Hi,
I've been having problems working with small organoids (around 100μm in diameter) for immunostaining. Whether I'm passaging them or fixing them for imaging, I consistently face the issue of losing organoids during processing. I have just one or two wells (24-well plate) at a time, I've been advised to use a 15ml tube for better handling. I typically spin them at 300g for 10 minutes, but after each spin, I struggle to locate the pellet, leading to potential loss. Given the small size of the organoids and the limited quantity, I'm seeking advice on the best approach. Many thanks
I'd try to mix the organoid suspension with molten agar or gelatine. After hardening, I'd do the processing like for tissue blocks.
As for finding small amounts of pellet after centrifugation, before spinning mark the tube where it faces the outside of the rotor.
First, it could help to place them in a smaller tube (e.g. 2ml Eppendorf). Suspending them in low melting point agarose will keep them together, but not sure if that is compatible with the immunostaining/imaging method you will be using.
Thus, my suggestion would be to place them in a small flow-through chamber/capsule, which can then be placed in a bigger vessel. There are such products eg. for TEM processing (freeze substitution/low temp embedding, critical point drying, or for grid staining), but you could also DIY.
Hi,
I was also used to include my organoid pellets in agarose before doing immunostaining. But then, you can only process then by cutting (to visualize the pellet I used a chromogenic dye before the agarose inclusion).
If you want to keep and visualize the entire organoid, you need to use small tubes, and very important thing, with V-shaped bottom. You can also slightly increase the speed of your centrifugation without damaging the cells. And of course, always keep around 100ul of supernatant to avoid aspiration of organoids. You can also try to use a chromogenic dye that would not interfere with your immunostaining, to vizualise your pellet.
Hope it helps,
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