceraldehyde 3-phosphate dehydrogenase (GAPDH) expression level and expressed relative to the LFDfed controls. UCP3: uncoupling protein 3, PGC1α: peroxisome proliferator activator γ coactivator-1α, OXPHOS: oxidative phosphorylation.

slices (P = 0.027; Fig. 8C–D), indicating the presence of increased fibrosis in HFD hearts.

3.9. HFD feeding affected cardiac calcium handling and contractile apparatus Increased ROS might stress sarco/endoplasmic reticulum, which could disturb calcium handling and the contractile apparatus. The expression of sarcoplasmic reticulum calcium ATPase2 (SERCA2) did not change in hearts of HFD compared with LFD mice (P = 0.84; Fig. 9A); however, there was a 60% decrease in the phosphorylation of phospholamban (PLB) (P = 0.018; Fig. 9B), a regulatory protein of SERCA2. HFD feeding also decreased the expression of contractile protein troponin T (cTNT; 19%, P = 0.020; Fig. 9C).

Obesity is often associated with abnormalities in cardiac morphology and function. In this study, we tested the hypothesis that obesityrelated cardiomyopathy is associated with impaired cardiac energy metabolism. In mice fed a HFD for 20 weeks, we evaluated cardiac function, and in vivo and ex vivo parameters of cardiac energy metabolism. In addition, we assessed whether HFD feeding induced myocardial lipid accumulation. Twenty weeks of HFD feeding resulted in diastolic dysfunction and increased LV mass, without changes in systolic function. In vivo cardiac PCr/ATP ratio and ex vivo oxygen consumption in isolated cardiac mitochondria were not reduced after HFD feeding, suggesting that the diastolic dysfunction was not caused by impaired cardiac energetics. HFD feeding promoted mitochondrial adaptations for increased utilization of FA, which was however not sufficient to prevent the accumulation of myocardial lipids and lipid intermediates. In addition, HFD feeding increased oxidative stress markers and collagen deposition, and reduced phosphorylation of phospholamban in the heart. Our data thus show that HFD-induced early stage cardiomyopathy in mice is associated with lipotoxicity-associated oxidative stress, fibrosis, and disturbed calcium homeostasis, rather than impaired cardiac energetics.

4.1. Effect of HFD feeding on cardiac function We showed that 20 weeks of HFD feeding was associated with early stage cardiomyopathy [39], marked by an increased LV mass and lower early peak filling rates, indicating impaired diastolic function, while systolic function was normal. This supports diet-induced obesity as a model of diastolic dysfunction. Similar to our findings, in a previous

Fig. 4. Myocardial lipid content measured using in vivo 1H MRS. (A) Short axis and 4-chamber long axis MR images showing the positioning of the 1 × 2 × 2 mm3 voxel used for 1H MRS in black. Representative in vivo 1H MR spectra from (B) a LFD-fed mouse and (C) a HFD-fed mouse. (D) Metabolite signal amplitude expressed as a percentage of the water signal. Data are means ± SD (n = 8 per diet group). White bars: LFD, black bars: HFD. Bold: resonating protons for the corresponding triglyceride (TG) peaks. **P b 0.01 and ***P b 0.001 vs. LFD.

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Fig. 5. Cardiac lipid intermediates (A) Triacylglycerols (TAG), diacylglycerols (DAG), monoacylglycerols (MAG), and cholesterol, and (B) cardiac acylcarnitine profiles. Data are means ± SD (n = 6–8 per diet group). White bars: LFD, black bars: HFD. *P b 0.05, **P b 0.01 and ***P b 0.001 vs. LFD.

study, 20 weeks of HFD feeding in mice did not affect systolic function, although no data on diastolic function was reported [40]. In contrast, another study in mice reported lower systolic function [17,41] and minor diastolic dysfunction [41] after 12 weeks of HFD feeding. In Wistar rats, posterior wall thickness increased and systolic function decreased after 8 weeks of HFD feeding [42]. In Sprague Dawley rats fed a HFD for 32 weeks, LV hypertrophy was observed without any cardiac dysfunction [43]. These different results might be caused by differences in experimental conditions: different types of animal models, differences in macronutrient composition of the diet and duration of the experiment, as well as differences in the sensitivity of the measurement techniques. 4.2. The role of cardiac energetics in HFD-induced cardiac dysfunction Continuous repetition of the cardiac contraction–relaxation cycle requires permanent supply of energy in the form of ATP. Therefore, an impairment of mitochondrial ATP production may underlie HFD-induced diastolic dysfunction. However, in vivo cardiac 31P MRS showed that 20 weeks of HFD feeding did not affect the cardiac PCr/ATP ratio, a measure of in vivo cardiac energy status. To our knowledge, we are the first to measure the cardiac PCr/ATP ratio in vivo in a mouse model of HFDinduced obesity. The finding that HFD feeding had no effect on the cardiac PCr/ATP ratio in vivo indicates that in this condition ATP production still meets the ATP demand. This was also supported by unaltered ATP,

