Sypro Ruby Dye Front - (Jan/15/2015 )

Hi all!

I did an SDS PAGE using 4-12% bis-tris gel (samples preped with LDS sample buffer) and stained my gel with Sypro Ruby. My bands come up nice and sharp but the really annoying thing is my dye front is really obvious on the blot! (ignore the broken gel lol)

Is there any way I can get rid of the dye front? Perhaps by not having a dye in my sample buffer or something?

-tofuj-

if you poured your own gel then 2 possibilities:

1- didn't form gradient, just low concentration on top of high concentration gel

2- it's a buffer front, not just the tracking dye.

if purchased gel then it is reason 2.

if reason 2 then complete the run, don't stop the gel halfway. it could also be caused by decomposing sds in the buffer. if you make the buffer, try a fresh lot of sds. if purchased then try a fresh batch of buffer.

-mdfenko-

No wonder because it was fine before and I just recently made up my own MES buffer because we ran out of the ones we bought.
My protein is only 6kda I am worried that it will run off the gel if I complete the run. And it also runs close to the dye front.

-tofuj-

when preparing your own buffers make sure you follow the recipe as precisely as possible. for example, with laemmli it says to add the sds, glycine and tris to make the running buffer but to not adjust the pH. if you adjust then you get artifacts in the run like yours so you have to be accurate in weighing for consistent results.

as long as your band of interest is behind the dye front then you can run the dye to the bottom of the gel.

-mdfenko-

This reminds of my late supervisor, who used to admonish me.. "the minute I leave you to your own devices.."

-CPRES-

We bought in some new buffers and I am still having the same issue with my gels! I am considering making up my own sample buffer without the dye in it to avoid that bright halo showing up. I am thinking of just using the recipe provided by life tech, but excluding the dye components in the recipe:

106 mM Tris HCl

141 mM Tris Base

2% LDS

10% Glycerol

0.51 mM EDTA

0.22 mM SERVA Blue G250 < OMIT

0.175 mM Phenol Red < OMIT

pH 8.5

Any suggestions on recipe for sample buffer?

-tofuj-

i would leave the phenol red in the loading buffer. serva blue g250 is coomassie blue g250 (or its equivalent) and may react with the protein. you could add bromphenol blue instead but it may migrate close to your protein of interest. phenol red migrates faster than bromphenol blue.

when you replaced the buffers, did you also replace the loading buffer? are you sure the lots are fresher than those you replaced?

-mdfenko-

- You can also make a loading buffer with less dye (1/5th dye)

- If you make a loading buffer with no dye, you should run an empty/neg control (sample buffer+ loading buffer with dye) in a well not too close but not too far away from your sample wells to know where things are running.

-CPRES-

@ mdfenko

I didn't replace the loading buffer but I am sure that the running buffer is a fresh one.

@CPRES

Thanks for the suggestion! Sounds like a good idea.

-tofuj-

tofuj on Tue Jan 20 22:21:06 2015 said:

@ mdfenko

I didn't replace the loading buffer but I am sure that the running buffer is a fresh one.

you should try a fresh lot of loading buffer, as well. the sds (or, in this case, lds) may be decomposing and causing the artifacts that you are seeing.

-mdfenko-