![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
|
|
|
|
|
||
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
CARDIOVASCULAR
Departments of Cardiology (A.A., J.W., C.B., C.H.) and Environmental Science (K.K.), University of Freiburg, Freiburg, Germany
Received November 15, 2007; accepted February 25, 2008.
 |
Abstract |
|---|
 Top Abstract Materials and MethodsResultsDiscussionReferences |
|---|
and 2- during reoxygenation. Overall, these results indicate that moderate doses of oxPFCs can improve cell survival during reoxygenation, causing deactivation of DAPK, up-regulation of AMPK, and HIF1- and 2- stabilization. These effects of oxPFCs are dose-dependent, and they lead to a stabilization of the mitochondrial membrane potential, decreased steady-state levels of superoxide, and pacification of mitochondrial activity.
). Oxygen, the final electron acceptor in oxidative phosphorylation, maintains ATP synthesis in the inner membrane of the mitochondrion. In contrast, oxidative phosphorylation is a central site of reactive oxygen species production in the heart (Cai and Jones, 1998; Chandel et al., 2000; Yellon and Hausenloy, 2007) that may induce irreversible cellular damage and cell death (Davies, 1995). Therefore, mitochondria have been recognized as central integration site at the crossroads of either cell survival or cell death (Murphy, 2004). Alterations of oxidative phosphorylation in the mitochondrion have a key function in cardiomyocyte survival after ischemia/reperfusion. Accordingly, opposite effects of oxygen delivery during reperfusion and increased ATP availability have been reported, including increased cell viability but also increased apoptosis (Weiss et al., 2003). These data raise the question of how the concept of normoxic reperfusion can be improved considering the fact that it triggers apoptosis.
Proapoptotic death-associated protein kinase (DAPK) is activated early during apoptosis (Schumacher et al., 2002a), and its inhibition is protective after ischemia of the brain (Velentza et al., 2003). It has been previously shown that DAPK is activated in human neuroblastoma cells by mitochondrial respiratory chain inhibitors that induce a decrease in the mitochondrial membrane potential and an increased steady-state level of superoxide (Shang et al., 2005). However, it is unclear, whether oxygen can influence DAPK expression by changing the mitochondrial membrane potential or by increasing superoxide production.
Antiapoptotic AMP-activated protein kinase (AMPK) serves as an indicator of ATP availability in cells, detecting changes in the AMP:ATP ratio and signaling an inadequate ATP supply by oxidative phosphorylation (Hardie et al., 1998; Baron et al., 2005). AMPK initiates a metabolic stress response that protects human cardiomyocytes against cell death (Spector et al., 2007), but it has never been shown whether AMPK activity is affected by an gradual increase in oxygen concentration.
HIF1- and 2- are both known to be expressed in ischemic cardiomyocytes (Jürgensen et al., 2004), and stabilization of HIF1- is known to be a cardiac survival factor during hypoxia (Lee et al., 2000). However, it is unclear whether factors other than NO production by inducible nitric-oxide synthase mimic a hypoxic response under normoxia and could stabilize HIF during reoxygenation (Sandau et al., 2001).
Chemically inert perfluorochemicals (PFCs) provide a unique tool for controlled delivery of oxygen (Rafikova et al., 2004). Perfluorochemicals possess a high oxygen-dissolving capacity that follows Henry's law, leading to oxygen solubility that is directly proportional to oxygen partial pressure. As a result, oxygen can be extracted rapidly and extensively from PFCs (Riess, 2005). Determining the conditions under which oxygen delivery by PFCs improves survival of ischemic cells and thereby preventing apoptosis in the surviving ischemic cells could result in an optimized regime of oxygen delivery during reperfusion.
Therefore, the aim of this study was to characterize the effects of delivering different dosing regimes of oxygenated (ox) PFCs with regard to changes in cell viability, oxidative phosphorylation, mitochondrial membrane potential, and superoxide production, and in particular to evaluate of the role of DAPK, AMPK, and HIF1- and 2- during reoxygenation.
|
Materials and Methods |
|---|
|
TopAbstractMaterials and MethodsResultsDiscussionReferences |
|---|
antibody was obtained from Cell Signaling Technology (Beverly, MA) and anti-HIF2- antibody was from Novus Biologicals, Inc. (Littleton, CO). Anti-phospho-AMPK-1-Thr-172 and anti-AMPK- antibodies were obtained from Cell Signaling Technology Inc. Anti-cardiac troponin I antibody was obtained from Accurate Chemical & Scientific (Westbury, NY) and anti-phospho-DAPK-Ser-308 antibody from Sigma-Aldrich (St. Louis, MO). Anti-DAPK was obtained from BD Biosciences Transduction Laboratories (Lexington, KY), and anti-manganese superoxide dismutase (MnSOD) antibody was obtained from Millipore Corporation (Billerica, MA).
