Keeping them still for hours (for lattice light sheet)

I’m working to completely immobilize L1 worms to image with a lattice light sheet microscope (for at least 6 hours). We’ve previously used varying concentrations of tetramisole hydrochloride, but are finding that they’re still quite active even at 40mM.

I’ve started trying sodium azide, but the worms continue to move until I hit them with about 8mM. While this solves the immobilization issue, I am afraid that this is also arresting development.

Any thoughts about a paralytic that we can use for several hours of imaging?

There are some previous threads that might help: 1, 2. Maybe others.

I wouldn’t recommend azide. It’s great for short exposures, but after 15-30 minutes the worms start falling apart.

Thanks for the response!

Unfortunately a lot of the suggestions I’ve seen on the forum are for brief amounts of time (15-40mins) or involve a coverslip to compress the worm against an agarose pad.

I should mention that we’re not able to use a coverslip on the lattice light sheet, and can’t use a high concentration of agar as it could create lensing effects.

Currently, I am using cell-tac to keep the worms adhered to the slide and covering them with 0.75% low-melt agar which has the paralytic mixed in. This slide is then submerged in buffer with paralytic for 6-10 hours during imaging.