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Protein stuck in the origin of resolving gel


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#1 tulip

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Posted 26 May 2015 - 12:54 PM

My protein size is 260kDa and I am using 5% stacking gel and 7.5% resolving gel. After wet tank transfer, I am seeing that my bands positive for my protein are way up at the origin of the resolving gel. Please provide suggestions to improve the run. thanks.



#2 bob1

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Posted 26 May 2015 - 01:38 PM

Lower your gel percentage. I would use a 6% gel for this.

 

Also check your running buffers, throw out old solutions and prepare fresh.



#3 labtastic

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Posted 27 May 2015 - 09:41 AM

Yeah use a 4% stacking gel.

 

It is possible your protein is aggregating in SDS. This is not unusual for membrane proteins if that is what you are working with. Are you boiling your samples prior to running them on a gel? If so, try not boiling. If you are not, try boiling. Also try with and without reducing agent.

 

Also consider sonicating your sample briefly prior to solubilization in SDS. This could be particularly helpful if you are running whole cells. Sometimes intact DNA can cause samples to be a little viscous causing them to run awkwardly on a gel.



#4 mdfenko

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Posted 28 May 2015 - 04:31 AM

also, instead of boiling you can heat at 60-70C for 10-20 minutes, sometimes proteins will aggregate when boiled in the presence of sds for too long.


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#5 tulip

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Posted 03 June 2015 - 03:10 PM

Thanks for all your suggestions. I repeated with 4% stacking and 6% resolving gel, tried boiling and also at 37C, still I see my protein at the origin of resolving gel, worse yet, the 250kDa protein std marker is also at the top. Please help. 



#6 labtastic

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Posted 04 June 2015 - 12:47 PM

If your standard 250kDa marker is stuck in the stacking gel, something is wrong with your gel, your running buffers, or your gel box.

 

What voltage are your running? What amperage?

 

What is the pH of your stacking and resolving gels?

 

Try borrowing someone else's gel box to see if that is the problem.

 

Or procure a commerically casted gel to see if your home-made gels are being made incorrectly.

 

Or double check that your running buffer has the right concentration of glycine, tris and SDS. Checking the pH is a quick way to do that. If it's not 8.3, then re-make it.


Edited by labtastic, 04 June 2015 - 12:48 PM.


#7 mdfenko

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Posted 05 June 2015 - 04:09 AM

how are you preparing the running buffer?

 

do you adjust the pH? if so then don't, this will introduce salts into the buffer and they will interfere with migration. laemmli running buffer should not be adjusted. just use the components as described in the protocol.


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#8 tulip

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Posted 05 June 2015 - 08:55 AM

No I usually don't adjust the pH of the running buffer.  I just re-checked the pH of the 1X running buffer it is 8.53. 


Edited by tulip, 05 June 2015 - 12:17 PM.


#9 mdfenko

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Posted 05 June 2015 - 10:08 AM

try preparing fresh running buffer. make sure you use the same form of tris, glycine and sds (in fact, you may want to use a newer lot of sds, the old one may have decomposed).

 

if you want to check the pH then do it before you add the sds.


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#10 tulip

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Posted 15 June 2015 - 01:30 PM

I prepared fresh running buffers, Tris buffers for stacking and resolving gel and still I see a faint 250kDa marker in the interface of stacking and resolving gel. I am waiting for commercial gels now. Please see this pic.

 

 

 

Thanks

 

 

Attached Thumbnails

  • Precision plus dual marker on 7.5% gel.JPG

Edited by tulip, 15 June 2015 - 01:32 PM.


#11 mdfenko

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Posted 16 June 2015 - 04:05 AM

what is the ratio of acrylamide to bisacrylamide? standard for protein gels is 37.5:1 (sometimes 29:1).

 

if you are using a greater crosslinker ratio (eg 19:1) then the gel will be more restrictive and will require even lower concentrations to allow high molecular weight proteins to migrate farther.


Edited by mdfenko, 16 June 2015 - 04:06 AM.

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#12 tulip

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Posted 16 June 2015 - 04:14 PM

It is 37.5:1 (2.6%C). I have no problem extracting the problem, or temperature, reducing agent, transfer,  it is just that my 260kDa protein is stuck on top as well as my 250kDa protein marker (at least a faint band). I really don't know what else to try. Any suggestions would be helpful at this point.



#13 mdfenko

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Posted 17 June 2015 - 04:20 AM

we look at a wide range of protein sizes ( we used to work with myosins), so we routinely use a 5-15% acrylamide gradient with a 4.5% stack.

 

if your only protein of interest is > 200 kDa then you may want to try a 5% gel with a 4 or 4.5% stack. be careful, the gel will be fragile and difficult to handle.


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