Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2014 May 1.
Published in final edited form as: Circ Heart Fail. 2013 Apr 9;6(3):550–562. doi: 10.1161/CIRCHEARTFAILURE.112.000177

PPARγ Activation Prevents Sepsis-Related Cardiac Dysfunction and Mortality in Mice

Drosatos et al: PPARγ Treats Septic Cardiac Dysfunction

Konstantinos Drosatos 1, Raffay S Khan 1, Chad M Trent 1, Hongfeng Jiang 1, Ni-Huiping Son 1, William S Blaner 1, Shunichi Homma 2, P Christian Schulze 2, Ira J Goldberg 1
PMCID: PMC3690188  NIHMSID: NIHMS481820  PMID: 23572494

Abstract

Background

Cardiac dysfunction with sepsis is associated with both inflammation and reduced fatty acid oxidation (FAO). We hypothesized that energy deprivation accounts for sepsis-related cardiac dysfunction.

Methods and Results

E. coli lipopolysaccharide (LPS) administered to C57BL/6 mice (WT) induced cardiac dysfunction and reduced FAO and mRNA levels of peroxisome proliferator-activated receptor (PPAR) α and its downstream targets within 6-8h. Transgenic mice in which cardiomyocyte-specific expression of PPARγ is driven by the alpha myosin heavy chain promoter (αMHC-PPARγ) were protected from LPS-induced cardiac dysfunction. Despite a reduction in PPARα, FAO and associated genes were not decreased in hearts of LPS-treated αMHC-PPARγ mice. LPS treatment, however, continued to induce inflammation-related genes, such as interleukin (IL)-1α, IL-1β, IL-6 and tumor necrosis factor α in hearts of αMHC-PPARγ mice. Treatment of WT mice with LPS and the PPARγ agonist rosiglitazone, but not the PPARα agonist (WY-14643), increased FAO, prevented LPS-mediated reduction of mitochondria and treated cardiac dysfunction, as well as it improved survival despite continued increases in the expression of cardiac inflammatory markers.

Conclusion

Activation of PPARγ in LPS-treated mice prevented cardiac dysfunction and mortality despite development of cardiac inflammation and PPARα downregulation.

Keywords: fatty acids, sepsis, PPAR, cardiac metabolism

Impaired cardiac contractility contributes to the hypotension and increased mortality that occur with sepsis1. A possible cause of sepsis-mediated cardiac dysfunction is reduced energy production due in part to compromised fatty acid oxidation (FAO)2-5 and glucose catabolism3, 6. Thus, it is likely that sepsis compromises cardiac energy production, which might be the major cause of cardiac dysfunction. Alternatively, sepsis induces the production of inflammatory cytokines, such as tumor necrosis factor (TNF) α, interleukin (IL)-1 and IL-6, and these might directly alter heart function7-9.

Intraperitoneal (i.p.) injection of lipopolysaccharide (LPS) has been extensively used to model many of the clinical features of sepsis, including elevated inflammation and cardiac dysfunction10. LPS leads to production of inflammatory cytokines7-9, 11 and also reduces cardiac energy utilization2, 3, 12.

Nuclear receptors, particularly peroxisomal proliferator-activated receptors (PPARs), regulate cardiac FAO. The PPAR family consists of three members, PPARα, PPARδ and PPARγ. PPARα increases FA storage in triglycerides13 and FAO in heart14 and induces expression of peroxisomal and mitochondrial enzymes. Besides PPARα, cardiac FAO can be increased by activation of PPARγ15 or PPARδ16. Cardiomyocyte-specific overexpression of PPARα14 or PPARγ17 leads to cardiac lipid accumulation, an indication that lipid uptake exceeds FAO. PPARγ-coactivator-1 (PGC-1) α and β18 enhance FAO and mitochondrial biogenesis19. Both PPARα and PGC-1 mRNA levels are markedly reduced in the heart by LPS administration2, 3, 12, 20, while PPARγ is not affected2.

Our group showed that maintenance of normal cardiac FAO via c-Jun-N-terminal kinase (JNK) inhibitor-mediated prevention of PPARα downregulation rescued cardiac function in septic mice despite elevated expression of cardiac inflammatory markers. In a similar context constitutive cardiac expression of PGC-1β prevented cardiac dysfunction that was caused by LPS-mediated sepsis3, an observation that was proposed to be due to improvement in cardiac FAO and attenuation of reactive oxygen species production. In the current study we show that constitutive cardiomyocyte-specific expression of PPARγ or systemic administration of the PPARγ agonist, rosiglitazone, increased cardiac FAO and prevented cardiac dysfunction in mice with LPS-induced sepsis, despite increased expression of cardiac inflammatory markers. In addition, we show that rosiglitazone-mediated activation of PPARγ prevents the loss of cardiac mitochondria that occurs in sepsis. Moreover, we show that restoration of cardiac FAO by rosiglitazone not only prevents but also treats LPS-induced heart dysfunction and improves survival. Thus the use of rosiglitazone is proposed as a potential treatment for septic cardiac dysfunction.

Methods

An expanded Methods section appears in the supplementary section. All procedures involving animals were approved by the Institutional Animal Care and Use Committee at Columbia University. Briefly, this study included i.p administration of LPS in mice for induction of sepsis. Cardiac function was assessed by 2D-echocardiography that was performed 6-8h post-LPS administration. Mice were sacrificed and pieces of myocardium were obtained for gene expression and protein analysis, measurement of FA oxidation, glucose oxidation, mitochondrial enzymes’ activity and ATP levels. Plasma and cardiac lipids were measured by enzymatic assays. Cardiac ceramide content was assessed by liquid chromatography and in tandem mass spectrometry (LC/MS/MS) and cardiac mitochondrial number and size were analyzed by electron microscopy and mitochondrial ATPase 6 DNA copy number measurement. Survival studies were performed over 72h and included a single i.p administration of a lethal dose of LPS and daily injections of rosiglitazone.

Statistical analysis

Comparisons between 2 groups were performed using unpaired 2-tailed Student’s t tests. Comparisons between mean values of 3 or more groups were performed using 1-way analysis of variance followed by Bonferroni post hoc tests, performing all pairwise comparisons. All values are presented as mean ± SE. Survival was assessed by the Kaplan-Meier method and compared between groups using the Mantel-Cox log rank test. Differences were considered statistically significant at p<0.05.

Results

LPS-mediated sepsis compromised cardiac function and FAO

Several studies2, 3, 12, 20-22 have shown that administration of LPS in mice reduces cardiac PPARα mRNA levels3, 12, 20-22, which may account for the reduction of cardiac FAO. We confirmed that LPS administration reduced both cardiac PPARα mRNA levels by 75% [Saline: 1.04 ± 0.08, LPS: 0.25 ± 0.03; p<0.0001, n=7] and heart function by 40% as shown by fractional shortening levels in C57BL/6 mice [Saline: 44.3% ± 2.1%, LPS: 25.9% ± 2.1%; p<0.0001, n=7]. Compromised cardiac function was associated with reduced cardiac FAO (Figure 1A) and ATP (Figure 1B) levels, as well as with increased phosphorylation of adenosine monophosphate kinase (AMPK) (Figure 1C and Suppl. Fig 1). Heart:body weight and lung:body weight ratios were not increased by LPS (Suppl. Figure 2A, 2B).

Figure 1. Constitutive expression of PPARγ in cardiomyocytes stimulates cardiac FAO and prevents LPS-mediated heart dysfunction despite elevated inflammation.

