I tried to detect my recombinant protein using dot blot. After spotting the cell lysate onto the membrane, I let it dry at room temperature for 1 hour, and then proceeded with blocking and incubation using antibodies similar to Western blot. However, it seems that my protein couldn't bind to the membrane. Before blocking, I could observe the spots where I dropped the lysate. After blocking, those spots smeared and disappeared. Upon visualization, there was no signal at all, only some smearing. I also dropped the His-tag protein marker as a positive control, but that spot also smeared. I tried with both nitrocellulose and PVDF membranes, but the results were the same.
It does sound like your protein is washing off the membrane during the blocking steps. Hmm, you could try doing a brief heat treatment to help the lysate adhere, or maybe even pre-coat the spot with something adherent like poly-lysine. Do you not have a traditional Western blot setup?
Amy Klocko Yes, I have. I used western blot to detect this recombinant protein before and it was successful. Now I want to use dot blot for screening purpose.
Ah, that makes sense. Maybe you could try a brief electronic transfer of your dotted membrane? You could mix your lysates with some of the loading/cracking buffer, spot them, then do a brief "transfer" run with your standard Western transfer rig so the proteins really stick to the membrane (hopefully). It would be quicker than a Western blot.
Amy KlockoThank you for your suggestion! I'll try. I have read several dot blot protocols, and all of them involve spotting the lysate onto the membrane and letting it dry or using a vacuum. I don't know why my samples couldn't stick to the membrane.
I used to do dot blots with a vacuum. In thinking back, if I added too much sample (in my case purified cell wall extract) then the whole disk of material would sometimes float up during the wash step. More diluted samples didn't have this issue. Maybe that might help your samples too?
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