You have no proteins in the gel at all or only some stay in wells?

I guess you're not interested in denaturing conditions if you want some interactions, are you?

Hi,

Did you check if your protein is active by other means? it seems that your proteins form big oligomers/aggregates that are too big to enter the pores of the gel. Or maybe simply they are not enough charged to migrate in electric field?

Re your proteins cationic? If so wouldn't migrate towards the anode on standard native PAGE.

One possible reason might be that your protein forms aggregates with SDS. You may try to acetylate your protein to help them to penetrate the stacking gel. More information can be found from: http://www.ncbi.nlm.nih.gov/pubmed/8874057

Hi it might be possible that ur protein have positive charge and thats why it will not migrates in the native PAGE, bcoz as we know DNA is negatively charge and thats why migrates towards positive charge in gel electrophoresis, i also did with my studies but didn't find any interaction, so might be this is the reason.............. more wl tell u latter wl try to find out

I think it depends on PH of the complex to enter inside gels. In native PAGE this is a common problem. use this conditions, It worked for my EMSA and several native PAGE very well. The main point is your protein after binding to DNA should be negatively charged. However, most of the time after addition of loading buffer (pH6.8), it becomes OK. Use HGMKE buffer (25 mM HEPES, pH 8 with 5% (v/v) glycerol, 10 mM MgCl2, 20 mM KCl and 0.1 mM EDTA) as a binding buffer.

Note; dont use stacking gel. It will be a continuous 6% resolving gel.

PAGE;

4X Tris-glycine buffer; 100 mM Tris-HCl (pH 8,6) and 800 mM Glycine

Acrylamide solution; 30 % acrylamide, 0.8 % bisacrylamide

APS; 10% (w/v)

Running buffer ;25 mM Tris-HCl (pH 8,6) and 200 mM Glycine

5X Sample buffer; 50 mM Tris-HCl (pH 6.8), 0.01% (w/v) bromophenol

blue, 25% (v/v) glycerol

composition of 6% gel;

4X Tris-glycine buffer; 2.5 ml

Acrylamide solution; 2.0 ml

Water; 5.4 ml

APS; 0.1 ml

TEMED; 0.008 ml

Run the gel at RT at low voltage not at 4 degrees.

Different reasons can explain that your protein doesn't migrate in the native gel :

- Multimerization or aggregation : complexes become too big to enter the gel, even at 6% acrylamide. To get rid of this problem, check for aggregation (UV, DLS...), and if it isn't aggregated, EMSA can also be made on agarose gels, which allows to deal with huge complexes.

- Your protein is too cationic : protein alone migrates in the wrong direction, and complexes with DNA aren't enough negatively charged to run in the gel. Eventually, if your protein is really too cationic, electric field can even dissociate the complex. To get rid of this problem, try using a buffer in the gel and in the tank of your electrophoresis apparatus with a pH not too different from the pI of your protein (Not more than one unit. For example use a buffer at pH 8 if the pI of your protein is 9).

Good luck

Hi the reason for this (if I understood the problem clearly) is that at the equilibrium not all the proteins bind the DNA but you have a mix of 3 species: the free (unbound DNA), the protein-DNA complex, and the free protein. Normally the protein concentration is much higher than the DNA concentration and so even if at a given concentration all the DNA is bound by the protein there will be a lot of protein still free in solution. If the pI of the protein is lower than the pH of the electrophoresis buffer then the protein will be negatively charge and move by itself (unbound) (in the direction of the DNA towards the "+") (in rare cases - never experience directly but I read about this specific event - if the charge of the protein is sufficiently high then the complex DNA-protein may migrate also very close if not lower than the unbound DNA (despite the complex being bigger its charge/mass ratio is higher and it moves faster). Otherwise the free protein will stay still (pI=pH) or move in the opposite direction (pI higher than pH).

If you use a competitor (rather than high salt concentration) to discriminate between specific and non-specific binding (like polydIdC) the protein should move anyway within the gel attached non-specifically to the competitor (invisible during detection anyway since it is not labelled like your target DNA).

