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Am J Physiol Endocrinol Metab 295: E420-E427, 2008. First published June 3, 2008; doi:10.1152/ajpendo.90329.2008
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Cardioselective dominant-negative thyroid hormone receptor ({Delta}337T) modulates myocardial metabolism and contractile efficiency

Outi M. Hyyti,1,2,3 Aaron K. Olson,1,3 Ming Ge,1,3 Xue-Han Ning,1,3 Norman E. Buroker,1,3 Youngran Chung,4 Thomas Jue,4 and Michael A. Portman1,3

1Division of Cardiology, Department of Pediatrics and 2Department of Radiology, University of Washington; 3Children's Hospital and Regional Medical Center, Seattle, Washington; and 4Department of Biological Chemistry, University of California, Davis, California

Submitted 2 April 2008 ; accepted in final form 27 May 2008


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Dominant-negative thyroid hormone receptors (TRs) show elevated expression relative to ligand-binding TRs during cardiac hypertrophy. We tested the hypothesis that overexpression of a dominant-negative TR alters cardiac metabolism and contractile efficiency (CE). We used mice expressing the cardioselective dominant-negative TRβ1 mutation {Delta}337T. Isolated working {Delta}337T hearts and nontransgenic control (Con) hearts were perfused with 13C-labeled free fatty acids (FFA), acetoacetate (ACAC), lactate, and glucose at physiological concentrations for 30 min. 13C NMR spectroscopy and isotopomer analyses were used to determine substrate flux and fractional contributions (Fc) of acetyl-CoA to the citric acid cycle (CAC). {Delta}337T hearts exhibited rate depression but higher developed pressure and CE, defined as work per oxygen consumption (MVO2). Unlabeled substrate Fc from endogenous sources was higher in {Delta}337T, but ACAC Fc was lower. Fluxes through CAC, lactate, ACAC, and FFA were reduced in {Delta}337T. CE and Fc differences were reversed by pacing {Delta}337T to Con rates, accompanied by an increase in FFA Fc. {Delta}337T hearts lacked the ability to increase MVO2. Decreases in protein expression for glucose transporter-4 and hexokinase-2 and increases in pyruvate dehydrogenase kinase-2 and -4 suggest that these hearts are unable to increase carbohydrate oxidation in response to stress. These data show that {Delta}337T alters the metabolic phenotype in murine heart by reducing substrate flux for multiple pathways. Some of these changes are heart rate dependent, indicating that the substrate shift may represent an accommodation to altered contractile protein kinetics, which can be disrupted by pacing stress.

glucose metabolism; free fatty acids

DISRUPTION OF THYROID HORMONE HOMEOSTASIS plays an important role in the pathogenesis of heart failure (4, 19, 28, 32). Low levels of circulating thyroid hormone occur during development of uncompensated or pathological hypertrophy. In some patients, triiodothyronine infusion reverses pathological cardiac remodeling (31). Additionally, cardiac myocytes from failing hearts exhibit shifts or deficiencies in thyroid hormone receptor (TR) expression (19–21). Particularly, failing or myopathic aging hearts show a reduction in ligand-binding TR{alpha}1 relative to TR{alpha}2, a product of alternative splicing, which lacks the ligand binding domain. Recent work demonstrates that TR{alpha}1 or TR{alpha}2 delivered to myocardium by adeno-associated virus prompts functional improvement in murine hearts with pressure overload-induced hypertrophy (2). The positive effect is linked to expression of sarco(endo)plasmic reticulum Ca2+-ATPase (SERCA), a determinant of contractile phenotype.

Numerous studies have focused on thyroid receptor regulation of contractile proteins, such as the myosin ATPase (28), or SERCA (2, 33). More recently, several investigators alternatively postulated that thyroid modifies cardiac function through control of cardiac substrate metabolism and ATP synthesis (1, 14, 29). Few prior studies have directly evaluated thyroid receptor control of these processes. Because the rates of activity for these ATPases determine contractile function and can influence energy expenditure in the intact heart, and ATP synthesis responds to energy demands, the hypothesis invoking thyroid receptor-mediated metabolic control of cardiac function remains difficult to evaluate.

