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J Thorac Cardiovasc Surg 2007;134:124-131
© 2007 The American Association for Thoracic Surgery
Cardiopulmonary Support and Physiology |
Cardiac Surgery Unit, Department of Cardiovascular Sciences, Glenfield Hospital, University of Leicester, Leicester, UK.
Received for publication September 8, 2006; revisions received December 7, 2006; accepted for publication December 28, 2006. * Address for reprints: Manuel Galiñanes, MD, PhD, FRCS, FECTS, Cardiac Surgery Unit, Department of Cardiovascular Sciences, University of Leicester, Glenfield Hospital, Groby Rd, Leicester, LE3 9QP UK. (Email: mg50{at}le.ac.uk).
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Abstract |
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Methods: The right atrial appendages (n = 8 per group) of patients without diabetes and patients with type 1 and 2 diabetes were subjected to 90 minutes of simulated ischemia and 120 minutes reoxygenation. Tissue injury was measured by the release of creatine kinase into the media, and cellular apoptosis and necrosis were assessed in tissue by the terminal transferase deoxyuridine triphosphate nick-end labeling assay and propidium iodide staining. Initiator and effector caspases activations were also measured.
Results: Apoptosis and necrosis were greater in the type 2 and type 1 diabetes groups than in the nondiabetes group both in fresh tissue and after simulated ischemia–reoxygenation. Activation of effector caspases was also higher in the diabetes groups than in the nondiabetes group after simulated ischemia–reoxygenation. Caspase-3 inhibition reduced apoptosis in all study groups without influencing necrosis; however, poly–adenosine diphosphate–ribose polymerase inhibition resulted in a similar reduction in apoptosis and in necrosis in all groups, whereas caspase-2 inhibition did not.
Conclusions: Diabetes increases both apoptosis and necrosis in human myocardium, both fresh and after being subjected to ischemia–reoxygenation, an effect that is mediated, at least in part, by caspase-3 and poly–adenosine diphosphate–ribose polymerase activation. These results may explain the increased cardiac-related morbidity and mortality associated with cardiac surgery in patients with diabetes.
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Introduction |
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and is also an important predictor of morbidity and mortality after cardiac surgery.2 Myocardial injury caused by ischemia and reperfusion during cardiac surgery may induce cell death, which is considered to be an important determinant in the development of left ventricular dysfunction and heart failure in ischemic cardiomyopathy.3 Animal studies have demonstrated that loss of myocytes as a result of necrosis leads to wall restructuring, side-to-side slippage of cells, mural thinning, chamber dilation, fibroblast activation, and myocardial scarring, which in turn result in depression of ventricular function.4 It is possible that diabetes may exacerbate these changes by increasing apoptosis and necrosis. Although patients with this condition are at an increased risk for ischemic events,5-7 the susceptibility of the diabetic myocardium to ischemic injury is controversial in the literature on animal studies,5,6,8,9 and the subject remains largely unexplored in human beings (for review see Galiñanes and Fowler10).
Caspases are a large protein family of cysteine proteases that have been specifically linked with apoptosis, and their inhibition has been shown to attenuate apoptosis and myocardial ischemic injury in nondiabetic rats11; however, the role of caspases in ischemic injury of the diabetic myocardium is unknown. More recently, poly–adenosine diphosphate–ribose polymerase (PARP), a nuclear protein that plays an essential role in DNA damage and repair, has also been shown to be linked to tissue damage in various pathologic conditions and to be associated with oxidant stress occurring in myocardial ischemic injury,12,13 stroke,14 and circulatory shock.15 It might be expected that PARP inhibition would reduce myocardial ischemic injury; however, the role of PARP in the presence of pathologic conditions such as diabetes needs to be clarified.
The first aim of these studies performed on the human myocardium was to elucidate the effect of diabetes on the degree of cell death by apoptosis and necrosis in both nonischemic and ischemic tissue. In addition, the roles played by caspases and PARP activation in ischemia reperfusion–induced cell death were investigated.