Culture of Human Cardiomyocytes. Monolayer cultures of human cardiomyocytes were prepared by modifying the method of Merante et al. (1998). In brief, heart tissue from the right atrium was obtained from male Caucasian patients with an average age of 76 years undergoing cardiovascular bypass surgery. The tissue was carefully dissected and further softened by digestion with 0.1% collagenase VII. Tissue specimens were washed in PBS and cultured in smooth muscle growth medium (PromoCell). After outgrowth of cells, cardiomyocyte colonies were transferred to new culture dishes. Cultures exhibiting >95% purity, as assessed by visual monitoring for rod-shaped cells and by fluorescent monoclonal antibody staining for troponin I during the first two passages, were used for subsequent studies.
Model of Hypoxia and Reoxygenation. Ischemia was mimicked by exposing cardiomyocytes (seeding density 3000 cells/cm2)to a low-oxygen atmosphere (pO2 < 40 mm Hg). Cardiomyocytes showed the most pronounced decrease of cell viability after increasing the period of hypoxia to 12 h. Hypoxia was maintained within the cell culture incubator by continuous infusion with a mixture of 95% N2/5% CO2 in an airtight chamber for a period of 12 h before reoxygenation. Just before reoxygenation, pure perfluorooctyl bromide was ventilated with 100% carbogen (95% oxygen, 5% carbon dioxide) and was kept in an airtight vessel. Cardiomyocytes were reoxygenated with different doses of 100% perfluorooctyl bromide that were added to the cell medium. Because of its high density and its highly hydrophobic nature perfluorooctyl bromide formed a film over the cardiomyocyte monolayer in the cell culture dishes. This perfluorooctyl bromide film was separated from the normoxic atmosphere by the overlying cell medium. Perfluorooctyl bromide was removed after 10 min, and dishes were incubated for additional 1, 3, and 6 h in normoxia (pO2 = 150 mm Hg), 5% CO2, and humidified atmosphere within the cell culture incubator. Each sample was run with a control that received reoxygenation entirely at normoxic conditions.
Volumetric Dosing of Perfluorooctyl Bromide. Oxygen solubility of 50 vol.% means that 1 volume of oxygen is dissolved in 2 volumes of perfluorooctyl bromide when 100% perfluorooctyl bromide is flushed with 
95% oxygen for 5 min at room temperature (Riess, 2005). The oxygen-carrying capacity of the perfluorooctyl bromide-supplemented cell medium obeys Henry's law of partial pressures (c [mol · l–1] = KO2·p [bar]), with KO2 being the oxygen specific Henry's law constant (KO2 = 1.3 x 10–3[mol · l–1 · bar–1]). We added 500 and 2000 µmol · l–1 oxygen to the already present 250 µmol · l–1 oxygen in the cell culture medium. Because 22.4 liters of oxygen make 1 mol, 1 mmol · l–1 oxygen equals approximately 25 µl of oxygen per milliliter of cells. Therefore, we added 18.75, 37.5, and 150 µl of oxygen dissolved in 39.5, 75, and 300 µl of oxPFCs in 3 ml of medium to cardiomyocytes cultured in a 60-mm dish (21.5 cm2). When we applied 6.25, 12.5, and 50 µl of oxygen per ml of cell medium, we increased the pO2 to superphysiological levels of 160, 320, and 1300 mm Hg. These oxygen levels of the cell medium were greater than the arterial plasma oxygen concentration (3 µl of oxygen per ml of plasma). The increase of oxygen in the cell culture medium was verified with a GMH 3610 analyzer (Greisinger Electronics, Regenstauf, Germany) for dissolved oxygen.
Measurement of Lactate Dehydrogenase. Lactate dehydrogenase (LDH) activity was measured in culture medium 6 h after reoxygenation. LDH was analyzed with the International Federation of Clinical Chemistry-recommended method (Schumann et al., 2002) using an LDH International Federation of Clinical Chemistry enzymatic reagent kit from Rolf Greiner BioChemica (Flacht, Germany). In brief, oxidation of lactate to pyruvate by LDH was measured as rate of increase in absorbance at 340 nm at 37°C.
