The authors have declared that no competing interests exist.
Conceived and designed the experiments: SK GS. Performed the experiments: S. Li SK S. Loganathan TR GV. Analyzed the data: S. Li SK KH AW EB PH YZ LT SP GV GS. Contributed reagents/materials/analysis tools: MK GS. Wrote the paper: SK S. Li TR GS.
Many donor organs come from youths involved in alcohol-related accidental death. The use of cardiac allografts for transplantation from donors after acute poisoning is still under discussion while acute ethanol intoxication is associated with myocardial functional and morphological changes. The aims of this work were 1) to evaluate in rats the time-course cardiac effects of acute ethanol-exposure and 2) to explore how its abuse by donors might affect recipients in cardiac pump function after transplantation.
Rats received saline or ethanol (3.45 g/kg, ip). We evaluated both the mechanical and electrical aspects of cardiac function 1 h, 6 h or 24 h after injection. Plasma cardiac troponin-T and glucose-levels were measured and histological examination of the myocardium was performed. In addition, heart transplantation was performed, in which donors received ethanol 6 h or 24 h prior to explantation. Graft function was measured 1 h or 24 h after transplantation. Myocardial TBARS-concentration was measured; mRNA and protein expression was assessed by quantitative real-time PCR and Western blot, respectively.
Ethanol administration resulted in decreased load-dependent (−34±9%) and load-independent (−33±12%) contractility parameters, LV end-diastolic pressure and elevated blood glucose levels at 1 h, which were reversed to the level of controls after 6 h and 24 h. In contrast to systolic dysfunction, active relaxation and passive stiffness are slowly recovered or sustained during 24 h. Moreover, troponin-T-levels were increased at 1 h, 6 h and 24 h after ethanol injection. ST-segment elevation (+47±10%), elongated QT-interval (+38±4%), enlarged cardiomyocyte, DNA-strand breaks, increased both mRNA and protein levels of superoxide dismutase-1, glutathione peroxydase-4, cytochrome-c-oxidase and metalloproteinase-9 were observed 24 h following ethanol-exposure. After heart transplantation, decreased myocardial contractility and relaxation, oxidative stress and altered protein expression were observed.
These results demonstrate acute alcohol abuse increases the susceptibility of donor hearts to ischemia/reperfusion in a rat heart transplant model even though the global contractile function recovers 6 h after ethanol-administration.
Episodic excessive alcohol consumption commonly referred to as “binge drinking” is common cause of accidental death, violent behaviour as well as suicide, and may be associated with compromised myocardial contractility
Currently, donors with a history of alcohol abuse, accounting for about 1/5th of all donors, are routinely accepted, despite existing evidence supporting the potential deleterious effect of donors' alcohol consumption on recipients' survival and higher rejection rate
Male Lewis rats (250 to 350 g; Charles River, Sulzfeld, Germany) were housed in a room at 22
The experimental procedure involved intraperitoneal injections (1 ml/100 g body weight) with either saline (0.9% NaCl) or alcohol. The dose was 3.45 g/kg (75 mmol/kg) body weight for ethanol. Controls were injected with identical volume of saline.
Rats (n = 74) were randomly divided into the following groups: (1) control groups received saline and were euthanized 1 h, 6 h or 24 h after saline injection respectively; (2) ethanol-1 h group (3) ethanol-6 h group and (4) ethanol-24 h group received ethanol and were euthanized 1 h, 6 h or 24 h after ethanol dosing, respectively. We found no difference between the groups receiving saline at different time points with any of the parameters measured and thus combined the values.