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Materials and Methods |
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Experimental Preparation and Solutions
The sectioning of the atrial muscle and the simulated ischemia–reoxygenation (SIR) preparation have been previously described.16 Briefly, the atrial muscles were immersed in cold (4°C) buffer solution and transferred within 2 minutes from the operating theater to the laboratory. The appendages were then mounted onto a ground glass plate with the epicardial surface face down and then sliced freehand with surgical skin graft blades (Swann-Morton Ltd, Sheffield, UK) to a thickness between 300 and 500 µm. The muscles (weight 30–50 mg) were transferred to conical flasks (25-mL Erlenmeyer flasks; SCHOTT Jenaer Glas GmbH, Jena, Germany) containing 10 mL oxygenated buffered solution (composition listed later), and the flasks were placed in a shaking water bath maintained at 37°C. After this, the muscles were equilibrated for 30 minutes in oxygenated (95% oxygen/5% carbon dioxide) Krebs–Henseleit N-2-hydroxyethylpiperazine-N-2-ethanesulfonic acid buffered solution containing the following: 118-mmol/L sodium chloride, 4.8-mmol/L potassium chloride, 27.2-mmol/L sodium hydrogen carbonate, 1.2-mmol/L magnesium chloride, 1.0-mmol/L potassium dihydrogen phosphate, 1.20-mmol/L calcium chloride, 10-mmol/L glucose, and 20-mmol/L N-2-hydroxyethylpiperazine-N-2-ethanesulfonic acid at a pH of 7.4. The buffer was supplemented with 10% fetal calf serum (Harlan SeraLabs No. S-0001A; Harlan Bioproducts for Science, Inc, Indianapolis, Ind). For the induction of simulated ischemia, the media were bubbled with 95% nitrogen/5% carbon dioxide (pH 6.80–7.00) in the absence of glucose.
Experimental Protocols
Study 1: Degree of apoptosis and necrosis in nonischemic fresh diabetic myocardium
Apoptosis, necrosis, and initiator (including caspases-2, –8, –9, and –10) and effector caspase (including caspases –3, –6 and –7) activations were assessed in nonischemic fresh atrial muscles from patients without diabetes and from those with type 1 and 2 diabetes (n = 8 per group).
Study 2: Effect of ischemia–reoxygenation in diabetic myocardium
The atrial muscles (n = 8/group) from patients without diabetes and with type 1 and 2 diabetes were equilibrated for 30 minutes and then subjected to the following two experimental protocols: aerobic perfusion for 210 minutes and 90 minutes of simulated ischemia followed by 120 minutes of reoxygenation (SIR). Creatine kinase (CK) release was measured in the incubation media during the 120 minutes of reoxygenation, or during the last 120 minutes of aerobic incubation in the control preparations, and the tissue was taken at the end of protocols for the assessment of tissue viability, cellular apoptosis and necrosis, and initiator and effector caspase activations.
Study 3: Roles of caspases and PARP in cell death in diabetic myocardium
Sections of atrial muscle (n = 6) from patients without diabetes and those with type 1 and 2 diabetes were equilibrated for 30 minutes and then subjected to the following protocols: aerobic perfusion for 210 minutes, 90 minutes of simulated ischemia followed by 120 minutes of reoxygenation (SIR), SIR with the caspase-3 inhibitor z.DEVD.fmk (70 nmol/L), SIR with different concentrations of the caspase-2 inhibitor z.VDVAD.fmk (0, 0.02, 0.2, and 2 µmol/L), and SIR with different concentrations of the PARP inhibitor PJ-34 (0, 0.07, 0.7, and 7 µmol/L). The caspase and PARP inhibitors were incubated with the muscles for the entire experimental period. As in the previous protocol, CK release was measured in the incubation media and cellular apoptosis and necrosis were assessed in the muscles at the end of the protocols.