Cell Viability Staining. Cell viability was analyzed by fluorescent staining as reported by Foglieni et al. (2001). In brief, we used a triple dye combination of acridine orange, Hoechst 33342, and propidium iodide that visualizes all cell viability states simultaneously on the basis of nuclear and cytoplasmatic signals. Consecutively, cardiomyocytes were incubated in 20 µM Hoechst 33342 for 20 min, in 0.67 nM acridine orange for 10 min and in 3 nM propidium iodide for 5 min at 37°C. Before and after each staining step, cells were washed with PBS. After fluorescent staining, cardiomyocytes were visualized with an Axioplan 2 imaging fluorescence microscopy under a long-pass fluorescein isothiocyanate filter from Zeiss (Goettingen, Germany). Healthy cells showed green colored cytoplasms; early apoptotic cells dark rust-yellow cytoplasms; necrotic cells showed orange cytoplasms. Axiovision 3.1 software and KS 300 3.0 software, both from Carl Zeiss (Goettingen, Germany), were used for quantification.
Mitochondrial Membrane Potential Visualization. Mitochondrial potential was assessed by the fluorescent dye MitoTracker Red 7513 (reduced chloromethyl-X-rosamine) as described previously by Gurevich et al. (2001). In brief, cardiomyocytes were incubated in cell medium supplemented with 0.5 µM chloromethyl-X-rosamine for 45 min within the cell culture incubator. Cells were fixed in 3.7% formalin for 15 min and in ice-cold acetone for 5 min. Before and after each staining step, cells were washed intensely with PBS. Axiovision 3.1 software and KS 300 3.0 software, both obtained from Carl Zeiss, were used for quantification.
Western Blot Analysis. Cardiomyocytes were washed and scraped off in ice-cold PBS supplemented with 
protease inhibitor cocktail and 1 mM phenylmethylsulfonyl fluoride. The cells were centrifuged for 10 min at 220g, and then they were lysed using the MEM-PER eukaryotic membrane protein extraction kit from Pierce Chemical (Rockford, IL), according to the manufacturer's instructions. The lysate was centrifuged for 10 min at 10,000g at 4°C. From the clear supernatants, protein concentration was determined by the Bio-Rad protein assay kit (Bio-Rad, Munich, Germany) using BSA as control. Protein aliquots were boiled for 3 min in 2x loading buffer and separated by electrophoresis on 7.5% SDS-polyacrylamide gel electrophoresis. Proteins were blotted onto polyvinylidene fluoride membranes (Millipore, Schwalbach, Germany), and then they were immunoblotted overnight with the indicated primary antibody followed by secondary antibody conjugated with alkaline phosphatase. The CDP-Star Reagent from New England Biolabs (Beverly, MA) was used for detection.
Superoxide Chemiluminescence. Superoxide production was analyzed by lucigenin chemiluminescence. In brief, after incubating cardiomyocytes with 50 mM diethyldithiocarbamate for 10 min (Omar et al., 1991), cells were analyzed in HEPES buffer containing lucigenin that emits light on reduction of superoxide (Pagano et al., 1995). Chemiluminescence was determined after a 2-s delay for 20 min, integrated over a 30-s period, and repeated every 2 min using a TD-20/20 luminometer (Turner Designs, Sunnyvale, CA).
Immunohistochemistry. Consecutively, cardiomyocytes were incubated in 1% formalin at 4°C overnight, in 0.2% Triton X-100 for 5 min, blocked in 5% BSA for 1 h, and finally stained in 5% BSA with the indicated primary antibody (1:25) for 1 h and fluorescence-labeled secondary antibody (1:200) for 4 h. Before and after each staining step, cells were washed with PBS. After fluorescent staining, cardiomyocytes were visualized with an Axioplan 2 imaging fluorescence microscopy under a fluorescein isothiocyanate filter from Carl Zeiss. Axiovision 3.1 software and KS 300 3.0 software, both from Carl Zeiss, were used for quantification.
ATP Determination. ATP was measured by the luciferin/luciferase method with a chemiluminescence kit (Promega) following the manufacturer's protocol. Chemiluminescence was determined in a TD-20/20 luminometer (Turner Designs), and data were analyzed in Excel (Microsoft, Redmond, WA). Alternatively, ATP was extracted from cardiomyocytes after reoxygenation treatment, separated by high-performance liquid chromatography, and quantified as described by Matoba et al. (1999).
