Alamethicin

We measured H₂O₂ production with the ROS-Glo™ H₂O₂ Assay system. All compounds were tested at 10 µM in PBS or MEM plus or minus 1 mM sodium pyruvate. Compounds were tested in triplicate and each experiment was performed a minimum of three separate times. Stock solutions were prepared as follows: 10 mM menadione in DMSO, 70 mM pyrogallol in DMSO, 40 mM benserazide in DMSO, 10 mM eseroline in DMSO, and 10.2 mM alamethicin in ethanol. Vehicle was present at 0.1% in these experiments. The H₂O₂ assay was performed as directed by the manufacturer as follows. The compounds were added to an opaque white 96 well plate in the desired media or PBS. The H₂O₂ substrate solution was then added, bringing the final volume to 100 µl. The plate was incubated at 37°C in a 5% CO₂ incubator for 60 min. 100 µl of the ROS-Glo detection solution was added to each well at the end of the incubation. After an additional 20 min incubation at room temperature, luminescence was recorded using a GloMax^® Multi Detection System luminometer.
To confirm the selectivity of ROS-Glo for H₂O₂, 35 U of catalase was included in some of the 100 μl reactions. In order to estimate the amount of H₂O₂ generated by these compounds, a standard curve was also performed at the same time as the corresponding experiment. Concentrations between 0 and 30 μM H₂O₂ were used to construct the standard curve. A standard curve in MEM media was used to quantitate the data using MEM media or MEM with pyruvate and a standard curve in PBS was used to quantitate the data in PBS. Due to the large difference in background between samples in MEM, samples in MEM with pyruvate, and those containing catalase, all data points and the standard curve were background corrected using the appropriate controls. The equation of the linear line of the curve was then used to determine the amount of H₂O₂ generated in each sample.
Assays to determine the production of H₂O₂ in the presence of cells were performed as described above with the following modifications. The day before the experiment, cells were plated in clear, tissue culture treated 96-well plates at a density of 10,000 MDA-MB-231 cells per well in a total volume of 70 µl. Wells with cells were paired with corresponding wells without cells (medium alone). After overnight incubation at 37°C in a 5% CO₂ incubator, 10 µl of compounds or their vehicle in the appropriate medium were added, followed by 20 µl of the H₂O₂ substrate solution for a total volume of 100 µl. After the 1 h incubation at 37°C in a 5% CO₂ incubator, 50 µl of each reaction mixture was transferred to a white 96-well plate and mixed with 50 µl of the ROS-Glo detection solution. The plate was then incubated 20 min at room temperature before reading luminescence. The standard curve for these experiments was performed analogously to the experiment. Briefly, the incubation of the H₂O₂ samples occurred in a cell culture treated plate, after which 50 µl was moved to a white luminometer plate and mixed with 50 µl of detection reagent.

A previously developed method for measurement of mitochondrial calcium retention capacity (described in detail in^{19 (link)}) was used to measure the time of yPTP opening after Ca²⁺ addition. Briefly, isolated mitochondria were diluted in CRC buffer (250 mM sucrose, 10 mM Tris-MOPS, 10 µM EGTA-Tris, 5 mM P_i-Tris, 1 µM Calcium Green-5N (Thermo Fisher), 0.5 mg/mL BSA, pH 7.4) to a concentration of 1 mg/mL. The reaction was started by adding 1 mM NADH and 5 µM of Calcium ionophore ETH129. After equilibration for 1 min, 200 µM CaCl₂ was added. The rapid increase in the fluorescence of Calcium Green-5N after sometime was attributed to the release of calcium ions from the mitochondrial matrix into the buffer, likely due to the opening of the permeability transition pore. Matrix swelling was evaluated by measuring optical density changes at 540 nm with a Parkin Elmer Lambda 925 UV–Vis spectrophotometer. Mitochondria (500 µg/mL) were suspended in 2 mL of swelling buffer (150 mM sucrose, 10 mM Tris–HCl, 2 mM KH₂PO₄, pH 7.4) and then, 2 mM CaCl₂ and 10 µM alamethicin were added^{51 (link)}.

Intracellular BH3 profiling was performed as described (77 (link)) on primary lung fibroblasts. Single-cell suspensions were obtained from enzymatically dispersed naive and fibrotic lungs. Lin^– cells were column enriched by incubating with CD45, CD31, and CD326 MicroBeads and purified off LS columns per manufacturer’s instructions (Miltenyi Biotec). Lin^– cells were stained with Live/Dead (1:100, L34965, Invitrogen) and anti-PDGFRα, -CD31, -CD45, and -EPCAM (1:100, Invitrogen, Thermo Fisher Scientific) for 30 minutes. Cells were permeabilized with 0.002% digitonin in MEB2 buffer (150 mM mannitol, 10 mM HEPES-KOH, 150 mM KCl, 1 mM EGTA, 1 mM EDTA, 0.1% BSA, 5 mM succinate in sterile distilled H₂O at a pH of 7.5) and exposed to BIM or BMF (100 μM, New England Peptides), DMSO, or 25 μM alamethicin (Enzo) for 60 minutes. Cells were fixed with 4% paraformaldehyde for 10 minutes, then solution neutralized (1.7 M Tris base, 1.25 M glycine, pH 9.1), stained with anti–cytochrome c (1:40, 11-6601-82, Invitrogen) in intracellular staining buffer (2% Tween 20 and 0.1 g/mL BSA in PBS) overnight at 4°C followed by FACS analysis on the LSRFortessa, and analyzed with FlowJo 2 software. Depolarization (% cytochrome c loss) was determined by measuring median fluorescence intensity (MFI) of the FITC (cytochrome c) gate and adjusting for positive and negative controls: depolarization = 1 – (MFI_sample – MFI_alamethicin)/(MFI_DMSO – MFI_alamethicin) (77 (link)).