Although studies in hypothyroid or hyperthyroid models indicate that this hormone modulates cardiac energy metabolism (14, 15, 30), the inability to effectively dissect out thyroid receptor-mediated regulation of metabolism from posttranscriptional thyroid mechanisms has hampered study in this area. Esaki et al. (5) attempted to evaluate thyroid receptor control of heart substrate metabolism by studying mice in vivo with non-organ-specific "knockin" (PV) mutations of the thyroid {alpha}- and β-receptors. Although those authors ostensibly evaluated glucose utilization in these mice, their study only determined cardiac uptake with radioisotope-labeled 2-deoxyglucose, with no analyses for other major substrates such as lactate, fatty acids, or ketones. The heterozygous TR{alpha} PV mutation showed a reduction in glucose uptake along with a decrease in heart size and mean arterial pressure. The homozygous TRβ PV mutant showed an increase in glucose uptake without change in heart size or mean arterial pressure, suggesting a modification unrelated to myocardial hypertrophy. These models are somewhat complex, because circulating thyroid hormone levels are elevated particularly in the TRβ mutation, and might be responsible for observed differences in glucose uptake. Thus the only study to date using a transgenic approach to modify thyroid receptor function and control of cardiac substrate metabolism provided suggestive but limited data.

Using an alternative approach, we tested the hypothesis that overexpression of a non-ligand-binding dominant-negative thyroid receptor alters cardiac metabolism and contractile efficiency (CE). We employed an isolated working heart model from transgenic mice expressing the cardioselective {Delta}337T thyroid receptor mutation. The {Delta}337T mutation occurs naturally in some humans with resistance to thyroid hormone syndrome and fulfills the requirements as a dominant-negative nonbinding TR (28). These mice exhibit a euthyroid general phenotype, while displaying specific hypothyroidism of the heart. The substrate utilization profile was determined with 13C substrate labeling and isotopomer analyses with protein analyses for key enzymes regulating the metabolic pathways in these isolated, perfused working hearts. The robust NMR method used in these studies allows analyses of fractional contribution (Fc) of acetyl-CoA to the citric acid cycle (CAC) from multiple substrates supplied to the heart simultaneously in physiological concentrations (14). These methods allowed analyses of the metabolic phenotype caused by a dominant-negative TR.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animals. Two groups of mice were used in this study: 1) mice expressing a cardioselective dominant-negative TRβ1 mutation, {Delta}337T (3, 28) (age 4–6 mo), and 2) age-matched male nontransgenic littermates as control. The {Delta}337T human mutation is inserted into the mouse genome and linked to the {alpha}-myosin promoter for cardiac selectivity. It does not bind triiodothyronine and inhibits DNA binding for all thyroid receptors in a dominant-negative manner. All animal procedures were in accordance with guidelines of Children's Hospital and Regional Medical Center and were approved by the University of Washington Animal Care Committee. The investigation conformed with the Guide for the Care and Use of Laboratory Animals published by the National Institutes of Health (NIH Pub. No. 85-23, revised 1996).

Isolated heart preparation. Mice were anesthetized with pentobarbital sodium (75 mg/kg ip) and heparinized (700 U/kg ip). The heart was rapidly excised and submerged in ice-cold physiological salt solution (PSS), pH 7.4, containing (in mmol/l) 118.0 NaCl, 25.0 NaHCO3, 4.7 KCl, 1.23 MgSO4, 1.2 NaH2PO4, 5.5 D-glucose, and 1.2 CaCl2.

The aorta of a spontaneously beating heart was first cannulated in a standard Langendorff mode and perfused with PSS. After 15 min the heart function stabilized, and the mode was changed to working heart perfusion with the inflow tube inserted into the left atrium. The perfusate was switched to PSS containing the following 13C-labeled substrates in addition to unlabeled glucose (5.5 mmol/l): [1,3-13C]acetoacetic acid (ACAC, 0.17 mmol/l), L-[3-13C]lactic acid (LAC, 1.2 mmol/l), and U-13C-labeled long-chain mixed free fatty acids (FFA, 0.35 mmol/l) bound to 0.75% (wt/vol) delipidated bovine serum albumin reconstituted with deionized water. The FFA mixture contains predominantly saturated and unsaturated fatty acids ranging from 14 to 22 carbons in length, with palmitic and linoleic acids as the most prominent. Details regarding this isotope preparation and labeling strategy were published previously (22).

Separate groups of hearts were perfused with PSS without unlabeled glucose and supplemented with [1-13C]glucose (5.5 mmol/l), [1,3-13C]ACAC (0.17 mmol/l), U-13C-labeled long-chain mixed FFA (0.35 mmol/l), and unlabeled LAC (1.2 mmol/l).

