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TRANSLATIONAL PHYSIOLOGY
is detrimental to cardiac recovery after ischemia
1Cardiovascular Research Group, Department of Pediatrics and Pharmacology, University of Alberta, Edmonton, Alberta, Canada; and 2Center for Cardiovascular Research, Department of Internal Medicine, Washington University School of Medicine, St. Louis, Missouri
Submitted 23 March 2005 ; accepted in final form 24 August 2005
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ABSTRACT |
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regulates transcription of several FAO enzymes in the heart, we examined the response of mice with cardiac-restricted overexpression of PPAR (MHC-PPAR) or whole body PPAR deletion including the heart (PPAR–/–) to myocardial ischemia-reperfusion injury. Isolated working hearts from MHC-PPAR and nontransgenic (NTG) littermates were subjected to no-flow global ischemia followed by reperfusion. MHC-PPAR hearts had significantly higher FAO rates during aerobic and postischemic reperfusion (aerobic 1,479 ± 171 vs. 699 ± 117, reperfusion 1,062 ± 214 vs. 601 ± 70 nmol·g dry wt–1·min–1; P < 0.05) and significantly lower glucose oxidation rates compared with NTG hearts (aerobic 225 ± 36 vs. 1,563 ± 165, reperfusion 402 ± 54 vs. 1,758 ± 165 nmol·g dry wt–1·min–1; P < 0.05). In hearts from PPAR–/– mice, FAO was significantly lower during aerobic and reperfusion (aerobic 235 ± 36 vs. 442 ± 75, reperfusion 205 ± 25 vs. 346 ± 38 nmol·g dry wt–1·min–1; P < 0.05) whereas glucose oxidation was significantly higher compared with wild-type (WT) hearts (aerobic 2,491 ± 631 vs. 901 ± 119, reperfusion 2,690 ± 562 vs. 1,315 ± 172 nmol·g dry wt–1·min–1; P < 0.05). Increased FAO rates in MHC-PPAR hearts were associated with a markedly lower recovery of cardiac power (45 ± 9% vs. 71 ± 6% of preischemic levels in NTG hearts; P < 0.05). In contrast, the percent recovery of cardiac power of PPAR–/– hearts was not significantly different from that of WT hearts (80 ± 8% vs. 75 ± 9%). This study demonstrates that chronic activation of PPAR is detrimental to the cardiac recovery during reperfusion after ischemia.
myocardial ischemia; reperfusion; peroxisome proliferator-activated receptor ; fatty acid oxidation; acetyl-CoA carboxylase
Fatty acid metabolism in the heart is regulated by substrate availability and allosteric modulation of enzyme activity at the level of transcription of enzymes involved in cellular uptake and oxidation of fatty acids. AMP-activated protein kinase (AMPK) is one enzyme that controls this process, secondary to phosphorylating and inhibiting acetyl-CoA carboxylase (ACC) (21). ACC produces malonyl CoA, which is a potent inhibitor of mitochondrial fatty acid uptake. After ischemia, FAO rates are high, in part because of AMPK activation, resulting in an increase in ACC phosphorylation and a decrease in malonyl CoA control of FAO (11, 21). Another important site of FAO control is peroxisome proliferator-activated receptor (PPAR). PPAR, a member of the nuclear receptor superfamily, is activated by lipid ligands including long-chain fatty acids. Recently we demonstrated (9) that mice lacking PPAR (PPAR–/– mice) have profoundly diminished cardiac FAO and increased glucose utilization. The expression of PPAR and its target genes is increased in the diabetic heart with an accompanying increase in FAO (14). Further, cardiac-specific overexpression of PPAR in transgenic mice (MHC-PPAR mice) exhibits a cardiac phenotype characterized by a marked increase in fatty acid uptake and oxidation rates with a concomitant decrease in glucose uptake and oxidation rates, a profile similar to that of the diabetic heart (12, 14, 17). Despite this link between PPAR and FAO, the role of cardiac PPAR pathway in response to ischemia-reperfusion injury is still unclear. We hypothesize that PPAR-driven increases in FAO rates, as occurs in the diabetic heart, lead to enhanced susceptibility to ischemic injury. To test this hypothesis, MHC-PPAR and PPAR–/– mice were subjected to ischemia-reperfusion in isolated working heart perfusions.
