I'm currently doing a western blot to identify smooth muscle actin in cardiac myofibroblast cells. Initially when I did the westerns I use to get nice strong clear band at the expected 43kDa however when I have been repeating westerns on new samples (processed the same as the initial samples) and the band appears around 70-100kDa very clearly and strongly compared to a very very faint band at 43kDa. My first thought was that it may be a dimer but I already use beta mercaptoethanol in my sample with the loading buffer. I have also tried changing the primary antibody concentrations but this hasn't helped. I am going to try heating the samples for 10mins rather than 5mins at 95degrees to see if that will make a difference. If you have any suggestions or have encountered this problem , any input will be greatly appreciated. Thank you in advance!
Hello Tarbit, i would say perhaps your protein is post translationally modified. For example, it might be highly glycosylated and this makes the molecular weight higher compare to the calculated value
Use 5% blocking solution either milk or B.S.A. Load around 25 to 30 ug of protein. Do primary at RT for 2 hours not more than that. Secondary should be done with 2% blocking. Also after harvesting the cells you should wash with pbs three times to remove FCS and other contaminants.
PTM is definitely an option, but I would be very surprised to see such a huge shift (to approx. dimer size!) just by that. We often see an unspecific ~67kDa band which probably is BSA (I wash 3x with PBS before harvest but still sometimes cannot get rid of everything.)
I have had a similar problem with an IgG-Fc linked protein that was not properly dissolved into monomeric subunits. It seems that repeated freezing and thawing decreases the strength of beta-mercaptoethanol to reduce disulfide bonds (so we store it at 4°C and add it fresh to self-made Laemmli sample buffer). However, as far as I am aware, actin is non-covalently linked in its polymer form, so that should not matter as much.
In addition to Aftab Alam's suggestion, if you use self-made gels and sample buffer, I suggest you check that your SDS concentration is still the same, and that the reagent has not changed, expired or been damaged in any way.
Side note: I use 5% milk for blocking and found it to work better than BSA at reducing background on my blots. I usually block 2h at RT, then put on the primary AB over night at 4°C in blocking buffer, and the secondary AB in TBST for 1-2h at RT.
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