Live-cell Imaging of Lysosomal Membrane Permeabilization During Necroptosis

This protocol describe a live-cell imaging method for monitoring lysosomal membrane permeabolization during necroptosis using fluorescent probes that detect change in lysosomal membrane integrity and acidification. The challenge in the field is to characterize the dynamic of organelle damaged during the cell death process, and to precisely define how this damage should drive, or contributes for cell demise. To begin, plate 5, 000 cells in two milliliters of antibiotic-supplemented DMEM media into a sterile glass-bottom dish.

Place the dish in an incubator set to 37 degrees Celsius with 5%carbon dioxide, and incubate the cells overnight. Add 20 microliters of dextran green beads to 25 micrograms per milliliter in fresh antibiotic supplemented DMEM media to dilute it. Remove the old medium from the dish and replace it with two milliliters of the beef-containing supplemented DMEM media.

Place the dish in an incubator set to 37 degrees Celsius with 5%carbon dioxide, and incubate for 24 hours to allow internalization of beads into lysosomes. Now add two microliters of one millimolar LysoTracker stock solution to two milliliters of fresh DMEM media. Replace the old medium with this staining solution.

Incubate the cells at 37 degrees Celsius with 5%carbon dioxide for two hours. Turn on the confocal system components by switching on the personal computer, microscope, scanner power, laser power, laser emission, and Leica power. Turn on the environmental controls, including the temperature controller, called the Cube, and the carbon dioxide gas mixer controller, called the Brick.

Open the carbon dioxide and oxygen supply. Set the temperature controller to 37 degrees Celsius. Set the gas and humidity controller to 5%carbon dioxide and 95%humidity.

Assemble the imaging chamber. After staining, wash the cells three times with two milliliters of PBS, then add two milliliters of fresh supplemented DMEM media. Apply immersion oil onto the objective lens before imaging.

Mount the dish on the microscope stage. Adjust the laser intensity up to 3%but not more than 5%Then adjust the detector gain in the microscope software to optimize the signal. Minimize the background by adjusting the contrast bar.

Acquire baseline images of untreated cells, which serve as the control condition. Return the plate to the laminar flow hood and add one milliliter antibiotic DMEM supplemented with cell death inducers. In the acquisition software, set the mode to XYT, format to 512 by 512, speed to 400, zoom to three, line average to one, line accumulation to three, frame accumulation to one, time interval to three minutes, duration to eight hours, and enable auto-focus.

Click start to begin the time-lapse acquisition. Initial imaging confirmed the presence of stable puncta of LysoTracker and fluorescein-labeled dextran, indicating intact lysosomes before treatment. Four hours after treatment with TSZ, LyoTracker fluorescence was noticeably reduced, while dextran began to show diffused cytosolic distribution, indicating initial lysosomal membrane permeabilization.

By eight hours after treatment, LysoTracker's signal had almost completely disappeared, while many cells retained green dextran puncta with diffused green cytosolic signal. We demonstrate that the lysosomal membrane permeabolization can be directly visualized in a live-cell undergoing necroptotic induction. This protocol demonstrate that the combination of both LysoTracker and dextran beads enables simultaneous monitoring of lysosomal pH and cargo release in real-time.

And important to advance of this live-cell image protocol is its ability to monitor a small pH change and lysosomal membrane permeabilization in real-time, capturing dynamic events that are missed by endpoint analysis such as flow cytometry and in monolayer staining.