Figure 1

Figure 1

Open in a new tab

(A, B) [3H]-palmitic acid oxidation (A) and ATP (B) levels in cardiac muscle of 10-12 weeks old C57BL/6 mice treated with 5 mg/kg LPS or combination of LPS and 30 mg/kg rosiglitazone (Rosi). Control mice were treated with saline; n = 5; *p<0.05, **p<0.01. (C) Western blot analysis of cardiac pAMPK and total AMPK of 10- to 12-week-old C57BL/6 mice treated with 5 mg/kg LPS. (D) Western blot analysis of cardiac pJNK, total JNK, pAMPK and total AMPK of 10- to 12-week-old αMHC-PPARγ mice treated with 5 mg/kg LPS. (E) PPARα, PPARγ, PPARδ, ERRα, Cpt1β, AOX, PGC-1α, PGC-1β, CD36, PLIN 2, PLIN 5 and PDK4 mRNA levels in hearts of C57BL/6 mice treated with 5 mg/kg LPS or a combination of LPS and 30 mg/kg WY-14643. Control mice were treated with saline; n=5; *p<0.05 vs CTRL, **p<0.01 vs. CTRL, #p<0.001 vs. CTRL. (F) Cardiac PPARα, PPARδ, ERRα, Cpt-1β, AOX, PGC-1α, PGC-1β, and PDK4 mRNA levels of 10- to 12-week-old αMHC-PPARγ mice treated with 5 mg/kg LPS. n = 5; *p<0.05, **p < 0.01, #p<0.0001. (G, H) Photographs of echocardiograms (G) and fractional shortening (H) of 10 weeks old αMHC-PPARγ mice treated with 5 mg/kg LPS; n = 7. (I-J) [3H]-palmitic acid oxidation (I) and ATP (J) levels in cardiac muscle of αMHC-PPARγ mice treated with 5 mg/kg LPS. n = 5. (K) IL-1α, IL-1β, IL-6 and TNFα mRNA levels in hearts of 10 weeks old αMHC-PPARγ mice treated with 5 mg/kg LPS. n = 5; **p < 0.01.

Administration of PPARα agonist does not rescue LPS-driven cardiac dysfunction and inhibition of FAO-related gene expression

To test whether PPARα agonists would alleviate septic cardiac dysfunction C57BL/6 mice were treated with combination of LPS (5 mg/kg) and the PPARα agonist, WY-14643 (30 mg/kg). Administration of WY-14643 alone improved fractional shortening (43%) as compared to control saline-treated mice (Suppl. Figure 3A, 3B). However administration of WY-14643 in LPS-treated mice did not prevent cardiac dysfunction (Suppl. Figure 3A, 3B). In addition, although WY-14643 alone stimulated the expression of FA metabolism-related genes, such as CD36, PGC-1α and PGC-1β, it did not prevent their downregulation by LPS. Moreover, WY-14643 did not prevent the LPS-mediated changes of PPARα, ERRα, Cpt-1β, AOX, PLIN2 and PDK4 mRNA levels (Figure 1E). Thus, administration of PPARα agonist cannot prevent cardiac dysfunction that occurs in sepsis, which might be due to the profound reduction of PPARα gene expression levels and eventually to weak PPARα activation by its agonist.

Constitutive cardiomyocyte-specific expression of PPARγ prevents reduction of cardiac FAO in LPS-treated mice despite PPARα downregulation

In order to assess whether PPARγ overexpression can compensate for the LPS-associated reduction of PPARα and can rescue FAO and cardiac function, we administered 5mg/kg LPS to mice with constitutive expression of PPARγ in cardiomyocytes (αMHC-PPARγ)17. Treatment of αMHC-PPARγ mice with LPS activated (increased phosphorylation) JNK (Figure 1D and Suppl. Figure 4) and reduced PPARα mRNA levels (Figure 1F) as was found in WT mice. However, cardiac function of the LPS-treated αMHC-PPARγ mice was not reduced. Specifically, 2D-echocardiography (Figure 1G) showed fractional shortening levels in LPS-treated αMHC-PPARγ mice that were comparable to saline treated αMHC-PPARγ and WT (Figure 1H). Thus, constitutive cardiomyocyte PPARγ expression prevented LPS-mediated cardiac dysfunction.

Opposite to the inhibitory effect of LPS on energy production in WT mice (Figure 1A, 1B), the levels of cardiac FAO (Figure 1I) and ATP (Figure 1J) were not reduced in LPS-treated αMHC-PPARγ mice as compared to saline treated αMHC-PPARγ mice. Phosphorylated AMPK (pAMPK) levels were reduced in the LPS-treated αMHC-PPARγ mice, as compared to saline-treated αMHC-PPARγ mice (Figure 1D and Suppl. Figure 4). Accordingly, the inhibitory effect of LPS on FAO-associated gene expression levels that was observed in WT mice2, 3, 12, 20 was ablated in septic αMHC-PPARγ mice. Specifically, AOX, PPARδ, PGC-1α and carnitine palmitoyl transferase (Cpt)-1β mRNA levels were increased over 10-fold each as compared to saline-treated αMHC-PPARγ mice (Figure 1F). αMHC-PPARγ mice have increased cardiac expression of AOX (3.5-fold)17 and Cpt-1β (30%)17, while PPARα, PPARδ and PGC-1α are not significantly altered, as compared to wild type mice (Suppl. Table 1). These changes occurred despite a reduced cardiac PGC-1β mRNA (76%) (Figure 1F), perhaps due to PGC-1α compensation. Pyruvate dehydrogenase kinase (PDK) 4 mRNA levels were increased by 5.4-fold in the hearts of LPS-treated αMHC-PPARγ (Figure 1F), suggesting that preserved cardiac function was unlikely due to greater glucose utilization. Basal PDK4 gene expression levels are 50% lower in αMHC-PPARγ mice, as compared to wild type mice (Suppl. Table 1). Thus, our data show that gene expression changes paralleled the prevention of FAO inhibition in LPS-treated αMHC-PPARγ mice.

Cardiac triglyceride stores are not increased in LPS-treated αMHC-PPARγ mice

Septic mice have increased cardiac triglyceride (TG) content3 perhaps as a consequence to increased plasma free fatty acid (FFA) and triglyceride levels23, 24 and reduced cardiac fatty acid utilization. We confirmed that LPS treatment led to accumulation of TGs in the hearts of C57BL/6 mice (Figure 2A) and this increase was associated with elevated plasma TG (Figure 2B) and FFA levels (Figure 2C). On the other hand LPS-treated αMHC-PPARγ mice did not show either cardiac (Figure 2D) or plasma (Figure 2E) TG increases. Plasma FFA levels were increased in LPS-treated αMHC-PPARγ mice (Figure 2F). This was likely due to balanced lipid uptake and FAO. Consistent with the lack of change in stored TG the expression levels of genes that are associated with lipid uptake and storage were not increased in LPS-treated αMHC-PPARγ mice. LPS reduced mRNA levels of cluster of differentiation (CD) 36 and fatty acid transport protein (FATP) 1 in both WT (Figure 2G) and αMHC-PPARγ (Figure 2H) LPS-treated mice. Basal CD36 and FATP expression levels are 50% higher17 and 50% lower (Suppl. Table 1), respectively, in αMHC-PPARγ mice as compared to wild type mice. Although lipoprotein lipase (LpL) gene expression levels were reduced in WT mice (Figure 2G) they were not significantly affected by LPS treatment in αMHC-PPARγ mice (Figure 2H), which have similar LpL basal gene expression levels, as compared to wild type mice (Suppl. Table 1). Angiopoietin-like protein (Angptl) 4, which inhibits LpL activity25, was increased in both WT (Figure 2G) and αMHC-PPARγ (Figure 2H) mice that were treated with LPS. The change in Angptl4 gene expression levels may account for the reported LPS-mediated suppression of cardiac LpL activity21. These changes might have modified the expected greater uptake due to increased circulating TG and FFA levels. Accordingly, gene expression levels of perilipin (PLIN) 2, which is associated with lipid droplet formation, were not modulated in the hearts of LPS-treated αMHC-PPARγ mice (Figure 2H) although cardiac PLIN2 mRNA levels were dramatically increased (7.7-fold) in the hearts of LPS-treated WT mice (Figure 2G). The expression levels of another lipid droplet-associated protein PLIN5, which might reduce fatty acid catabolism26, were not modulated in LPS-treated WT (Figure 2G) mice but decreased (50%) in αMHC-PPARγ mice treated with LPS (Figure 2H). Thus, while LPS-treated WT mice accumulate lipid droplets associated with decreased FAO and expression of PLIN2, LPS did not alter the balance between lipid uptake, oxidation, and storage in mice with constitutive expression of PPARγ in cardiomyocytes, and did not induce cardiac dysfunction.