ADP, and AMP concentrations in hearts of HFD compared with LFD mice, as determined biochemically. These data are in agreement with observations in perfused heart setups, where unaffected PCr and ATP concentrations have been reported in mouse hearts after 20 weeks of HFD feeding [40]. The PCr/ATP ratio measured in this study (0.85 ± 0.15 in LFD mice) is lower compared to literature values of healthy mice [44]. The values are also generally lower than values reported for humans [45,46], which is likely due to methodological differences [47]. It has been recognized that many aspects of localized 31P-MRS acquisition and quantification can contribute to differences in PCr/ATP values [47]. Although the method we used (cardiac-triggered, respiratory-gated 3D ISIS) may underestimate the PCr/ATP values, our method has been shown to be sensitive to detect a decrease in PCr/ATP in the hearts of fasted long-chain acyl-CoA dehydrogenase (LCAD) knockout mice compared with controls [29]. In vitro assessment of mitochondrial function revealed that HFD feeding increased oxidative phosphorylation capacity with the FA βoxidation substrate palmitoyl-L-carnitine, while oxidative phosphorylation capacity for pyruvate was unaffected, showing that cardiac metabolism adapted to the dietary lipid oversupply. This is in agreement with previous findings in perfused heart studies demonstrating that HFD feeding shifts cardiac substrate use from glucose to FA [40,48,49], while myocardial energetics is maintained [40,49]. A shift in cardiac substrate use from glucose to FA has been associated with impaired

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Fig. 6. Markers for the production of ROS. Expression of antioxidant enzymes (A) Cu/Zn SOD (n = 3 per diet group) and (B) Mn SOD (n = 6 per diet group), normalized to GAPDH. (C) Representative immunoblot images and (D) the quantification of protein oxidation using OxyBlot™ (n = 3 per diet group). SOD: superoxide dismutase, GAPDH: glyceraldehyde 3phosphate dehydrogenase, NC: negative control, DNPH: 2,4-dinitrophenylhydrazine. Data are means ± SD. White bars: LFD, black bars: HFD. *P b 0.05, **P b 0.01, ***P b 0.001 vs. LFD.

cardiac function [50,51]. HFD feeding had no effect on the basal (state 4) oxygen consumption rate in isolated cardiac mitochondria or on the expression of cardiac UCP3, suggesting unaltered ion permeability of the inner mitochondrial membrane/mitochondrial uncoupling. In contrast, a previous study in rats fed with HFD showed increased mitochondrial uncoupling, which was associated with reduced cardiac energetics [15]. The maintenance of in vivo cardiac energetics upon HFD feeding in the present study was also supported by unaltered cardiac citrate synthase activity, a mitochondrial marker enzyme, unaltered expression of PGC1α, a key regulator of mitochondrial biogenesis, and unaltered

expression levels of OXPHOS complexes. Studies with other obesity and diabetes mouse models, such as leptin-deficient (ob/ob) mice and leptin receptor-deficient (db/db) mice, showed decreased expression of oxidative phosphorylation proteins in association with impaired cardiac energetics and function [16,17,52], suggesting that genetic obesity has more profound detrimental effects on cardiac energy metabolism compared to HFD-induced obesity. Taken together, our data show that HFD feeding has a subtle effect on cardiac energy metabolism, manifesting in the activation of signal transduction pathways that shift cardiac metabolism toward increased

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Fig. 7. Apoptosis markers. Expression of (A) procaspase-9, (B) caspase-9, and (C) procaspase-3, normalized to GAPDH (n = 3 per diet group). (D) Visualization of TUNEL staining, in which TUNEL-positive cells appear brown, while TUNEL-negative cells appear purple. Black arrows indicate TUNEL-positive cells. (E) The quantification of TUNEL-positive cells (LFD n = 3, HFD n = 4). GAPDH: glyceraldehyde 3-phosphate dehydrogenase, TUNEL: Terminal deoxynucleotidyl transferase dUTP nick end labeling. Data are means ± SD. White bars: LFD, black bars: HFD. *P b 0.05, **P b 0.01 vs. LFD.