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METHODS |
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mice (C57BL/6 x CBA/J F1) were produced with FLAG epitope-tagged PPAR cDNA with cardiac -myosin heavy chain (MHC) promoter as described previously (13). Of the three different lines described previously, we used a low-expressing MHC-PPAR line (line 404-4) because this level of overexpression exhibited a metabolic phenotype without any differences in cardiac function (14). Production of PPAR-null (PPAR–/–) and wild-type (WT) control (PPAR+/+) mice of the SV129 strain was described previously (23, 24). Isolated Mouse Working Heart Perfusion
Isolated mouse working heart perfusions were based on a previously described procedure (7, 9). Adult male mice (11–12 wk old with 25–30 g body wt) fed ad libitum were heparinized (100 U ip) 10 min before anesthesia. Animals were then deeply anesthetized with pentobarbital sodium (5–10 mg ip), and hearts were excised and placed in a ice-cold Krebs-Henseleit bicarbonate (KHB) solution (in mM: 118 NaCl, 25 NaHCO3, 4.7 KCl, 1.2 KH2PO4, 2.5 CaCl2, and 5.0 glucose, pH 7.4). Hearts were cannulated first via the aorta with a 15-gauge plastic cannula and perfused retrogradely by the Langendorff method with KHB solution (gassed with 95% O2-5% CO2). During this time, the left atrium was cannulated through a pulmonary opening with a 16-gauge steel cannula. Once the cannulation for working heart mode was complete, the Langendorff line was closed and the left atrial and aortic lines were opened and perfused with KHB solution containing 5.0 mM glucose, 1.2 mM palmitate bound to 3% fatty acid-free BSA and 600 pM insulin with a preload pressure of 11.5 mmHg and an afterload pressure of 50 mmHg.
The perfusion protocol included a 30-min aerobic perfusion of spontaneously beating hearts followed by an 18-min global ischemia and then a 40-min reperfusion with oxygenated buffer solution. The spontaneous recovery of heart function was monitored. To determine palmitate oxidation, glucose oxidation, and glycolytic rates, trace amounts of [9,10-3H]palmitate (0.1 µCi/ml) and [U-14C]glucose (0.1 µCi/ml) or [5-3H]glucose (0.1 µCi/ml) were used, respectively. Functional measurements were acquired for 10 s every 10 min with the MP100 system from AcqKnowledge (BioPac Systems, Santa Barbara, CA). Cardiac output and aortic flows were measured with inline flow probes (Transonic Systems, Ithaca, NY) placed in the left atrial line and the aortic afterload line, respectively. Coronary flow was calculated as the difference between cardiac output and aortic flows. Heart rate and pressures were measured with a pressure transducer (TSD104A, BioPac Systems) placed in the aortic line at the level of the heart. Cardiac work was calculated as the product of peak systolic pressure and cardiac output. Stroke volume was calculated as the cardiac output divided by the heart rate, whereas stroke work was calculated as the stroke volume times the peak systolic pressure. Cardiac power was calculated as the product of developed pressure and cardiac output. A conversion factor of 1.33 x 10–4 was used to convert cardiac power values from millimeters of mercury per milliliter to joules (7).
At the end of each perfusion, hearts were frozen immediately in liquid nitrogen and tissue homogenates were used for various biochemical measurements. Heart tissue was also used for determining dry-to-wet weight ratio.
Measurement of FAO, Glucose Oxidation, and Glycolysis
For palmitate oxidation [9,10-3H]palmitate was used as a tracer, and the by-product 3H2O that was produced during palmitate oxidation was separated with a water vapor exchange method. Briefly, 200 µl of perfusate was placed in a capless microfuge tube and kept inside a scintillation vial containing 500 µl of distilled water. The scintillation vial was closed tightly and kept in a 50°C oven for 24 h to facilitate vapor exchange between the perfusate and the distilled water. The vial was then kept in a 4°C cold room to allow the vapor condensation to occur for a subsequent 24 h. At the end of this period, microfuge tubes containing the perfusate were removed, making sure that all the water condensed on the outer surface was drained into the scintillation vial. Scintillation cocktail was added to the vials and counted. An equivalent volume of standard buffer containing a known amount of 3H2O was used each time to calculate vapor exchange efficiency. Palmitate oxidation rates were calculated from 3H2O production, taking into account the vapor exchange efficiency. This assay procedure has been standardized in our lab with commercially available standard 3H2O and found to be linear within the range of samples. The values were found to be comparable to those obtained by the chloroform-methanol extraction method that were described previously (7).
