Transfer your fly cultures every 14-21 days. (Occasionally stocks with deleterious mutations may require longer to mature; in these cases, you can of course make exceptions.) Do not keep old cruddy cultures around. This encourages mites and other parasites to destroy our stocks.

Wipe down your dissecting area and the fly-pushing area frequently with ethanol. This also discourages parasitic infestation. (Wiping down with a few drops of concentrated acid gets rid of saline corrosion spots on the benchtop.)

If you are using daily saline perfusion, rinse ~20ml of distilled water through your setup at the end of every experimental day. Every month, replace the tubing connecting your saline reservoir with your recording stage. (You may have to do this more often if you use lipophilic drugs.) This prevents algae from growing in your tubing.

Before putting fresh saline in jugs, inspect for algae. Scrub interior of jug with detergent if needed, and rinse copiously. (Soap isn’t very good for neurons, either.)

Spilled salts on the balance will eventually corrode the top surface. If you spill salts, clean them up right away.

Objectives should be cleaned with lens paper and distilled water. Anhydrous methanol can also be useful for this task.

A handy trick for removing salt corrosion (noticeable as a chromatic aberration) from your condenser lens, etc. (if regular ol’ water and ethanol haven’t done the deed) is to try 0.1M HCl. Apply a few drops then gently wipe lens with lens cleaning tissue. Remember to blow off any potential grit with the canned air first so as not to scratch the lens.