I opted bodipy due to its smaller stoke’s shift (493/503) but my images do not have consistent results. in the same well few fields are clean without background and other fields have background. It looks where the cells are in group in the co-culture astrocyte and neurons there is no background. these cells are plated and fixed in 96 well and imaged in opera Phenix.
I
What is the catalog number of your BODIPY dye?
What is your fixation and permeabilization protocol?
How did you mount the sample, or is it imaged in buffer?
Bodipy dye catalogue no: D3922
Protocol:
1) Wash the cells after treatment and fix them in 4% methanol-free PFA (EMS: 15710) for 15 min.
2) Wash, permeabilize and block 10% NGS in 1xPBS 0.1% Saponin.
3)Add primary antibody [O/N 4 deg], wash and secondary antibody [1 hr RT].
4) Wash and incubate HOECHST [1:1000] for 15 min.
Bodipy prep:
In-room temperature PBS, add bodipy of 1uM concentration just before adding.
5) Vortex thoroughly and add immediately to the wells.
6) Incubate for 3min at RT and wash three times for 5-10mins.
7) Image in 1xPBS.
*I have tried adding bodipy to live cells and fixing them. Still has the background.
* Since I use 30 wells for imaging, I thought even slight drying could cause the issue. So, after removing the bodipy, I immediately added the PBS within a second and still have the background.
I am unable to figure out this strange phenomenon.
Now I realize that adding saponin is also not good because I am imaging the lipid droplets, and saponin removes the cholesterol molecules in the membrane.
As you suspected, in your last comment, BODIPY dyes are lipophilic, and as such, any delipidation of the membranes will lead to inconsistent labeling or remove labeling entirely. So use of detergents, even mild ones like saponin, can cause this.
Also, they are really only useful for live samples. Fixation with formaldehyde can also lead to inconsistent labeling, though it isn't as bad as detergents.
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