Rats were anesthetized with sodium pentobarbital (60 mg/kg, i.p.) and kept in a supine position on heating pads maintaining their core temperature (measured via a rectal probe) at 37°C. Standard 12-lead electrocardiograms were recorded using needle electrodes placed subcutaneously in each fore leg and hind legs, and six around the chest. All leads were connected to a standard direct-writing recorder (Mortara Instrument, WI, USA). The paper speed was set at 50 mm/sec and the sensitivity at 10 mm/mV. ST-segment elevation and the length of the QT-interval measured in lead-II were the focuses of electrocardiographical analysis. The QT interval was defined as the segment from the onset of the QRS complex to the end of the T wave, defined as the intersection point with the isoelectric line. QT interval was corrected using normalized Bazett's formula adjusted for rats (nQTc = QT/(RR/f)1/2)
Rats were placed on controlled heating pads, core temperature was measured via a rectal probe and was maintained at 37°C, tracheotomized, intubated and artificially ventilated. To assess cardiac function, LV pressure-volume analysis was performed with a 2F pressure-volume conductance catheter (SPR-838, Millar Instruments, Houston, TX, USA). With the special pressure-volume analysis program (PVAN, Millar Instruments, Houston, TX, USA), heart rate, systolic and diastolic blood pressures, mean arterial pressure, LV end-systolic pressure, LV end-diastolic pressure, maximal slope of the systolic pressure increment (dP/dtmax), maximal slope of the diastolic pressure decrement (dP/dtmin) as load dependent hemodynamic parameters were calculated. LV pressure-volume relations were measured by transiently occluding the inferior vena cava (reducing preload) under the diaphragm with a cotton-tipped applicator. The slope of dP/dtmax/end-diastolic volume (dP/dtmax/EDV) relationship was calculated as load independent index of LV contractility. The slope of the end-diastolic pressure-volume relationship (EDPVR) was calculated as a reliable index of LV stiffness.
At the end of each experiment, 0.1 ml hypertonic saline was injected into the jugular vein, and from the shift of pressure-volume relations parallel conductance volume was calculated by pressure-volume analysis software (PVAN, Millar Instruments, Houston, TX, USA) and used for correction for cardiac mass volume. After completing the hemodynamic measurements, blood samples were collected from the inferior vena cava. The volume calibration of the conductance system was performed as described previously
Blood collected from the rats in lithium with heparin and gel tubes (S-Monovette®, Sarstedt, Nümbrecht, Germany) was immediately centrifuged and plasma was separated. Levels of plasma cardiac troponin-T and glucose were determined.
Hearts from rats in each experimental group were fixed in buffered paraformaldehyde solution (4%) and embedded in paraffin. Then, 5-µm thick sections were placed on adhesive slides and stained with hematoxylin and eosin (H&E). Cardiomyocyte cross-sectional areas were calculated on a microscope (×400) using the Cell∧A software (Olympus Soft Imaging Solutions GmbH, Germany). Histological evaluation was conducted by an investigator unaware of treatment attribution of the animals.
For identification of cardiomyocytes, immunohistochemical staining for cardiac troponin I has been performed. Sections were de-paraffinized with xylene and passaged through decreasing concentrations of ethanol, washed with distilled water and heated with Tris- ethylenediaminetetraacetic acid (EDTA) buffer (pH = 9) for 30 min to retrieve antigenic epitopes. Then, sections were washed with phosphate buffer saline (PBS, 1×) for 3×5 min. After permeabilization by 0.3% Triton X-100 for 10 min and blocking with 1% bovine serum albumin (BSA) and 0.1% Triton X-100, sections were incubated at room temperature for 1 h with the primary rabbit polyclonal antibodies directed against troponin-I (1∶200, Abcam, Cambridge, UK). The incubation with undiluted FITC-conjugated goat anti-rabbit IgG polyclonal secondary antibody (Abcam, Cambridge, UK) was at 37°C for 30 min. After washing with PBS (1×) for 3×5 min, the sections were incubated with 50 µl of Terminal deoxynucleotidyl Transferase (TdT) enzyme and TUNEL Reaction mixture for 1 h at 37°C in the dark (Roche Diagnostics, Mannheim, Germany). The sections were then washed with PBS (1×) for 3×5 min. The slides were mounted using 4′, 6-diamidino-2-phenylindole (DAPI)-Fluoromount-G™ (SouthernBiotech, Birmingham, USA), covered with cover glass and analyzed under a fluorescence microscope. The number of TUNEL-positive cells was expressed as the ratio of DAPI-TUNEL double-labeled nuclei to the total number of nuclei stained with DAPI. Cells were counted in four fields characterizing each specimen), and an average value was calculated for each experimental group. The evaluation was conducted by an investigator blinded to the experimental groups.
