Isolation of neonatal rat cardiomyocytes and H/Re experiments
Primary cultures of neonatal rat cardiomyocytes were performed as previously described . Newborn Wistar rats (1-3 days) were used for this study. The rats were handled in accordance with the Guide for the Care and Use of Laboratory Animals published by the China National Institutes of Health. Briefly, hearts from male Wistar rats (1-3 days old) were minced and dissociated with 0.25% trypsin. Dispersed cells were seeded at 2 × 105 cells/cm2 in 60-mm culture dishes with Dulbecco's modified Eagle medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and then cultured in a 5% CO2 incubator at 37°C. Hypoxic conditions were produced using D-Hanks solution (mM: 5.37 KCl, 0.44 KH2PO4, 136.89 NaCl, 4.166 NaHCO3, 0.338 Na2HPO4, 5 D-glucose, pH 7.3-7.4 at 37°C) saturated with 95% N2 and 5% CO2. The pH was adjusted to 6.8 with lactate to mimic ischemic conditions. The dishes were put into a hypoxic incubator that was equilibrated with 1% O2/5%CO2/94%N2. After hypoxic treatment, the culture medium was rapidly replaced with fresh DMEM with 10% FBS (10% FBS/DMEM) to initiate reoxygenation .
At 72 h post-culturing with 10% FBS/DMEM, the cells were randomly divided into six groups: (1) control group: cells were continuously cultured for 9 h with 10% FBS-DMEM; (2) H/Re: cells were placed in hypoxic culture medium for 3 h and then reoxygenated for 6 h by replacing hypoxic culture medium with fresh DMEM containing 10% FBS; (3) CaCl2 + NiCl2 + CdCl2-H/Re (Ca + Ni + Cd-H/Re): neonatal rat cardiomyocytes were treated with CaCl2 (2.2 mM), NiCl2 (1 mM) and CdCl2 (200 μM) for 30 min in hypoxic medium and then reoxygenated for 6 h by replacing hypoxic culture medium with fresh DMEM containing 10% FBS (CaCl2 is an activator of CaR, NiCl2 is an inhibitor of the Na+-Ca2+ exchanger, CdCl2 is a inhibitor of the L-type calcium channel; these drugs do not affect cardiomyocyte viability); (4) NPS-2390 + CaCl2 + NiCl2 + CdCl2-H/Re (NPS-2390 + Ca + Ni + Cd-H/Re): neonatal rat cardiomyocytes were treated with NPS-2390 (10 μM) for 40 min, and the following steps were the same as for group 3 (NPS-2390 is an allosteric antagonist of the group 1 metabotropic glutamate receptors); (5) 2-APB + CaCl2 + NiCl2 + CdCl2-H/Re (2-APB + Ca + Ni + Cd-H/Re): neonatal rat cardiomyocytes were treated with 2-APB (3 μM) for 40 min, and then other steps were the same as in group 3 (2-APB or 2- aminoethoxydiphenyl borate is a membrane permeable IP3R inhibitor); (6) Ruthenium red + CaCl2 + NiCl2 + CdCl2-H/Re (Ru + Ca + Ni + Cd-H/Re): neonatal rat cardiomyocytes were treated with Ruthenium red (10 μM) for 40 min, and then underwent the same steps as in group 3 (Ruthenium red is an inhibitor of mitochondrial calcium uniporter ).
Cardiomyocytes were fixed in 10% formaldehyde in phosphate-buffered saline (PBS) for 10 min, permeabilized with 0.1% Triton X-100, washed three times in PBS and blocked in PBS containing 5% bovine serum albumin, 5% horse serum and 0.05% Triton X-100 for 1 h at room temperature (RT). Specific subtype anti-IP3R rabbit polyclonal antibodies were incubated overnight at 4°C at 1:200 or 1:100 (Santa Cruz). FITC-conjugated anti-rabbit IgG was used as a secondary antibody. As indicated, some cells were stained with 4-6- diamidino-2-phenylindole (25 μg/ml) (DAPI, Roche) for 1 h. The results of immunocytochemical staining were read and recorded with a laser confocal scanning microscope (Olympus, LSM, Japan).