Assessment of Tissue Injury and Viability
CK release into the perfusate during the 120 minutes of reoxygenation was measured as an index of tissue injury. The enzyme activity was measured with a linked-enzyme kinetic assay according to manufacturer instruction (DG147-K; Sigma Chemicals Pty Ltd, Perth, Australia) and expressed as international units per gram of wet weight.
Tissue viability was assessed by the mitochondrial reduction of 3-[4, 5 dimethylthiazol-2-y1]-2, 5-diphenyltetrazolium bromide (MTT) to an insoluble purple formazan dye (M2128; Sigma Chemicals).17 The absorbance of the blue formazan product was measured on a plate reader (Benchmark; Bio-Rad Laboratories Inc, Hercules, Calif) at 550 nm, and the results were expressed as micromol of formazan per gram of wet weight. A reduction in the MTT values was considered to represent decreased tissue viability.
Assessment of Apoptosis and Necrosis
First, the muscles were incubated for 10 minutes on ice with 5-µmol/L propidium iodide (PI) in 0.1-mol/L trisodium citrate and 20-mmol/L phosphate-buffered saline solution at pH 7.4 to identify the necrotic nuclei. Sections were then fixed twice, initially for 30 minutes with 4% paraformaldehyde in 30% sucrose and then with 20-mmol/L phosphate-buffered saline solution overnight on ice at pH 7.4. After this, serial 10-µm sections were cut with a Bright cryomicrotome (model OTF; Bright Instrument Co Ltd, Huntingdon, UK) at –25°C in tissue-embedding matrix (Tissue Tek OCT compound; Sakura Finetek USA, Inc, Torrance, Calif). The cryopreserved tissue sections were washed with 20-mmol/L phosphate-buffered saline solution at pH 7.4 for 2 minutes, then made permeable in 0.02-mg/mL proteinase K for 10 minutes at 37°C and presensitized for 1 minute in a microwave oven at 800 W in 0.1% Triton X-100 (The Dow Chemical Company, Midland, Mich) and 0.1-mol/L sodium citrate at pH 6.0. To assess apoptosis, terminal deoxynucleotidyl transferase was used to incorporate deoxyuridine triphosphate oligonucleotides labeled with fluorescein isothiocyanate to DNA strand breaks at the 3'-OH termini in a template-dependent manner (terminal transferase deoxyuridine triphosphate nick-end labeling [TUNEL] assay) with a commercially available kit (1684795; Roche Diagnostics Division, Basel, Switzerland). With this labeling procedure sequence, the nuclei could be stained either with PI or by TUNEL but not both. An 8-µm section of the mirror specimens opened up the cellular membrane of all the cells to enable all the nuclei to be stained. Before the TUNEL labeling of muscles, positive control specimens were treated with DNase I, and negative control specimens were obtained by adding the label solution of the kit without the enzyme solution. The fluorescein isothiocyanate fluorescence emission (range 600–630 nm) was measured with argon-ion fluorescence excitation at 488 nm and detected by laser confocal epifluorescence microscopy with a x10 oil-immersion objective. The PI-labeled nuclei were excited with helium–neon laser light at 543 nm, and fluorescence was detected at an emission range of 680 to 730 nm to abolish fluorescence bleed through from fluorescein isothiocyanate–labeled nuclei. Analysis was done with National Institutes of Health Image software (Scion Corporation, Frederick, Md) with the Cavalieri-3 macro (G. MacDonald, University of Washington, Seattle, Wash), which allows placement of point-counting templates over an image to perform stereologic estimates. Fluorescent signals with areas greater than 16 µm2 were counted to ensure that only nuclei were taken into account and to avoid the inclusion of artifact. Absolute numbers of green fluorescent apoptotic and necrotic red fluorescent nuclei in any given image field were determined by dividing by the total number of PI-labeled nuclei in the next serial or mirror section. The absolute percentage of apoptotic cells was given by dividing apoptotic nuclei by total PI-labeled nuclei and multiplying by 100%, whereas the percentage of necrotic cells was obtained by dividing necrotic nuclei by total PI-labeled nuclei and multiplying by 100%. The automatic counting was combined with regular manual inspection to ensure that artifacts were not represented in the data.