Figure 2. Lipid profiles in LPS-treated WT and αMHC-PPARγ mice.

Figure 2

Figure 2

Open in a new tab

(A-F) Cardiac triglycerides (A and D), plasma triglycerides (B and E) and plasma non-esterified fatty acids (C and F) of C57BL/6 mice (A-C) treated with 5 mg/kg LPS or a combination of LPS and 30 mg/kg rosiglitazone (Rosi) or αMHC-PPARγ mice (D-F) treated with 5 mg/kg LPS. Control mice were treated with saline; n=4-8, *p<0.05, **p<0.01, #p<0.001, ¤p<0.05 vs. non-LPS control. (G-H) Cardiac CD36, FATP, LpL, Angptl4, PLIN 2 and PLIN 5 mRNA levels of C57BL/6 mice treated with 5 mg/kg LPS or a combination of LPS and 30 mg/kg rosiglitazone (Rosi) (G) or αMHC-PPARγ mice (H) treated with 5 mg/kg LPS. Control mice were treated with saline; n=5-10; ¤p<0.05 vs. CTRL, ¶p<0.01 vs CTRL, ±p<0.001 vs. CTRL, ¥p<0.0001 vs CTRL, ÷p<0.05 vs LPS, ~p<0.01 vs. LPS, *p<0.05 vs. CTRL, **p<0.01 vs. CTRL. (I-J) Cardiac total ceramide (I) and ceramide species (J) content of C57BL/6 mice treated with 5 mg/kg LPS or a combination of LPS and 30 mg/kg rosiglitazone (Rosi). Control mice were treated with saline; n=4, *p<0.05, #p<0.001, ¤p<0.05 vs. non-LPS control.

Prevention of LPS-induced cardiac dysfunction in αMHC-PPARγ mice was not associated with alleviation of inflammation

Inflammation is a major component of sepsis and its alleviation has been proposed as a therapeutic approach in sepsis based on studies in animal models with deletions of inflammation-related genes7-9, 11. However, prevention of LPS-related cardiac dysfunction in αMHC-PPARγ mice was not accompanied by alleviation of cardiac inflammation, as shown by cardiac mRNA levels of IL-1α, IL-1β, IL-6 and TNFα (Figure 1K) that were elevated by >10 fold by LPS treatment. Thus, the beneficial effect of PPARγ in the prevention of cardiac dysfunction during sepsis cannot be accounted for by direct anti-inflammatory effects on the heart.

Pharmacologic activation of PPARγ improves cardiac FAO and prevents heart dysfunction in LPS-treated C57BL/6 mice

In order to assess whether pharmacologic activation of PPARγ prevents septic cardiac dysfunction, WT mice were treated i.p. with the PPARγ agonist, rosiglitazone (30 mg/kg) 30 min prior to the administration of LPS. Although LPS treatment activated JNK (Figure 3A and Suppl. Figure 5) and reduced PPARα mRNA levels in both control saline-treated and rosiglitazone-treated animals, rosiglitazone prevented LPS-mediated cardiac dysfunction (Figure 3B, 3C, 3D). Rosiglitazone treatment did not prevent increases in plasma TG (Figure 2B) and FFA (Figure 2C) levels, which usually occur by LPS treatment23, 24. Opposite to the LPS-treated mice, cardiac FAO (Figure 1A) and ATP levels (Figure 1B) were not reduced in mice receiving both LPS and rosiglitazone.

Figure 3. Rosiglitazone-mediated PPARγ activation improves cardiac function by preventing LPS-induced reduction in cardiac FAO despite elevated inflammation and suppressed glucose catabolism.

Figure 3

Figure 3

Open in a new tab

(A) Western blot analysis of cardiac pAMPK, total AMPK, pJNK and total JNK of C57BL/6 mice treated with 5 mg/kg LPS or a combination of LPS and 30 mg/kg rosiglitazone (Rosi). Control mice were treated with saline. (B, C) Photographs of echocardiograms (B) and fractional shortening (C) of 10-12 weeks old C57BL/6 mice treated with 5 mg/kg LPS or a combination of LPS and 30 mg/kg rosiglitazone (Rosi). Control mice were treated with saline; n = 5; #p<0.0001. (D) Cardiac PPARα, PPARγ, PPARδ, ERRα, AOX and Cpt1β mRNA levels of C57BL/6 mice treated with 5 mg/kg LPS or a combination of LPS and 35 mg/kg rosiglitazone (Rosi). Control mice were treated with saline; n=5-10; ¤p<0.05 vs. CTRL, ¶p<0.01 vs CTRL, ±p<0.001 vs. CTRL, ¥p<0.0001 vs CTRL, ÷p<0.05 vs LPS, ~p<0.01 vs. LPS. (E) Cardiac IL-1α, IL-1β, IL-6 and TNFα mRNA levels of C57BL/6 mice treated with 5 mg/kg LPS or a combination of LPS and 35 mg/kg rosiglitazone (Rosi). Control mice were treated with saline; n=5; *p<0.05, **p<0.01, ***p<0.001, #p<0.0001 vs. non-LPS CTRL. (F) [14C]-glucose oxidation levels in cardiac muscle of C57BL/6 mice treated with 5 mg/kg LPS or a combination of LPS and 30 mg/kg rosiglitazone (Rosi). Control mice were treated with saline; n=6-7, **p<0.01, ***p<0.001. (G) Cardiac GLUT4, GLUT1 and PDK4 mRNA levels of C57BL/6 mice treated with 5 mg/kg LPS or a combination of LPS and 35 mg/kg rosiglitazone (Rosi). Control mice were treated with saline; n=5-10; *p<0.05, **p<0.01, ***p<0.001, #p<0.0001 vs. non-LPS CTRL. (H) Western blot analysis of cardiac pAkt, total Akt and β-actin of C57BL/6 mice treated with 5 mg/kg LPS or a combination of LPS and 30 mg/kg rosiglitazone (Rosi). Control mice were treated with saline.

The expression profile of several FAO-related genes correlated with the prevention of LPS-mediated suppression of FAO and was similar to the changes found in the LPS-treated αMHC-PPARγ mice. Although PPARα mRNA levels were reduced (75%) by treatment of WT mice with LPS (Figure 3D), as they were in αMHC-PPARγ mice, Cpt-1β mRNA was increased by 35 % (Figure 3D) and Cpt activity tended to increase in rosiglitazone-treated septic mice as compared to LPS-treated mice [Saline: 2.02 ± 0.54, Rosi: 1.49 ± 0.36, LPS: 1.09 ± 0.18*, LPS + Rosi: 1.61 ± 0.49 nmol/min/mg; n=5, *p<0.05 vs saline]. Similarly, medium- and very long-chain acyl-CoA dehydrogenase (MCAD and VLCAD) mRNA levels were reduced by 30% and 25% respectively in LPS-treated mice, while they were not affected in mice treated with LPS and rosiglitazone (Suppl. Figure 6). LCAD mRNA levels were not modulated (Suppl. Figure 6). Phosphorylated AMPK that was increased in LPS-treated mice did not change significantly in mice treated with combination of LPS and rosiglitazone, as compared to septic mice (Figure 3A and Suppl. Fig 5).