For glucose oxidation, [U-14C]glucose was used as a tracer. 14CO2 produced as a result of glucose oxidation was captured with a 1 M hyamine hydroxide trap as described previously (38). 14CO2 that was dissolved in the perfusate was also liberated with 9 N H2SO4 in a test tube sealed with hyamine hydroxide-soaked filter paper (to absorb 14CO2 that was liberated during acidification). Radioactivity from both sources was combined to determine the glucose oxidation rates.
For glycolysis, [5-3H]glucose tracer was used and the 3H2O that was produced at the triosephosphate isomerase and enolase steps of the glycolytic pathway was separated from the parent label with the method described above for palmitate oxidation rates. Glycolysis was performed in separate series of hearts for which [9,10-3H]palmitate tracer was not included in the perfusate buffer.
Biochemical Measurements
Detection and quantification of nucleotides were performed by extracting them from powdered tissue into 6% perchloric acid and measuring with a modified high-pressure liquid chromatography procedure, as described previously (30). Triglycerides were extracted from heart tissue according to the method of Folch et al. (15) and measured by an enzymatic method with the Wako L-Type TG H kit (Wako Pure Chemical Industries, Richmond, VA) as detailed previously (4, 9).
For Western blot analysis for phospho-AMPK (p-AMPK), phospho-ACC (p-ACC), AMPK, and ACC expression, the following antibodies were used: anti-p-AMPK (Thr172) and AMPK (Cell Signaling Technology, Beverly, MA); p-ACC (Ser79) and ACC (Upstate Technology, Lake Placid, NY); and goat anti-rabbit antibody (Santa Cruz Biotechnology, Santa Cruz, CA). Western blots were performed according to standard methods, using antibody dilutions as described previously (9). Results are expressed as a percentage of the optical density of a standard sample loaded on all membranes.
Statistical Analysis
Values are expressed as means ± SE. A Student's t-test or one-way ANOVA on ranks was performed to determine the statistical difference between groups. In some cases repeated-measures ANOVA was performed. P < 0.05 was considered significantly different.
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RESULTS |
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Mice
Glucose oxidation and glycolysis.
Figure 1, A and B, shows the steady-state glucose oxidation and glycolytic rates from hearts isolated from MHC-PPAR and nontransgenic (NTG) control mice. The substrate utilization rates were linear throughout the aerobic and reperfusion perfusion periods (data not shown). Both glycolysis (Fig. 1B) and glucose oxidation (Fig. 1A) rates were markedly lower compared with NTG hearts during preischemic aerobic perfusion. During reperfusion, although glucose oxidation and glycolytic rates recovered back to the preischemic values in both groups, the rates remained lower in MHC-PPAR mouse hearts.
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mouse hearts compared with NTG controls. During reperfusion after the global ischemia, palmitate oxidation recovered back to significantly higher rates compared with the NTG littermate group.
The relative contributions of glucose oxidation, glycolysis, and palmitate oxidation to the total production of ATP are shown in Fig. 1D. ATP production was calculated based on the stoichiometry that each mole of glucose produces 30 moles of ATP during complete oxidation and 2 moles of ATP during glycolysis and each mole of palmitate produces 105 moles of ATP during complete oxidation. Although the theoretical level of ATP does not account for any uncoupled oxidation that might have occurred, it gives a good perspective on the relative contribution of different substrates toward total ATP pools in the myocardium. In NTG control mouse hearts, during preischemic aerobic perfusions palmitate oxidation contributed 55% of total ATP (90.2 ± 15.0 µmol·min–1·g dry wt–1), with the rest of the ATP originating from glucose oxidation and glycolysis (74.4 ± 6.0 µmol·min–1·g dry wt–1). In MHC-PPAR hearts, >90% of ATP (237.7 ± 33.5 µmol·min–1·g dry wt–1) was derived from palmitate and <10% from glucose metabolism (15.2 ± 2 µmol·min–1·g dry wt–1). During reperfusion after global ischemia, the relative contributions of glucose and palmitate to total ATP production did not change in either NTG control or MHC-PPAR hearts compared with their preischemic levels (NTG: ATP from glucose 86.3 ± 6, from palmitate 77.5 ± 9.0; MHC-PPAR: ATP from glucose metabolism 19.8 ± 2, from palmitate oxidation 148.8 ± 60 µmol·min–1·g dry wt–1). When the cardiac metabolic efficiency was calculated as a ratio of total ATP produced from FAO and glucose oxidation over cardiac power, MHC-PPAR hearts exhibited a poor efficiency compared with that of NTG littermates during aerobic (MHC-PPAR 2.4 ± 0.5, NTG 1.4 ± 0.2 µmol·g dry wt–1·J–1) and postischemic (MHC-PPAR 6.0 ± 1.7, NTG 2.1 ± 0.3 µmol·g dry wt–1·J–1) periods.