Total RNA was isolated from the hearts with the RNeasy Fibrous Tissue Mini Kit (Qiagen, Hilden, Germany). RNA concentration and purity were determined photometrically (230 nm, 260 nm and 280 nm). Reverse transcription was performed with the QuantiTect Reverse Transcription Kit (Qiagen) using 800 ng RNA in a total volume of 20 µl. Quantitative real-time PCR was performed with the Light-Cycler480 system using the LightCycler480 Probes Master and Universal ProbeLibrary probes (Roche, Mannheim, Germany). The conditions for PCR were as follows: 95°C for 10 min (1-cycle), 95°C for 10 s, 60°C for 30 s (single; 45-cycle quantification), 40°C for 10 s (1-cycle). Efficiency of the PCR-reaction was confirmed with standard curve analysis. Sample quantifications were normalized to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) expression. Primers were obtained from TIB Molbiol (Berlin, Germany), their sequences and UPL probes used are represented on
Assay | Sequence | UPL probes |
Cytochrome c oxidase | F: |
|
GAPDH | F: |
|
GPx4 | F: |
|
Cav1.2, α1c-subunit | F: |
|
Na+-K+-ATPase(α1-subunit) | F: |
|
SERCA-2 | F: |
|
SOD-1 | F: |
|
MMP-9 | F: |
|
TNF-α | F: |
|
Inducible NOS | F: |
|
Glyceraldehyde 3-phosphate dehydrogenase (GAPDH), glutathione peroxidase (GPx)-4, L-Type calcium-channel (α1c-subunit; Cav1.2), sodium-potassium Adenosine Triphosphatase (Na+-K+-ATPase), sarco(endo)plasmic Ca2+-ATPas (SERCA)-2, superoxide dismutase (SOD)-1, matrix metalloproteinase (MMP)-9, tumor necrosis factor (TNF)-α, inducible nitric oxide synthase (NOS).
Rats (n = 72) were randomly divided into 3 groups for each protocol. First study: (1) 1 h ischemia/1 h reperfusion group (1 h I/R); (2) ethanol-treatment 6 h before explantation followed by 1 h ischemia/1 h reperfusion (6 h ethanol+1 h I/R), and (3) ethanol-treatment 24 h before explantation followed by 1 h ischemia/1 h reperfusion groups (24 h ethanol+1 h I/R).
In the second study, the influence of ethanol exposure on long-term has been investigated: (1) 1 h ischemia/24 h reperfusion group (24 h I/R); (2) ethanol-treatment 6 h before explantation followed by 1 h ischemia/24 h reperfusion (6 h ethanol+24 h I/R), and (3) ethanol-treatment 24 h before explantation followed by 1 h ischemia/24 h reperfusion groups (24 h ethanol+1 h I/R).
Transplantations were performed in isogeneic Lewis to Lewis rat strain, therefore no organ rejection can be expected. The donor-rats of the ethanol groups received a single intraperitoneal injection of ethanol (3.45 g/kg) either 6 h or 24 h prior to explantation. The donor-rats of the control groups received saline instead.
The experimental model was established according to the reported method
One hour or 24 h after transplantation a 3F latex balloon catheter (Edwards Lifesciences Corporation, Irvine, CA, USA) was introduced into the left ventricle via the apex for administration and withdrawal of fluid to determine dP/dtmax, dP/dtmin, developed pressure, LV systolic pressure and LV end-diastolic pressure by a Millar micromanometer (Millar Instruments, Houston, TX, USA) at different LV volumes. From these data LV pressure-volume relationships were constructed using PVAN3.6 software (Millar Instruments, Houston, TX, USA).
Heart TBARS concentration was measured by a commercial kit (Zeptometrix Corporation, Buffalo, New York, USA). Briefly, the homogenate was mixed with sodium dodecyl sulfate (SDS) solution and thiobarbituric acid/Buffer Reagent with thorough shaking, and heated for 60 min at 95°C. The samples were then cooled to room temperature in an ice bath for 10 min. The absorbance in the supernatant after centrifugation at 3000 rpm for 15 min was measured at 532 nm using a spectrophotometer (Thermo Electron Corporation, Waltham, Massachusetts, USA).
Myocardial proteins were extracted into a solution containing 8 M urea, 5 mM EDTA, 0.002% trasylol, 0.05 mM phenylmethanesulfonylfluoride (PMSF), 0,003% TritonX-100 containing protease inhibitors (Roche, Mannheim, Germany). Protein concentration was determined by a commercial kit according to the manufacturer's protocol (bicinchoninic acid, BCA protein assay kit; Thermo Scientific, Rockford, USA). Total protein homogenates (30 µg) were denatured, separated on sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) gradient gels (Invitrogen, Darmstadt, Germany) and transferred to polyvinylidene fluoride (PVDF) membrane (Invitrogen, Darmstadt, Germany). The membranes were blocked with 5% milk in Tris-Buffered Saline Tween 20 before incubation overnight at 4°C with primary antibodies specific to superoxide dismutase (SOD)-1 (1∶10000, Abcam, Cambridge, UK), cytochrome-c oxidase (1∶1000, New England Biolabs GmbH, Frankfurt am Main), glutathione peroxidase (GPx)-4 (1∶10000, Abcam, Cambridge, UK), metalloproteinase (MMP)-9 (1∶100, Santa Cruz, Biotehnology, Heidelberg, Germany) and cardiac troponin I (1∶5000, Abcam, Cambridge, UK). After washing blots to remove excessive primary antibody binding, blots were incubated for 1 h with horseradish peroxydase conjugated secondary antibody (1∶5000, Santa Cruz Biothechnology, Heidelberg, Germany). The immunoreactive protein bands were developed using Enhanced Chemiluminescence system (PerkinElmer, Rodgau-Juegesheim, Germany). The intensity of immunoblot bands was detected with a Fujifilm LAS-3000 Imager.