Prevention of LPS-induced cardiac dysfunction by PPARγ agonist was not associated with alleviation of cardiac inflammation

Although PPARγ activation exerts anti-inflammatory effects by inhibiting the expression of pro-inflammatory genes in macrophages27, rosiglitazone-mediated improvement of cardiac function in LPS-treated WT mice was not due to attenuation of cardiac inflammation. Specifically, injection of rosiglitazone in LPS-treated-mice did not prevent the increases in IL-1α, IL-1β, IL-6 and TNFα gene expression levels (Figure 3E). Thus, the improvement in cardiac function by rosiglitazone was not associated with attenuation of expression of LPS-triggered inflammatory markers in the hearts.

Improvement in cardiac function by rosiglitazone was not associated with increased glucose utilization

Rosiglitazone is an insulin sensitizer that increases glucose catabolism. Thus we tested, whether rosiglitazone-mediated improvement in cardiac function of septic mice was accompanied by increased cardiac glucose catabolism. Glucose oxidation was reduced by 70% in C57BL6 mice that were treated with LPS and this defect was not attenuated by administration of rosiglitazone (Figure 3F). Accordingly, glucose transporter (GLUT) 4 mRNA levels were reduced (60%) in LPS-treated mice and were not increased by co-administration of rosiglitazone (Figure 3G). GLUT1 mRNA levels did not change by any of the treatments (Figure 3G). PDK4 was increased 8-fold by LPS treatment and 4-fold in mice that received LPS and rosiglitazone as compared to saline-treated mice (Figure 3G), also indicating suppressed cardiac glucose utilization. In addition, phosphorylated Akt levels were reduced in the hearts of LPS-treated mice, indicating inhibition of the insulin signaling pathway, which was not reactivated by rosiglitazone treatment (Figure 3H and Suppl. Figure 7). Plasma glucose levels were reduced in mice treated with either LPS or combination of LPS and rosiglitazone [CTRL: 130 ± 10.6 mg/dl, Rosi: 93.6 ± 4.9 mg/dl**, LPS: 43 ± 4.2 mg/dl****, LPS+Rosi: 50.3 ± 2.5 mg/dl¶; n=5, **p<0.01 vs. CTRL, ****p<0.0001 vs. CTRL, p<0.01 vs. Rosi]. Thus, the rosiglitazone-mediated improvement in cardiac function of septic mice was not accompanied by restoration of glucose utilization.

Rosiglitazone-mediated improvement of cardiac function in LPS-treated mice was not associated with cardiac lipid changes

Rosiglitazone administration along with LPS was still associated with increased cardiac TG content, although to a slightly less extent than was found in mice treated with LPS alone (Figure 2A). This likely reflected the increased FAO driven by rosiglitazone, as circulating TG and FFAs were similar to those measured in LPS-treated WT mice (Figure 2B, 2C). CD36 mRNA levels were increased (7-fold) in mice treated with LPS and rosiglitazone, as compared to mice treated with LPS only (Figure 2G). Moreover, these data suggest that lipid accumulation, which is toxic in other circumstances17, 28-30, was not the reason for LPS-mediated heart dysfunction.

Further evidence was obtained by measuring intracellular toxic lipids. Impaired utilization and eventual intracellular accumulation of fatty acids are often accompanied by increased ceramide levels, which may account for cardiac dysfunction31, 32. Despite accumulation of TGs in the hearts of LPS-treated C57BL/6 mice, liquid chromatography tandem mass spectrometry (LC/MS/MS) analysis of cardiac lipids showed that neither total ceramide levels (Figure 2I) nor any of the ceramide species that we tested (Figure 2J) were increased in septic animals. In addition, combination of LPS and rosiglitazone did not affect cardiac ceramide levels as compared to LPS-treated mice (Figure 2I, 2J). Thus, both LPS-driven cardiac dysfunction and rosiglitazone-mediated improvement of cardiac function in LPS-treated mice were not associated with changes in ceramide levels.

Rosiglitazone-mediated improvement in cardiac function of septic mice is associated with increased cardiac mitochondrial number and size

Mitochondria are the major site of substrate oxidation and ATP production in cardiomyocytes. Electron microscopy (EM) analysis (magnification: 6,000×) revealed reduced number of mitochondria in cardiomyocytes of LPS-treated mice (Figure 4A, Suppl. Fig 8A). This observation was confirmed by quantitation of ATPase 6/β-actin DNA ratio, which was 30% lower in hearts of LPS-treated mice (Figure 4B), an additional indication of reduced number of mitochondria. Both EM (Figure 4A) and DNA analyses (Figure 4B) showed that combined treatment of mice with LPS and rosiglitazone prevented reduction of mitochondria. Analysis of the activity of mitochondrial enzymes showed that citrate synthase activity was reduced by 35% in LPS-treated mice and was restored to normal levels in septic mice that were treated with rosiglitazone (Figure 4C). Similarly, a 25% reduction was observed in cardiac complex IV activity by LPS treatment that was prevented in mice treated with LPS and rosiglitazone. (Figure 4D). However, when complex IV activity was normalized to citrate synthase activity no significant change was observed among all animal groups (Figure 4E). This observation indicates that the reduction in complex IV activity was likely due to the reduction in mitochondrial number. Thus, EM, DNA analysis and mitochondrial enzyme activity assays showed that rosiglitazone prevented LPS-driven reduction of mitochondria.

Figure 4. Rosiglitazone prevents LPS-mediated autophagy and reduction of mitochondrial number and size.

Figure 4

Figure 4

Open in a new tab

(A) Electron microscopy analysis (6000×) of cardiomyocytes of C57BL/6 mice treated with 5 mg/kg LPS, 30 mg/kg rosiglitazone (rosi) or combination of LPS and rosi. Control mice were treated with saline. (B-E) Ratio of cardiac mitochondrial gene DNA (ATPase6) to nuclear gene DNA (β-actin) (B), citrate synthase activity (C), complex IV activity (D), complex IV activity normalized to citrate synthase activity (E) and PGC1-α, PGC1-β and mitochondrial transcription factor A (mtTFA) mRNA levels (F) C57BL/6 mice treated with 5 mg/kg LPS, 30 mg/kg rosiglitazone (rosi) or combination of LPS and rosi. Control mice were treated with saline; n = 5-14, ¤p<0.05 vs. CTRL, ±p<0.001 vs. CTRL, ÷p<0.05 vs LPS, ~p<0.01 vs. LPS. (G) Electron microscopy analysis (20000×) for the assessment of mitochondrial size in cardiomyocytes of C57BL/6 mice treated with 5 mg/kg LPS, 30 mg/kg rosiglitazone (rosi) or combination of LPS and rosi. Control mice were treated with saline. (H) Cardiac Atg3, Atg5, Atg7, Atg10 and Bcn1 mRNA levels of C57BL/6 mice treated with 5 mg/kg LPS or combination of LPS and 35 mg/kg rosiglitazone (Rosi). Control mice were treated with saline; n=5; *p<0.05 vs. CTRL. (I) Western blot analysis of cardiac pAMPK, total AMPK, LC3B, Atg7 and β-actin of C57BL/6 mice treated with 5 mg/kg LPS or combination of LPS and 30 mg/kg rosiglitazone (Rosi). Control mice were treated with saline. Each panel was obtained by a separate SDS-PAGE and Western Blotting assay.

