Author Topic: Background problems - IPG buffer? How do I avoid it in DIGE without fixing?  (Read 11508 times)

Bevan

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Hi,
I've been having a problem with a dark smearing effect in all of my recent gels.  I haven't encountered this before and was wondering if anyone could help.  I've had a look around the forum and seen a similar thing with excess ampholytes, and I have recently switched from using 0.5% to 2% to counteract the effect of Tris in the buffer, which is also a new thing for me - but essential for DIGE, which is what I intend to use these samples for.  However, the IPG buffer I'm using is within the recommended amount.

Lysis buffer: 7M urea 2M thiourea, 4% CHAPS, 30mM Tris pH 8.5.  Sonication for 2x 10sec bursts on ice.
Clean up using Amersham 2D kit and resuspension in 7M urea, 2M thiourea, 4% CHAPS, 30mM Tris, pH 8.5.
Quant using Amersham kit.
Samples are then diluted to give 110ug per 11cm strip in 7M urea, 2M thiourea , 4% CHAPS, and 50mM DTT, 2% IPG buffer and 0.01%Bromophenol blue solution are added.
Strips are loaded with in-gel active rehydration for 12hr before IEF with 4 steps:
1. gradual ramp up to 250V for 1hr
2 gradual ramp up to 6000V for 2Hr
3. 6000V for 25000vHr
4 1000V until removed from IEF
Total of 32000-35000 Vhr

Strips are equilibrated 2.5ml/strip, for 15 min in equilibration buffer (50mM Tris-HCl pH 8.8, 6M Urea, 30% glycerol, 2% SDS, trace bromophenol blue) with 10mg/ml DTT then 15 minutes with 25mg/ml Iodoacetamide.

2nd Dimension is in pre-cast 11cm Criterion XT gels with MOPS XT buffer using constant power of either 3.75W or 7.5W per gel for approx 1hr45 or 2hr30.  Previously I had been using constant volts, but changed after reading comments on this forum.

Gels were washed 3 times in dH2O then stained with Coomasie Biosafe (BioRad) for 1hr, as per manufacturers instructions.  Destaining was with dH2O over night.


Modified by Bevan at 7:52 AM 8/26/2007

Modified by Bevan at 8:34 AM 8/28/2007


Bevan

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Re: background problems (Bevan)
« Reply #1 on: August 25, 2007, 05:00:00 PM »
Some more gels. This time pH 3-11NL

millennium

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Re: background problems (Bevan)
« Reply #2 on: August 25, 2007, 05:00:00 PM »
Some comments:

Perhaps it is the excess ampholytes, so that is something to consider... it doesn't look necessarily like dirty glass to me. If so... why do you use tris in the lysis buffer and the resuspension buffer following clean-up? Are your proteins of interest extremely sensitive to pH even in a denaturing solution? If you're compensating the tris with increased ampholytes, then removing Tris may be a way to return to your previous state of clean looking gels.

Random comment: i'm not familiar with pre-cast gels, but do the criterion gels use bis-tris in the resolving gel (given you're using a mops running buffer)? We cast our own bis-tris sds gels and we've had good results substituting the Tris in the DTT / IAA incubations with Bis-tris, pH 6.4. Subjectively (!) the results appear a little better to me, but at the least, you would have some theoretical consistency in your protocol :-)


Bevan

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Re: background problems (millennium)
« Reply #3 on: August 26, 2007, 05:00:00 PM »
Thanks for the comments - I'm running strips with 0.5, 1 and 2% IPG buffer to see if that is the root of my problem.  I'll have a look at Bis-Tris in the equilibration buffer too.  
The reason I've been using Tris in the sample buffer is to replicate the conditions I'll have to use for DIGE.  If I can't get good resolution with 2D using that protocol then DIGE won't work either.  After samples are dissolved and quanted they are diluted with sample buffer that doesn't contain Tris - bringing the concentration down to <10mM.

Does anyone know if glycerol could produce this effect?  Although I used 30% - my equilibration buffer does seem a bit viscous.