3-(4,5-dimethyl thiazol-2yl)-2,5-diphenyltetrazolium bromide(MTT) assay
In the current study, cardiomyocytes were planted in 96-well plates. The MTT assay was performed as described previously . Briefly, MTT (Sigma) was added into the cell cultures at a final concentration of 0.5 mg/mL and the mixture was incubated for 4 h at 37°C. Subsequently, the culture medium was removed and DMSO was added to each well to dissolve the resulting formazan crystals. The absorbance was measured at a wavelength of 570 nm using a microplate reader (Bio-Tek Instruments Inc., Richmond, Va). Background absorbance of medium in the absence of cells was subtracted . Percent viability was defined as the relative absorbance of treated versus untreated control cells.
Apoptotic cells were identified by the distinctive condensed or fragmented nuclear structure in cells stained with the chromatin dye Hoechst 33342 (Sigma). Cells were fixed with 4% paraformaldehyde for 10 min at room temperature and were washed twice with phosphate buffer solution (PBS). Cells were then incubated with 5 μg/mL Hoechst 33342 for 15 min. Next, the cells were washed three times and photographed using fluorescence microscope (Leica DFC500 System; Leica Microsystems, Bannockburn, Ill). At least 500 nuclei from randomly selected fields in each group were analyzed for each experiment, and the percentage of apoptotic cells was calculated as the ratio of the number of apoptotic cells versus the total cells counted.
Neonatal rat cardiomyocytes loaded with Fluo-4 AM, Fluo-5N AM and X-rhod-1 AM and cell permeabilization
[Ca2+]i was determined as previously described . Briefly, cells were seeded on the culture slides. After experimentation, cells were loaded with fluo-4 AM in 1% working solution at 37°C for 1 h, washed three times with Ca2+-free PBS to remove extracellular fluo-4 AM, and diluted to the required concentration. The reagents were added in Ca2+-free solution (145 mM NaCl, 5 mM KCl, 1.0 mM EGTA, 1 mM MgCl2, 10 mM HEPES-Na, 5.6 mM glucose, pH 7.4). Fluorescence measurement of Ca2+ was performed using a laser confocal scanning microscope (Olympus, LSM, Japan) at an excitation wavelength of 485 nm for [Ca2+]i and an emission wavelength of 530 nm for [Ca2+]i, using the equation [Ca2+]i = Kd[(F -Fmin)/(Fmax - F)], where K d is the dissociation constant (345 nM for fluo-4), F is the fluorescence at intermediate Ca2+ levels (corrected from background fluorescence), Fmin is the fluorescence intensity of the indicator in the absence of Ca2+and is obtained by adding a solution of 10 mM EGTA for 15 min, and Fmax is the fluorescence of the Ca2+-saturated indicator and is obtained by adding a solution of 25 μM digitonin in 2.2 nM CaCl2 for 15 min. Final values for [Ca2+]i are expressed in nanomoles.
To determine [Ca2+]SR, cardiomyocytes were treated with Fluo-5N acetoxymethylester (10 μM) for 2 h and deesterified for 1.5 h. For intact myocytes, the superfusate contained (in mM) 140 NaCl, 4 KCl, 1 MgCl2, 2 CaCl2, 10 HEPES, and 10 glucose (pH 7.4, 23°C). For permeabilization, myocytes were exposed to solution (in mM: 0.1 EGTA, 10 HEPES, 120 K-aspartate, 1 free MgCl2, 5 ATP, 10 reduced glutathione, and 5 phosphocreatine; pH 7.4) and then permeabilized using saponin (50 μg/ml) for 20 seconds. Excitation was set at 488 nm and emission was measured at 530 nm at room temperature . Images of fluorescence reflecting [Ca2+]i and [Ca2+]SR were recorded using a laser confocal scanning microscope (Olympus, LSM, Japan). There were more than 10 cells to be analyzed in each view and quantified using the analysis software for the microscope.
Recent study showed that the mitochondrial Ca2+ concentration ([Ca2+]m) consistently increases during reoxygenation . Therefore, [Ca2+]m was measured at 60 min post-reoxygenation. [Ca2+]m was determined according to the manufacturer's instructions (Molecular Probes). In brief, the cultured cardiomyocytes (1 × 106 cells/sample) were initially washed with HEPES buffer containing (in mM) 130 NaCl, 4.7 KCl, 1.2 MgSO4, 1.2 KH2PO4, 10 HEPES, 11 glucose, and 0.2 CaCl2 at pH 7.4 and then stained with 5 μmol/L X-rhod-1 AM for 30 min at room temperature. To avoid deesterification of intracellular X-rhod-1 AM in the cytosolic compartment, which would interfere with the detection of [Ca2+]m, the cardiomyocytes were rinsed and incubated with 100 μM MnCl2-HEPES for an additional 20 min to quench the cytosolic Ca2+ signal . Fluorescence measurement was determined using a fluorescence plate reader (CytoFluor II; PerSeptive Biosystems; Framingham, MA) at an excitation wavelength of 580 nm and an emission wavelength of 645 nm for [Ca2+]m. To validate the measurement of [Ca2+]m, the cultured cardiomyocytes were transferred into a slide chamber after X-rhod-1 AM staining and were placed on the stage of a fluorescence microscope (×50 objective; Olympus). The images from the slides were captured using a digital camera connected to Image-Pro Plus software (Media Cybernetics; Silver Spring, MD). There were more than 10 cells to be analyzed in each view.