Perfusates were equilibrated with 95% O2-5% CO2 at 37°C and passed twice through filters with 3.0-µm pore size. Perfusion pressure was maintained at 70 mmHg with Langendorff mode. Preload was 10 mmHg, and afterload was 50 mmHg with working mode. The entire perfusion system was jacketed and maintained at 37°C.

An SPR-PV-Catheter (SPR-869 or -839, Millar Pressure-Volume Systems) was inserted into the left ventricle through the apex for continuous measurement of left ventricular pressure (LVP). Recordable parameters from the left ventricle in addition to LVP (mmHg) were heart rate (beats/min), rates of left ventricular contraction and relaxation (±dP/dt; mmHg/s), and left ventricular volume (µl). Calculated parameters were stroke volume, cardiac output, work, power, CE (power/oxygen consumption), pressure-rate product (PRP), Vmax, and P-V loop, etc., although only some of these are presented here.

The pulmonary artery was cannulated to enable collection of coronary flow, which with aortic flow was measured with a flowmeter (T403; Transonic Systems, Ithaca, NY). The caudal vena cava, cranial vena cava, and azygous vein were ligated. Pacing leads were attached to the right atrial appendage.

To characterize cardiac function, left ventricular developed pressure (LVDP) was defined as peak systolic pressure minus end-diastolic pressure. Myocardial oxygen consumption (MVO2) was calculated as MVO2 = CF x [(PaO2 – PvO2) x (c/760)] x dw, where CF is coronary flow (ml·min–1g wet tissue–1), (PaO2 – PvO2) is the difference in the partial pressure of oxygen (PO2, mmHg) between perfusate and coronary effluent, c is the Bunsen solubility coefficient of O2 in perfusate at 37°C (22.7 µl O2·atm–1·ml–1), and dw is a previously determined conversion factor from heart wet weight to dry weight (1/0.18). PO2, PCO2, and pH were determined with a ABL800 blood gas analyzer (Radiometer, Copenhagen, Denmark).

Experimental protocol. Hearts were divided into three groups: control hearts at rest (n = 8), {Delta}337T hearts at rest ({Delta}337T, n = 8), and {Delta}337T hearts with pacing ({Delta}337T-P, n = 6). All hearts were perfused first for a 15-min equilibration period in Langendorff mode, after which working heart mode was established. Functional parameters were recorded continuously with the Millar Pressure-Volume Systems, and outflow samples for blood gas analyses were taken every 10 min. Baseline recording was taken 15 min into working heart mode. Hearts with developed pressures <80 mmHg or >140 mmHg were eliminated and not subjected to further experiments or protocols. After 15 min of baseline recording, {Delta}337T-P hearts were atrial paced at 330 beats/min. After 30 min of 13C-labeled substrate perfusion nonventricular tissue was removed, and hearts were immediately freeze clamped with copper tongs that had been chilled in liquid nitrogen.

Extraction. Briefly, freeze-clamped hearts were ground into fine powder under liquid nitrogen, extracted with 0.6 M perchloric acid, and neutralized with cold KOH to pH 7.4. The final supernatant was lyophilized overnight at –50°C for later NMR analysis. Separate groups of hearts were used for glycogen analyses of control and {Delta}337T hearts (see below).

13C NMR and isotopomer analyses. Substrate metabolism was determined with 13C-labeled substrates and glutamate isotopomer analyses in conjunction with NMR detection. Computer analyses of the labeling patterns provide Fc of acetyl-CoA to the CAC from up to three differentially labeled substrates and the unlabeled component (16–18, 24). The unlabeled and anaplerotic component can also be determined through algebraic computations in the software.

To perform these NMR detection experiments, lyophilized heart extracts were dissolved in 99.8% 2H2O for NMR spectral acquisition. Decoupled 13C NMR spectra of the samples were acquired on Bruker DMX 750-MHz and 500-MHz spectrometers with a 45° pulse and a 4-s recycle delay. Free-induction decays were baseline corrected, zero-filled, and Fourier transformed. All of the labeled carbon resonances (C1–C5) of glutamate were integrated with the Lorentzian peak fitting subroutine in the acquisition program (NUTS; Acorn NMR, Livermore, CA). The individual integral values were used as starting parameters for the CAC analysis-fitting algorithm tcaCALC, kindly provided by Dr. C. R. Malloy and Dr. F. M. Jeffrey (University of Texas Southwestern, Dallas, TX). This algorithm provided the Fc for each substrate to the acetyl-CoA pool entering CAC.