Statistical Analysis. Data are normalized for the amount of protein or number of cells engaged. All experiments were repeated at least three times. Data are given as percentage from normoxic value, which was set at 100%, and they are presented as mean ± S.E. Comparisons were made using unpaired Student's t test or one-way analysis of variance when appropriate. Results were considered to be significantly different when P < 0.05.
|
Results |
|---|
|
TopAbstractMaterials and MethodsResultsDiscussionReferences |
|---|
15%; P < 0.05) after reoxygenation with oxygenated PFCs in moderate concentrations (750 µMO2 in 0.1 µM PFCs). After exposing ischemic cardiomyocytes to reoxygenation with oxygenated PFCs in moderate concentrations (750 µMO2 in 0.1 µM PFCs) and in high concentrations (2250 µMO2 in 0.4 µM PFCs) and to normoxic reoxygenation (250 µMO2), we investigated the effect of supplemental oxygen delivery on cell viability during reoxygenation (Fig. 3). Ischemic cells show a significant increase in cell viability ( 30% at 1 and 3 h and 20% at 6 h; P < 0.05), when O2 concentrations were elevated transiently to 750 µMO2 dissolved in 0.1 µM PFCs (Fig. 4). No further increase was seen when hypoxic cardiomyocytes were reoxygenated with a higher O2 concentration of 2250 µMO2 dissolved in 0.4 µM PFCs (Fig. 5). On the contrary, ischemic cardiomyocytes reoxygenated with 2250 µMO2 in 0.4 µM PFCs showed a significant decrease in cell viability (40% at 3 h and 100% at 6 h; P < 0.05).
|
|
|
|
|
Deactivation of DAPK by Moderate Doses of oxPFCs during Reoxygenation. An increase in mitochondrial membrane potential after reoxygenation with addition of moderate concentrations of oxPFCs (750 µMO2 in 0.1 µM PFCs) compared with normoxic reoxygenation seemed to be mediated by phosphorylation, and, consequently, by deactivation of DAPK. As shown in Fig. 6A, phosphorylated DAPK (pDAPK) was found almost at background level at normoxia (250 µMO2) and maximal after addition of 750 µMO2 in a moderate dosage of oxPFCs (P < 0.05). In contrast to this finding, DAPK phosphorylation did not increase significantly, when ischemic cardiomyocytes were exposed to 2250 µMO2 by a high dosage of oxPFCs (Fig. 6B).
|
Stabilization of Mitochondrial Membrane Potential after oxPFC-Supplemented Reoxygenation. After normoxic reoxygenation (250 µMO2), a loss of mitochondrial membrane potential occurred as redistribution of the fluorochrome chloromethyl-X-rosamine from mitochondria to cytosol (Fig. 7A). When cardiomyocytes were exposed to oxPFCs in moderate concentrations (750 µMO2 in 0.1 µM PFCs), mitochondrial membrane potential stabilized and cardiomyocytes displayed a mitochondrion-selective fluorochrome staining (Fig. 6B). An increase in mitochondrial membrane potential as well as a diminished loss of mitochondrial membrane potential was observed, when O2 concentrations were elevated to 750 µMO2 by 0.1 µM oxPFCs (Fig. 7C). Six hours after O2 concentrations were moderately elevated, the stabilization of the mitochondrial membrane potential was significant (40%; P < 0.01) compared with normoxic reoxygenation.
|
15% at 3 h and 20% at 6 h; P < 0.05) compared with normoxic reoxygenation. This decrease disappeared when hypoxic cardiomyocytes were reoxygenated with 2250 µMO2 in 0.4 µM PFCs (Fig. 8B). Hypoxic cardiomyocytes reoxygenated with a concentration of 2250 µMO2 showed a cumulative increase in superoxide production that became significant at 6 h (50%; P < 0.05).
|
Mitochondrial Manganese Superoxide Dismutase Expression Decreases after Reoxygenation with Addition of oxPFCs in Moderate Concentrations. After reoxygenation applying 750 µMO2 in 0.1 µM PFCs, mitochondrial MnSOD expression decreased significantly in comparison with normoxic reoxygenation (26%; P < 0.05). When ischemic cardiomyocytes were reoxygenated with hyperoxic oxygen concentrations of 2250 µMO2, MnSOD expression was increased (154%; P < 0.05; Fig. 9) compared with normoxic reoxygenation.
