Western Blots on Membrane Proteins
Posted 11 November 2010 - 01:15 PM
I have been able to visualize GAPDH loading control perfectly, which makes me think that my procedure is not completely wrong. I have introduced protease inhibitors to my samples, and keep them on ice at all times to avoid protein degradation. I have beta-mercaptoethanol and SDS in my loading buffer. I have tried varying boiling times for about 5-10 min at 95C. I have tried incubating the primary antibody in blocking buffer, without blocking buffer. I have changed blocking buffers as well from BSA to Normal Goat serum, also filtering this solution. Also, since it is a rather large protein, I have only 10% methanol in my transfer buffer.
I have tried using multiple cell lines, and different lysates to ensure that my samples were not bad. I have also tried different primary antibodies, with the same results.
I am really at a loss of where to go next. All of my exposures end up looking like weak signal in a laddering sort of fashion with non-specific binding, and nothing where I expect the band. Do any of you think this has to do with the protein being a transmembrane protein? Is there any way you treat them differently?
If you have any ideas or suggestions I would VERY MUCH appreciate them!
Posted 11 November 2010 - 04:41 PM
One other transmembrane protein (CFTR) that I can think of doesn't do well with boiling, I think it is denatured at 70 deg C for 15 minutes in denaturing buffer.
How is your antibody stored, and how old is it? Do you have a positive control?
Posted 11 November 2010 - 04:57 PM
I have tried not boiling the samples as well but that gave similar results, but I could possibly try a less aggressive temperature like 70C. By denaturing buffer do you mean the electrophoresis loading buffer?
My antibody is only about 3 mos old, stored at 4C, per manufacturer instructions. I also attempted a different antibody that was brand new, which gave the same pattern on my blots. I think the next step would be the positive control lysate that the manufacturer (Santa Cruz) recommends.
What about playing with the reducing agent in my loading buffer. Any thoughts on the exclusion of that and if it could help?
Thanks so much!
Posted 11 November 2010 - 11:52 PM
Posted 12 November 2010 - 06:29 AM
Posted 12 November 2010 - 09:33 AM
As you explained, it really seems to me like the problem is your primary antibody. I routinely blot 250 kda proteins and boil them for 5 mins at 95 with absolutely no degradation.
Posted 12 November 2010 - 10:12 AM
I think that the boiling question is less about the size of the protein, but the fact that it is membrane bound. This is becoming a very puzzling dilemma....
Does anyone have an opinion on if my reducing agent (I've tried DTT and BME) is having an effect in all of this?
Thanks for the input!
Posted 12 November 2010 - 11:56 AM
And since you say that a lot of people do their WB with homemade antibodies, I think you have found the problem ^^
The ab from Santa Cruz can probably be optimized to give some results. But you could also try to get a sample antibody from another lab, or do your own. I would prefer the second option.
Posted 12 November 2010 - 01:30 PM
you may need to find another antibody that is recommended for immunoblotting.
genius does what it must
i do what i get paid to do
Posted 04 January 2011 - 06:01 PM
I had similar troubles until a labmate told me some membrane-bound proteins are better detected using TCA to extract the lysate and DON'T SPIN DOWN! You'll have to sonicate the lysate to make it clear before you run it, but changing to 10% TCA made all the difference for me.
Boiling, DTT/b-mercaptoethanol, etc. are important only if you're concerned about the state of the protein that the antibody recognizes--native, reduced. If you're spinning down, then you're losing the membrane fraction and this could account for your low signal.
Edited by NicoleValenz, 04 January 2011 - 06:04 PM.