The absolute flux for the CAC and oxidative flux for individual substrates were calculated from MVO2 and the stoichiometric relationships between oxygen consumption and citrate formation from the various substrates as described by Jeffrey et al. (18). The calculated value accounts for changes in oxidative rates, as well as the anaplerotic contribution to the CAC. Briefly, MVO2/CACflux = FcFFARFFA + FcLACRLAC + FcACACRACAC + FcendRend + yRa, where the Fcs are fractional contributions for each substrate determined by isotopomer analysis and R is an assumed respiratory quotient [RFFA = 2.8, RLAC = 3, RACAC = 2, and Rend (end = endogenous substrates) = 2.9]. yRa represents the anaplerotic component. The calculated CACflux was normalized for each substrate by dividing the total CACflux by the number of acetyl-CoA esters yielded per molecule of that substrate (FFA = 8.5, LAC = 1, ACAC = 2) and multiplying with the corresponding Fc.

Glycogen. Samples for glycogen were taken from {Delta}337T mice and littermate controls (n = 7 each). Heart tissue for glycogen analyses was extracted at the protocol point before perfusion in order to determine the glycogen pool available at baseline in these experiments. Perchloric acid extracts of muscle were assayed for glycogen by the amyloglucosidase method (27).

Immunoblotting. Fifty micrograms of total protein extracts from mouse heart tissue was electrophoresed along with one lane of molecular weight size markers (Chemichrome Western control, Sigma) in a 4.5% stacking gel and a 7.5%, 10%, or 12% running SDS-polyacrylamide gel depending on the molecular weight of the protein of interest. The gels were then electroblotted onto polyvinylidene fluoride plus membranes. Western blots were prepared with standard techniques. The primary antibodies used in the study are glucose transporter (GLUT)-4 (SC-7938), GAPDH (SC-25778), hexokinase (HK)-2 (SC-6521), and succinate dehydrogenase subunit-{alpha} (SDHA) (SC-27992) obtained from Santa Cruz Biotechnology. The primary antibodies liver (L)- and muscle (M)-carnitine palmitoyltransferase (CPT) I and pyruvate dehydrogenase kinase (PDK)-2 and PDK-4 were obtained as personal gifts from Gebre Woldegiorgis (Oregon Health Science University, Beaverton, OR) and Robert Harris (Indiana University School of Medicine, Indianapolis, IN), respectively. The GAPDH was used to verify protein lane loadings.

Statistical analyses. Reported values are means ± SE in Figs. 1–3, 5, and 6. Data were analyzed with repeated-measures analysis of variance (ANOVA) within groups and single-factor ANOVA across groups (StatView 4.5, Abacus Concepts, Berkeley, CA), as well as Fisher's test and unpaired t-tests when appropriate. Paired t-test and Mann-Whitney were also used when appropriate for glycogen levels as described in RESULTS. The criterion for significance was P < 0.05 for all comparisons.


Figure 1
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  		Fig. 1. Cardiac functional parameters during 13C-labeled substrate perfusion. Developed pressure was significantly higher in {Delta}337T hearts vs. control (Con) hearts, pacing causing a decrease. Heart rate was lower in {Delta}337T hearts. Paced {Delta}337T ({Delta}337T-P) hearts were paced to control levels, which caused a decrease in maximum rate of ventricular contraction (+dP/dtmax), although the maximum rate of ventricular relaxation (–dP/dt) was lower in both transgenic groups. Values are means ± SE. *P < 0.05 vs. Con, {dagger}P < 0.05 vs. {Delta}337T.

 

Figure 3
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  		Fig. 3. Fractional contributions of acetyl-CoA from substrates to the citric acid cycle (CAC). Values are means ± SE. *P < 0.05 vs. Con, {dagger}P < 0.05 vs. {Delta}337T. ACAC, acetoacetate; LAC, lactate; FFA, free fatty acids; YS, anaplerotic component.

 

Figure 5
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  		Fig. 5. Calculated flux rates (µmol·g–1·min–1) for the total CAC and each individual substrate. Values are means ± SE. *P < 0.05 vs. Con. LAC, lactate.