|
2-fold; P < 0.01) after treatment of ischemic cardiomyocytes with oxPFCs in moderate concentrations (750 µM O2). However, oxPFCs (0.4 µM) delivering oxygen in the toxic concentration of 2250 µMO2 did not increase but decreased AMPK activation significantly (30%; P < 0.01), compared with normoxic reoxygenation (Fig. 10B).
|
Based on the results of AMPK activation, we investigated whether the observed increase of AMPK activation with oxPFCs in moderate concentrations (750 µM O2 in 0.1 µM PFCs) correlated with changes in cellular metabolism, compared with normoxic reoxygenation. We detected a small (15%) but significant (P < 0.05) decrease in ATP content of cardiomyocytes after transient exposure to moderate concentrations of oxPFCs delivering 750 µMO2.
Moderate Doses of oxPFCs during Reoxygenation Stabilize HIF1- and HIF2-. We investigated whether a transient treatment of ischemic cardiomyocytes with hyperoxia induced a change in HIF1- and HIF2- stabilization compared with normoxic reoxygenation (250 µMO2). One hour after reoxygenation with a brief hyperoxia of 750 µM O2, cardiomyocytes showed a weak HIF1- expression signal (data not shown) that became prominent 3 h after reoxygenation with 750 µMO2. Three hours after the oxygen concentration was elevated transiently to 750 µM O2, ischemic cardiomyocytes showed a significant increase in HIF1- stabilization (45%; P < 0.05), as shown in Fig. 11A. This increase in HIF1- stabilization disappeared 6 h after reoxygenation with 750 µMO2, when the HIF1- signal was not visible any longer. A particularly intense signal was obtained for HIF2- after applying 750 µMO2 (130%; P < 0.05), as shown in Fig. 11B. However, when ischemic cardiomyocytes were reoxygenated with the higher oxygen concentration of 2250 µMO2, HIF1- stabilization was not detectable at any time (data not shown).
|
|
Discussion |
|---|
|
TopAbstractMaterials and MethodsResultsDiscussionReferences |
|---|
), but further studies using different administration routes and carrier doses resulted in conflicting data on infarct reduction (Wall et al., 1994). Recent studies using intracoronary oxygen perfusion with an aqueous oxygen solution have been reported to reduce infarct size and to improve myocardial contractility (Glazier, 2005). Although promising, reperfusion strategies that supplement oxygen require fine-tuning to gain the desired cardioprotective effect and to limit toxic side effects. So far, it has never been specifically addressed how reoxygenation regimes differing in oxygen concentrations influence cell viability in ischemic cardiomyocytes.
We have found that the cell viability of human cardiomyocytes after prolonged ischemia depends on the regime of reoxygenation. Although reoxygenation with oxPFCs in high concentrations results in decreased cell viability compared with reoxygenation at normoxic conditions, reoxygenation with oxPFCs in moderate concentrations improves cell survival significantly by reducing necrosis and apoptosis. Our experiments show that the regenerative effect of oxPFC is a dose-dependent oxygen effect, whereas addition of PFCs alone had no effect on cell survival.
Proapoptotic DAPK has been suggested to serve as a sensor of mitochondrial membrane potential because mitochondrial respiratory chain inhibitors cause DAPK activation by decreasing mitochondrial membrane potential and increasing steady-state levels of superoxide (Shang et al., 2005). Furthermore, DAPK plays an important role during the recovery phase after brain injury (Schumacher et al., 2002b), and it has been shown that a selective DAPK inhibitor is neuroprotective in a model of brain ischemia (Shamloo et al., 2005). We show that an increase in mitochondrial membrane potential after reoxygenation with moderate doses of oxPFCs may cause deactivation of DAPK, but normoxic reoxygenation does not alter DAPK, although this result is methodically limited by a change of cellular milieu by the fluorescent dye chloromethyl-X-rosamine. Toxic doses of oxPFCs lead to an increase in apoptosis and reduced viability of cardiomyocytes, and they do not deactivate DAPK. Thus, the dosing regime of oxygen is an important modulator of cell viability during reoxygenation.
