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Western blotting for proteins in vesicles?


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#1 Choux

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Posted 18 December 2009 - 12:20 PM

Hello,
I am trying to measure total protein for a receptor which is membrane bound, either at the cell surface or stored in vesicles--not cytoplasmic. (The protein is P-selectin, stored in Weibel-Palade bodies in primary endothelial cells.) When using lysis buffer, our lab usually spins down the lysate to remove debris and also uses protein AG beads to remove Ig. I used the same lysis buffer but did not spin the lysate because I didn't want to lose the membrane fraction. Before loading the lysates, I sonicated for 2 minutes. The problem is that the signal was still very weak, and I don't think I am able to see real changes in protein amount this way.
I don't want to isolate the vesicles (I've read those protocols, they're too complicated for what I'm doing). I just want to make sure I'm not losing the membrane fractions by using the wrong lysis buffer. Someone mentioned doing a crude extract with TCA? Or just lysing directly with loading buffer? Does anyone have other suggestions for how I can make sure I retain a membrane bound receptor in the lysate?

Thanks!

#2 jangajarn

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Posted 09 February 2010 - 08:58 PM

If the protein is bound by actin or something you can try TritonX100 extraction to separate soluble and insoluble fractions. You protein if bound by actin would be in the insoluble fraction, and if not would be in the insoluble fraction. Either way, you win.

#3 NicoleValenz

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Posted 07 December 2010 - 07:41 PM

Hello,
I am trying to measure total protein for a receptor which is membrane bound, either at the cell surface or stored in vesicles--not cytoplasmic. (The protein is P-selectin, stored in Weibel-Palade bodies in primary endothelial cells.) When using lysis buffer, our lab usually spins down the lysate to remove debris and also uses protein AG beads to remove Ig. I used the same lysis buffer but did not spin the lysate because I didn't want to lose the membrane fraction. Before loading the lysates, I sonicated for 2 minutes. The problem is that the signal was still very weak, and I don't think I am able to see real changes in protein amount this way.
I don't want to isolate the vesicles (I've read those protocols, they're too complicated for what I'm doing). I just want to make sure I'm not losing the membrane fractions by using the wrong lysis buffer. Someone mentioned doing a crude extract with TCA? Or just lysing directly with loading buffer? Does anyone have other suggestions for how I can make sure I retain a membrane bound receptor in the lysate?

Thanks!


If you just want total P-selectin, and don't need to differentiate between membrane bound and WPb/intracellular, then definitely TCA extraction will work. Don't spin down because you'll lose the membrane fraction, but make sure to sonicate thoroughly so the lysate runs smoothly.
A problem that I encountered in blotting for P-selectin is that the antibodies I tried (from Santa Cruz and somewhere else) recognize nonreduced protein only, though the data sheet didn't stress that fact. If your signal is really weak (in my case, it was weak and there were multiple bands around the right molecular weight and a strong one that seemed too "small") try leaving out b-mercaptoethanol or any other reducing agent. You'll have to play with the SDS concentration to get a sharp band, but running nonreduced lysate totally solved my problem and I got a strong band around 140kDa.
One more consideration is that people have reported downregulation of P-selectin with increased passage number in primary endothelial cells. If they're high passage, then maybe they have very little P-selectin expression.
Good luck!

Edited by NicoleValenz, 07 December 2010 - 07:43 PM.





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