I am studying DNA binding of a protein using EMSA. The reaction mixture contains 5 nM of DNA substrate while I vary the protein concentration from 2 to 50 nM. I see an increase in band shift with progressive increase in the protein concentration ultimately reaching binding saturation at 50 nM protein. However, it does not lead to 100 percent binding. What might be the reason behind the binding curve not reaching 100%?
It is true that the gel-shift assay is not an equilibrium measurement, since any ligand that dissociates can become separated from the receptor as they move through the gel at different rates. Because of this, gel-shift assays can produce smears for weakly interacting systems, in which the off-rate is relatively fast. For tight-binding interactions, such as the low-nM Kd system in this case, discrete bands are seen because the off-rate is low compared to the separation rate. Thus it is fair to consider it as approximating an equilibrium system and measure a Kd. One must also consider factors such as binding stoichiometry and cooperativity of binding.
Dear Adam. I have attached the gel picture for your kind view.I shall be looking forward to your reply.
Hi Andrei
I wish to state here that my protein is not sequence specific. It is AP endonuclease. So would the probable mutation in the oligos affect the protein binding ? Unless the abasic site isn't intact!!
I'm not sure there is really a problem here. It requires a very large excess of a binding partner over the Kd to 100% deplete a ligand. At the highest protein concentration (50 nM), the DNA is almost completely bound. The Kd appears to be about 4-8 nM. Under these conditions, I would expect a small amount of DNA still to be unbound at 50 nM protein.
Hi Taran,
Note also that gel-shift is not an assay at equilibrium: many things can happen during sample concentration within the well, entry in the gel and during electrophoresis, and these things can depend on the protein and the DNA you are using. I am not saying that it is the reason of your problem, but you must keep that in mind while interpreting results from gel-shifts.
Olivier
It is true that the gel-shift assay is not an equilibrium measurement, since any ligand that dissociates can become separated from the receptor as they move through the gel at different rates. Because of this, gel-shift assays can produce smears for weakly interacting systems, in which the off-rate is relatively fast. For tight-binding interactions, such as the low-nM Kd system in this case, discrete bands are seen because the off-rate is low compared to the separation rate. Thus it is fair to consider it as approximating an equilibrium system and measure a Kd. One must also consider factors such as binding stoichiometry and cooperativity of binding.
Thanks Adam, Oliver and Andrie for your answers. I shall take into account all points that have been discussed here.
However I wish to state here that my DNA is fluorescently labelled at 5' terminus and 4–6 nM fluorescein end-labeled nucleic acid is needed to achieve sufficient signal- for a quantitative measurement. Since, it has been suggested in various studies that for proteins that bind to their target sequences with an apparent Kd
Hi Taran, You are correct to apply the quadratic tight-binding equation. Unlike the simpler binding equation in which the receptor and ligand concentration factors are for the free species, in the quadratic equation you use the total concentration of each. AT and BT would be the total concentrations of DNA and protein, respectively, and AB would be the concentration of DNA-protein complex.
One thing that might cause trouble in fitting this equation accurately to get Kd would be if the fluorescence intensities of the bound and free DNA are not the same. You can test this out by adding the intensities of the 2 bands in each lane. If the bound and free intensities are the same, the total intensity of both bands should be the same for all the wells. It would be a good idea to use triplicates for each condition.
If you have access to a spectrofluorometer or plate reader capable of fluorescence polarization measurements, you can do this experiment more easily in solution, without the gel electrophoresis, and therefore without having to worry about whether or not there is a true equilibrium. Binding is measured as an increase in fluorescence polarization.
Hi Taran, I agree with Adam's comment that there may not really be a problem, but it is not obvious to decide on that without quantifying the amounts in each band and determining the apparent binding constant lane per lane - so not by curve-fitting - as we did with the 'Densitometric Image Analysis Software'. Note that this software may help you to accurately quantify the bands in your EMSA but should however not be used to determine binding constants in this case, because of the trace assumption. After quantitation of the bound and unbound fraction, you can determine the concentrations of unbound and bound DNA based on the originally added concentration, and therefore, you also know the concentrations of bound and unbound protein, and thus the binding constant corresponding to each lane. If you would observe a significant increase in observed binding constant with increasing protein concentrations, then you can conclude that there is a problem.
Article The 'Densitometric Image Analysis Software' and Its Applicat...
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