Protein Migration Problem [S.O.S.] - Moves in SDS-PAGE but not in Native (Dec/28/2010 )

Hi All,

I had some proteins with my PAGE gel, I run my samples in both NuPAGE (4-12%, MES buffer, Coomassie Blue staining) and discontinuous Native Page (10% Separation, 5% sample, both pH ~7.0, Bis-Tris & tricine, silver staining, Invitrogen NativeMark ladder)

All my samples (4 samples, each with 3 dilutions, almost of same sizes in NuPAGE) run fine in NuPAGE, however, my last protein was not moving in native page. Could it be due to the addition of glycerol in my separation gel?

Actually I intend to separate the protein and elute it from the gel since I do not have a liquid chromatography or column with me. I also wish to check for the protein subunits.

Thanks for your kind help. Appreciate any comments and suggestions.
Adrian

-adrian kohsf-

if your protein was not moving in native page then it was either uncharged (or nearly so) or reverse charged. the pH of your native page may have been too close to the pI of the protein.

ornstein-davis native page runs at a higher pH (9.5) and may work better for at least that protein.

you can find the formulation here.

-mdfenko-

the Ornstein and Davis formulation:

8X resolving gel buffer:
48 ml 1N hcl
36.3 gm tris base
0.23 ml temed (you can leave this out if you wish to add later)
dw to 100 ml
(pH should be 8.8-9.0)
use 37.5:1 acrylamide:bisacrylamide ratio (30% acrylamide solution- 30:0.8 acryl:bis)

8X stacking gel buffer:
48 ml 1N hcl
5.98 gm tris base
0.46 ml temed (see above)
dw to 100 ml
(pH should be 6.6-6.8)
stacking gel should be photopolymerized with riboflavin and fluorescent light
acrylamide for stacking gel should be 10% acryl, 2.5% bisacryl (will polymerize cloudy or nearly milky white), high crosslinker concentration is for mechanical stability.

10X electrode buffer (might be 1x but my protocol calls it "10x (as used)" and is probably correct):
3.0 gm tris base
14.4 gm glycine
dw to 1L

the standard acrylamide concentration is 7% but you can use what you need (or a gradient might be nice).
-mdfenko-

Hi mdfenko,
Thanks for your reply again. I owe you a beer. ^_^

is there any reason for the stacking buffer for having acidic pH rather the alkaline pH as is in the resolving gel?
Also, can I use any other chemicals as substitution for riboflavin?

Million thanks again.

-adrian kohsf-

the stacking buffer has a lower pH to retard the mobility of the sample until the electrode buffer front passes through. this sharpens the banding pattern (the process is called "stacking").

if you look at the formulation for most page buffer systems you will see that the stacking gel buffers are at a pH different from the running gel buffers. they are called discontinuous buffer systems or discontinuous page.

riboflavin (and light) is used so that no aps will be left in the stacking gel when the sample is applied. when you have a stacking gel with a different buffer you can't prerun the gel to eliminate troublesome compounds.

-mdfenko-

Hi mdfenko,
I had tried the native page with some modification where I changed the glycine to tricine as I do not have glycine in hand. I still use APS and TEMED because I dont have riboflavin yet.
I use a 4% stacking and 10% resolving. Run at 200V for 70 minutes as the bromophenol blue and ponceau S had reach to the bottom. the current flows from ~30 till ~70 and drop back to ~50 in the end.

however when i stain my gel, I find that my proteins and ladder were still in the top stacking gel, I can see there are separations of ladders but is very little. my both tracking dye had reach the bottom of the gel. how is this possible to happen, is it because of current leakage or because i mixed the 2 tracking dye together?

thanks

-adrian kohsf-

tricine is not a direct replacement for glycine. tricine is used to resolve low molecular weight proteins and peptides (with or without sds).

i'm only making an educated guess, you may be able to substitute boric acid for glycine (adjust the pH of the tris with boric acid). however, it would probably be better for you to borrow glycine from another lab.

did you use a native ladder or one prepared for sds-page? if it already has sds in it then it can't be used for native page.

-mdfenko-

mdfenko on Fri Jan 7 15:38:44 2011 said:

Thanks mdfenko,
Perhaps I have to beg other labs for some glycine this time... lol....
I will make all new fresh buffer again, and will use a new stock of native-page ladder.
Let's see how it goes on monday.

Cheers and happy new year mdfenko.

-adrian kohsf-

Hi mdfenko, I had tried the native page formula using glycine. It really works well. Million thanks again.

However, in SDS-PAGE my protein had a size around 12kDa, but in native-PAGE it shows ~260kDa.

Is this means my protein forming a multimers or does it aggregates?

How you all think?

Million thanks again.

-adrian kohsf-

it could be either or neither. in native page, charge may be a more important factor than size for mobility. if the electrode buffer pH is too close to the pI of the protein then it will move slower than if the pH is farther away.

if you want to determine molecular weight in a non-denaturing system then i would recommend gel filtration (which you already said is not available to you).

if you are feeling a little adventurous then you could try running your protein in blue native page (it uses coomassie blue, in the electrode buffer, to equalize charge density instead of sds). i don't know if it will maintain activity if your protein is an enzyme but should be otherwise intact (disclaimer, i have not run blue native page, i've only read about it).

you could also try native page at other pHs (i have formulations for neutral pH and acid page, which should be in the archives). which you may use depends upon the pI of your protein.

ps- happy birthday

-mdfenko-