3 replies to this topic

[icome]

My target proteins must stay in the native form for primary antibody detection. Hence, I always prepare my samples in a non-reduced and non-boiled conditon. However, during the detection of internal control--the beta-actin, the antibodies for actin are inapt to recognize the native form, resulting in weak and inconsistent signals. This antibodies work well on denatured samples. Dose anyone know the way to denature the proteins that were transferred on a membrane? I tried washing the membrane using 6M urea-PBST for 30min at R.T, but the signals were still weak. Is there another method to overcome this problem? Thanks for your replay!

[HomeBrew]

Interesting problem...  Assuming there's not a better antibody for beta-actin, could you load two samples before running and transferring the gel -- one that is boiled and reduced, and another that is not?

[icome]

HomeBrew, on Oct 7 2009, 09:08 PM, said:

Interesting problem...  Assuming there's not a better antibody for beta-actin, could you load two samples before running and transferring the gel -- one that is boiled and reduced, and another that is not?

Thanks for your suggestion. But I think if the internal control is carried out separately, it would not be a loading control for my target protein. They should be performed on the same lane.
Today I tried a high stringent condion. Wasing the membrane with 8 M urea-PBST plus 10% beta-ME for 1 hr, the result is no signal detected. It seems that all the proteins shed away.
Maybe it is impossible to do that. To denature proteins may also disrupt the binding between proteins and a membrane.  
I will look for another antibody working well on native samples.

[HomeBrew]

If you had a lane of denatured sample and a lane of non-denatured sample on the same membrane, and probed the western blot with both antibodies, you could do a ratio of signal strengths for each of the antibodies in each of the lanes and normalize.