Measurement of mitochondrial membrane potential
Mitochondrial membrane potential (△ψm) was measured with a unique cationic dye of 5,5',6,6'-tetrachloro 1,1'3,3'-tetraethylbenzimidazolcarbocyanine iodide (JC-1), as previously described . Briefly, cells were seeded on culture slides and treated according to experimental protocols. Previous data demonstrated that [Ca2+]m might continuously increase during the process of reoxygenation and result in mitochondrial △ψm collapse , so we detected △ψm at 1 h after reoxygenation. At the end of the above-described treatments, cells were stained with JC-1 (1 μg/ml) at 37°C for 15 min and then rinsed three times with PBS. Observations were immediately made using a laser confocal scanning microscope. In live cells, the mitochondria appear red due to the aggregation of accumulated JC-1, which has absorption/emission maxima of 585/590 nm (red). In apoptotic and dead cells, the dye remains in its monomeric form, which has absorption/emission maxima of 510/530 nm (green). More than 100 areas were selected from each image. The average intensity of red and green fluorescence was determined. The ratio of JC-1 aggregate (red) to monomer (green) intensity was calculated. A decrease in this ratio was interpreted as a decrease in the △ψm, whereas an increase in this ratio was interpreted as a gain in the △ψm.
Identification of bax/bak translocation to the mitochondria and assay for cytochrome c release from mitochondria
Western blotting of cellular fractions was used to quantify changes in cytochrome c, bax and bak distribution within cells, as previously described . Briefly, 1 × 107 rat cardiomyocytes were homogenized in ice-cold Tris-sucrose buffer (in mM: 350 sucrose, 10 Tris-HCl, 1 ethylenediaminetetraacetic acid, 0.5 dithiothreitol, and 0.1 phenylmethanesulfonylfluoride; pH 7.5). After 10 min of incubation, cardiomyocyte homogenates were initially centrifuged at 1000 × g for 5 min at 4°C, and the supernatant was further centrifuged at 40,000 × g for another 30 min at 4°C. The supernatant was saved as the cytosolic fraction. The precipitate was re-suspended in the above buffer (containing 0.5% v/v Nonidet P-40) and saved as the mitochondrial fraction. The mitochondrial fractions were blotted with a primary rat anti-bax, bak and cytochrome c monoclonal antibody (Santa Cruz Inc.). The volume of specific bands was measured using a Bio-Rad Chemi EQ densitometer and Bio-Rad QuantityOne software (Bio-Rad laboratories, Hercules, USA).
Western blot analyses were performed as previously described . In brief, the protein concentration of samples was first determined using the Bio-Rad DC protein assay kit (Bio-Rad Laboratories, Hercules, CA). A total of 20 μg of protein was electrophoresed on a 12% SDS-polyacrylamide gel and transferred to nitrocellulose membranes (Amersham International, Amersham, UK). The membranes were blocked with 10% skim milk in TBST buffer (10 mM Tris, pH 7.6, 150 mM NaCl, and 0.1% Tween 20) for 1 h at room temperature and then incubated with a rabbit anti-BAP31 polyclonal antibody (1:500 dilution, sc-48766, Santa Cruz Biotechnology) overnight at 4°C. HRP-conjugated anti-rabbit IgG (1:3000 dilution, Bio-Rad Laboratories) was used as a secondary antibody. Specific bands were visualized with a chemiluminescent substrate (ECL kit, Amersham International).
Significance was evaluated using student's t-test, and p < 0.05 was considered statistically significant. Data are expressed as mean ± standard error of the mean (S.E.M.) and are representative of at least three independent experiments. [Ca2+]i data were obtained from 2-3 experiments, and 10-12 images were analyzed in each group.