We are trying to image isolated live RBC on a glass-bottom dish in a WF incubated system.
Unfortunately, we have problems like that after several hours, we see round accumulations in the RBC that are fluorescent in all four channels (dapi/gfp/mcherry/far-red), and the cells deform and die.
I run out of ideas of how to make it work, and therefore I would appreciate it if someone could share protocols/tips on live RBC imaging. And specifically, what to use to attach the cells to the cover glass (0.01 polylysine deform them), tips on keeping them alive, and making them feel comfortable.
Are all the cells undergoing that transition or only the ones that are being imaged? It’s helpful to make sure you’re solving the right problem :).
If all the cells are doing that, I would first suspect (a) osmotic stress from evaporation and/or (b) pH stress if they are in bicarb media without CO2.
If you think you can get away with an anoxic environment for a little while, I find that suspension cells work great in a doublestick tape chamber. The confined space massively reduces lateral drift. https://openwetware.org/wiki/Koch_Lab:Protocols/Flow_cell
With live samples I prefer sealing the ends with something inert rather than nail polish, though do the experiment. Vaseline (put in 5ml syringe with p200 tip for careful application) or VALAP (1:1:1 by weight VAseline, LAnolin and Parafin wax) are my usuals, but I know folks who use dental wax and others.
The ibidi u-slides are a commercial options with similar properties. Before buying polymer bottom check that it works with your immersion oil (list in the product sheet).
Let me know if you have any follow up questions. Good luck!
Hi - I’ve used polylysine, but also selectin and a product called “CellTak” that is (was?) sold by BD. Like the other reply, be sure to address pH stability, which maybe you already are, minimize intensity, etc. Good luck!