Fluorescent microscopy of neutral lipids in yeast

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Fluorescent microscopy of neutral lipids in yeast (S.cerevisiae)

This method describes how to visually analyze lipid droplets in baker’s yeast. The stained neutral lipids are analyzed by fluorescent light microscopy.

Step 1:
Grow yeast cells to the desired optical density (OD).

Step 2:
Prepare the following solutions:
    4% Paraformaldehyde fixative:
        - 100ml H2O + 700µl 6M NaOH, warmed up to 50-60°C, but not more
        - add 4g Paraformaldehyde powder (stored at 4°C)
        - after dissolving add 1,36g KH2PO4 and 100µl 1M MgCl2

        - filter solution; use fresh, or freeze in liquid nitrogen and store at -20°C

    Nile Red solution:               
        - Dissolve Nile Red in ethanol to final concentration of 1mg/ml and store at +4°C

Step 3:
Take 1,5 OD units of cells and spin down the cells at 4000 rpm for 4 min.

Step 4:
Resuspend cells in 100µl fixative and add 1µl Nile Red solution. Shake this suspension 20 min at room temperature in the dark.

Step 5:
Spin cells down (optional: wash 2 times with 1 ml PBS) and resuspend in 20µl PBS.

Step 6:
Microscopy: observe stained lipid droplets in GFP channel (Nile Red has less background in this channel).

Nile Red has a rather broad excitation and emission spectrum. In case of double staining, check for fluorescent interference with Nile Red of the used dyes, fluorophores. In addition, Nile Red is bleaching rapidly, even when exposed to UV light. Therefore, use bright-field imaging or any other wavelength, not touching the Nile Red spectrum, to set the focus plane.
Store the stained cells not more than 2 days.

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