Hamish

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(No subject)
« Reply #4 on: August 27, 2007, 05:00:00 PM »
Hi Bevan,
Recently I had this exact same problem (I'm 99.9% sure it's the same dark smear I saw). The problem was fixed when I fixed the gel in 45% methanol, 1% acetic acid overnight. Then I did the water washes and the coomassie stain, and it was fine.
I'm pretty sure any variation will work too, like ethanol instead of methanol, and TCA or phosphoric acid instead of acetic acid.
I'll put a response in my thread so people know it worked..
All the best,
Hamish

Bevan

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Re: (Hamish)
« Reply #5 on: August 27, 2007, 05:00:00 PM »
Hi Hamish
Were you using colloidal coomasie?

I'm using Biosafe, from Bio-Rad and the instructions specifically mention that no fixing is necessary. I just wonder if using ethanol/methanol-acetic acid would disturb the Biosafe chemistry.  I'll contact Bio-Rad and post their reply.


Hamish

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(No subject)
« Reply #6 on: August 27, 2007, 05:00:00 PM »
Hi Bevan,
Yeah I was using Invitrogen's Simplyblue colloidal coomassie stain, which is probably a very similar product. The protocol says to do 3 x 5 min water washes then add stain, similar to your protocol. Simplyblue's instructions say the same thing, that fixing is not necessary.
Hopefully you'llbe pleasantly surprised:)

Bevan

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Re: (Hamish)
« Reply #7 on: August 27, 2007, 05:00:00 PM »
Thanks, I'll give it a go.

Bevan

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Re: (Bevan)
« Reply #8 on: August 27, 2007, 05:00:00 PM »
Here's my comparison between three different IPG 3-11NL buffer concentrations.  The top is with 2%, middle with 1% and bottom with 0.5%.  As you can see - IPG concentration does make a difference, although even with 0.5% IPG there's still background smearing.  Next I'll try fixing vs no fixing.

Hamish

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Re: (Bevan)
« Reply #9 on: August 27, 2007, 05:00:00 PM »
That's interesting. Did you notice whether more IPG buffer improves the gels at all? And I'm interested to know if fixing solves it, so post your results if you can.
Cheers

Bevan

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Re: (Hamish)
« Reply #10 on: August 27, 2007, 05:00:00 PM »
I only ran 1 gel from each concentration because 2D is so expensive and my budget is tight.  I haven't bothered comparing them using Image Master because, as you know, 2D gels are notoriously bad for gel-gel variation and comparing gels without replicates wouldn't produce anything meaningful.   Saying that, and being purely subjective, in my opinion the 0.5% buffer produced sharper and more spots in the lower pH range and the 2% buffer produced sharper and more spots in the higher range, but there isn't much in it, so it's six and two threes when deciding which to use.  If I remember rightly Amersham suggest 2% IPG for one piece of IEF equipment and 0.5% for another.  I'm unsure why there's a difference.  I'm using a Bio-Rad IEF cell, which didn't come with any recommendations, so it's all down to trial and error.

I will post the results of my fixing test though.  


Bevan

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Re: (Bevan)
« Reply #11 on: August 27, 2007, 05:00:00 PM »
Re-reading through the DIGE manual it says that gels should not be fixed before scanning, as acid reduces the signal strength.  

How then am I going to remove the IPG background contamination when I come to do DIGE!!!  


Sjouke Hoving

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Re: (Bevan)
« Reply #12 on: August 29, 2007, 05:00:00 PM »
"as you know, 2D gels are notoriously bad for gel-gel variation"

if you know how to make a good 2D gel, the variation for gel to gel is very acceptible.

the ampholyte concentration might disturb a silver or coomassie stain, but for direct fluorescence scanning when using DIGE they do not contribute to background smears.

sjouke


Bevan

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Re: (sjouke)
« Reply #13 on: August 29, 2007, 05:00:00 PM »
Does IPG have any effect on mass spec?

Sjouke Hoving

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Re: (Bevan)
« Reply #14 on: August 29, 2007, 05:00:00 PM »
you mean IPG buffers or ampholytes? I have no idea, but during the digestion protocol the gel piece is washed so extensively that no IPG buffers are present anymore.

sjouke