 

Figure 6
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  		Fig. 6. Protein content relative to GAPDH for enzymes involved in substrate metabolism. Values are means ± SE. *P < 0.05 vs. Con. L-CPT I, liver-carnitine palmitoyltransferase I; M-CPT I, muscle-carnitine palmitoyltransferase I; PDK, pyruvate dehydrogenase kinase; HK, hexokinase; GLUT, glucose transporter; SDHA, succinate dehydrogenase subunit-{alpha}.

 

   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Cardiac function and oxygen consumption. Heart weights for the {Delta}337T and control mice were 119.0 ± 8.12 and 118.8 ± 3.37 mg, respectively; body weights 25.6 ± 2.8 and 24.7 ± 1.5 g; and heart/body weight (x10–3) 0.46 ± 0.05 and 0.49 ± 0.02, showing no significant differences between groups. Functional parameters (Fig. 1) and MVO2, power, CE (defined as cardiac power per unit of MVO2), and CF (Fig. 2) are reported for 10-min intervals during 13C-labeled substrate perfusion. These parameters did not deviate significantly over the time of the protocol, thereby demonstrating the required functional and metabolic steady state for the tcaCALC algorithms. Developed pressure was significantly higher in the {Delta}337T group compared with control, although pacing caused it to decrease (Fig. 1). Pacing the heart to rates similar to control also caused a significant decrease in +dP/dtmax (Fig. 1). However, –dP/dtmax, representing ventricular relaxation speed, was significantly depressed in both {Delta}337T and {Delta}337T-P groups (Fig. 1), consistent with previous literature in hypothyroid models (7). Also, spontaneous heart rate (Fig. 1) and MVO2 (Fig. 2) were lower in the {Delta}337T group throughout the protocol. Interestingly, the lower MVO2 was independent of heart rate as shown by the pacing experiments, where higher heart rate did not increase MVO2 during the experiment. Power and CE were significantly higher in {Delta}337T hearts, although stress induced by pacing brought them to control levels (Fig. 2).


Figure 2
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  		Fig. 2. Myocardial oxygen consumption (MVO2) was reduced in {Delta}337T hearts independent of heart rate compared with Con hearts, as was coronary flow. Power and cardiac efficiency (power/MVO2) were higher in {Delta}337T hearts, although stress induced by pacing brought them to control levels. Values are means ± SE. *P < 0.05 vs. Con, {dagger}P < 0.05 vs. {Delta}337T.

 
Substrate selection. Acetyl-CoA enters the CAC either through acyl-CoA synthase or pyruvate dehydrogenase (PDH). Fc of acetyl-CoA into CAC for each substrate during the 30-min steady-state working period were determined from glutamate peak areas with NMR and 13C isotopomer analysis (18) and are shown in Fig. 3. Figure 4 demonstrates the signal to noise and resolution for a spectrum obtained from mouse heart extract. It also identifies each component of the spectrum and the derivation of the peaks from their original labeled substrate (see Fig. 4 legend).


Figure 4
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  		Fig. 4. Representative proton-decoupled 13C NMR spectra from a {Delta}337T-P heart showing C4 of glutamate. A: spectrum obtained from hearts perfused with [1-13C]glucose with [1,3-13C]ACAC, U-13C-labeled long-chain mixed FFA, and unlabeled lactate. B: spectra obtained with our standard labeling strategy: [1,3-13C]ACAC, L-[3-13C]lactic acid, U-13C-labeled long-chain mixed FFA, and unlabeled glucose. The singlet (S) and doublet (D34) peaks are derived from labeling lactate at C3 or glucose at C1. The quartet (Q) and the D45 are generated from labeling FFA. The ACAC labels the C5 glutamate (not shown) and does not label C4. Comparison of A with B shows that labeled glucose generates a much smaller S and D34 than does labeled lactate. Labeling for these peaks increases substantially when L-[3-13C]lactic acid and unlabeled glucose are used (B).