Ethanol absolute, sodium chloride, Triton X-100 were bought from Sigma-Aldrich (Steinheim, Germany). Custodiol was purschased from Dr Franz Köhler Chemie GmbH (Alsbach-Hähnlein, Germany), sodium pentobarbital from Merial GmbH (Halbergmoos, Germany), heparin sodium from Ratiopharm GmbH (Ulm, Germany), buffered paraformaldehyde 4% from Carl Roth GmbH (Karlsruhe, Germany), PBS from Genaxxon Bioscience GmbH (Ulm, Germany), Tris-Buffered Saline Tween 20 from Thermo Fisher Scientific (Cheshire, UK), BSA from Invitrogen Corporation (Auckland, New Zealand), xylene from National Diagnostics (Atlanta, USA), and EDTA from Applied Chem (Darmstadt, Germany).
All data are expressed as means ± standard error of the mean (S.E.M). Means between groups were compared by 1-way ANOVA followed by an unpaired t test with Bonferroni correction for multiple comparisons. A value of p<0.05 was considered statistically significant.
Whereas ethanol exposure had no effect on heart rate, systolic and diastolic blood pressures, mean arterial pressure and LV end-systolic pressure were significantly reduced at 1 h and 6 h and recovered 24 h after ethanol administration compared with the control group (
Assessment of (A) load-dependent (dP/dtmax) and (B) load-independent (dP/dtmax-EDV) contractility parameters revealed a significant decline only at 1 h in response to acute ethanol when compared with the control group. Whereas (C) maximal slope of the diastolic pressure decrement (dP/dtmin) 1 h and 6 h after ethanol administration was significantly decreased, (D) end-diastolic pressure-volume relationship (EDPVR) was significantly increased in rats treated with ethanol after 1 h, 6 h and 24 h. Eth indicates ethanol. *p<0.05 versus control.
Parameters | Control | Ethanol 1 h | Ethanol 6 h | Ethanol 24 h |
Heart rate, beats/min | 345±16 | 353±23 | 399±38 | 390±28 |
SBP, mmHg | 112±4 |
|
|
104±7 |
DBP, mmHg | 83±5 |
|
|
77±7 |
MAP, mmHg | 93±4 |
|
|
86±7 |
LVESP, mmHg | 100±5 |
|
|
86±7 |
LVEDP, mmHg | 7.6±0.5 |
|
6.5±1.1 | 7.9±1.1 |
SBP: systolic blood pressure; DBP: diastolic blood pressure; MAP: mean arterial pressure; LVESP: left-ventricular end-systolic pressure; LVEDP: left-ventricular end-diastolic pressure.
p<0.05 versus control;
p<0.05 versus ethanol 1 h.
Ethanol-treated rats showed a marked elevation in ST-segments at 24 h and elongated corrected QT interval at 6 h and 24 h when compared with the control group (
(A) Representative electrocardiogram tracings. Rats treated with ethanol showed a marked (B) ST-segment elevation at 24 h and (C) elongated corrected QT interval at 6 h and 24 h when compared with the control group. Eth indicates ethanol. *p<0.05 versus control; #p<0.05 versus ethanol 1 h.
Control | Ethanol 1 h | Ethanol 6 h | Ethanol 24 h | |
Cardiac troponin-T, pg/ml | 76±11 |
|
|
|
Glucose, mg/dl | 174±5 |
|
227±98 | 182±20 |
Following ethanol injection, glucose levels began to increase after 1 h and returned towards baseline values after 6 h and 24 h, cardiac troponin-T levels remained elevated after 1 h, 6 h and 24 h.
p<0.05 versus control.
