Hello Dear colleagues,
I need some help with NFATc2 (NFAT1) westernblot in Jurkat T cells.
I induced Jurkat T cell activation with CD3+CD28 antibody and goat-anti-moue IgG then collected the cells at different time points and checked NFATc2 with WB.
I lysis the cells using my lysis buffer (1% triton X-100, 150 mM NaCl, 50 mM Hepes pH7.5, 10% glycerol with protease inhibitor), then use supernatant to do WB.
With same samples I can detect NFATc1 and another kinase very well. But the NFAT1 bands seems odd. I got 5 major bands 100 kD, 80 kD and 50 kD. But according to the datasheet of the company, the band should be around 110-130. I did get some weak bands at around 120 kD, but it seems not the major band.
btw, I am using the NFATc2 antibody from Santa Cruz 4G6-G5, which is widely used to detect NFATc2.
Any suggestion will be appreciated.
Does your antibody recognizes phospho or dephospho NFAT? If you are not using phosphatase inhibitor you will only detect the lower (dephospho) band around 100kDa.
Hi Andre,
I think the antibody I am using (4G6-G5) doesn't differentiate the phospho-and dephospho-NFAT1.
You think the band I got around 100 kD is the dephospho-NFAT1? I didn't use the phosphatase inhibitor, and I will include next time.
Thanks for the pic.
Hello! I am using an anti-NFATc1 Santa Cruz antibody (7A6, sc-7294). The company predicts the molecular weight for the isoforms is 90; 110 and 140 kDa. My problem is that I hardly get to see these bands, and I have a very strong band between 37 and 50 kDa.
On the other hand, Sigma for its anti-NFATc1 shows a band between 40 and 65 kDa, although it predicts that the band is at 78 kDa.
Did something similar happen to someone?
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