Myocardial Substrate Utilization During Pre- and Postischemic Aerobic Perfusions in PPAR–/– Mice
As expected, the steady-state glucose oxidation rates were significantly higher in PPAR–/– hearts compared with those of WT controls. This increase in glucose oxidation was observed during both the pre- and postischemic aerobic perfusions (Fig. 2A). On the other hand, palmitate oxidation was significantly lower in PPAR–/– mice during pre- and postischemic perfusions compared with WT littermates (Fig. 2B). Calculated ATP production from glucose and palmitate oxidation indicates that the PPAR–/– mouse heart derives its energy mainly from glucose (Fig. 2C). The contribution of palmitate to total ATP during reperfusion remained low in PPAR–/– hearts (ATP from glucose 96.8 ± 20.2, from palmitate 26.4 ± 3.2 µmol·min–1·g dry wt–1). WT control hearts derived their energy equally from glucose and palmitate metabolism both during the pre- and postischemic perfusions, similar to the NTG control hearts in the MHC-PPAR study (during preischemia ATP from glucose 32.4 ± 4.3 and from palmitate 57.0 ± 9.7 and during postischemia ATP from glucose 47.3 ± 6.2 and from palmitate 44.6 ± 4.9 µmol·min–1·g dry wt–1). Metabolic efficiency in PPAR–/– mice was comparable to that in WT littermates both during preischemic (PPAR–/– 1.0 ± 0.3 and WT 1.1 ± 0.2 µmol·g dry wt–1·J–1) and postischemic (PPAR–/– 1.3 ± 0.2 and WT 1.7 ± 0.4 µmol·g dry wt–1·J–1) perfusions.
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and PPAR–/– Hearts After Ischemia-Reperfusion
The levels of triglycerides (TG) were not statistically different between the hearts of NTG and MHC-PPAR mice (36.8 ± 4.9 vs. 33.4 ± 5.2 µmol/g dry wt). However, the cardiac TG content was significantly decreased in PPAR–/– mice compared with WT (12.66 ± 0.73 vs. 27.8 ± 2.4 µmol/g dry wt; P < 0.05). AMPK is induced in response to metabolic stress. Previous studies have shown that AMPK could be a key player in the metabolic alterations during ischemia-reperfusion (22, 32, 36). Therefore, we measured phosphorylation of AMPK in the heart homogenates frozen at the end of reperfusion. Figure 3A shows that the ratio of p-AMPK to total AMPK (tot-AMPK) was greater, although not statistically significant, in cardiac tissues of MHC-PPAR compared with NTG (798 ± 63 vs. 701 ± 38) littermates. Ser79 phosphorylation of ACC, the downstream target of AMPK, was increased by 2.7-fold in MHC-PPAR hearts. The ratio of p-ACC to total ACC (tot-ACC) was 169.7 ± 29.5 in MHC-PPAR compared with 63.9 ± 10.9 in NTG (Fig. 3B, left; P < 0.05) hearts. In PPAR–/– hearts, p-AMPK/tot-AMPK was decreased by 57% compared with WT hearts (293 ± 32 vs. 513 ± 67; P < 0.05; Fig. 3A, right) without a significant change in percentage ratio of p-ACC to tot-ACC compared with that of WT littermates (51 ± 12 vs. 155 ± 56; P > 0.05; Fig. 3B).