Histological examination revealed that at 24 h following ethanol administration, cardiomyocyte transverse cross-section area was significantly increased in the H&E-stained sections compared with the control-group and there was no sign of myocardial inflammation (
(A) Hematoxylin and eosin (H&E) staining micrographs of transverse sections of myocardium (magnification ×400; scale bar: 20 µm) and (B) quantitative analysis of cardiomyocyte cross-sectional area using measurements of ∼20 cardiomyocytes in each group. Twenty-four hours after ethanol administration, the cardiomyocyte transverse cross-section area was significantly increased in the H&E staining sections compared with the control group. Eth indicates ethanol. *p<0.05 versus control.
Representative photomicrographs of (A) cardiomyocytes stained with cardiac troponin I (green) and nuclei with 4′, 6-diamidino-2-phenylindole (DAPI, blue), (B) nuclei with fragmented DNA visualized by TUNEL staining (red), and (C) merged image. Magnification ×400; scale bar: 20 µm. White arrows indicate TUNEL-positive cells (not all are marked) (D) Quantification of TUNEL-positive cells. We found DNA damage in the myocardium of rats treated with ethanol at 24 h. Eth indicates ethanol. *p<0.05 versus control.
After ethanol injection, plasma glucose levels began to increase after 1 h and returned towards baseline values after 6 h and 24 h (
Administration of ethanol significantly increased SOD-1, GPx-4 and cytochrome-c oxidase relative mRNA expression after 24 h (
Quantitative real-time PCR from myocardium RNA extracts revealed that mRNA levels for (A) superoxide dismutase (SOD)-1, (B) glutathione peroxydase (GPx)-4, (C) cytochrome-c oxidase were significantly increased after 24 h and (D) matrix metalloproteinase (MMP)-9 following 6 h and 24 h compared with the control group. However, mRNA-levels of (E) L-Type calcium-channel (Cav1.2), (F) sarco(endo)plasmic Ca2+-ATPase (SERCA)-2 and (G) sodium-potassium Adenosine Triphosphatase (Na+/K+-ATPase) remained unchanged following ethanol administration compared to the control group. Eth indicates ethanol. *p<0.05 versus control,#p<0.05 versus ethanol 1 h, $p<0.05 versus ethanol 6 h.
Densitometric analysis of bands for SOD-1 showed a significant increase at 6 h and 24 h following ethanol administration, and at 24 h for GPx4 (
Densitometric analysis of bands for (A) superoxide dismutase (SOD)-1 showed a significant increase after 6 h and 24 h ethanol administration, and (B) glutathione peroxidase (GPx)-4, (C) cytochrome-c oxidase (cyto-c oxi) and (D) MMP-9 following 24 h, compared with the control group. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH). Eth indicates ethanol. *p<0.05 versus control; #p<0.05 versus ethanol 1 h, $p<0.05 versus ethanol 6 h.
One hour and 24 h after transplantation, in which donors received ethanol 6 h or 24 h prior to explantation, significantly lower dP/dtmax and dP/dtmin were observed when compared with the control-group, indicating decrease myocardial contractility and relaxation (
One hour and 24 h after transplantation, in which donors received ethanol 6 h or 24 h prior to explantation, significantly lower (A, E) dP/dtmax (maximal slope of the systolic pressure increment) and (B, F) dP/dtmin (maximal slope of the diastolic pressure decrement) were observed when compared with the control-group. Although (C) LV systolic pressure (LVSP) and (D) LV developed pressure, as indexes of myocardial contractile function, were significantly decreased in the 24 h ethanol+1 h I/R group 1 h after transplantation. However, 24 h after transplantation, (G) LVSP and (H) LV developed pressure were significantly decreased in both 6 h ethanol+24 h I/R and 24 h ethanol+24 h I/R groups compared with the control-group. I/R indicates ischemia/reperfusion, LVV left-ventricular volume. *p<0.05 versus 1 h I/R group.
One hour and 24 h after transplantation, there was a significant increase in graft TBARS concentration in the group receiving ethanol 24 h prior to explantation when compared with the control- and 6 h ethanol+I/R groups (
One hour and 24 h after transplantation, there was a significant increase in graft TBARS concentration in the group receiving ethanol 24 h prior to explantation when compared with the control- and 6 h ethanol+24 I/R groups. I/R indicates ischemia/reperfusion, Eth ethanol. *p<0.05 versus I/R groups; #p<0.05 versus 6 h eth + I/R groups.