Every specific DNA-binding protein in fact also binds non-specifically (lower affinity anyway) to ant given DNA because it is by this scanning mechanism that they find their binding site quickly (moving on a 2D rather than a 3D space). Because of this the Kd increases with the lenght of your DNA (labelled) normally.

Max.

Sorry the Kd decreases increasing the lenght of the DNA containing the binding site (i.e. the affinity increases).

I've recently been optimizing some of my EMSAs. Depends on the acrylamide...you can use 19:1, 29:1 or even 75:1 acrylamide:bis acrylamide in your 40% stock solutions. The higher that ratio, the more porous the gel. I would also suggest trying different concentrations of agarose gels.

I may not have understood clearly the problem since it is not very clear if the DNA which migrates is only the unbound or both (bound and unbound).

If only the unbound DNA migrates and the "shift" is missing there is likely too much protein in the well. I would reduce the amount of protein or as I said above use a DNA competitor like polydIdC and titrate the amount of protein as well. Usually (as a starting point could be ok) I was using 2nM DNA (labelled, 100-1000ng polydIdC and protein concentration from 0.1 to 5 micromolar in 20 microliter binding reactions.

Thanks to all of you for valuable suggestions. I have not a single protein. I am using the nuclear protein extract and incubate it with 250 bp DNA fragment which is PCR purified product, So I cant estimate the charge on protein. May be multiple proteins bind to DNA fragment. On 6% native PAGE I loaded the different conc. of proteins but kept the DNA conc. constant and there is decrease in the intensity of DNA when there is increas ein protein concentration but protien remains in wells.

I am attaching an image so that you can understand what i am trying to say.

I am using SYBR Green and SYPRO Ruby EMSA kit from Invitrogen

Dear Tomáš Hluska I am using nondenaturing conditions. When gel is stained with SYPRo Ruby, there is flourescence at wells. and this is increased with increasing protein concentration.

Anju Singh

Thanks for your response... I am using a portion of promoter so there are chances that proteins will bind with them and size is 260 bp.

Birendra Singh ·

Thank u so much for your suggestions. I will try thisI am thinking to move to Agarose from PAGe.

No I am not uisng Coomassie BLue for staining. I am using Sypro Ruby

Try using 2.5% stacking gel instead of 4%. I guess it should work.

But its handling will be difficult. 2.5 % is very delicate...:(

Though it is delicate, in our lab we routinely use 2.5% and it gives satisfactory results.

ok..Thanks.I will try

How was it ?

Farah

I also faced the same problem working with a purified protein from staph.

i used a gradient of regulatory protein to check 240bp prove.

the binding seems to start from 100nM but in none of the case the complex entered the gel. various standardizations were tried but still i found no improvements.

Recently i started using triton x 100 in binding buffer and found that the affinity increased 10 times. still the protein dna complex never enterd the gel

Sukhendu Mandal

What is the recipe of your binding buffer?

I am using the nuclear protein extract not a single protein..:( so its difficult to calculate the nM concentration of proteins, I can run ng or ug of proteins.Please suggest how much conc should I use . Now I am trying to do EMSA on Agarose gel

What is the composition of your running buffer?

Hi Farah! I think you should bear in mind that if your are using total cell extract, the fraction of your protein that will bind to the DNA probe is very small and likely you will not be able to see anything meaningful unless you radiolabel your probe or use other non-radioactive methods (DIG, biotin-dUTP, etc…). It is normal if you are using native conditions that a lot of protein may remain in the well, others will migrate in the opposite direction. Remember that in native conditions the migration of the proteins will be determined by their net charge at the pH of the running buffer.

Running buffer that i used is Tris-Glycine buffer for native PAGE and 1X strength was used.

Hello Farah,

I would say use first check the pI of your protein because that will give you idea about the net charge. Then use 2% agarose gel and cast the wells in the center of the gel for capturing the either side protein mobility. Gud luck.

Raj