I'm trying to separate out low Mw proteins using 15% gels, however I'm getting a persistent problem where the stacking gel separates from the resolving gel during the run and I lose all my sample as it diffuses out from the interface. Any that does make it through doesn't run straight, and in any case the amount that makes it across in each case is variable between wells which makes it very unreliable to draw any conclusions. I've tried running it at low voltage (80V) to get it past the border but still get the same problem. I don't think it's a polymerisation issue as both stack and separating gels appear solid enough when I pry the plates apart at the end of the run. It also don't see the problem with my 10 or 12% gels.
I tend to cast the day before and leave them overnight wrapped in damp paper towels and cling film to keep things hydrated, and use isopropanol or butanol to level them off. I've tried washing the gel with Tris prior to adding the stack in case it's down to trace isopropanol/butanol but without luck.
I'm wondering if anybody can shed some light on what I'm doing wrong? Thanks!
When I cast the gels I simply use 1ml dH2O to level the running gel(because if you pipette on the top of it it won't mix well with the higher-gravity unpolymerised gel). After that you can just pipette it off from the gel, and the residual i-prop/butanol won't interfere.
When I met this problem I usually raise the SDS cc. by 1/5-1/4 to focus the gel.
I hope that will help!
Its not clear if you are adding the stacking gel on the same day. If not, then can try adding the stacking on the same day, like an hour later (for small or midi format gels) and then leave for overnight. Recheck your both tris buffer pH. Water saturated butanol is good enough for leveling off.
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