Hey there,
Due to limited resource, we are trying to assess the interaction of a 50mer aptamer with a 25kDa protein domain. The protocol we are using is unusual in that the gel is 2.5% agarose with the running buffer being 0.5x TB, run at 20 V/cm for 10-30 mins. The intention of this was to increase resolution so we can get an approximation of Kd through gel image analysis software.
In the binding buffer (PBS with 0.55mM MgCl2), the aptamer is heated at 90, then put on ice before being held at room temperature to allow for folding. Following this, our target (diluted in our binding buffer from a 7.4 pH buffered 6M urea stock suspension) is added in increasing concentrations and held at room temperature for 30mins to allow binding to happen.
After electrophoresis and staining and destaining with RedSafe, we see the decreasing gradient of free oligo you’d expect to see, but no heavier band showing a gradient in inverse proportion to the free oligo band, indicating the presence of the complex.
Where‘s my complex at?
1. Is it possible that protein-aptamer complexes did not enter the gel, stayed in the gel pocket and were washed away? This could happen if the isoelectric point of the protein (or protein-aptamer complex) and the pH of the buffer are the same.
2. Cant't interaction of aptamer with protein interfere with RedSafe staining?
I would suggest to do EMSA in 5-6% native PAGE (19 acrylamide : 1 bis-acrylamide) in the same conditions you mentioned. You can label the DNA with phosphorus or order fluorescent oligo (its quite cheap).
How many binding sites are on your DNA molecule? Maybe the complex is just to heavy? If so, then maybe it would be good to shorten the oligo?
Yeshveer Singh Pavel Khramtsov good suggestions, theoretical isoeletric point is at least a pH point higher than the buffer so I don't think that's the issue. My fear is that the aptamer is not able to be stained when in complex. Will try a prestain now. Anton Slyvka yes, PAGE would be ideal, unfortunately we don't have the facility to cast PA gels yet and there are no native precast gels that will fit our PAGE tank. Only 1 binding site btw, I don't think complex weight is an issue.
Can you afford having fluorescent oligo? Would make you life easier. Also, if there is only one binding site there is no way that 25kDa protein needs 50mer for binding. I think that the footprint is max. 10nt. If you add another 10 for stability -> 20nt is sufficient for your experiment.
Another point: what about ion requirements of the binding? I see you add Mg2+ to the binding buffer. I would try adding it to the gel and running buffer too.
And out of curiosity: why do you have to keep your protein in 6M urea? It seems very denaturing...
Anton Slyvka the oligo does bind in its tertiary structure, it's called an aptamer and it requires the specific 50nt sequence to form that structure, this particular interaction having been shown previously. To fold into this structure and to bind to the protein requires the Mg2+, adding that to the running buffer is a good suggestion, but based on the gradient we've seen it seems that the binding is happening. Its 5nm Kd so it should stick regardless of the ionic composition of the running buffer. Regarding the 6M urea, that is a question I'd like to ask our supplier :P. My guess is it's how they purified it but it is buffered to neutral pH though so it should be okay.
Your right though, our budget and time is quite scarce but getting a fluorescent oligo would be our best bet at detection. I'll have to research whether the tag would interfer with binding or folding before hand.
Fergus Morris I see.
Have you tried to run it in cold buffer? If the secondary structure is what's being recognized then the high temperature (which always rises during the run) might be compromising it. As a result - no complex observed, even though the DNA disappears.
Based on what you've told so far and given your available equipment I would do the following:
1) supply the gel and TB (not TBE!) buffer with Mg2+; Remember, the DNA and metal ions have the opposite charges, so if they get separated in the field, your structure can get destabilized.
2) do the biding at +4*C or on ice;
3) run the gel with cold (chilled on ice) TB+Mg2+
I wouldn't add the dye before the binding, because as an intercalator it might interfere with aptamer folding.
Additionally, I would also try to stain the protein with coomassie (20% methanol, 10% acetic acid, coomassie) at room temperature. Might work, might not, but easy to check.
Please, let know what has worked. I find it potentially useful for my future experiments too :-)
Cheers
Anton Slyvka so we just ran the gel again with the same concentration of MgCl2 in the running buffer and gel. We also kept the running buffer nice and cool, initially chilled to 4*C with the electrophoresis tank being surrounded by ice-packs. Not only was the resolution of the poststain much higher, but we have been able to detect our complex! Thank you very much for your suggestion. This was done without the binding reaction occuring at low temp, as it has been previously shown that the complex forms at RT. Now, we just need to optimise the ratio of oligo and protein.
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