Preparing cut colonies for fluorescence and confocal microscopy
1) Spread 200-300 cells on a standard agar plate (e.g. 2% potassium acetate, 0.5 % Yeast extract, 2% agar). Use a razor blade to cut colonies (~ 2 mm diameter) plus an underlying 3 mm2 cube of agar from the plate and place the block face up to a glass slide.
2) Overlay the colony/block with ~2 ml 2% agar kept at 42° C. Colonies can be kept in a humid slide box for at least several hours until the next step.
3) Transfer the agar embedded colonies to a rectangular glass coverslip (Fisher Scientific catalog#12-544-D).
4) Use a razor blade attached to micromanipulator w to slowly cleave the colony and block in half from top to bottom.
5) Use a second razor blade to pull away one side of the block, then manually trim agar containing colony to a 3mm cube. Tip the exposed face of the colony face down on the same coverslip. Spot 6ul of 50% glycerol on each end of the coverslip. Next, place a 100 mm X15 mm petri dish containing a 21mm x 21mm hole in its center over the slide and “glue” the plate to the slide as a result of the glycerol.
6) Samples were imaged using an inverted conventional fluorescence microscope or an inverted confocal microscope. Confocal mages were collected using LAS AF LITE software.
Useful Variations:
1) Staining GFP colonies with propidium iodide. In step 2 above, add 16 μg/ml (final concentration) to 2% agar at 60° C. Immediately overlay the agar block with this mixture. Incubate for 24 hr. at 30° C in a humid slide box. Cut and visualize colonies as above
2) Staining LacZ + GFP colonies with X-gal: In step 2 above, add 120 μg/ml of X-gal, 0.1%SDS and 6% DMF (final concentrations) to 2% agar at 60° C. Immediately overlay the agar block with this mixture. Incubate for 24 hr. at 30° C in a humid slide box. Cut and visualize colonies as above
