I am working on electro-mobility shift assay using unlabeled DNA that is a purified PCR product of a promoter sequence, and use nuclear extract for the binding reaction. For the nuclear extraction protocol I used these buffer recipes and followed an abcam provided protocol: Buffer A: 10 mM HEPES, 1.5 mM MgCl2, 10 mM KCl, 0.5 mM DTT, 0.05% Igepal, pH 7.9. Buffer B 5 mM HEPES, 1.5 mM MgCl2, 0.2 mM EDTA, 0.5 mM DTT, 26% glycerol (v/v), pH 7.9. I also used buffer B as the binding buffer. I need to optimize the DNA and proteins concentration and binding reaction condition. I incubated the binding reaction at 23 degrees for half an hour before loading on 1% agarose gel and on 6% TBE PAGE (with no loading dye). There was no shift, I also repeated the reaction with more protein and buffer concentration, another issue was that the protein had DNA contamination that interfered with the target DNA binding. Any suggestion for the binding reaction optimization would be very helpful and appreciated.
Dear Aya Aladdin,
In my opinion, EMSA is usually performed with the single purified protein/or protein complex to see sequence or structure specific protein binding to the DNA of interest. What is the goal of your experiment? I think, that there are many better methods for this - e.g. pull-down assay followed by 2D electrophoresis or mass spectrometry, if you are interested in which proteins bind your promoter sequence.
Best regards
Martin
But if you really interested in doing EMSA with nuclear extracts, I may recommend this article:
http://iai.asm.org/content/70/4/2238.full
especially this paragraph:
"Gel shift reaction conditions.
Nuclear protein extracts were thawed and desalted on P-6 columns prior to use. Each reaction mixture was assembled on ice and consisted of 5 μg of protein extract, 0.5 μg of nonspecific carrier DNA [poly(dI-dC) · poly(dI-dC)] and 10,000 cpm of labeled oligonucleotide in binding buffer (40 mM HEPES, pH 7.6, 10 mM NaCl, 1.5 mM MgCl2, 2% glycerol, and protease inhibitor cocktail lacking phenylmethylsulfonyl fluoride and dithiothreitol). The binding specificity was assessed with 100-fold excess cold specific or nonspecific oligonucleotide. For electrophoresis, 1 μl of gel loading buffer (25 mM Tris, pH 7.5, 4% glycerol [final]) was added and the entire reaction mixture was loaded onto a 4% acrylamide (60:1 acrylamide/bisacrylamide ratio) native gel in 0.5× Tris-borate-EDTA (1× Tris-borate-EDTA is 45 mM Tris, 45 mM boric acid, pH 8.3, 1 mM EDTA) containing 2.5% glycerol. Gels were run at a constant rate of 10 V/cm for 3 h at 4°C with a comb width of 0.5 cm."
Hope this help
Martin
Dear Martin,
Thank you for your answer. The problem is that I don't know which protein I am looking for, I need to know if anything binds to a target sequence then I will try to isolate the protein and identify it. I am currently optimizing the protocol using a promoter sequence (that is 500 bp long) as a positive control as it would bind transcription factors and make a shift. But since I am using unlabeled DNA, I cannot find a protocol to follow since I need to use higher concentrations of everything due to the low sensitivity of ethidium bromide compared to radio-labeling or chemiluminiscence. I tried three different DNA concentrations (5, 10 and 50 ng) in a 20 ul binding reaction, and 100 ng of protein that was not enough, I raised the protein concentration but that increased the gDNA contamination with it and caused another problem.
Dear Aya,
instead EtBr I use SYBR Gold, it is much more sensitive stain and not expensive (about 200 euro for 500 ul 10 000x stock solution - it is enough for staining of app. 50 agarose minigels, i. e. you take 10 ul stock solution and solve in 100 ml 1x TAE and stain your gel)
https://www.thermofisher.com/order/catalog/product/S11494
SYBR Gold is about 50x more sensitive than ethidium bromide.
Hope this help
Martin
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