Mitochondrial biogenesis is controlled by mitochondrial transcription factor A (mtTFA)33, 34, which is regulated by PGC-135. PGC-1 is important for the maintenance of healthy cardiac function under stressed conditions3, 36 and is a central regulator of mitochondrial biogenesis19. PGC-1α and PGC-1β mRNA levels were reduced by 70% and 60%, respectively, in LPS-treated mice and restored by rosiglitazone treatment of septic animals (Figure 4F). Specifically, rosiglitazone administration in LPS-treated mice increased PGC-1α (5.3-fold) and PGC-1β (4-fold) mRNA levels, as compared to mice treated with LPS only (Figure 4F). Moreover, PGC1-α mRNA levels were increased by 2.4-fold in mice that were treated with rosiglitazone alone, as compared to saline-treated mice (Figure 4F). Accordingly, cardiac mtTFA mRNA levels were reduced by 50% in LPS-treated mice and treatment with rosiglitazone restored normal levels of mtTFA in septic mice (Figure 4F). Further analysis of the EM photos showed that cardiomyocytes of LPS-treated mice had less area occupied by mitochondria as compared to either saline-treated mice or mice treated with LPS and rosiglitazone (Suppl. Figure 8B). LPS reduced average mitochondrial size, which tended to be reversed in LPS-treated mice that received rosiglitazone (Figure 4G, Suppl. Figure 8C). Thus, LPS reduces cardiac mitochondrial number and size and the mitochondrial phenotype is rescued by rosiglitazone.

Autophagy is a cellular process that affects mitochondrial turnover37 and is induced by LPS administration in cardiomyocytes38. Thus, we tested whether markers of autophagosome formation are modulated in the hearts of mice treated with LPS or combination of LPS and rosiglitazone. LPS administration increased mRNA levels of autophagy-related gene (Atg) 3 and Atg5 (Figure 4H), while administration of rosiglitazone in LPS-treated mice normalized Atg3 and Atg5. On the other hand Atg7, Atg10 and Beclin (Bcn) 1 mRNA levels were not significantly modulated in any of the tested treatments (Figure 4H). Protein analysis of LPS-treated hearts showed that septic mice had increased levels of Atg7 and microtubule-associated protein 1, light chain 3, beta (LC3B) II, markers of autophagosome formation (Figure 4I, Suppl. Fig 9). These changes were prohibited when LPS-treated mice were co-administered rosiglitazone (Figure 4I, Suppl. Fig 9). Thus, autophagy, which may affect mitochondrial number, is induced in the “starved” hearts of septic mice and is normalized in mice treated with LPS and rosiglitazone.

Administration of rosiglitazone after induction of sepsis with LPS treats cardiac dysfunction

In order to assess whether rosiglitazone administration is a potential treatment for established sepsis-related cardiac dysfunction, C57BL/6 mice were injected i.p. with 5 mg/kg LPS and 30 mg/kg rosiglitazone was administered i.p. 4h post-LPS injection, when cardiac dysfunction was confirmed by 2D-echocardiography. Rosiglitazone restored normal cardiac function in all mice tested (Figure 5A, 5B). On the other hand, cardiac function of LPS-treated mice that received vehicle (DMSO) either did not improve or was even more aggravated (Figure 5C, 5D). Rosiglitazone did not alter cardiac function of saline-treated mice (Figure 5E, 5F).

Figure 5. Administration of rosiglitazone in C57BL/6 mice with LPS-mediated cardiac dysfunction corrects cardiac function and improves survival.

Figure 5

Open in a new tab

(A-F) Photographs of echocardiograms and fractional shortening of individual 10-12 weeks old C57BL/6 mice treated with LPS and rosiglitazone (30 mg/kg) 4h post-LPS administration; p<0.01 (A, B) or 5 mg/kg LPS and DMSO (vehicle) 4h post-LPS administration (C, D) or saline and rosiglitazone (30 mg/kg) 4h post-saline administration (E, F). (G) Survival curves of C57BL/6 mice treated with saline (n=8), 30mg/kg/day rosiglitazone (n=8), 15 mg/kg LPS (n=15) or combination of rosiglitazone and LPS (n=19); p<0.0001.

Treatment with rosiglitazone improved survival of LPS-treated animals

C57BL/6 mice were administered a lethal dose of LPS (15 mg/kg, i.p.) or combination of LPS and daily injections of rosiglitazone (30 mg/kg), and were monitored for 72h. Mice treated with LPS alone started dying 24h post-injection with the vast majority of lethal events occurring between 48h and 72h post-injection. Daily administration of rosiglitazone reduced mortality (Figure 5G) and improved mobility and general appearance of the LPS-treated mice (Supplementary video files).

Discussion

Sepsis is a major cause of death in intensive care units. Approximately 25% of patients with severe sepsis have cardiovascular complications39 and 30% of them are terminal1. Cardiac FAO is suppressed during sepsis as shown by analysis of myocardial pieces2, isolated perfused hearts3, 5 and cardiac homogenates4. Opposite to afterload-induced HF that FAO is reduced and glucose oxidation is increased40, our work and previous studies3, 6 showed that LPS-mediated reduction cardiac FAO is not compensated by increased glucose catabolism. Thus, sepsis is accompanied by cardiac energy deficiency. Since either PPARα or PPARγ can stimulate cardiac FAO15, 41, we tested whether PPARγ activation compensates for LPS-mediated PPARα reduction and restores FAO and cardiac function. Constitutive cardiomyocyte expression and rosiglitazone-mediated activation of PPARγ improved FAO and prevented cardiac dysfunction in mice that were treated with LPS, although cardiac inflammation was not alleviated.

The high expressing transgenic line of αMHC-PPARγ has impaired cardiac function17. Therefore, we used the medium-expressing line that has normal cardiac function to test whether constitutive cardiomyocyte expression of PPARγ prevents the detrimental effects of LPS on cardiac function. Despite the downregulation of PPARα by LPS, αMHC-PPARγ mice had normal cardiac FAO, ATP and increased FAO-associated gene expression levels, as well as normal heart function. Similarly, rosiglitazone increased cardiac FAO and ATP, prevented cardiac dysfunction in LPS-treated mice and improved survival when given after confirmation of septic cardiac dysfunction. Prevention of septic cardiac dysfunction in αMHC-PPARγ mice is associated with increased PGC-1α, Cpt-1β, PPARδ and AOX mRNA levels. Rosiglitazone-mediated improvement in cardiac function of LPS-treated C57BL/6 mice is accompanied by increased PGC-1α, CD36 and Cpt-1β mRNA levels. The slightly different profile in gene expression changes between the two animal models may reflect differences between chronic (genetic model) and acute (pharmacological model) PPARγ activation in sepsis. It also may be due to the fact that pharmacological treatments are not totally specific and may also affect other tissues that in turn may alter cardiac gene expression profile. The gene expression profile was consistent with our previous findings showing that constitutive PPARγ expression can substitute for PPARα to maintain FAO15. PPARγ activation increases FAO more potently when PPARα is downregulated but the underlying cellular mechanism remains unknown.

Although chronic PPARγ agonist administration has anti-inflammatory effects especially in macrophages42, the PPARγ-mediated improvement in cardiac function of LPS-treated mice was not associated with attenuation of the expression of inflammatory markers in the heart during our short-term studies. This finding is consistent with our previous study showing that improvement in cardiac function by JNK inhibition was not associated with alleviation of local cardiac inflammation2. These observations indicate that inflammation is not the major aggravating factor for sepsis-triggered cardiac dysfunction, at least at the early stages of the disease.