In addition, our experiments show that the superoxide production of human cardiomyocytes after hypoxia depends on the regime of reoxygenation. Reoxygenation with toxic doses of oxPFCs results in a cumulative increase in superoxide production compared with normoxic reoxygenation. However, moderate doses of oxPFCs lead to a decrease in steady-state levels of superoxide. Our results underscore the "oxygen paradox," the mitochondrial electron-transport chain of higher eukaryotic aerobic organisms generates superoxide that seems to be responsible for oxygen toxicity (Davies, 1995).
MnSOD is the principal antioxidant enzyme located in the mitochondrial matrix (Muller et al., 2004) that detoxifies mitochondrial superoxide into hydrogen peroxide. We show that MnSOD expression decreases after reoxygenation with oxPFCs in moderate concentrations. As mitochondrial superoxide is formed during oxidative phosphorylation from the mitochondrial electron transport chain (Babior et al., 1973), the decrease in MnSOD expression implies a decrease in oxidative phosphorylation and superoxide production, underscoring the pacifying effect of moderate doses of oxPFCs on mitochondrial activity.
AMPK adjusts the cardiac energy metabolism to ischemia (Xing et al., 2003), and it prevents postischemic cell death (Russell et al., 2004). We show that AMPK activation increases after reoxygenation of ischemic cardiomyocytes using moderate doses of oxPFCs compared with normoxic reoxygenation. In addition, we observed a decrease in cellular ATP content with moderate doses of oxPFCs. Our findings corroborate previous findings that AMPK serves as an indicator of ATP availability in cells, detecting changes in the AMP:ATP ratio and signaling an inadequate ATP supply by oxidative phosphorylation (Hardie et al., 1998; Baron et al., 2005). Reoxygenation with high doses of oxPFCs does not increase but rather decreases AMPK activation, indicating that toxic oxygen doses of oxPFCs may unhinge the protective capacity of antiapoptotic AMPK, leading eventually to necrosis and apoptosis.
We have demonstrated a dose-dependent effect of oxygen on stabilization of HIF1- and HIF2- in ischemic cardiomyocytes during reoxygenation. We were able to show a regenerative effect of moderate doses of oxPFCs that is lost when the dose of oxPFC is further increased. Although reoxygenation with oxygen in moderate concentrations (750 µMO2) increases HIF1- and HIF2- stabilization, compared with reoxygenation at normoxic conditions, reoxygenation with oxygen in higher concentrations (2250 µMO2) has no effect on HIF stabilization.
In conclusion, we report for the first time that treatment of ischemic cardiomyocytes with oxPFCs in moderate concentrations results in improved cell viability by stabilization of the mitochondrial membrane potential and reducing steady-state levels of superoxide in an in vitro model of ischemia and reperfusion. The gain in mitochondrial membrane potential is consistent with the pattern of increased AMPK expression after reoxygenation with moderate doses of oxPFCs, which signals a reduced rate of oxidative phosphorylation. DAPK senses an increase in mitochondrial membrane potential mediated by reduced steady-state levels of superoxide and becomes deactivated. We propose that increased viability of ischemic cardiomyocytes results from a pacifying effect of moderate doses of oxPFCs on the mitochondrial activity, superoxide steady-state levels and the stabilizing effect on HIF1- and HIF2- during reoxygenation. We propose that HIF1- and HIF2- stabilization could mediate a protective effect on ischemic cardiomyocytes during reperfusion. Accordingly, reperfusion strategies may offer advantages over conventional normoxic reperfusion when applying optimal oxygen concentrations to the ischemic myocardium.
|
Footnotes |
|---|
ABBREVIATIONS: DAPK, death-associated protein kinase; AMPK, AMP-activated protein kinase; HIF, hypoxia-inducible factor; PFC, perfluorochemical; ox, oxygenated; p, phosphorylated; BSA, bovine serum albumin; PBS, phosphate-buffered saline; LDH, lactate dehydrogenase; MnSOD, manganese superoxide dismutase; 
m, mitochondrial membrane potential.
Address correspondence to: Dr. Amina Arab, Department of Cardiology, University of Freiburg, Hugstetterstrasse 55, D-79106 Freiburg i. Br, Germany. E-mail: arab{at}medizin.ukl.uni-freiburg.de
|
References |
|---|
|
TopAbstractMaterials and MethodsResultsDiscussionReferences |
|---|
Babior BM, Kipnes RS, and Curnutte JT (1973) Biological defense mechanisms. The production by leukocytes of superoxide, a potential bactericidal agent. J Clin Invest 52: 741–744.[Medline]
Baron SJ, Li J, Russell RR III, Neumann D, Miller EJ, Tuerk R, Wallimann T, Hurley RL, Witters LA, and Young LH (2005) Dual mechanisms regulating AMPK kinase action in the ischemic heart. Circ Res 96: 337–345.
