Hello! I am having trouble staining for plasma membrane localization of a transporter. This transporter also resides intracellularly, however, I am only interested in staining its plasma membrane location. The primary antibody that I used supposedly targets the extracellular/transmembrane portion of the transporter. My staining protocol:
(Staining in primary astrocytes)
1. Fix with 2% PFA 15min
2.Wash PBS 3x5min
3.Block 10% NGS
3. Primary incubation overnight in PBS-BSA
4. Wash 3x10min PBS-BSA
5.Secondary incubation in PBS-BSA
6. Wash 2x10min PBS-BSA
7. DAPI staining
I have attached the imaging results below. As you can see the red signal is low and there are a lot of dots. Does anyone have any suggestions?
1) PFA is a very poor fixative if you want intact membranes. I'd try using just glutaraldehyde or glutaraldehyde and PFA mixed.
2) You must ensure that there are no large osmotic differences between the cells you will fix and the fixative you use (osmotic gradients).
Hello,
Chemical fixation permeabilize cell membrane. You could try to label the membrane on live cells kept at 4°C on ice to avoid membrane movement then fix them.
Use low glutaraldehyde concentration. With high concentration, you may lose the recognition site for your antibody and you will have high green fluorescence background.
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