 
Fc for unlabeled substrates was significantly higher in {Delta}337T than in control hearts. Fc for ACAC was lower in the {Delta}337T group at 0.24 ± 0.01 compared with control (0.32 ± 0.02) (Fig. 3), thereby representing a 25% reduction. However, pacing the {Delta}337T hearts ({Delta}337T-P) to a heart rate approximating the rate in control hearts decreased unlabeled fraction to control levels. FFA contribution was substantially higher in paced {Delta}337T hearts (0.47 ± 0.07) than in control (0.39 ± 0.02) and nonpaced (0.38 ± 0.02) hearts. Having shown that substantial changes in Fc for unlabeled substrates occurred in the transgenic hearts and then reversed with pacing, we attempted to identify the source. Glucose represented the most likely candidate, although our previous studies in rats with similar substrate perfusion showed that glucose was a minor contributor (22). In Fig. 4, we illustrate the difference between using labeled glucose and lactate in these experiments. In Fig. 4B, the labeling in carbon-4 glutamate by L-[3-13C]lactic acid is shown by D34 doublet and singlet (S). These peaks are dramatically decreased in area, when [1-13C]glucose (Fig. 4A) was used instead, indicating a marked relative decrease in labeling through [3-13C]pyruvate. In all transgenic hearts (n = 4; 2 paced) glucose contributed <4% acetyl-CoA to the CAC with Fc 0.01 ± 0.01 for nonpaced {Delta}337T hearts and 0.04 ± 0.01 for paced hearts. These values are not substantially different from the unlabeled Fc in the control and paced hearts, thereby confirming that glucose contributes only a minor portion of oxidative substrate under these conditions. The data also indicate that the elevated Fc for unlabeled fraction observed in {Delta}337T hearts at baseline heart rate is not caused by elevated glucose oxidation relative to the paced hearts and that ~10–15% of acetyl-CoA is derived from endogenous unlabeled sources.

Substrate flux. The total CACflux and individual substrate flux rates are presented in Fig. 5. Lactate (LAC) flux indicates directional flux from lactate toward acetyl-CoA and entry into CAC. As expected, the {Delta}337T mutation reduced the total CACflux at rest and during stress. Accordingly, individual flux rates for ACACflux and LACflux were significantly lower in {Delta}337T hearts compared with control hearts independent of heart rate. However, FFAflux was only significantly decreased in {Delta}337T hearts at rest. Pacing reversed the FFAflux change caused by {Delta}337T but did not significantly alter flux for LAC and ACAC.

Glycogen. {Delta}337T hearts contained significantly greater amounts of glycogen than paired nontransgenic littermates: 5.14 ± 0.83 vs. 3.62 ± 1.02 µmol glucose derived/g wet wt (P = 0.03 by paired t-test and confirmed by Wilcoxon signed rank test).

Metabolic regulatory proteins. The changes in Fc of substrates into the CAC in the transgenic hearts prompted protein analyses for enzymes involved in these pathways. We analyzed protein concentration for several key enzymes involved in regulation of substrate oxidation and SDHA, a biomarker for mitochondrial biogenesis. Results for each analyzed protein appear in Fig. 6. An increase was noted in {Delta}337T hearts for both PDK-2 and PDK-4 and a marked decrease in HK-2 and GLUT-4, with no change in L-CPT I and M-CPT I or SDHA.


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Our study demonstrates that insertion of a cardioselective and dominant-negative thyroid receptor alters the metabolic phenotype in murine heart. The {Delta}337T hearts showed reductions in flux for multiple substrate oxidative pathways, indicating that the decrease in total CAC flux is not caused by deficits in a single pathway. As noted previously, limited prior data exist regarding thyroid receptor modulation of myocardial metabolic phenotype. Studies in the hypothyroid rat heart demonstrate important transcriptionally mediated regulation of cardiac metabolism by thyroid hormone (14). Experiments in isolated perfused heart from euthyroid or hypothyroid rats indicate that triiodothyronine also exerts rapid modulation of cardiac substrate metabolism including enhancement of fatty acid flux (14, 22). The rapidity of these latter actions implies that they do not occur through transcriptional processes controlled by thyroid receptor. Because alterations in circulating triiodothyronine and thyroxine modify tissue TR expression, full assessment of the role for these receptors in metabolic control in rat heart has remained difficult. Mice displaying tissue ubiquitous mutations in TR, obtained from humans with resistance to thyroid syndrome, also have abnormal circulating thyroid hormone levels (5). We avoided the complexities of other models employing noncardiac selective TR mutations and exhibiting substantial abnormalities in circulating thyroid hormone levels by using the systemic euthyroid cardioselective {Delta}337T mice.

