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Native PAGE problem - not entering gel enough

native page resolution

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#1 Missle

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Posted 27 March 2014 - 04:19 AM

Hi.  I'm new to running native PAGE gels and am having a problem.  I'm using BioRad 4-20% Tris-HCl gels, BioRad Native PAGE Running Buffer, & BioRad Native PAGE 2x Sample Buffer and running the gel at 200V for ~30 minutes.  The proteins (have looked at 3 in various states) are not entering the gel far enough to get any information on aggregates - which was my purpose.  An IEF has been run on the proteins to determine pI and they were ~7.1, 7.5, & 7.8, (150kD, 150kD, & 30kD). I thought if a protein was too basic (>8 - 9) then it wouldn't enter so I thought my protein would be okay.

 

Does anyone have any suggestions of how to fix this?  Should I reverse the electrodes when running the gel?  How does gel % effect native gel migration?

 

Thanks!



#2 mdfenko

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Posted 27 March 2014 - 04:57 AM

forgive me if i appear to write "down"

 

native page is used for charge/size separation. gel (and crosslinker) percentage is adjusted to make the gel more or less restrictive.

 

4-20% should allow most proteins to migrate, at least part way, through the gel. only the largest proteins may not enter the gel.

 

charge on the protein is influenced by the pH of the buffers used in the gel and the electrode (most important).

 

the electrode buffer for the biorad system is pH 8.3. that is very close to your proteins of interest, their charges will small so they won't migrate well through the gel (as you have seen). just reversing electrodes will not help, in fact, it will make matters worse. the proteins will still have a net negative charge so will still migrate to the positive electrode (out of the gel).

 

three things you can do:

 

1) run for a longer time and/or higher current (careful not to let the system get too hot), until the proteins migrate far enough to suit you (not the best idea but will work, if you don't mind other proteins running out of the gel, although with 20% at the bottom many proteins will just bunch up).

 

2) switch to an acid pH gel. then you can switch electrodes and get proper migration (you will, however, have to cast your own gel). i think i posted the buffer formulations somewhere in these fora, you can find them with a search.

 

3) raise the pH of the electrode buffer. you can use the ornstein-davis buffer formulation for the electrode buffer (this should also be available within these fora, i think i posted it also). this will give you a sort of hybrid protocol. the o-d electrode buffer runs at pH 9.5. this should solve your problem and resolve your proteins.


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#3 Missle

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Posted 27 March 2014 - 06:08 AM

No forgiveness necessary - information is what I was after...

 

Option 3 seems like the best fit for my situation but I have another question.  The orstein-davis buffer formulation I found on this fora as well as elsewhere seems to be pH 8.3 like what I had used (0.025M Tris, 0.192M Glycine pH 8.3).  Are you suggesting that the pH of that buffer be increased to 9.5?



#4 mdfenko

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Posted 27 March 2014 - 06:52 AM

no. the pH of ornstein-davis buffer is higher than 8.3.

 

this archived thread gives all of the buffer formulations for o-d gels: http://www.protocol-...osts/18064.html

 

as you can see, the pH is not adjusted in any of the buffers. just prepare the 10x electrode buffer as shown and it should work fine when diluted.


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#5 Missle

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Posted 27 March 2014 - 07:28 AM

I'm sorry to be a pain but the electrode buffer outlined in that link (seen below) has a pH of ~8.3 (I just made it to confirm).  I hate to come across as dense but I appear to be missing something....

 

"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"



#6 mdfenko

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Posted 27 March 2014 - 07:35 AM

not a pain.

 

did you dilute to 1x?

 

still doesn't matter. in an electric field the components will separate and pH will be different.

 

try it.

 

do you have the equipment to pour your own gel?

 

if so then you can use the entire buffer system.


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#7 Ya yesh

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Posted 28 August 2014 - 08:00 PM

When I run my protein samples in native gel, part of the sample struck on the well and part of the sample ran just below the stacking gel. I could see my dye front has run already.

 

May I know what could be the reason ? My protein supposed to form dimer 



#8 mdfenko

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Posted 29 August 2014 - 03:35 AM

When I run my protein samples in native gel, part of the sample struck on the well and part of the sample ran just below the stacking gel. I could see my dye front has run already.

 

May I know what could be the reason ? My protein supposed to form dimer 

we need more information. what is the size of the protein? gel percentage? pI of the protein and pH of the gel system?

 

in general, proteins that stay in the well are aggregates. proteins that just enter the running gel are too big to migrate in the gel at that percentage or their pIs are too close to the running pH of the gel system to migrate far.


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#9 Ya yesh

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Posted 29 August 2014 - 08:31 PM

Hi,

 

Thanks for your suggestions.

My protein is 12 Kda and I use 15 % native gel.(http://www.assay-pro...sis/native-page)

My protein supposed to form dimer when it is added with ligand of 500Da. 

 

Attached is the picture of my native gel. The first 3 lanes are the protein, which supposed to be monomer. But the protein is not entering into resolving gel.

 

The other lanes are protein /ligand mixture, which supposing form dimer when mixed together.

I am not able to predict anything from my gel.

 

Could you give suggestions on this.

 

Appreciate your help.1.jpg

 

 



#10 mdfenko

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Posted 02 September 2014 - 04:11 AM

15% is too high. you can try a 10% gel for 12-25 kDa.

 

what is the pI of your protein? it may be too close to the running pH of the gel.

 

if it still doesn't work then you can try the buffer system of ornstein and davis (see this archived post for the formulation), it runs at a pH of 9.5.


talent does what it can
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