Isoelectric focusing of peptides.

1D, 2D, HPLC, SDS PAGE, etc. Talk about it here.
Artur
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Isoelectric focusing of peptides.

Postby Artur » Sun Oct 23, 2011 11:19 am

Wchich buffer sytem do you reccommend?
Will 1M urea with ampholytes be ok? (cysteines was blocked prior to digestion)
What resolution do you observe?

Toxic
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Postby Toxic » Tue Oct 25, 2011 12:14 am

I've tried focusing in plain water and it works OK. 1M urea works much better. Peptides are pretty soluble, even at their isoelectric point, so more urea is probably unnecessary, but wouldn't hurt. I always pass the peptides I extract from the IPG strip through SPE before going near the LC/MS/MS. I perform SPE on the peptides before IEF to remove salts etc and don't use any acid in the solvents for SPE, just water and ACN.

I don't use ampholytes much in IEF except for narrow range strips when trying to focus really basic proteins (between pH 9-11). I haven't tried them in peptide IEF using IPG's.

As for resolution, most peptides are in the acidic end of a 3-10 IPG, but that maybe the bacteria I work on, whose proteins in the majority, focus in the 4-7 range on 2D-PAGE. I cut the strip into 10 x ~1cm pieces using an 11cm IPG and run the peptides from each on a 90min gradient from 5-30% ACN in LC/MS/MS.

Artur
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Postby Artur » Tue Oct 25, 2011 3:05 am

Thanks for help,
What protocol do you use for peptide extraction and subsequent SPE prom IPG strip pieces?

Toxic
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Postby Toxic » Tue Oct 25, 2011 4:32 pm

After slicing the IPG up with a scalpel and scraping the gel off into a tube, I add 50% ACN, 0.1% TFA and sonicate them in a waterbath for 10 mins. Leave them on the bench a while, remove the solution and repeat. I then vac the ACN off and use 100uL OMIX C18 tips to clean the sample, pretty much to the manufacturer's instructions except I load the sample through the top of the tip to make sure it all passes through the resin. I elute in 75% ACN, vac that and load it all into an autosampler vial, then into a michrom peptide CapTrap. Then into the C18 into the QSTAR.

Artur
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Postby Artur » Thu Oct 27, 2011 12:31 am

How do you perform digestion to avoid excessive salt content in the sample before focusing? Or is it desalted?

Toxic
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Postby Toxic » Sun Oct 30, 2011 11:29 am

I just do normal in-solution digests. Solubilise in 8M Urea in 100mM ammonium bicarb (or 7M Urea, 2M Thiourea, 1% C7BzO, 40mM Tris), reduce and alkylate with TBP and acrylamide monomers, quench with DTT and dilute to 1M urea with 100mM ammonium bicarb before adding trypsin at a 1:100 ratio. Incubate overnight at 37.

The digest then goes through a 1cc OASIS HLB syringe equilibrated in water, no acid or modifiers. Wash the salt and other stuff away with more water and then elute in 75% ACN in water with no acid. I have accidently used 0.1% TFA in these before and done IEF and it still works. The focusing just hits the 50uA current limit I set and stays there for 8 or so hours until the acid all runs out into the strip, then the current drops at the voltage gets to 10kV. I focus overnight for 100kVhours.

The TFA may have effected the pI of the peptides, but I haven't checked to see if the peptides were found in the strip at a pH corresponding to their calculated pI.

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Postby machina267 » Sun May 20, 2012 3:28 am

Dear Toxic. Your help was very useful for peptide IEF. Currently I am also working on this procedure and have a question regarding the IEF procedure. How do you remove the mineral oil from the strip after IEF? Blotting with kimwipe help a little bit, but some mineral oil does remain on the strip and get carried to the down stream procedure and that is quite annoying me.

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Postby Toxic » Sun May 20, 2012 11:52 pm

I simply hold it with a pair of forceps with the end of the plastic backing strip on a KimWipe and let it drain. I move it around on the tissue to get the oil to collect at dry parts of the tissue. I try and do this for at least a minute or until the tissue is not taking up oil when moving to a dry spot. Then I just what I described above and have had no issue with oil. I forgot to mention that I focus under parrafin oil (Shell Ondina) rather than mineral oil and that might be the difference.

