Dear all,
I'm currently trying to detect the signals of intracellular LC3 in isolated eosinophil from human blood. After being purified (the purity is ~ 99%) and treated, eosinophils are harvested, permeabilized immediately (with 0.1% Triton-X) and incubated with PE-LC3 antibody, and resuspended in 1X HBSS. Recently I got two problems:
1/ Another cell populations shift towards SSC high/FSC high. This makes it harder to gate the general population.
2/ In some samples, signals of PE-LC3 were very high. So I think maybe it's due to unspecific staining.
Is there anyone has encountered these problems? I appreciate any help you may provide.
Do you fix your cells? If not, I guess your permeabilization would just cause them to swell badly, if not completely destroy fairly soon.
Also, I guess you have some surface marker to identify eosinophils... if so, even if the populations shift, you would be able to clean up your gate with surface marker staining.
To prevent non-specific binding, begin with blocking your cells with some protein/Ig containing solution (BSA, FBS, serum of the same origin as your antibody etc).
You also don't need to stain in presence of triton-x, as your permeabilization is permanent (as opposed to perming with saponin-containing buffers).
Dear Belkina and Lekishvili:
Thanks for your suggestions. Previously I tried to avoid fixation step (2% Paraformaldehyde, 10 min, RT) since I observed cell clumped together in SSC/FSC gate. Can you please tell me what's your protocol for fixation?
After antibody incubation, I washed cells once with staining buffer (we used HBSS 1X + 5% FBS). From your experience, is it appropriate?
Thank you,
Tu
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