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and in PPAR–/– Mice
Table 1 lists various parameters of cardiac function in MHC-PPAR and NTG control mouse hearts perfused at 50-mmHg afterload and 11.5-mmHg preload. The values are the averages of 10-, 20-, and 30-min measurements of aerobic perfusion. Under the aerobic perfusion conditions, there were no significant differences in heart rate, peak systolic pressure, developed pressure, cardiac output, coronary flow, and rate-pressure product (heart rate x peak systolic pressure) between MHC-PPAR and NTG littermates (Table 1). As summarized in Table 2, except for cardiac output and cardiac power, other hemodynamic parameters like peak systolic pressure, developed pressure, coronary flow, and rate-pressure product were not different in PPAR–/– mice compared with WT littermates during aerobic perfusion.
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Mice and PPAR–/– Mice
During 18 min of no-flow ischemia, the heart ceases to function. When reperfusion begins after ischemia, heart function starts to recover spontaneously from zero function. Except for coronary flow and developed pressure, recovery of heart rate, peak systolic pressure, cardiac output, and rate-pressure product was significantly lower in the MHC-PPAR group compared with NTG control hearts (Table 1). Recovery of cardiac power was markedly reduced compared with NTG controls during reperfusion after global ischemia (Fig. 4A). Because changes in heart rate could have contributed to the decrement in cardiac work, we calculated stroke work (g x m), which is the product of stroke volume and peak systolic pressure. Table 1 shows that the mean stroke work for 40 min of postischemic reperfusion was significantly lower in MHC-PPAR hearts. Although mean cardiac power recovered back to 71 ± 6% and stroke work to 75 ± 7% of preischemic performance in NTG controls, MHC-PPAR hearts recovered to 45 ± 9% and 52 ± 10% of cardiac power and stroke work, respectively, during reperfusion.
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–/– mice compared with WT controls (Table 2 and Fig. 4B; P < 0.05). The mean rate of recovery of stroke work during 40 min of reperfusion was not significantly different between the two groups (Table 2). Recovery of heart rate was 75 ± 9% of preischemic levels in WT controls, whereas recovery was 80 ± 8% of preischemic levels in PPAR–/– hearts (Table 2). Although not statistically significant, PPAR –/– mice exhibited a better recovery of other parameters like peak systolic pressure and coronary flow compared with age-matched WT controls. It is interesting to note that mean cardiac power over the 40-min reperfusion period was significantly greater in PPAR–/– hearts compared with MHC-PPAR hearts (95 ± 8 vs. 37 ± 8 J/min; P < 0.05, one-way ANOVA followed by Fisher's least significant difference multiple comparison). There is no significant difference between respective control groups (NTG littermates 71 ± 6 vs. WT littermates 63 ± 6 J/min). |
DISCUSSION |
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controls FAO by regulating the expression of genes involved in the uptake and oxidation of fatty acids. Despite this role of PPAR in the regulation of cardiac fatty acid metabolism, the role of PPAR in ischemia-reperfusion injury is not clear. The existing literature on the cardioprotective effects of PPAR agonists in ischemia-reperfusion is inconsistent (1, 8, 34, 44). Whereas some studies have shown that PPAR agonists are cardioprotective (8, 43, 44), other studies have not demonstrated any beneficial effect during postischemic recovery (1, 34). The differences may be related to the effects of specific agonists, duration of treatment, and differences in conditions to which the heart is exposed (i.e., the presence or absence of high levels of fatty acids), duration of ischemia, and type of ischemia (global vs. local). In the present study, we have clearly demonstrated that cardiac-restricted PPAR overexpression leads to increased FAO rates during aerobic and postischemic reperfusion, coinciding with a significant decrement in functional recovery of heart function during reperfusion. This decrease in heart function after ischemia in PPAR-overexpressing hearts was due to both a decreased recovery of heart rate and a decreased recovery of stroke work. Conversely, when the hearts from the mice lacking PPAR were subjected to ischemia-reperfusion, recovery of stroke work was improved compared with control hearts subjected to similar insults. This suggests that alterations in PPAR expression in the heart can have profound effects on functional recovery of ischemic hearts. Not surprisingly, hearts from PPAR–/– mice metabolized carbohydrates more favorably than fatty acids even during reperfusion after no-flow ischemia. These observations corroborate the already existing large body of literature that elevated FAO rates are indeed detrimental to the heart during reperfusion and that partial inhibition of FAO could be beneficial in the recovering heart.
