Bands dark on sides and light in middle - (Sep/27/2012 )
I'm having trouble getting a solid band for one of my proteins when I run a western. The edges of the bands are dark, and the center is light, leaving me with bands that look like a dumbbell, or like two dots with nothing in between.
Has anyone ever had this problem, or have any idea about what might be causing it? I've tried using lower and higher percentage gels, running at a lower voltage, making all new buffers, trying different types of loading buffers, etc., but nothing seems to solve the problem. Please help!
It is probably more related to your transfer. What size is the protein, is it basic/acidic, and what transfer buffer are you using?
I usually use this guide when referring to westerns: http://www.bio-rad.com/LifeScience/pdf/Bulletin_2895.pdf
The protein is fibronectin, so it's about 250 kDa, and my transfer buffer is 25mM Tris, 192 mM Glycine and 20% MeOH. If it's a transfer issue, I don't understand why it would more readily transfer the protein that was on the edges of the well, but not the protein in the center of the band?
How much protein per well? Antibody titers used? I think it may be local depletion of substrate around the area of the bands due to excessive secondary conj.
You can try to do a colorimetric development using TMB and see if you get better resolved bands. Or you can ponceau stain the blot before blocking and see if the bands look normal there.
From my cell lysate, I load about 5 ug total protein per well, so there's less of the protein I'm blotting for. Other people in my lab have titered the antibodies, so I am using the optimal antibody concentration.
I get the same results when I use different primary and secondary antibodies, so I don't think it's the antibodies. Also, when I do longer exposures, I will get solid, dark bands, but they're overexposed. I need my bands to be in the linear range so I can quantify them, but when they're in the linear range, they have this dumbbell shape, which makes them difficult to quantify.
Caribou on Sat Sep 29 01:04:45 2012 said:
The protein is fibronectin, so it's about 250 kDa, and my transfer buffer is 25mM Tris, 192 mM Glycine and 20% MeOH. If it's a transfer issue, I don't understand why it would more readily transfer the protein that was on the edges of the well, but not the protein in the center of the band?
Isn't fibronectin ~400kDa in size? From my knowledge, it's a huge glycoprotein. Would be good if you double checked your antibody specification sheet.
How did your loading control turn out? I.e. housekeeping proteins such as tubulin, actin or GAPDH?
What kind of SDS-PAGE gel did you use? homemade or commercial?
Yes, it's a disulfide-linked dimer of about 500 kDa, so the monomers run at 250 kDa when reduced.
My GAPDH looks fine, with solid bands even at the lightest exposures. All of the bands in my marker look normal too. In fact, every other protein I run yields normal-looking bands, except for this one.
We make our own gels. Because of the large size of the protein, I usually run a 5%/3% or 6%/4% resolving/stacking gel, and I always make sure the stacking gel is at least 1X the height of the well. I sometimes run a 8%/4% gel, and the edges of the bands are still noticeably darker than the middle, though it's less pronounced.
I would say try running your transfer with 10% methanol or transfer O/N at 30V if using a wet system.