use of FA as a fuel, rather than a gross impairment of energy producing function. 4.3. The role of lipotoxicity in HFD-induced cardiac dysfunction The storage of lipids in cardiomyocytes, which occurs when cardiac FA uptake exceeds the rate of oxidation, is a protective mechanism against the formation of intermediate lipid metabolites, such as diacylglycerols, ceramides, and long chain fatty acyl-CoA esters [53,54]. However, prolonged myocardial lipid overload could contribute to cardiac dysfunction through increased ROS production and oxidative stress [55], apoptosis [6,56], fibrosis [7,8], and induction of endoplasmic reticulum (ER) stress [57]. We showed that 20 weeks of HFD feeding resulted in a strong accumulation of myocardial neutral lipids as well as lipid intermediates, such as acylcarnitines and DAG. DAG may interfere with insulin signaling [58], which supports the reduction in cardiac insulin sensitivity as observed in mice as early as after 1.5 weeks of HFD feeding [59]. The accumulation of myocardial lipids suggests that FA supply and uptake exceed the rate of oxidation, despite the increase in mitochondrial FA oxidation capacity [60]. In this respect, we have earlier observed in HFD-fed rats that the main cardiac FA transporter CD36 relocates from intracellular membrane compartments to the sarcolemma [61]. This CD36 relocation occurs in the absence of changes in the expression of this transporter, and causes a chronically increased influx of FA into the heart, followed by insulin resistance, and finally contractile dysfunction [62]. Furthermore, the accumulation of cardiac acylcarnitines indicates that the increased FA oxidation in the HFD mouse hearts is

associated with elevated incomplete FA oxidation [63], which can be explained by the lack of a concomitant increase in OXPHOS capacity as shown by the unchanged glucose oxidation capacity and protein expression of OXPHOS complexes. The increase in FA oxidation without a concomitant increase in OXPHOS capacity may result in the loss of electrons from the ETC and subsequently, excessive stimulation of ROS production. In the present study, we observed upregulation of antioxidant enzymes Cu/ZnSOD and MnSOD and increased protein oxidation in response to HFD feeding, which indeed suggests increased production of ROS. 4.3.1. Lipotoxicity and apoptosis Increased ROS production may lead to the activation of the apoptosis pathway [56,64], as well as lower cardiac efficiency due to the induction of mitochondrial uncoupling, a mechanism to reduce ROS formation at the expense of ATP production [11]. In the present study, HFD feeding had no effect on the coupling of cardiac mitochondria but it indeed induced activation of the apoptosis pathway, as indicated by upregulation of procaspase-9 as well as increased formation of the mature caspase-9. However, we did not observe increased processing of procaspase-3 to its mature form, which would ultimately lead to the execution of apoptosis. Supporting this data, TUNEL staining did not show differences in the amount of apoptotic cells in LFD and HFD heart slices. This is not surprising, since the processing of procaspase-3 to its active form catalyzed by caspase-9 requires additional mitochondrial signals, such as the release of cytochrome c from mitochondria [65] and a drop in cytosolic ATP level [66] due to the loss of mitochondrial integrity and

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Fig. 8. Fibrosis markers. Expression of (A) procollagen I and (B) collagen I, normalized to GAPDH (n = 3 per diet group). (C) Representative images of Sirius Red staining from 3 different mice per diet group, and (D) the quantification of Sirius Red-enhanced areas (LFD n = 3, HFD n = 4). Collagen appears red on yellow cytoplasm background. GAPDH: glyceraldehyde 3phosphate dehydrogenase. Data are means ± SD. White bars: LFD, black bars: HFD. *P b 0.05 vs. LFD.

impairment of mitochondrial function. We showed that ATP concentration was similar in LFD and HFD hearts, and the integrity of cardiac mitochondria was not affected by HFD as indicated by unaltered oxygen consumption rate in state 4, implying that the extent of damage did not yet lead to the actual execution of apoptosis. 4.3.2. Lipotoxicity and fibrosis Excessive production of toxic lipid intermediates such as DAG and ROS stimulates the production of pro-collagens and collagens, through the activation of protein kinases PKC-β or PKA [67]. Collagens, primarily collagen type I, make up the extracellular matrix (ECM) of the heart [67]. The increase in ECM, a hallmark of fibrosis, has been shown to increase myocardial stiffness [68]. The stiffening of myocardium reduces LV compliance (i.e. the change in LV pressure needed to allow the LV filling), and therefore the ability of the muscle to relax (i.e. reduced LV relaxation/diastolic function). Indeed, it has been shown that collagen content was increased in rats with pre-diabetes or upon HFD feeding, together with reduced LV diastolic function [7,8]. In addition, treatment using anti-diabetic drug pioglitazone was shown to normalize LV diastolic function, which was associated with reduced collagen content [8]. In human studies, the serum levels of breakdown products of procollagen I and collagen I (i.e. carboxy-terminal propeptide of procollagen I (PICP) and carboxy-terminal telopeptide of collagen I (CITP)) were shown to be higher in relation to the severity of diastolic dysfunction [69], providing evidence for the relevance of fibrosis in the development of diastolic dysfunction. In the present study, we found higher expression of collagen I levels in HFD compared with LFD mouse hearts. This indication of excess formation of fibrous connective tissue was supported by an increase in the fibrotic area in Sirius Red-stained HFD heart slices. Therefore, our data suggest that