Rosiglitazone increased PGC-1α in saline and LPS-treated C57BL/6 mice. PGC-1 is a coactivator of cellular energy metabolism-related transcription factors including PPARs43 and a regulator of mitochondrial biogenesis19. A recent study showed that constitutive PGC-1 expression protects cardiomyocytes from LPS-mediated consequences3 via improvement in FAO and reduction in reactive oxygen species production. Furthermore, our data show that the number and size of mitochondria were reduced in hearts of LPS-treated mice and rescued in septic animals that were treated with rosiglitazone. Reduction mitochondrial number, has also been reported in hearts of septic rats44. Mitochondrial turnover that is induced in LPS-treated cardiomyocytes38, occurs via autophagy37, which is triggered by nutrient deprivation45 and subsequent AMPK activation46 that we also observed. Restoration of cardiac mitochondrial number in septic mice treated with rosiglitazone was associated with reduced levels of autophagy markers. Thus, the beneficial effect of rosiglitazone in the prevention of mitochondrial loss may be due to autophagy suppression.

In conclusion, we have found that PPARγ activation prevents LPS-mediated reduction in cardiac FAO and mechanical dysfunction despite increased expression of inflammatory markers in the heart. Systemic administration of PPARγ agonist improves survival and overall appearance of LPS treated mice. The findings of this and our previous study2 suggest that compromised energy production rather than cardiac inflammation appears to be a critical factor for reduced cardiac function within the first hours of disease progression. A schematic representation of the proposed pathway that mediates the effects of sepsis on cardiac FAO and heart function, as well as the protective role of PPARγ is in Figure 6. LPS induces the production of inflammatory cytokines and suppresses PPARα and PGC-1 leading to compromised FAO and cardiac dysfunction. Impaired FAO is associated with triglyceride accumulation, increased autophagy, which in combination with PGC-1 downregulation, leads to reduction of mitochondrial number. Activation of PPARγ compensates for LPS-mediated downregulation of PPARα, and preserves FAO and cardiac function despite continued inflammation. Besides heart, sepsis affects the function of multiple other organs47. The importance of defective metabolism versus inflammation in sepsis-induced dysfunction in these organs is unknown. The beneficial effects on heart function of septic mice by the acute treatment with rosiglitazone, which we observed in our study, contrast with those associated with chronic use of PPARγ agonist in patients with diabetes48. In these situations PPARγ agonists are associated with increased heart failure, either because they increased salt and water retention49 or because they directly affect heart function perhaps due to lipid toxicity17, 50. Nonetheless, our data implicate acute PPARγ-mediated induction of FAO as a treatment of sepsis that can be used in combination with anti-inflammatory therapies to prevent dysfunction of the heart and possibly other organs.

Figure 6. Proposed model about the role of PPARγ activation in the prevention of LPS-mediated cardiac dysfunction.

Figure 6

Figure 6

Open in a new tab

(A) LPS reduces PPARα and PGC-1, which cause fatty acid oxidation inhibition, triglyceride accumulation and cardiac dysfunction. Nutrient deprivation induces autophagy, which in combination with PGC-1 downregulation, reduces mitochondrial number. Inflammatory pathways are also activated. (B) Activation of PPARγ induces PGC-1 and CD36 expression and prevents sepsis-mediated down-regulation of cardiac fatty acid utilization and energy production, thus it protects cardiac function in sepsis despite increased levels of inflammatory cytokines. Sufficient nutrient supply prohibits autophagy and thus maintains normal mitochondrial number.

Supplementary Material

1
Download video file (6.6MB, mp4)
2
Download video file (6.8MB, mp4)
3
Download video file (6.5MB, mp4)
4
Download video file (6.6MB, mp4)
5
Download video file (6.5MB, mp4)
6
Download video file (6.7MB, mp4)
Supp

CLINICAL SUMMARY.

Sepsis is a major cause of death in critically ill patients and has been associated with impaired cardiac contractility and diastolic dysfunction that contribute to hypotension and increased mortality. Approximately 25% of patients with severe sepsis have cardiovascular complications and 30% of them are terminal. Cardiac dysfunction in sepsis is associated with both inflammation and reduced fatty acid oxidation (FAO). In contrast to chronic left ventricular dysfunction secondary to pressure-overload or ischemia, which show reduced FAO and increased glucose oxidation rates, septic cardiac dysfunction is characterized by general suppression of both FA and glucose utilization for energy homeostasis. We hypothesized that cardiac dysfunction during sepsis is due, at least in part, to cardiac energy deficiency. Using an animal model of lipopolysaccharide (LPS)-induced cardiac dysfunction we show that left ventricular dysfunction is prevented by genetic or pharmacological activation of peroxisome proliferator-activated receptor (PPAR)-γ, treatments that improve FAO while cardiac expression of inflammatory markers was not affected. Moreover, administration of rosiglitazone, a PPARγ agonist, reduces mortality and improves overall frailty of septic mice. The beneficial effects of rosiglitazone are associated with improved cardiac FAO and reduced mitophagy. In conclusion, activation of PPARγ in animals with sepsis increases myocardial energy production and improves cardiac function and survival.

Acknowledgments

We thank Zoi Drosatos-Tampakaki for the EM figures’ analysis, and Yunying Hu and Shuiqing Yu for animal care.

Sources of Funding

This study was supported by National Heart, Lung, and Blood Institute (NHLBI) Grants HL45095, 73029 (I.J.G), “Pathway to Independence” Award 1K99HL112853-01 by the NHLBI (K.D.), a postdoctoral grant of the American Heart Association (AHA #10POST4440032) (KD) and T32 HL007343 (RK).

Footnotes

Disclosures

None.

Journal Subject Codes : [107] Biochemistry and metabolism, [130] Animal models of human disease

