Hello, I am new to designing flow cytometry experiments.
I have some cells transfected with EGFP using mRNA and want to quantify its percentage expressed. I also wish to differentiate between viable and dead cells, which i'm not sure which method yet.
Here is my current idea:
1. Trypsinze 1 x 10^6 cells, transfer to a tube for centrifugation at 300g, 5 mins
2. Wash cells by removing the supernatant and resuspend in PBS
3. Centrifuge at 300g, 5 mins, remove supernatant
3. Add 500µl 2%PFA to the cells at 4°C for 10 mins
4. Wash away PFA by centrifugation
5. Load it on the flow machine and detect at 510nm
Do I need primary and secondary antibodies?
How would the protocol look like if I chose not to fix my cells?
There is no need to fix the cells, just trypsinize them and resuspend them at an appropriate cell density in medium or PBS/BSA. EGFP is fluorescent itself so there is also do need to apply any antibody.
For the identification of dead cells you can add propidium iodide.
A collection of protocols you can find e.g. here https://cyto.med.ucla.edu/protocols.html
As mentioned by Sabine, there is no need for antibody treatment as you are using EGFP. additionally, you don't need to fix the cells unless you want to stop at this step (you can stock the fixed cells in PBS at 4°C for a few days) and then resume the other steps on another day.
Remember to wash the cells with PBS after the fixation step at least once.
Before moving the samples to the FACS tubes, make sure that you resuspend your cells in FACS buffer (ex: PBS, 0.5-1% BSA).
Mix the cells in the FACS tubes before loading them in the FACS Calibur (by gentle tabbing on the tube).
Good luck.