The secondary antibodies I am using are:
1.Goat anti-rabbit (Alexa Fluor 488)
2.Goat anti-mouse 594 (Alexa Fluor 594)
The protocol I have been following:
- 0.2% Tween in PBS (10 mins x 3 with mild rocking)
- 5% BSA/10% Normal Goat Serum in Tween-PBS (blocking for 1 hr)
- Overnight incubation with 1% BSA in PBS @ 4C
- PBS wash (5 mins x 5 with mild rocking)
- Secondary (1:1000) in 1% BSA solution (*started of with 1 hr incubation in the dark @ RT)
- PBS wash as above
- DAPI counterstain
For now, I would suggest to use the blocking buffer to do the antibody dilutions.
More I formation is needed to understand where the problem is.
1. Is the sample cultured cells or tissue section?
2. If it is cultured cells, is it intracellular antigen to be stained.
3. What is the fixing agent?
4. If it is the tissue section, what is the thickness.
5. Do you have an unstained control? And do you see auto fluorescence?
Hello Rama, thanks for the suggestion. I'll try diluting using the blocking buffer too. The answers to your questions are:
1. They are cells on scaffolds.
2. I am looking for both intracellular and secreted proteins.
3. 10% Neutral Buffered Formalin for 10 mins.
4. Thickness of the section is 5 um
5. Yes, slightly because of the fixation protocol however, the intensity is the same throughout the sample which suggests that the cells do not show high autofluorescence.
I did not find any other problems in your protocol. You may have to try different detergents. Do you see this nonspecific in do get with both the second dart antibodies?
The following article may help you better. Good luck.
https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3216515/
Thanks UD. I have tried a couple of things you suggested here. For instance, I diluted the secondary antibody to my blocking solution which is 5% BSA/10% NGS in TPBS. Also, I tried incubating it for only 30 mins. Unfortunately, it gave the same result. Lastly, I tried a secondary from a different batch and it's still the same. May be I should ask the company for a replacement.
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