Measurement of Caspase Activities
The muscle sections that had been stored at –800°C until analysis were thawed in 400 µL cell lysis buffer (100 mmol/L N-2-hydroxyethylpiperazine-N-2-ethanesulfonic acid, 10% sucrose, 0.1% 3-[3-(cholamidopropyl)dimethylammonio]-1-proanesulfonate, and 10-mmol/L dithiothreitol) in the presence of a cocktail of enzyme inhibitors (P2850; Sigma Chemicals) at a pH of 7.0 to release the intracellular contents. The muscle was diced finely and then homogenized (Ultra-Turrax homogenizer; Janke & Kunkel GmbH & Co, Staufen, Germany) at 13,000 rpm for 1 minute on ice. This was followed by centrifugation (PK121R; ALC International, Cologno Honzese, Italy) at 14,000 rpm for 30 minutes. Subsequently, the protein concentration of the soluble supernatant (cellular lysate) was measured with a detergent-compatible Bio-Rad assay (23225; Pierce & Warriner [UK] Ltd, Chester, UK). Aliquots of cellular lysate were then tested for caspase activity by the addition of a caspase-specific peptide or substrate, D(OMe)-E(OMe)-V-D(OMe) (DEVD), conjugated to the chromophore (fluorescent reporter molecule) 7-amino-4-trifluoromethyl coumarin (AFC). The cleavage of the peptide DEVD from DEVD.AFC (final concentration 20 µmol/L; Alexis Corporation, San Diego, Calif) releases AFC, which when excited by light at 400 nm emits fluorescence at 505 nm. The level of caspase activity in the cellular lysate was detected by fluorescence signal obtained with a fluorometer (FLUOstar P401; BMG LABTECH GmbH, Offenburg, Germany). The amount of caspase-3–like activity was measured by using the effector caspase inhibitor z.DEVD.fmk at a final concentration of 10 µmol/L in the well of the reader plate and by subtracting the fluorescence obtained by the total fluorescence measured in the absence of the inhibitor. The results were expressed as arbitrary units of fluorescence activity per gram of wet weight.
Chemicals
The caspase-2 inhibitor z.VDVAD.fmk (FMK003; R&D Systems, Minneapolis, Minn) was used at different concentrations (0, 0.02, 0.2, and 2 µmol/L). The caspase-3 inhibitor z.DEVD.fmk (FMK004) was used at the concentration of 70 nmol/L, which was shown to be the optimal dose by previous experiments in our laboratory.18 The PARP inhibitor PJ-34 (L10210; Alexis Corporation) was also used at different concentrations (0, 0.07, 0.7, and 7.0 µmol/L).
Statistical Analysis
All results are expressed as mean ± SEM. To compare the overall statistical significance among no diabetes and type 1 and 2 diabetes groups, the nonparametric analysis of variance (Kruskal–Wallis H) technique was used. The comparisons between the independent groups are based on nonparametric Mann–Whitney test.
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Study 3: Role of Caspases and PARP in Cell Death in Diabetic Myocardium
Apoptosis and necrosis
Figure 4
(A and B) shows that in the presence of the caspase-3 inhibitor z.DEVD.fmk the apoptosis caused by SIR was almost completely abolished in the nondiabetes group, but necrosis was unaffected. In contrast, in the muscles from patients with diabetes, caspase-3 inhibition reduced apoptosis by only 50%, although also without effect on necrosis.
