This is my third time western blotting, and it hasn't really improved with minor adjustments. I'm trying to blot the 97 kDa Fibroblast activation protien (FAP) in PC-3, DU-145, and LNCaP cells, with GAPDH as a loading control. Here is a short run through of my protocol:
-I seed 1 million cells into a 6-well plate and use 200 uL of RIPA buffer with protease/phosphatase inhibitors, but I keep getting a concentration of 1-1.5 ug/uL. This is all done on ice/4 degrees so that proteins remain stable.
-I load around 20-25 ug into each well, with 8 uL of Laemelli loading buffer and 1 uL DTT. This is run with tris-glycine running buffer for 90 minutes at 120V.
-I activate my PVDF membrane for 30 seconds with methanol, and then soak everything (sponges, filter paper, membrane) in tris-glycine transfer (+10% methanol) buffer for 10 minutes before assembling the transfer stack. Transfer is 1 hour at 100V, with ice surrounding the chamber to keep it cool.
-I'm using 5% BSA for 1 hour at room temperature to block, with a 1:1000 primary antibody dilution overnight at 4 degrees, and a 1:10,000 secondary antibody dilution the next day after washing.
My question is, why are the bands so faint, is there any way to improve this? And why did the last three wells run high?
Hay Allison,
I would recommend the following points:
1. Make the sample in whole-cell lysis buffer
(HEPES,pH7.4, NaCl, EDTA, DTT, NP-40)
2. Increase the loading amount to 40ug
3. Dont run the gel in 120V, instead,, run it within 100V
4. Increase transfer time to 90 minutes under 95V
5. Also activate the membrane for a longer duration depending upon the brand protocol
6. Use primary ab according to the manufacturing protocol
7. Use fewer secondary antibody as your image contains too much nonspecificity; you can use 5%Non fat dry milk for blocking
best,
Subhajit
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