PPAR controls the transcription of numerous genes involved in cardiac fatty acid metabolism (uptake and utilization) (5). Cardiac-restricted overexpression of PPAR (MHC-PPAR mice) results in stimulated expression of genes involved in cardiac fatty acid uptake (fatty acid transport protein 1 and CD36), thioesterification (fatty acyl-CoA synthetase 1), triacylglycerol synthesis (glycerol-3-phosphate acyltransferase and diacylglycerol acyltransferase), FAO (carnitine palmitoyl transferase 1 and acyl-CoA dehydrogenases), and malonyl-CoA decarboxylase (13). Furthermore, MHC-PPAR mice hearts exhibit a profound increase in palmitate uptake and oxidation and significantly decreased glucose utilization (13), which was confirmed during preischemic aerobic perfusion in this study. During postischemic reperfusion, the oxidation rates for palmitate were significantly increased in MHC-PPAR hearts compared with NTG littermates. Glucose oxidation also recovers almost to preischemic levels in control hearts during reperfusion, which is in contrast to earlier reports on rat heart substrate utilization (18). Mouse hearts in our hands seem to be more responsive to insulin (100 µU/ml) in the perfusate than rat hearts. Thus, despite rapid recovery of FAO during reperfusion, the glucose oxidation rates also resume preischemic levels in reperfused hearts and contribute to 40–50% of total energy production in the control hearts. Nevertheless, in MHC-PPAR hearts the majority of ATP (>80%) comes from palmitate during reperfusion, overriding the favorable effects of insulin on glucose metabolism. During postischemic reperfusion, the oxidation rates for palmitate were increased in MHC-PPAR hearts compared with NTG controls. During reperfusion after ischemia, MHC-PPAR hearts also exhibited significantly higher phosphorylation of ACC at Ser79. This phosphorylation is consistent with the increase in FAO seen in these hearts. Although Ser79 of ACC is a downstream target of AMPK, phosphorylation of AMPK was not significantly altered in MHC-PPAR heart homogenates. This suggests that kinases other then AMPK (such as protein kinase A) may be stimulated in MHC-PPAR hearts. Phosphorylation of ACC at Ser79 inhibits this enzyme, leading to diminished synthesis of malonyl-CoA, a known negative modulator of FAO. Thus PPAR could also mediate allosteric control of FAO via the ACC pathway in addition to the direct role of PPAR in upregulating other fatty acid oxidizing genes.
Despite the presence of insulin in our perfusions, glucose oxidation and glycolysis rates remained low during both the pre- and postischemic perfusions in MHC-PPAR hearts. This decrease in glucose metabolism is probably due to decreased glucose uptake by MHC-PPAR hearts and fatty acid inhibition of glucose oxidation (13). Although we did not measure GLUT4 in this study, we showed previously (14) that PPAR overexpression downregulates GLUT4 mRNA and protein expressions. ATP production calculations showed that 90% of total ATP was derived from palmitate oxidation and the remaining 10% was from glucose metabolism in MHC-PPAR hearts. Recently, Hopkins et al. (17) reported a much higher contribution of glucose oxidation (20–30%) to the total ATP pool, whereas FAO contributed the remaining 70–80% of ATP in MHC-PPAR hearts. It is important to note that the present study used a higher concentration of palmitate (1.2 mM) in the perfusion buffer, which increases FAO rates contributing to 
90% of the total ATP production. However, glucose oxidation was not severely affected in the present study because of the presence of a high concentration of insulin in the perfusate. In the control hearts, palmitate and glucose metabolism contributed equally to the total ATP production in during preischemic aerobic perfusion. Conversely, PPAR–/– mouse hearts derived a preponderance of their ATP from glucose utilization pathways. Previous and present studies demonstrated that palmitate oxidation rates in these hearts were remarkably reduced and glycolysis and glucose oxidation rates were significantly elevated (9). The above metabolic profiles in PPAR–/– mouse hearts conform to the previous findings that the various lipid metabolizing genes in these hearts were not upregulated (24). On the basis of theoretical calculation of the total number of ATP molecules derived from fatty acid and glucose oxidation over cardiac power, it is apparent that MHC-PPAR hearts exhibited the least metabolic efficiency among all the groups.
