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Alkylation vastly changes the western blot signal of my protein

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#1 litrallis



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Posted 30 June 2014 - 09:56 AM

Hey guys.


I have this protein called STIM1, an ER calcium sensor. I am trying to run an oxidation assay on it, but I am only just beginning to tweak the protocol. This protein comes with 5-6 sulfur containing amino acid residues, and 600-700 total amino acids.


The sulfur residues exist at various states of oxidation, and so by using various alkylation-labeling and biotin methods, I aim to be able to measure oxidation of this protein under different conditions.



I am doing the following:


1a. Lysing Jurkat cells in the presence of iodoacetamide.

1b. Lysing another set of Jurkat cells in the absence of iodoacetamide.



The problem: Even when referenced to actin, there is a HUGE difference in antibody detection between these two lysis conditions.


I have tried three different polyclonal antibodies to 3 different epitopes (c-terminus, a residue near the middle, and the n-terminus) and each gives identical results.


Any ideas why?



(Rest of the protocol:)


2. Sonicating, spinning down to transfer the supernatant, then running 20ug of lysate mixed with Laemmlis buffer on a 10% SDS-PAGE gel, followed by wet transfer, and room transfer blocking, and incubating the primary overnight. I then do 3x TBST washes, do secondary in blocking buffer for 1 hour at room temperature, then do 2x TBST washes, and 1x TBS wash.


I'm imaging with a Licor Odyssey system, and my blocking buffer is a proprietary Licor buffer. It works for everything else.


My lysis buffer contains NP-40, glycerol, Tris, as well as aprotinin, leupeptin, pepstatin, and microcystin added immediately prior to lysis.

Edited by litrallis, 30 June 2014 - 09:59 AM.

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