Barth E, Stammler G, Speiser B, and Schaper J (1992) Ultrastructural quantitation of mitochondria and myofilaments in cardiac muscle from 10 different animal species including man. J Mol Cell Cardiol 24: 669–681.[CrossRef][Medline]
Cai J and Jones DP (1998) Superoxide in apoptosis. Mitochondrial generation triggered by cytochrome c loss. J Biol Chem 273: 11401–11404.
Chandel NS, McClintock DS, Feliciano CE, Wood TM, Melendez JA, Rodriguez AM, and Schumacker PT (2000) Reactive oxygen species generated at mitochondrial complex III stabilize hypoxia-inducible factor-1alpha during hypoxia: a mechanism of O2 sensing. J Biol Chem 275: 25130–25138.
Davies KJ (1995) Oxidative stress: the paradox of aerobic life. Biochem Soc Symp 61: 1–31.[Medline]
Foglieni C, Meoni C, and Davalli AM (2001) Fluorescent dyes for cell viability: an application on prefixed conditions. Histochem Cell Biol 115: 223–229.[Medline]
Forman MB, Perry JM, Wilson BH, Verani MS, Kaplan PR, Shawl FA, and Friesinger GC (1991) Demonstration of myocardial reperfusion injury in humans: results of a pilot study utilizing acute coronary angioplasty with perfluorochemical in anterior myocardial infarction. J Am Coll Cardiol 18: 911–918.[Abstract]
Glazier JJ (2005) Attenuation of reperfusion microvascular ischemia by aqueous oxygen: experimental and clinical observations. Am Heart J 149: 580–584.[CrossRef][Medline]
Gurevich RM, Regula KM, and Kirshenbaum LA (2001) Serpin protein CrmA suppresses hypoxia-mediated apoptosis of ventricular myocytes. Circulation 103: 1984–1991.
Hardie DG, Carling D, and Carlson M (1998) The AMP-activated/SNF1 protein kinase subfamily: metabolic sensors of the eukaryotic cell? Annu Rev Biochem 67: 821–855.[CrossRef][Medline]
Jürgensen JS, Rosenberger C, Wiesener MS, Warnecke C, Horstrup JH, Grafe M, Philipp S, Griethe W, Maxwell PH, Frei U, et al. (2004) Persistent induction of HIF-1alpha and -2alpha in cardiomyocytes and stromal cells of ischemic myocardium. FASEB J 18: 1415–1417.
Lee SH, Wolf PL, Escudero R, Deutsch R, Jamieson SW, and Thistlethwaite PA (2000) Early expression of angiogenesis factors in acute myocardial ischemia and infarction. N Engl J Med 342: 626–633.
Matoba S, Tatsumi T, Keira N, Kawahara A, Akashi K, Kobara M, Asayama J, and Nakagawa M (1999) Cardioprotective effect of angiotensin-converting enzyme inhibition against hypoxia/reoxygenation injury in cultured rat cardiac myocytes. Circulation 99: 817–822.
Merante F, Mickle DA, Weisel RD, Li RK, Tumiati LC, Rao V, Williams WG, and Robinson BH (1998) Myocardial aerobic metabolism is impaired in a cell culture model of cyanotic heart disease. Am J Physiol Heart Circ Physiol 275: H1673–H1681.
Muller FL, Liu Y, and Van RH (2004) Complex III releases superoxide to both sides of the inner mitochondrial membrane. J Biol Chem 279: 49064–49073.
Murphy E (2004) Primary and secondary signaling pathways in early preconditioning that converge on the mitochondria to produce cardioprotection. Circ Res 94: 7–16.
Omar HA, Cherry PD, Mortelliti MP, Burke-Wolin T, and Wolin MS (1991) Inhibition of coronary artery superoxide dismutase attenuates endothelium-dependent and -independent nitrovasodilator relaxation. Circ Res 69: 601–608.
Pagano PJ, Ito Y, Tornheim K, Gallop PM, Tauber AI, and Cohen RA (1995) An NADPH oxidase superoxide-generating system in the rabbit aorta. Am J Physiol Heart Circ Physiol 268: H2274–H2280.