Twenty-four hours after transplantation, densitometric analysis of bands for SOD-1 showed a significant increase at 6 h and 24 h following ethanol administration, at 6 h for cytochrome-c oxidase, and at 24 h for MMP-9 compared with the control group (
Twenty-four hours after transplantation, densitometric analysis of bands for (A) superoxide dismutase (SOD)-1 showed a significant increase at 6 h and 24 h following ethanol administration, at 6 h for (C) cytochrome-c oxidase (cyto-c oxi), and at 24 h for (D) matrix metalloproteinase (MMP)-9 compared with the control group. Moreover, 24 h after transplantation, there was a significant increase in protein expression for (B) glutathione peroxydase (GPx)-4 in the group receiving ethanol 24 h prior to explantation when compared with the control- and 6 h ethanol+24 I/R groups. I/R indicates ischemia/reperfusion, Eth ethanol, glyceraldehyde-3-phosphate dehydrogenase (GAPDH). *p<0.05 versus 24 h I/R group; #p<0.05 versus 6 h Eth+ 24 h I/R group.
One hour and 24 h after transplantation, graft Troponin I expression was significantly increased in the groups receiving ethanol 6 h or 24 h prior to explantation when compared with the respective control groups (
(
The major finding of this present study is that acute alcohol abuse increases the susceptibility of donor hearts to ischemia/reperfusion injury in a rat model of heart transplantation even though global heart contractile function recovers 6 h after ethanol administration.
In the present study, we focussed our attention on acute ethanol exposure in which the total consumption is compressed into a short period of time to mimic binge drinking. A bolus of ethanol at a standard dose of 3.45 g/kg body weight, intraperitoneally ensures a complete bioavailability and succeeded in producing rapid rises in circulating level of ethanol. The resulting plasma ethanol concentrations were approximately 375, 185 and 0 mg/100 ml at 1 h, 6 h and 24 h respectively and were similar to pathological levels occurring clinically
The principal indicator of myopathic alteration following ethanol exposure is characterized by compromised myocardial contractility
The donor's cardiac function has been shown to be an important prognostic factoring the clinical outcome of heart transplantation. In the present study we showed elevated circulating plasma cardiac troponin-T levels, the sensitive marker of myocardial injury in heart donors after alcohol intoxication even when hemodynamic measurement showed no evidence of impaired contractile function. A clinical study showed cardiac troponin-T >0.1 µg/l in the serum of heart donors to be predictive of early impaired graft function after transplantation
Excessive consumption of alcohol over a short period of time (binge drinking) induces a systemic inflammatory reaction
Accumulation of reactive oxygen species in response to ethanol exposure is believed to play an important role. Ethanol or acetaldehyde, the primary metabolic product of ethanol, is known to trigger both oxidative stress and apoptosis
The use of hearts from donors with a history of “alcohol abuse” remains uncertain
In summary, in a model of potential organ donor, we demonstrated that after 1 h ethanol induced myocardial contractile dysfunction and elevated plasma glucose concentration. Although these parameters returned to near normal levels 6 h and 24 h after ethanol administration, morphological and molecular changes at the level of myocardium began to appear only at 24 h. Moreover, diastolic dysfunction is also observed following acute ethanol administration and sustained during 24 h. One hour and unexpectedly 24 h after heart transplantation, in which donors received ethanol 6 h or 24 h prior to explantation, decreased myocardial contractility and relaxation were observed even though the global contractile function of the donor hearts recovers 6 h after ethanol-administration. Oxidative stress, apoptosis, and mitochondrial dysfunction could predispose the donor hearts pump function in recipient to increased myocardial susceptibility to ischemia/reperfusion injury after transplantation. Further study is warranted to unveil the impact of acute-on-chronic alcohol ingestion on the outcome of ischemia/reperfusion injury after heart transplantation.
The rat model of heterotopic heart transplantation was selected to be a suitable model to evaluate global ischemia/reperfusion injury. However, this model has certain limitations. In particular, the left ventricle beats in an unloaded condition (e.g. the ventricles are perfused via the coronary circulation, but they do not eject) which on one hand allows a faster recovery after ischemia/reperfusion and on the other hand leads to a time-dependent mechanical deterioration and atrophy. Nevertheless, in detailed characterization studies with this model, it has been shown that major deterioration does not occur until at least 24 h after implantation
The expert technical assistance of Patricia Kraft, Karin Sonnenberg, Lutz Hoffmann and Nadine Weiberg is gratefully acknowledged. The authors thank Alice Lehner for her help with histological studies and Hermine Azadeh Harrison for critically reading the manuscript.