Postischemic recovery of cardiac function was very poor in MHC-PPAR hearts compared with control hearts. The poor functional recovery can be attributed to elevated levels of FAO rates during perfusions before and after ischemia. High levels of FAO have been shown to inhibit glucose oxidation at the level of pyruvate dehydrogenase (6, 25). This can lead to lactate and proton accumulation, which are detrimental to the myocardium. Myocardial damage can also occur because of generation of excessive amounts of reactive oxygen species as a result of enhanced FAO. It was shown that mitochondria from ischemic myocardium could be a source of toxic reactive oxygen species due to defective complex I activity (35). Indeed, previous studies in MHC-PPAR mice demonstrated that high fat feeding resulted in significant increases of markers of oxidative stress (e.g., hydrogen peroxide) (12). Thus poor functional recovery of MHC-PPAR mouse hearts during ischemia-reperfusion could therefore be due to a high FAO-induced oxidative stress.
The low-overexpressing MHC-PPAR (line 404-4) hearts in the present study did not exhibit hypertrophy (body wt/dry heart wt 136 ± 7 vs. 130 ± 4 for NTG littermates). Previously, Finck et al. (14) reported that low levels of PPAR overexpression caused moderate hypertrophy without alterations in cardiac function. This discrepancy between the two studies could be due to differences in age and or differences in litters that were used in the above-described studies. On the basis of our data, although we can eliminate contributions of hypertrophy to impaired functional recovery, we still must establish the causal relationship between impaired postischemic function and increased FAO. Further interventional studies are under way to examine the underlying mechanism(s).
PPAR–/– hearts performed better during both aerobic and postischemic perfusions compared with WT littermates. On the contrary, previous studies (9, 34) did not observe any alterations in basal cardiac function. This is probably due to the differences in perfusion conditions. For example, Panagia et al. (34) used 11 mM glucose without palmitate or other fatty acids whereas Campbell et al. (9) perfused the hearts with 0.4 mM palmitate and 5.0 mM glucose. In the present study, we used 1.2 mM palmitate and 5.0 mM glucose in the perfusion buffer. Postischemic functional recovery of PPAR–/– hearts was also greater compared with WT controls, and the percent recovery was significantly higher compared with postischemic recovery of MHC-PPAR hearts. In PPAR–/– mouse hearts, FAO rates remained low and glucose oxidation rates remained high during reperfusion after ischemia. The high rates of glucose oxidation were probably due to a compensatory mechanism in PPAR–/– hearts such that their AMP-to-ATP ratio (data not shown) and AMPK and ACC activities were not altered compared with perfusion-matched WT controls, indicating that these hearts were not energetically compromised.
In summary, cardiac-specific overexpression of PPAR favors lipid oxidation at the expense of glucose metabolism during pre- and postischemic perfusion and conversely affects functional recovery of the heart after global ischemia. Our results seem to contradict reports on the beneficial effects of PPAR agonist on ischemia-reperfusion injury to the heart (20, 44). In contrast to studies using acute systemic activation with PPAR ligands, our model is a chronic, cardiac-specific activation of PPAR that was previously characterized to exhibit a phenotype that resembles the diabetic heart (12). Therefore, although PPAR agonists by their systemic effects (both metabolic and anti-inflammatory) may improve myocardial performance, localized activation of PPAR in the heart may have deleterious effects on ischemic myocardium, especially in the presence of clinically relevant high levels of free fatty acids (3, 27). This evidence suggests that cardiac PPAR antagonism could be a potential therapeutic target to treat ischemia-reperfusion injury.
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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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 S. E. Nissen, S. J. Nicholls, K. Wolski, D. C. Howey, E. McErlean, M.-D. Wang, E. V. Gomez, and J. M. Russo Effects of a Potent and Selective PPAR-{alpha} Agonist in Patients With Atherogenic Dyslipidemia or Hypercholesterolemia: Two Randomized Controlled Trials JAMA, March 28, 2007; 297(12): 1362 - 1373. [Abstract] [Full Text] [PDF]
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B. N. Finck The PPAR regulatory system in cardiac physiology and disease Cardiovasc Res, January 15, 2007; 73(2): 269 - 277. [Abstract] [Full Text] [PDF]
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S. E. Nissen, K. Wolski, and E. J. Topol Effect of Muraglitazar on Death and Major Adverse Cardiovascular Events in Patients With Type 2 Diabetes Mellitus JAMA, November 23, 2005; 294(20): 2581 - 2586. [Abstract] [Full Text] [PDF]
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