To begin, take one milliliter of a mid-log phase budding yeast cell culture, expressing mitochondria-targeted HyPer7. Centrifuge the cells at 6, 000 G for 30 seconds, and remove the supernatant, leaving 10 to 20 microliters in the tube. Re-suspend the cell pellet by gently mixing it with residual media using a micro-pipette.
Next, use an air blower or lint-free tissue to clean a glass microscope slide, and add 1.8 microliters of the cell suspension to the slide. Finally, cover the cells with number 1.5 glass cover slip by lowering it slowly at an angle to avoid introducing bubbles. Then place the slide on the microscope stage.
To image the cells, set up the acquisition conditions to ensure a detectable signal and acceptable resolution in each fluorescence channel. For example, on a spinning disc confocal microscope with an sCMOS camera use 2x2 binning, 20 to 40%laser power, and 200 to 600 milliseconds exposure. Next, examine the image histogram.
In a 12-bit image ensure the total dynamic range of the pixel values is at least several hundred gray levels without saturation. Additionally, ensure the range is one order of magnitude higher than the noise level. To calculate the noise level, measure the standard deviation of pixel values in a cell-free area of an image.
Then collect time-lapse images with no delay between acquisitions to evaluate the effects of the imaging conditions on signal stability and oxidative stress in mitochondria. Using the microscope acquisition software or ImageJ, measure the average pixel value in each fluorescence channel to ensure signal stability. If the experiment does not involve time-lapse imaging, ensure that the fluorescence is stable over two or three repeated Z-stacks.
Acquire images with a Z interval of 0.5 micrometers in a total stack depth of six micrometers for budding yeast. If collecting Z-stacks ensure that the Z interval is the same for all images in the dataset and include the entire cell. Also acquire transmitted light images to document cell boundaries.