This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

References

  • 1.Angus DC, Linde-Zwirble WT, Lidicker J, Clermont G, Carcillo J, Pinsky MR. Epidemiology of severe sepsis in the united states: Analysis of incidence, outcome, and associated costs of care. Crit Care Med. 2001;29:1303–1310. doi: 10.1097/00003246-200107000-00002. [DOI] [PubMed] [Google Scholar]
  • 2.Drosatos K, Drosatos-Tampakaki Z, Khan R, Homma S, Schulze PC, Zannis VI, Goldberg IJ. Inhibition of c-jun-n-terminal kinase increases cardiac ppar{alpha} expression and fatty acid oxidation and prevents lps-induced heart dysfunction. J Biol Chem. 2011;286:36331–36339. doi: 10.1074/jbc.M111.272146. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3.Schilling J, Lai L, Sambandam N, Dey CE, Leone TC, Kelly DP. Toll-like receptor-mediated inflammatory signaling reprograms cardiac energy metabolism by repressing peroxisome proliferator-activated receptor γ coactivator-1 signaling. Circ Heart Fail. 2011;4:474–482. doi: 10.1161/CIRCHEARTFAILURE.110.959833. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Liu MS, Spitzer JJ. Fatty acid and lactate metabolism by heart homogenates from alloxan-diabetic dogs. Horm Metab Res. 1978;10:114–117. doi: 10.1055/s-0028-1093455. [DOI] [PubMed] [Google Scholar]
  • 5.Wang X, Evans RD. Effect of endotoxin and platelet-activating factor on lipid oxidation in the rat heart. J Mol Cell Cardiol. 1997;29:1915–1926. doi: 10.1006/jmcc.1997.0430. [DOI] [PubMed] [Google Scholar]
  • 6.Tessier JP, Thurner B, Jungling E, Luckhoff A, Fischer Y. Impairment of glucose metabolism in hearts from rats treated with endotoxin. Cardiovasc Res. 2003;60:119–130. doi: 10.1016/s0008-6363(03)00320-1. [DOI] [PubMed] [Google Scholar]
  • 7.Carlson DL, Willis MS, White DJ, Horton JW, Giroir BP. Tumor necrosis factor-alpha-induced caspase activation mediates endotoxin-related cardiac dysfunction. Crit Care Med. 2005;33:1021–1028. doi: 10.1097/01.ccm.0000163398.79679.66. [DOI] [PubMed] [Google Scholar]
  • 8.Fallach R, Shainberg A, Avlas O, Fainblut M, Chepurko Y, Porat E, Hochhauser E. Cardiomyocyte toll-like receptor 4 is involved in heart dysfunction following septic shock or myocardial ischemia. J Mol Cell Cardiol. 2010;48:1236–1244. doi: 10.1016/j.yjmcc.2010.02.020. [DOI] [PubMed] [Google Scholar]
  • 9.Knuefermann P, Nemoto S, Misra A, Nozaki N, Defreitas G, Goyert SM, Carabello BA, Mann DL, Vallejo JG. Cd14-deficient mice are protected against lipopolysaccharide-induced cardiac inflammation and left ventricular dysfunction. Circulation. 2002;106:2608–2615. doi: 10.1161/01.cir.0000038110.69369.4c. [DOI] [PubMed] [Google Scholar]
  • 10.Doi K, Leelahavanichkul A, Yuen PS, Star RA. Animal models of sepsis and sepsis-induced kidney injury. J Clin Invest. 2009;119:2868–2878. doi: 10.1172/JCI39421. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11.Ohlsson K, Bjork P, Bergenfeldt M, Hageman R, Thompson RC. Interleukin-1 receptor antagonist reduces mortality from endotoxin shock. Nature. 1990;348:550–552. doi: 10.1038/348550a0. [DOI] [PubMed] [Google Scholar]
  • 12.Feingold K, Kim MS, Shigenaga J, Moser A, Grunfeld C. Altered expression of nuclear hormone receptors and coactivators in mouse heart during the acute-phase response. Am J Physiol Endocrinol Metab. 2004;286:E201–207. doi: 10.1152/ajpendo.00205.2003. [DOI] [PubMed] [Google Scholar]
  • 13.Banke NH, Wende AR, Leone TC, O’Donnell JM, Abel ED, Kelly DP, Lewandowski ED. Preferential oxidation of triacylglyceride-derived fatty acids in heart is augmented by the nuclear receptor pparalpha. Circ Res. 2010;107:233–241. doi: 10.1161/CIRCRESAHA.110.221713. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Finck BN, Lehman JJ, Leone TC, Welch MJ, Bennett MJ, Kovacs A, Han X, Gross RW, Kozak R, Lopaschuk GD, Kelly DP. The cardiac phenotype induced by pparalpha overexpression mimics that caused by diabetes mellitus. J Clin Invest. 2002;109:121–130. doi: 10.1172/JCI14080. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Son NH, Yu S, Tuinei J, Arai K, Hamai H, Homma S, Shulman GI, Abel ED, Goldberg IJ. Ppargamma-induced cardiolipotoxicity in mice is ameliorated by pparalpha deficiency despite increases in fatty acid oxidation. J Clin Invest. 2010;120:3443–3454. doi: 10.1172/JCI40905. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Cheng L, Ding G, Qin Q, Huang Y, Lewis W, He N, Evans RM, Schneider MD, Brako FA, Xiao Y, Chen YE, Yang Q. Cardiomyocyte-restricted peroxisome proliferator-activated receptor-delta deletion perturbs myocardial fatty acid oxidation and leads to cardiomyopathy. Nat Med. 2004;10:1245–1250. doi: 10.1038/nm1116. [DOI] [PubMed] [Google Scholar]
  • 17.Son NH, Park TS, Yamashita H, Yokoyama M, Huggins LA, Okajima K, Homma S, Szabolcs MJ, Huang LS, Goldberg IJ. Cardiomyocyte expression of ppargamma leads to cardiac dysfunction in mice. J Clin Invest. 2007;117:2791–2801. doi: 10.1172/JCI30335. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.Vega RB, Huss JM, Kelly DP. The coactivator pgc-1 cooperates with peroxisome proliferator-activated receptor alpha in transcriptional control of nuclear genes encoding mitochondrial fatty acid oxidation enzymes. Mol Cell Biol. 2000;20:1868–1876. doi: 10.1128/mcb.20.5.1868-1876.2000. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Lehman JJ, Barger PM, Kovacs A, Saffitz JE, Medeiros DM, Kelly DP. Peroxisome proliferator-activated receptor gamma coactivator-1 promotes cardiac mitochondrial biogenesis. J Clin Invest. 2000;106:847–856. doi: 10.1172/JCI10268. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20.Maitra U, Chang S, Singh N, Li L. Molecular mechanism underlying the suppression of lipid oxidation during endotoxemia. Mol Immunol. 2009;47:420–425. doi: 10.1016/j.molimm.2009.08.023. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.Bagby GJ, Spitzer JA. Lipoprotein lipase activity in rat heart and adipose tissue during endotoxic shock. Am J Physiol. 1980;238:H325–330. doi: 10.1152/ajpheart.1980.238.3.H325. [DOI] [PubMed] [Google Scholar]
  • 22.Lu B, Moser A, Shigenaga JK, Grunfeld C, Feingold KR. The acute phase response stimulates the expression of angiopoietin like protein 4. Biochem Biophys Res Commun. 2010;391:1737–1741. doi: 10.1016/j.bbrc.2009.12.145. [DOI] [PubMed] [Google Scholar]
  • 23.Kaufmann RL, Matson CF, Beisel WR. Hypertriglyceridemia produced by endotoxin: Role of impaired triglyceride disposal mechanisms. J Infect Dis. 1976;133:548–555. doi: 10.1093/infdis/133.5.548. [DOI] [PubMed] [Google Scholar]
  • 24.Gouni I, Oka K, Etienne J, Chan L. Endotoxin-induced hypertriglyceridemia is mediated by suppression of lipoprotein lipase at a post-transcriptional level. J Lipid Res. 1993;34:139–146. [PubMed] [Google Scholar]
  • 25.Sukonina V, Lookene A, Olivecrona T, Olivecrona G. Angiopoietin-like protein 4 converts lipoprotein lipase to inactive monomers and modulates lipase activity in adipose tissue. Proc Natl Acad Sci U S A. 2006;103:17450–17455. doi: 10.1073/pnas.0604026103. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Wang H, Sreenevasan U, Hu H, Saladino A, Polster BM, Lund LM, Gong DW, Stanley WC, Sztalryd C. Perilipin 5, a lipid droplet-associated protein, provides physical and metabolic linkage to mitochondria. J Lipid Res. 2011;52:2159–2168. doi: 10.1194/jlr.M017939. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Guyton K, Zingarelli B, Ashton S, Teti G, Tempel G, Reilly C, Gilkeson G, Halushka P, Cook J. Peroxisome proliferator-activated receptor-gamma agonists modulate macrophage activation by gram-negative and gram-positive bacterial stimuli. Shock. 2003;20:56–62. doi: 10.1097/01.shk.0000070903.21762.f8. [DOI] [PubMed] [Google Scholar]