Consistent with previous studies in systemic hypothyroid models, contractile energy efficiency is increased in {Delta}337T mice without substantial reduction in function. Contractile efficiency (CE) is the relationship of cardiac work to oxygen consumed and can be modified by several factors. Some data suggest that switching of myosin isoforms toward a slower and predominantly V3 phenotype through hypothyroidism or genetic manipulation raises cardiac contractile energy efficiency (11, 13, 25). This myosin shift in isolation induces bradycardia. The {Delta}337T mutation alters myosin isoform profile, producing a marked predominance of β-myosin heavy chain (MHC) (>70% total MHC) in {Delta}337T mice compared with wild type (< 10%), thereby confirming the presence of a slow MHC in these transgenic mice (28). Hypothyroidism and/or ubiquitous TR{alpha}1-knockout also cause bradycardia by decreasing expression for mRNAs coding for potassium channels involved in action potential repolarization, such as KV1.5, KV4.2, and KV4.3, and cyclic nucleotide-gated channels HCN2 and HCN4 (9). A genomewide molecular array analysis in {Delta}337T revealed that only KV4.2 among these genes showed a substantial (>25%) and significant (P < 0.01) decrease in expression for multiple sequences (3). Thus the data suggest that the bradycardia in {Delta}337T results from changes in regulation of multiple proteins at the transcriptional level, and that energy metabolism shifts to accommodate these changes.

In the present study, we considered that the energetic advantage for {Delta}337T hearts occurs through an alteration in substrate utilization, which could yield a higher free energy of ATP hydrolysis ({Delta}GATP). The relationship of substrate preference to cardiac efficiency has been previously established in isolated working mouse hearts (12). In prior studies, raising palmitate concentration in perfusate decreased the ratio of glucose to fatty acid oxidation and concomitantly decreased cardiac efficiency (12). Those results can be explained in part by the greater ATP-to-O ratios shown for glucose oxidation than for fatty acid oxidation (26). However, the major difference in substrate utilization between {Delta}337T hearts operating at intrinsic heart rate and wild-type hearts is the increase in Fc through unlabeled substrates, which occurs along with a small reduction in Fc for ketones in the form of ACAC. These substrate preference differences along with the energetic advantage can be abrogated by increasing heart rate with pacing. This study was not designed to directly identify various sources of unlabeled substrate. However, we eliminated glucose as a source with our labeled glucose perfusions by showing that this substrate's contribution is similar between {Delta}337T mice at intrinsic heart rate and with pacing to heart rates comparable to control mice. Additionally glucose Fc obtained from mice perfused with 13C-labeled glucose approximated the unlabeled fraction in paced {Delta}337T hearts perfused with unlabeled glucose. Accordingly, the remaining two candidates for contribution as unlabeled fraction in the nonpaced {Delta}337T hearts are the endogenous substrates glycogen and triglycerides (10, 34, 35). Addition of ACAC to the perfusate for isolated working rat hearts enhances glycogen content and glucose incorporation into glycogen, suggesting that preferential oxidation of this ketone inhibits glycogenolysis (34). Therefore, our observed reductions in ACAC Fc and flux in {Delta}337T hearts and the concomitant increase in cardiac efficiency are all consistent with a reciprocal increase in glucose 6-phosphate derived from glycogen, as the principal unlabeled substrate. Oxidation of an alternative source of unlabeled substrate, the endogenous triglyceride pool, would be expected to decrease CE because of the lower P-to-O ratio inherent with fatty acids. Although endogenous triglycerides can contribute a substantial portion of oxidative substrate under specific conditions (35), they only provide a minor portion compared with glycogen in the presence of multiple substrates such as those supplied in our perfusate (34). Our analyses using the amyloglucosidase assay revealed that heart extraction procedures do not deplete glycogen. Furthermore, glycogen stores are significantly elevated in the {Delta}337T hearts at baseline in these experiments. These data strongly support the contention that glycogen is the source of the unlabeled acetyl-CoA entering the CAC in the transgenic hearts under intrinsic heart rate conditions.

