Precast gels should be run for 90mins at 90V or 60 mins at 120-150V. The loading before separates to a lower band of 65bp and an upper band of around 235bp. The above times are to run the 65bp band to the bottom of the gel. Make sure to remove white strip at bottom of the pre-cast gels!
Home-made gels (TBE!-no SDS!) should be pre run for at least 1 hour before loading. Also, lanes must be washed thoroughly using a syringe or 1ml Gilson pipette to remove unpolymerized acrylamide.
The buffer for running and transferring pre-cast gels is 0.5X TBE.
When transferring, ensure sponges are thoroughly washed before and after in fresh buffer. Place a sheet of PVDF or other spare membrane between the sponges and first piece of blotting paper- this prevents and dirt being transferred from the sponges to membrane, which would otherwise ruin the blot (or just buy new sponges). Transferring takes 1hr at 30V (can’t run it higher than that).
Remove membrane from transfer apparatus. Place on blotting paper and crosslink by UV treatment on the top side (with DNA/protein attached) twice at level C3. If not developing immediately, place in clear wrap plastic and leave in the fridge or freezer until needed. Developing- use waterbath or hot tap to heat blocking buffer (20mls) and wash buffer (25mls) to 60 degrees before use (dissolves the precipitant).
Steps 1-4 should be carried out on the belly dancer 1. Cover the membrane with blocking buffer - leave shaking for 20mins 2. Add 66.7ul of HRP conjugate to the blocking buffer- leave for 20mins. 3. Prepare 100mls (per blot) of 1X wash buffer (in ddH20). Wash blot briefly (1 min) then 4 more times for 5mins each (20mls at a time). Don’t try shorten this!!!! 4. Remove wash buffer and replace with 20mls (room temp) substrate equilibriation buffer-leave for 5 mins
5. Place the blot in clear wrap plastic. Mix 2mls of both of the HRP activation solutions in a sterilin. Cover the blot with the mixed solution and leave for 5 mins (exactly!) 6. Place blot in fresh plastic and develop immediately. (develop/wash/fix/wash)
General rules: These come simply from my personal experience so feel free to ignore them. DNA protein interaction usually take place at room temperature for 20-30 mins. After this, loading dye is added, the sample mixed and loaded. If your reaction buffer contains glycerol (~10%) it probably wont need loading dye to sink into the wells and will only be used as a visual aid. I avoid the use of loading dye where possible as the mixing step may cause some dissociation of bound protein from the DNA. Loading dye should be added to empty lanes for visualization of DNA markers during electrophoresis. The above protocol is for use with biotinylated probes which allows for the use of very small (pg) quantities of probe and for a reduction in the amount of protein used. Another methods is to stain by ethidium bromide. This is very convenient and quick but is not as sensitive as using biotinylated probes. More DNA (~1ng) must be used but samples can be run on 1% TBE gels (instead of acrylamide). Staining is with 1ug/ml EtBR and can be between 10-30mins. These are a great place to start if you just want to make sure your protein is functional and identify potential binding sites. Competative bandshifts alleviate the need for non-specific inhibitors such as poly-dIdC or salmon sperm DNA since the other probe competes with the bound probe. Non specific binding will occur at higher protein concentrations (all bands will start to shift) but a specific shift should be visible before this.