non specific bands - (Aug/27/2011 )

Hi everyone,

I am writing out of complete frustration.

I am trying to detect changes in expression of a protein from my cell line with western blot. I tried one antibody from abcam and used the standard western blotting method, and there was a band at 36 kDa instead of the expected 21 kDa. Then I called them up and they gave another antibody to try, and this time there was no band at all. They had three source of antibody from the protein but there was no review listed in the website for the protein, and apparently all three of them are 'low-seller'. Then I tried antibody from R&D since they do have pictures of the blot run on different samples and it looked promising. I tried it and this time used 2% milk to dilute my primary and secondary solution. Then finally I have bands on the region of 21 kDa, but the problem is there was also one major non specific binding at about 100 kDa, and LOTS of minor ones. Imagine ponceau red on the blot.. that's how it looked like. If I just ignored the non specific bindings on everything above 21 kDa, then I can conclude that my treatment had no effect on the protein expression.

Another time I blocked in 2.5% milk for both primary and secondary, and instead of incubating for one hour of primary at room temperature, I did overnight at 4C, and the non specific binding was decreased much, but the band that I wanted also became faint. Today then, I tried using 2.5% BSA for both primary and secondary, and I got a pretty dark background, but I can still see my bands, again at the right place but with huge amount of non specific bindings above the 21 kDa, and lots of minor ones across the vertical lane.

I am wondering therefore, if one can even trust that there is no effect of treatment considering the non specific bindings?

Any opinions, or suggestions on how to solve this problem, is appreciated.

zienpiggie on Sun Aug 28 00:29:54 2011 said:

Hi Cassandra,

Yes, I loaded paired samples of before and after treatment and all the banding patterns are the same, as well as the intensity of my protein of interest. I have not titrated my primary antibodies. I used a concentration that is within the range of what was recommended in the sheet from R&D. I should also mention that I loaded 100 ug of my protein. I realize that it is a lot of protein, but this is what I have seen of a leading author in the area has done. They loaded up to 150 ug of protein and the band is not super thick or saturated or whatever, which makes me think that it is low abundance protein to begin with. So I thought it made sense if the protein is present at lower amount, and I am using only 67% of what these authors loaded (from the same cell line), that I should start with relatively high dilution of my primary antibody. Do you think if I lower my primary antibody concentration then it will be more specific?

Titrating the antibody is always a good move when you first get it in. Often this will reduce non-specific binding. You should also titrate your secondary, if this is too concentrated it will tend to give high background. You can also make the binding more specific by making up the blocking solution in detergent solutions such as TBS with 0.1-0.5 % tween-20 (TBS-T) and by incubating at 4 deg C. Washing in TBS-T is also a good idea.

Some antibodies have non-specific binding no matter what you do, for example, BRCA1 antibodies have a massive non-specific binding above the actual protein band that is so strong that people cut the top off the membrane to eliminate this.

Hi Bob1, thank you very much for your input. I will try optimizing the primary and secondary antibody and see what happens. I am glad to know that the non-specific bindings above the protein has been observed before with other antibodies and may be it's how it's supposed to be.

hi zienpiggie...at which concentration of primary did you do your O/N incubation? As Bob suggested, longer incubation at 4 increases the specificity and you can also use a more diluted solution. Have you tried incubating the blots with just the secondary alone as a control? If it reacts and especially if it's the same bands appearing, then perhaps it's worth switching to another secondary....

as for the amount of protein loaded, that should be determined by the physical characteristics of the gel (i.e.- size, thickness, etc).

if you are going to use the same amount of protein as in the literature then you have to match the gel used.

Hi Cassandra and mdfenko,

Thank you for your inputs. As per the primary antibody, I used 10 ul per 10 ml of solution. The dilution was to follow R&D recommended ratio of 1 ug/mL for western blot. For the secondary antibody, I used 20 ul for 20 mL solution. I have used this secondary before for a different primary antibody and it was working fine at a pretty standard 30 ug of protein per lane. As for incubating at 4C O/N. I have tried that but I was losing my band too.

Regarding the physical characteristic of the gel, it did not say the size or thickness, but the gel percentage was similar and I have seen these authors loading from 75 to 150 ug of protein. BUT I was re-reading the article and then I realized they blocked at 2% skim milk and primary in TBST (3h, RT, no indication of the use of milk or BSA), and secondary for 1 hour in RT. Another point to mention, is they used antibody from santa cruz and I have learned from experience to avoid them.

I think I do need to titer my primary antibody and secondary and see if I can improve specificity. Otherwise I think I will conclude that my treatment has no effect and move on to the next enzyme in the list to look at.

Again, thank you everyone for your inputs.