Hi all !
I've used 30:1 acrylamide/bisacrylamide solution (Bio-Rad) for making WB gel.
Once I run the gel, the line in separating gel suddenly showed up (please see the attached figure).
It really looks like the line between stacking and separating gel.
To deal with this problem, I already changed all the reagent including Tris-buffer, SDS, APS.
What should I do for this gel?
Please answer or comment is find. Tnx
I guess it didn't affect you protein migration, am i right? Just that you would like to find out the reason for it. Did you used isopropanol or ethanol to stack on the gel during the polymerization of the separating gel? Probably the line visible due to improper removal of the organic solvent. Hope this help.
I can't really tell from your photos, but there is a salt front that migrates as the gel is running. You can see it move while the gel is running and it is normal. It is continuously horizontal right across the gel, looks like a clear refractive line (like the interface between stacking and separating gels), but the salt front generally runs ahead of proteins, which does not seem to be the case here, judging by the stained markers on the right. If you changed buffers partway through the run you might get a second salt front. Isopropanol and ethanol are uncharged and would no tmove through the gel.
If the line (visible refraction boundary) you are seeing is the salt front as described by Dr. Kartner above, it normally stains a different colour to the rest of the gel in colloidal Coomassie stain and appears as a diffuse / smudgy band running at around 5-10 kDa in a 15-16% gel.
If your protein runs close to this boundary and you cannot use a different gel composition (for instance - Tris-Tricine gels for lower molecular weight proteins), then you can try a trick we used to use with sequencing gels 20 years ago : pre-run your gel empty for 15-20 minutes *before* you load any of your proteins and molecular weight marker. This moves the salt front 10-15mm down the gel before your proteins load, and I find that it helps enormously with visualising proteins running at ~10kDa on 15% SDS- Glycine gels that otherwise I would need to use a Tris Tricine gel to resolve cleanly.
What I referred to loosely as a salt front is in fact two fronts, a leading chloride front and a trailing glycine front (a tricine front for tricine gels). In the stacking gel, while running, these two closely spaced fronts concentrate the protein sample between them into a tight band, then when they hit the separating gel, which has a basic pH, the glycine (or tricine) front overtakes the proteins and both fronts run ahead of the proteins. The proteins are then separated by size and charge (usually contributed mainly by bound SDS) in the separating gel. The salt front is visible in the running gel due to the refraction of the concentrated chloride and glycine fronts, but it does not stain. The small ionic species that make up the fronts diffuse away as soon as you put them in the staining buffer. If a continuous band is staining behind the salt front it is due to an impurity in the gel or top running buffer. Another point: If you prerun your gel you lose all the advantages of discontinuous SDS-PAGE, because you have turned it into a continuous system (the concentrating fronts are long gone before you load your samples). Continuous systems have their place... they are better for heterogeneously charged proteins, like many glycoproteins, and for some small proteins, but you get no stacking, so they only give sharp bands if your sample depth is kept to a minimum in the sample wells (meaning you need very concentrated samples, or a more sensitive staining method). It is pointless to pour a stacking gel if you are doing this, and you are better off using a bona fide continuous buffer system like the Fairbanks system, rather than the Laemmli system which was intended for discontinuous PAGE. If you are a beginner at doing SDS-PAGE, it is well worth the time reading one of the many reviews on theory of SDS-PAGE.
Hi everyone,
We're having the same problem. We see two transparent lines during the running and we also note that 25 and 20 MW band get stack in the upper transparent band. When we do the Coomassie staining we get a blue bar that is between the transparent lines we see during the running (Image attached).
We use this protein marker (DUAL COLOR) http://www.bio-rad.com/es-ar/product/prestained-protein-standards
I am aslo having the same problem as Julia. Did you manage to find out what it was?
The pink line might simply be a contaminant of the loading dye with somewhat higher electrophoretic mobility. Dyes are mass-produced for technical purposes, and everything but analytical grade. Looking at a thin-layer chromatogram of laboratory dyes can be a quite sobering experience (why then are they so @*&$ expensive?!).
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