I have always used TBE or tris acetate EDTA for EMSA, but I would not hesitate to use TGE: I see no reason for it not working, provided that the gel itself (and the buffer, of course) does not contain SDS.

The reason you can run SDS-PAGE on protein is because SDS binds the protein and denatured it and furthermore negatively charged the protein. Thus it can be separated only by size. Your protein binds DNA only when it is "alive". If you denature it, it will release the bound DNA. Try some other methods. Pull-down assay (based on histrap, ion-ex or other affinity column you used to purify the protein), ITC, fluorescence titration or anisotropy, NMR and so on.

Quian Wang is right that SDS will strip the protein off your DNA/oligo, but at least some of the precast tris glycine gels (for example TGX from BioRad - you need to check for your own gels), do not contain SDS, so they would be suitable for EMSA.

Our preferred method is to have the oligo target tagged with a fluorophore such as HEX or SIMA, but we found that Cybr gold is also good for both double and singlestranded oligos (~200fmoles sensitivity) if you have a laser gel scanning machine.

According to Geoff Margison's idea, he suggested a gel without SDS, I guess it is a native gel. If it is a native gel, then there might be a problem of running it. Because your protein is not highly negatively charged. Maybe it is even positively charged which you have to be careful.

Another quick and dirty way to do this is: assume you purify your protein with HisTrap column. You load you protein onto the column and then inject your DNA sample. Elute your protein after a bunch volume of flushing by the binding buffer. Test the UV280/260 ratio. If the ratio is smaller than before (

Quian Wang is again correct that mobility in a native gel will depend on the protein's charge. But the DNA to which the protein might (or might not!) bind is negatively charged. So what we see with oligos binding our proteins in native gels is retardation of the oligo migration, and that can be in multiple steps if there is cooperative binding. Of course we dont know the extent to which retardation is a function of the size of the oligo-protein complex and/or neutralisation. But since our protein by itself does migrate in these gels (as shown by blotting and probing with an antibody) it is probably more a function of complex size.

But we obtained good binding data (on rates, off rates, KD) using surface plasmon resonance, which has the same principle as the method described by Quian Wang, except we bound our oligos to the SPR chips via biotin.

I agree with Geoff Margison. SPR is a better way to quantify the binding rates. Make sure the biotin and your protein won't compete and bind the same site of your DNA. Otherwise, hopefully not, if biotin binds DNA tighter than your protein in the same site, you will not see any binding signal. Best of luck.

My tris-glycine are without SDS of course.

I do not have access to SYBR Gold, so I will be using GelRed which I'm hoping will give the same results ( I have access to a GelDoc imaging machine).

Thank you for your input !

Gel Red works well for DS oligos (and DNA) - sensitivity down to about 200fmoles with our phosphorimager. Not good at all with SS oligos, especially short ones (~20 or less). Are you using oligos as a target?

Just to clarify an earlier point: for SPR, our oligos were synthesised with a 5'biotin tag. If thats not possible for you you can add a 3'biotin using terminal transferase and dideoxy-biotin UTP to DNA or oligos. SPR is really informative!!

If you're going to use a gel shift I agree with the comments above that say you should omit SDS. However, this then leads to the issue of the protein migrating inefficiently or in the opposite direction, as also eluded to above - which becomes a problem when you are running a no-DNA control, or when you have samples that don't bind the DNA present.

In the absence of SDS the protein's migration will depend on its inherent charge; if it is net zero it may remain in the well, or if positively charged it may migrate out of the well entirely but rather than into the gel it could be lost to the running buffer.

A trick to overcome this problem is to load your samples into the wells dry (ie without having first immersed the gel in running buffer) using a dense loading buffer and then float over the top of them (in the well) a high percentage (higher than your main gel) polyacrylamide mix. Allow this to set so that it forms a plug above the sample in your well. Then add the running buffer as you would normally and perform the electrophoresis. Proteins that (in the absence of DNA binding) would otherwise migrate into the running buffer and be lost will instead become trapped in the high percentage acrylamide plug. This allows you to visualise them, and if desired make an assessment of their relative amount.

I used this trick a lot in the past to perform EMSAs on a range of DNA binding proteins.

Alternatively you could try pouring a native acrylamide gel in a DNA (agarose electrophoresis) mold - to generate a horizontally orientated gel. In this case you could place your well forming combs mid gel, providing you with the ability to track migration of any species in either direction (ie towards either pole).

This is something else I've done in the past, which also works nicely, but which did generate necessarily thicker gels.

Hope that helps.

Uptade :

My first test wasn't too bad. I am getting a good quality gel with crisp DNA bands in both my DNA ladder and samples. However, I do not see any difference between my negative controls (DNA without protein) and my samples (DNA with different concentrations of my protein).

Also, there might have been a problem with the oligos annealing step since I am seing 2 or 3 bands depending on the template (my DNA templates are different branched structures made from annealing 2-4 partially complementary oligos together). I will verify this hypothesis by agarose electrophoresis (without protein).

Do you know what the molar ratio of oligo to protein is? And do you know that the protein is active? How long are you giving the protein to bind to the targets, and at what temperature? Do you have any positive controls for protein binding? Are you running the gel surrounded by iced water and with cold running buffers?

Your oligos sound like a recipe for multimer formation!!

Geoff Margison I am using 30uM of hybridized oligos with 0-5uM of purified protein. My protocol is based on a paper that performed EMSAs using only the C-terminal domain of a protein containing the same functional domain as the protein I am studying. Since my protein is very small and only has that one functional domain, I am hoping to replicate the results they had.

My goal is to have multimers actually. I am trying to replicate branched DNA structure, for example a holliday junction by annealing 4 oligos that were designed to hybridize in a holliday junction shape. The point is to check wether my protein has a binding specificity for different DNA structures.

Unfortunately, I do no have a positive control, but I am expecting a binding activity in most of the DNA templates I use since this is what the article has shown, so my experiment is kind of a positive control itself.

Martin,

Interseting project!

All sort sof questions! How is the protein purified? If by affinity tag, was this cleaved off? Is the residual (or purified) protein exactly that predicted by the ORF (or with additional amino acids)? Any post translational modifications needed (i.e. what system was it expressed)? Does the protein need Zinc or other cofactor for DNA binding? My proteins lose a lot (sometimes, most!!) of functional activity (I know not why) on purification: is 5uM of your protein enough to compensate for that? Is it possible for you to make the protein from the original paper as a postitive control? Apart from structural features, does the original protein have any sequence proclivity (or do your targets have exactly the same sequence as in the original paper?).

The thing is, my protein is uncharacterized (yet), and it's so small that it only has one functional domain, which is shared with several endonucleases.

We have already proven that 4uM of this protein can nick dsDNA with magnesium as a cofactor. This was from a His-tag purified protein stock which I now use for EMSA. We basically just want to make a more complete "story" by showing it recognizes (binds) to specific DNA structures, which would make sense.

I will try to look for possible positive controls that would work against my targets.