myocardial fibrosis plays a key role in HFD-induced impairment of cardiac function in mice. 4.3.3. Lipotoxicity and calcium homeostasis Finally, the increase in lipotoxicity-induced oxidative stress may impair calcium homeostasis [70,71], which may also play a role in HFDinduced diastolic dysfunction [72], since diastolic dysfunction is characterized by a prolonged relaxation time [73]. Cardiac muscle relaxation after contraction is induced in response to the lowering of the sarcoplasmic calcium concentration due to active re-uptake of calcium into the sarcoplasmic reticulum, catalyzed by sarcoplasmic reticulum calcium ATPases (SERCA). SERCA activity is crucially regulated by the phosphorylation of PLB [74] and a reduction in PLB phosphorylation may be associated with lipotoxicity-induced ER stress as shown in a previous study in HFD mice [70]. In the unphosphorylated state, PLB binds to SERCA, inhibiting calcium pump activity. The phosphorylation of PLB releases this inhibition, which has been shown to result in an increase in calcium transport up to 4-fold or greater [73]. PLB levels were shown to be linearly correlated with the rates of contraction and relaxation in isolated perfused hearts [75] and in hearts of intact mice [76], suggesting that the remarkable 60% decrease in PLB phosphorylation in the HFD mouse hearts may have contributed to the diastolic dysfunction observed in the present study. Supporting our results, a recent study reported decreased PLB phosphorylation, impaired calcium regulation, and contractile dysfunction in cardiomyocytes of HFD-fed mice [70]. In addition, we also observed reduced cardiac troponin T (cTNT) in HFD mouse hearts. cTNT is essential in regulating calcium sensitivity and magnitude of force production during the contraction. Cardiac troponin T mutation or deletion has been associated with diastolic dysfunction in mice [77,78], and a decrease in cTNT expression has been observed in

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D.A., B.W. and J.J.P. are supported by a VIDI grant (project number 700.58.421) from the Netherlands Organisation for Scientific Research (NWO). J.C. is supported by the NWO-funded Groningen Systems Biology Center for Energy Metabolism and Ageing. This work was supported by the Dutch Technology Foundation STW (grant number 10191), the Applied Science Division of NWO and the Technology Program of the Ministry of Economic Affairs. M.W.N. is supported by a VENI grant (project number 916.14.050) from NWO.

References

Fig. 9. Calcium handling and contractile protein. Expression of (A) SERCA2 (n = 3 per diet group), (B) phosphorylated PLBThr17 (n = 3 per diet group), and (C) troponin T (n = 6 per diet group). The expression of SERCA2 and troponin T were normalized to GAPDH. Phosphorylated PLBThr17 was expressed relative to total PLB. SERCA2: sarcoplasmic reticulum calcium ATPase2, GAPDH: glyceraldehyde 3-phosphate dehydrogenase, pPLBThr17: phosphorylated phospholamban, PLB: phospholamban. Data are means ± SD. White bars: LFD, black bars: HFD. *P b 0.05, **P b 0.01 vs. LFD.

response to HFD feeding in mice [52]. Taken together, our data show that disturbed calcium regulation may contribute to the observed diastolic dysfunction.

5. Conclusion In conclusion, 20 weeks of HFD feeding led to cardiac dysfunction in mice as indicated by increased LV mass and reduced diastolic function. HFD feeding induced mitochondrial adaptations promoting the utilization of FA. However, HFD-induced cardiac dysfunction was not caused by abnormalities in cardiac energetics, but rather was associated with lipotoxicity-induced myocardial oxidative stress, fibrosis, and disturbed calcium homeostasis. Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.bbalip.2014.07.016.

Acknowledgements We thank Leonie Niesen and David Veraart for their assistance in animal handling, Albert Gerding for the assistance with cardiac acylcarnitine quantification, Will Coumans for the assistance with biochemical determination of intramyocellular lipids, and Abdallah Motaal for the reconstruction of high temporal resolution cine MRI data.

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