Hi everyone.
So I have tried to optimize my phospho-antibodies for western Blotting, however after several attempts the one antibody, Phospho-4E-BP1, keeps binding to the wrong protein.
The antibodies I have been using are Phospho-4E-BP1 (Thr37/46) Antibody (CST #9459) and
Phospho-p70 S6 Kinase (Thr389) (108D2) (CST #9234S).
1. I have used a 10% acrylamide gel as well as the 12% gel. Only difference I saw is that the smaller mw protein of the ladder(Precision Plus- Western C) does resolve better using the 12%. The binding of the antibody remains the same ( at around 100kDa - 150kDa)
2. I have tested various blocking buffers 1% Casein/TBS vs 3-5% BSA/TBS, and the Casein/TBS produced better results between the two. A little bit more background than the BSA, however not nearly as much non-specific bands as with the BSA.
3. I have tested both antibodies, and it looks like the p70S6K antibody binds to the correct place, however the 4E-BP1 not.
4. For the primary antibody dilution I dilute 1:1000 in 1% Casein/TBS. Diluting it in BSA/TBS either gives me a ton of non-specific bands or ghost bands.
5. For the secondary antibody I make use of Goat Anti-Rabbit IgG HRP (ab97051) + StrepTactin HRP (for the ladder) with a dilution of 1:10000 in 1% Casein/TBS. Using BSA also causes a lot of non-specific binding.
6. For the 10% gel I ran it at 150V for 52min (only p70S6K was resolved and 4E-BP1 bonded to the incorrect place. I tried again using the P-4E-BP1 only on the 10% gel and run it at 200V for 36min. Wrong binding position again, but a much cleaner blot compared to the previous trial runs. Then I tried a 12% gel again (P-4E-BP1 only) running it at 300V for 23min @ 4 degrees celsius. Still binding to the same place as all the other blots.
7. The samples that I am using are liver samples and both protease as well as phosphatase inhibitors have been added to the lysis buffer before hand.
8. The membrane I am using is a 0,2 micrometer PVDF membrane.
I have attached a file showing one of my previous blots. Details of the blot is on the image. It is not my latest blot using 12% (could not download it from the lab's computer as of yet), however it does not look much different.
Please let me know if anyone might know what is causing this problem and how I can solve this issue.
Suggestions;
Decrease primary Ab dilution go up to 1:2500 atleast and 1:10000 dilution for secondary.
Increase concentration of blocking buffer as well as time upto 2 hr.
Washing steps after blocking, primary staining and secondary staining should strictly be done no less than 5 times.
Your antibody is supposed to crossreact with two other sites (4E-BP2 and 3) as well, so check their size as well.
in your fig:
I found a bright/white band at the size your protein should be. decrease the 2nd antibody and develop time. the bright/white band means too much HRP consume the substrate too quick, so, no signal there (lower than the overall background)
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