Previous studies have shown inherently lower heart rates in {Delta}337T mice than in nontransgenic control mice (28). Accordingly, we paced {Delta}337T hearts to comparable control levels in order to obviate any differences in energy metabolism induced by the lower heart rate alone. In fact, pacing did eliminate the cardiac efficiency advantage. The nontransgenic working mouse heart demonstrates limited heart rate reserve (23) and shows a marked decrease in function, cardiac output, and oxygen consumption during pacing. In contrast, the {Delta}337T mutant baseline heart rate is ~60% of control and tolerates robust increases induced by pacing. Although pacing in {Delta}337T hearts achieves an increase in cardiac output, these hearts were unable to increase overall work, indexed by left ventricular power. The heart rate elevation in these transgenic hearts prompted an increase in fatty acid acetyl-CoA contribution to the CAC and a reversal of the inhibition of fatty acid flux. However, the {Delta}337T hearts exhibited an inability to increase oxygen consumption, suggesting a limitation in carbohydrate oxidation capacity and involving pathways leading to or including the PDH complex.

The shifts in flux and Fc during pacing are consistent with the proteomic findings for the enzymes regulating pivotal junctures in carbohydrate oxidation. As noted, {Delta}337T shows a decrease in overall GLUT-4 content. Translocation of GLUT-4 into the plasma membrane from its intracellular storage promotes glucose transport and oxidation while inhibiting fatty acid oxidation. Therefore, it is tempting to suggest that the reduced overall GLUT-4 content bears some responsibility for the apparent reduction in carbohydrate oxidation capacity noted in {Delta}337T hearts. Recent data indicate that a GLUT-4 deficit alone does not limit stress-induced glucose oxidation (6). However, concomitant changes in HK-2, responsible for glucose phosphorylation, the step subsequent to transport, can modify maximal glucose oxidation capacity. Partial deletion of the HK-2 gene (HK+/–) that results in a 50% reduction in HK-2 activity in heart does not alter carbohydrate metabolism at rest but does prevent the normal increase in myocardial glucose uptake during exercise (6). We found a similar magnitude of reduction in HK-2 protein content in {Delta}337T mice, suggesting that a glucose phosphorylation deficit in conjunction with a reduction in transport capacity was partly responsible for the inability of these hearts to increase oxygen consumption with pacing.

A surge in lactate oxidation generally accounts for a major portion of increased cardiac oxygen consumption during elevations in work, when this substrate is proffered (22). The finding that lactate oxidation did not respond to pacing in these {Delta}337T hearts also suggests that carbohydrate oxidation is limited or inhibited along the glycolysis pathway beyond formation of pyruvate. The PDKs regulate both glucose and lactate oxidation by inhibiting PDH complex through enzyme phosphorylation. We observed elevation in content for two distinct PDK isoforms in the {Delta}337T hearts, supporting the premise that thyroid receptors regulate cardiac metabolism via PDH. To our knowledge, the response of PDK-2 and PDK-4 to thyroid hormone in mice has not been previously evaluated. In rat heart, the PDK-4 response has been highly variable, with separate reports of increased protein accumulation in both hyperthyroid (36) and hypothyroid (14) states. Because PDK-4 mRNA is not elevated in these {Delta}337T hearts (14), the protein response likely represents an adaptation to the overall metabolic phenotype caused by the {Delta}337T mutation.

In summary, these data show that insertion of a dominant-negative TR alters metabolic phenotype in heart. Elevations in cardiac efficiency can be eliminated by raising heart rate, which also causes a shift in substrate preference to FFA. The results suggest that substrate metabolism plays a role in the pathogenesis of heart failure associated with disturbances in thyroid hormone homeostasis and corresponding increases in relative expression of dominant-negative thyroid receptors. TR{alpha}2 is the predominant dominant-negative isoform, appearing in heart during noncompensated hypertrophy (19). Although this isoform differs in sequence from the {Delta}337T TRβ1 mutation, they are functionally similar. Metabolic modulation of heart through thyroid receptor manipulation, such as an increase in ligand-binding isoforms, or repression of the dominant-negatives, represents a potential area for research in the future.

These modifications in substrate utilization as well as limitations in metabolic response to stress may play a role in pathogenesis of the heart.


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This study was funded by National Heart, Lung, and Blood Institute Grant R01-HL-60666 to M. A. Portman and a fellowship award from the American Heart Association to O. M. Hyyti.


   FOOTNOTES
 

Address for reprint requests and other correspondence: M. A. Portman, Children's Hospital and Regional Medical Center MSW 4841, 4800 Sand Point Way NE, Seattle, WA, 98105 (e-mail: michael.portman{at}seattlechildrens.org)

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


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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 

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