Hi everyone,
I have been attempting Electrophoretic mobility shift assay to detect the formation of DNA:DNA:RNA triplex structure and to standardise it the assay, I have tried setting up binding between a previously reported triplex forming 16-nt dsDNA and 20-nt RNA. Briefly, I end-label ssDNA oligo with T4 PNK in presence of gamma-32P ATP. This is then annealed with the complimentary DNA oligo. I initially set up annealing at 1:2.5 molar ratio of labeled:unlabeled oligo and observed two bands in my probe only lane. This was resolved when I set up annealing at 1:1 molar ratio of the two oligos.The labelled probe is subjected to purification by passing through sephadex G-25 column. Then triplex binding reaction is set up with 10,000cpm of dsDNA in buffer containing 10mM Tris pH7.5, 25mM NaCl and 10mM MgCl2 in the absence (probe only) or presence of increasing molar concentrations of Triplex forming RNA oligo that has been heated at 70degree celsius for 10 mins and immediately transferred to ice. The reaction is incubated for 2 hours at RT and then resolved on 10% native PAGE in 1XTBE plus 10mM MgCl2 at 150V for 3 hours. What I expect to see is a reduction in the mobility of DNA:DNA:RNA complex as compared to my dsDNA only probe. However, what I observe is a band with increased mobility ahead of my free probe along with free probe in all reactions containing TFO RNA but no lower mobility triplex complex. It was suggested that the high mobility band could be labelled oligo that is getting displaced by RNA oligo from the DNA duplex. If true, is there any way I can stabilise the dsDNA duplex? If not, what could the problem be? Any suggestions would be helpful.