  • 28.Yagyu H, Chen G, Yokoyama M, Hirata K, Augustus A, Kako Y, Seo T, Hu Y, Lutz EP, Merkel M, Bensadoun A, Homma S, Goldberg IJ. Lipoprotein lipase (lpl) on the surface of cardiomyocytes increases lipid uptake and produces a cardiomyopathy. J Clin Invest. 2003;111:419–426. doi: 10.1172/JCI16751. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 29.Chiu HC, Kovacs A, Blanton RM, Han X, Courtois M, Weinheimer CJ, Yamada KA, Brunet S, Xu H, Nerbonne JM, Welch MJ, Fettig NM, Sharp TL, Sambandam N, Olson KM, Ory DS, Schaffer JE. Transgenic expression of fatty acid transport protein 1 in the heart causes lipotoxic cardiomyopathy. Circ Res. 2005;96:225–233. doi: 10.1161/01.RES.0000154079.20681.B9. [DOI] [PubMed] [Google Scholar]
  • 30.Chiu HC, Kovacs A, Ford DA, Hsu FF, Garcia R, Herrero P, Saffitz JE, Schaffer JE. A novel mouse model of lipotoxic cardiomyopathy. J Clin Invest. 2001;107:813–822. doi: 10.1172/JCI10947. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31.Drosatos K, Bharadwaj KG, Lymperopoulos A, Ikeda S, Khan R, Hu Y, Agarwal R, Yu S, Jiang H, Steinberg SF, Blaner WS, Koch WJ, Goldberg IJ. Cardiomyocyte lipids impair beta-adrenergic receptor function via pkc activation. Am J Physiol Endocrinol Metab. 2011;300:E489–499. doi: 10.1152/ajpendo.00569.2010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.Park TS, Hu Y, Noh HL, Drosatos K, Okajima K, Buchanan J, Tuinei J, Homma S, Jiang XC, Abel ED, Goldberg IJ. Ceramide is a cardiotoxin in lipotoxic cardiomyopathy. J Lipid Res. 2008;49:2101–2112. doi: 10.1194/jlr.M800147-JLR200. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33.Garesse R, Vallejo CG. Animal mitochondrial biogenesis and function: A regulatory cross-talk between two genomes. Gene. 2001;263:1–16. doi: 10.1016/s0378-1119(00)00582-5. [DOI] [PubMed] [Google Scholar]
  • 34.Larsson NG, Wang J, Wilhelmsson H, Oldfors A, Rustin P, Lewandoski M, Barsh GS, Clayton DA. Mitochondrial transcription factor a is necessary for mtdna maintenance and embryogenesis in mice. Nat Genet. 1998;18:231–236. doi: 10.1038/ng0398-231. [DOI] [PubMed] [Google Scholar]
  • 35.Wu Z, Puigserver P, Andersson U, Zhang C, Adelmant G, Mootha V, Troy A, Cinti S, Lowell B, Scarpulla RC, Spiegelman BM. Mechanisms controlling mitochondrial biogenesis and respiration through the thermogenic coactivator pgc-1. Cell. 1999;98:115–124. doi: 10.1016/S0092-8674(00)80611-X. [DOI] [PubMed] [Google Scholar]
  • 36.Riehle C, Wende AR, Zaha VG, Pires KM, Wayment B, Olsen C, Bugger H, Buchanan J, Wang X, Moreira AB, Doenst T, Medina-Gomez G, Litwin SE, Lelliott CJ, Vidal-Puig A, Abel ED. Pgc-1beta deficiency accelerates the transition to heart failure in pressure overload hypertrophy. Circ Res. 2011;109:783–793. doi: 10.1161/CIRCRESAHA.111.243964. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Tolkovsky AM. Mitophagy. Biochim Biophys Acta. 2009;1793:1508–1515. doi: 10.1016/j.bbamcr.2009.03.002. [DOI] [PubMed] [Google Scholar]
  • 38.Yuan H, Perry CN, Huang C, Iwai-Kanai E, Carreira RS, Glembotski CC, Gottlieb RA. Lps-induced autophagy is mediated by oxidative signaling in cardiomyocytes and is associated with cytoprotection. Am J Physiol Heart Circ Physiol. 2009;296:H470–479. doi: 10.1152/ajpheart.01051.2008. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39.Levy MM, Fink MP, Marshall JC, Abraham E, Angus D, Cook D, Cohen J, Opal SM, Vincent JL, Ramsay G. 2001 sccm/esicm/accp/ats/sis international sepsis definitions conference. Crit Care Med. 2003;31:1250–1256. doi: 10.1097/01.CCM.0000050454.01978.3B. [DOI] [PubMed] [Google Scholar]
  • 40.Neubauer S. The failing heart--an engine out of fuel. N Engl J Med. 2007;356:1140–1151. doi: 10.1056/NEJMra063052. [DOI] [PubMed] [Google Scholar]
  • 41.Cha BS, Ciaraldi TP, Park KS, Carter L, Mudaliar SR, Henry RR. Impaired fatty acid metabolism in type 2 diabetic skeletal muscle cells is reversed by ppargamma agonists. Am J Physiol Endocrinol Metab. 2005;289:E151–159. doi: 10.1152/ajpendo.00141.2004. [DOI] [PubMed] [Google Scholar]
  • 42.Abdelrahman M, Sivarajah A, Thiemermann C. Beneficial effects of ppar-gamma ligands in ischemia-reperfusion injury, inflammation and shock. Cardiovasc Res. 2005;65:772–781. doi: 10.1016/j.cardiores.2004.12.008. [DOI] [PubMed] [Google Scholar]
  • 43.Finck BN, Kelly DP. Pgc-1 coactivators: Inducible regulators of energy metabolism in health and disease. J Clin Invest. 2006;116:615–622. doi: 10.1172/JCI27794. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 44.Suliman HB, Welty-Wolf KE, Carraway M, Tatro L, Piantadosi CA. Lipopolysaccharide induces oxidative cardiac mitochondrial damage and biogenesis. Cardiovascular research. 2004;64:279–288. doi: 10.1016/j.cardiores.2004.07.005. [DOI] [PubMed] [Google Scholar]
  • 45.Kanamori H, Takemura G, Maruyama R, Goto K, Tsujimoto A, Ogino A, Li L, Kawamura I, Takeyama T, Kawaguchi T, Nagashima K, Fujiwara T, Fujiwara H, Seishima M, Minatoguchi S. Functional significance and morphological characterization of starvation-induced autophagy in the adult heart. Am J Pathol. 2009;174:1705–1714. doi: 10.2353/ajpath.2009.080875. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 46.Takagi H, Matsui Y, Hirotani S, Sakoda H, Asano T, Sadoshima J. Ampk mediates autophagy during myocardial ischemia in vivo. Autophagy. 2007;3:405–407. doi: 10.4161/auto.4281. [DOI] [PubMed] [Google Scholar]
  • 47.de Montmollin E, Annane D. Year in review 2010: Critical care--multiple organ dysfunction and sepsis. Crit Care. 2011;15:236. doi: 10.1186/cc10359. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 48.Kaul S, Bolger AF, Herrington D, Giugliano RP, Eckel RH. Thiazolidinedione drugs and cardiovascular risks: A science advisory from the american heart association and american college of cardiology foundation. Circulation. 2010;121:1868–1877. doi: 10.1161/CIR.0b013e3181d34114. [DOI] [PubMed] [Google Scholar]
  • 49.Guan Y, Hao C, Cha DR, Rao R, Lu W, Kohan DE, Magnuson MA, Redha R, Zhang Y, Breyer MD. Thiazolidinediones expand body fluid volume through ppargamma stimulation of enac-mediated renal salt absorption. Nat Med. 2005;11:861–866. doi: 10.1038/nm1278. [DOI] [PubMed] [Google Scholar]
  • 50.Marfella R, Portoghese M, Ferraraccio F, Siniscalchi M, Babieri M, Di Filippo C, D’Amico M, Rossi F, Paolisso G. Thiazolidinediones may contribute to the intramyocardial lipid accumulation in diabetic myocardium: Effects on cardiac function. Heart. 2009;95:1020–1022. doi: 10.1136/hrt.2009.165969. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

1
Download video file (6.6MB, mp4)
2
Download video file (6.8MB, mp4)
3
Download video file (6.5MB, mp4)
4
Download video file (6.6MB, mp4)
5
Download video file (6.5MB, mp4)
6
Download video file (6.7MB, mp4)
Supp
NIHMS481820-supplement-Supp.pdf (857.9KB, pdf)

RESOURCES