Rafikova O, Sokolova E, Rafikov R, and Nudler E (2004) Control of plasma nitric oxide bioactivity by perfluorocarbons: physiological mechanisms and clinical implications. Circulation 110: 3573–3580.
Riess JG (2005) Understanding the fundamentals of perfluorocarbons and perfluorocarbon emulsions relevant to in vivo oxygen delivery. Artif Cells Blood Substit Immobil Biotechnol 33: 47–63.[Medline]
Russell RR III, Li J, Coven DL, Pypaert M, Zechner C, Palmeri M, Giordano FJ, Mu J, Birnbaum MJ, and Young LH (2004) AMP-activated protein kinase mediates ischemic glucose uptake and prevents postischemic cardiac dysfunction, apoptosis, and injury. J Clin Invest 114: 495–503.[CrossRef][Medline]
Sandau KB, Fandrey J, and Brune B (2001) Accumulation of HIF-1alpha under the influence of nitric oxide. Blood 97: 1009–1015.
Schumacher AM, Velentza AV, and Watterson DM (2002a) Death-associated protein kinase as a potential therapeutic target. Expert Opin Ther Targets 6: 497–506.[CrossRef][Medline]
Schumacher AM, Velentza AV, Watterson DM, and Wainwright MS (2002b) DAPK catalytic activity in the hippocampus increases during the recovery phase in an animal model of brain hypoxic-ischemic injury. Biochim Biophys Acta 1600: 128–137.[Medline]
Schumann G, Bonora R, Ceriotti F, Clerc-Renaud P, Ferrero CA, Ferard G, Franck PF, Gella FJ, Hoelzel W, Jorgensen PJ, et al. (2002) IFCC primary reference procedures for the measurement of catalytic activity concentrations of enzymes at 37 degrees C. Part 3. Reference procedure for the measurement of catalytic concentration of lactate dehydrogenase. Clin Chem Lab Med 40: 643–648.[CrossRef][Medline]
Shamloo M, Soriano L, Wieloch T, Nikolich K, Urfer R, and Oksenberg D (2005) Death-associated protein kinase is activated by dephosphorylation in response to cerebral ischemia. J Biol Chem 280: 42290–42299.
Shang T, Joseph J, Hillard CJ, and Kalyanaraman B (2005) Death-associated protein kinase as a sensor of mitochondrial membrane potential: role of lysosome in mitochondrial toxin-induced cell death. J Biol Chem 280: 34644–34653.
Spector NL, Yarden Y, Smith B, Lyass L, Trusk P, Pry K, Hill JE, Xia W, Seger R, and Bacus SS (2007) Activation of AMP-activated protein kinase by human EGF receptor 2/EGF receptor tyrosine kinase inhibitor protects cardiac cells. Proc Natl Acad Sci U S A 104: 10607–10612.
Velentza AV, Wainwright MS, Zasadzki M, Mirzoeva S, Schumacher AM, Haiech J, Focia PJ, Egli M, and Watterson DM (2003) An aminopyridazine-based inhibitor of a pro-apoptotic protein kinase attenuates hypoxia-ischemia induced acute brain injury. Bioorg Med Chem Lett 13: 3465–3470.[CrossRef][Medline]
Wall TC, Califf RM, Blankenship J, Talley JD, Tannenbaum M, Schwaiger M, Gacioch G, Cohen MD, Sanz M, Leimberger JD, et al. (1994) Intravenous Fluosol in the treatment of acute myocardial infarction. Results of the Thrombolysis and Angioplasty in Myocardial Infarction 9 Trial. TAMI 9 Research Group. Circulation 90: 114–120.
Weiss JN, Korge P, Honda HM, and Ping P (2003) Role of the mitochondrial permeability transition in myocardial disease. Circ Res 93: 292–301.
Xing Y, Musi N, Fujii N, Zou L, Luptak I, Hirshman MF, Goodyear LJ, and Tian R (2003) Glucose metabolism and energy homeostasis in mouse hearts overexpressing dominant negative alpha2 subunit of AMP-activated protein kinase. J Biol Chem 278: 28372–28377.
Yellon DM and Hausenloy DJ (2007) Myocardial reperfusion injury. N Engl J Med 357: 1121–1135.
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
 |
 |
 |
|
 |
 |
 |