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Discussion |
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Cell Death in Fresh, Nonischemic Diabetic Myocardium
Cell death by apoptosis and necrosis is a feature of end-stage heart failure,19 but it also occurs in the healthy myocardium.20 Here we have demonstrated that the occurrence of apoptosis and necrosis in human atrial tissue are greater in patients with diabetes than without diabetes, a finding supported by those of Frustaci and colleagues21 in ventricular biopsies. Because myocytes rarely proliferate in adult cardiac muscles, the increased loss of cardiac muscles in the diabetic myocardium may lead to a reduction in cardiac mass and to elevated interstitial and perivascular fibrosis, causing a decrease in myocardial performance and ventricular dilatation, a sequence of events that may be responsible for the increasing cardiovascular mortality and morbidity in patients with diabetes.22
A possible explanation for the increase in cell death in patients with diabetes may be a greater oxidative stress observed with this condition,23 which in turn may be responsible for the activation of effector caspases seen in our studies. There is experimental evidence that caspases are activated in diabetes11 and also that caspases are activated by oxidative stress in both patients without20 and with24 diabetes. Indeed, high glucose causes oxidative stress that results in caspase-3 activation in human endothelial cells through stress activated protein kinase/c-Jun NH2-terminal kinases activation.25 Activating of effector caspases in diabetes can also be induced by the abnormal accumulation of ß-hydroxyl fatty acid, as seen in mice, which alters the permeability of the mitochondrial membrane and causes the release of cytochrome c and the activation of the downstream caspases.26 Our findings that effector caspases are activated in the fresh, nonischemic human atrial myocardium suggests a central role of these proteins in the increase in apoptosis seen in the hearts of patients with diabetes. It is clear, however, that further investigation is required to elucidate the precise molecular mechanism involved.
Susceptibility of Diabetic Myocardium to Ischemia–Reoxygenation Injury
These studies also show that diabetic myocardium is more susceptible to ischemia–reoxygenation injury than is nondiabetic myocardium when the PI and TUNEL techniques to test for apoptosis and necrosis are used. These results in the human myocardium are supported by experimental animal studies in rats27 and dogs28 showing that diabetes makes the heart more susceptible to ischemia–reoxygenation injury. Other experimental studies,29 however, have shown diabetic myocardium in fact to be more tolerant to injury than nondiabetic tissue. The reasons for these divergent results are not clear, but it is possible that duration and severity of the diabetic state and differences in the experimental preparations used may be involved.10 Another potential explanation could be a lack of correspondence between different end points, as shown by our studies of CK release and MTT reduction.
The observation that the activity of the effector caspases is more elevated in diabetic than nondiabetic myocardium in response to an ischemic insult suggests that this class of enzymes is responsible for the greater occurrence of apoptosis in diabetes. Caspase-3 inhibition reduced apoptosis in diabetic myocardium to a lesser extent than in nondiabetic myocardium, however, and inhibition of caspase-2, an enzyme that shares sequence homology with initiator caspases like caspase-9 and –130 but has cleavage specificity closer to the effector caspases caspase-3 and –7,31 did not effect apoptosis in either groups, suggesting that the increase of myocardial apoptosis in diabetes is not dependent on greater activation of the effector caspases alone. Our finding that the induction of apoptosis by ischemia–reoxygenation in human nondiabetic myocardium is caspase-3 dependent is supported by in vivo animal experimental studies11; however, the finding that caspase-3 inhibition only partially reduced apoptosis in diabetic myocardium suggests that a caspase-independent pathway causing apoptosis also exist in the diabetic myocardium.
In contrast with the results on inhibition of caspases-3 activity, the inhibition of PARP similarly reduced apoptosis in a dose-dependent manner in both diabetic and nondiabetic ischemic myocardium, with almost complete abolition at the highest concentration of the inhibitor used (7-µmol/L PJ-34). This suggests that both caspase-dependent and -independent pathways of apoptosis converge in activation of PARP. It is possible that the reported increase activation of PARP seen in the diabetic rat myocardium32 is responsible for the greater susceptibility of this tissue to apoptosis. The role of PARP in ischemic injury has been disputed, however, and whereas in vitro33 and in vivo13 studies have demonstrated limitation of cellular injury by PARP inhibition, other investigators have suggested that PARP activation is not indispensable to apoptosis.30,34
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Acknowledgments |
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Footnotes |
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