I stripped several western blots with 62.5mM Tris-HCl (pH 6.8), containing ß-mercaptoethanol and 2% SDS for 1h at 50°C. After this the FLAG epitope bound to my protein was not detectable anymore although the tag was previously proven to appear on the same blots. It seems, that the stripping somehow interferes with the FLAG tag, as other tags or proteins on the blot are not affected by stripping in their detection.
Has anybody experienced this before or knows an article where this did work.
Hi,
I usually analyze the flag-protein first. But your stripping might be to strong. You could use mild stripping buffer ( https://www.researchgate.net/post/What_is_the_best_stripping_solution_recipe_and_method_for_Nitrocellulose_and_PVDF_membrane ). I use the buffer only once for 5 minutes, then wash 10 min with PBS, and 10 min with TBST. I still get detection of proteins even after 4 strippings.
Thank you, I'm trying the mild stripping at the moment. Its just odd that with the stripping method I applied, i can detect all my proteins and other tags even after 3 strippings but FLAG is missing after the first one completely.
You say that you usually detect FLAG first is there a specific reason why FLAG first? And did you ever experience good signals with FLAG after stripping the membrane with your method?
From my experience, these are strong stripping conditions. When I do need to use strong stripping buffer containing β-mercapoethanol, the membranes are incubated at 50°C for 20-25 min. However, I usually use mild stripping buffer: (for 1 litre buffer, pH 2.2: Glycine, 15g; SDS, 1g (or 10ml of 10%SDS); Tween 20, 10ml). Incubate membrane (shaking) in this buffer at room temperature for 5-10 min, repeating this step with fresh buffer. Wash membrane twice in PBS for 10min. Then wash twice in PBS/Tween for 5min. Proceed to blocking buffer. Using mild stripping conditions, I have probed membranes for FLAG and V5 epitopes. Anti-FLAG M2 antibody has been used throughout our studies.
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