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Postby machina267 » Sat May 26, 2012 3:50 am

Thanks for your reply. Now the mineral oil problem seems to be gone, but I now realize that there are actually multiple problems with my experiment.
Long story short. I have tried the protocol with BSA. No problem getting separation on pH 4-7 strips. MALDI-TOF spectrum seems clean. Then I try to apply the protocol to serum sample, checking the quality of peptide with MALDI-TOF along the way.
Then I get into trouble. To solubilize the serum protein after acetone precipitation, I used 8M urea and adjust concentration down to 1M during trypsin digestion, and then Waters HLB cartridge clean up, speed vac, then adjust volume with water and apply to IPG strip. But the eluate after Waters HLB clean up showed huge white crystal precipitate, presumably residual urea. So it appears that Waters HLB cartridge cannot get rid of the urea in the system before IEF focusing. This is my first problem.
Some protocol also use urea during strip rehydration and IEF so I guess residual urea won't hurt, but that would be problematic for MALDI-TOF MS analysis. Because in our lab we dun have nanoLC and Q-TOF yet, so at this moment we have to stick with MALDI first. One dimension IEF is not perfect, but at least it should give me some protein ID (I assume). But after IEF separation I can only see very weak signal and cannot ID anything with MS/MS analysis. So is it because of the residual urea in the IPG fractions that interfere with MALDI, or because of the insufficient fractionation by IPG-IEF alone?
Sorry for the long post. Thanks in advance for your attention and advice!

Toxic
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Postby Toxic » Mon May 28, 2012 11:45 am

You are essentially doing exactly what I do. However, if there is still urea in the sample after you elute the HLB cartridge, you need to wash it more. The urea shouldn't be binding to the resin. I use the 1cc syringe columns and fill them with 1mL of solutions and let them drip through under gravity (except the sample volume is whatever it is and the elution volume is 400uL). 1mL of water should be enough to get rid all of the urea, but it can't hurt putting another 1mL through. You are correct that the urea won't affect the focusing. All I can suggest is that you try further desalting the samples after IEF and peptide extraction with a ZipTip or similar of good capacity. Vac off any ACN from the extraction first.

I can't say beyond that. If your loading 100ug of protein/peptide on the IPG and we assume equal distribution across the strip during focusing (highly unlikely, but anyway), then there should be 10ug in each cm of an 11cm strip (I use a 3-10). That should be enough to detect, unless you have too many components or too high a concentration and its causing ion suppression in the MALDI. Your post reads as if your going straight from IEF to MALDI without further fractionation of the peptides by reversed phase and the sample is likely still too complex and you're getting ion suppression. If you haven't got a nanoLC you could try sequentially eluting a ZipTip with different concentrations of ACN to form a step gradient. Maybe do 5, 10, 15, 20, 25 and 30% ACN, where peptides normally elute in a C18 gradient, and then 80% to get off any remaining. Time consuming, but may solve the problem.

machina267
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Postby machina267 » Mon May 28, 2012 6:38 pm

I think there is something seriously wrong with my HLB procedure, because I have already washed my cartridge with 4 ml MilliQ water. Occasionally I forced the liquid through the cartridge with pipette, so maybe it is the problem with it.
Well yes, you are correct, I went straight from IEF to MALDI. Definitely not a good choice, but at this moment we have to make do with it. I will try the tedious ziptip procedure you described. Thanks for your advice!

Toxic
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Postby Toxic » Mon May 28, 2012 11:11 pm

Forced the liquid through? Did you centrifuge the sample at 16,000g or so before loading it in the HLB? You may have particulates and are blocking it. I can drip through 1mL of water in a couple of minutes (I've never timed it as you can run the HLB dry without issue). My HLB steps are
1) 1mL of 100% ACN.
2) 1mL of 2% ACN or water.
3) Load centrifuged sample.
4) Wash with 1mL of 2% ACN or water.
5) Elute with 400uL of 75% ACN.
6) Speedvac to remove ACN.

machina267
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Postby machina267 » Wed Jun 13, 2012 6:06 am

Thanks for your advice. Now I am able to grab a nano-LC for the project and do the protein ID properly. However, the number of proteins identified is very low, only about dozen. Also the chromatogram only gives a few peaks with low intensity. So I suspected that there are two reasons: loading amount of LC analysis and MS/MS parameter.

Will you load all the IEF eluate through the OMIX tip for LC analysis? What would be your usual loading amount to get decent protein ID? Is there any recommended "minimum" in order to get protein ID working?

Toxic
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Postby Toxic » Thu Jun 14, 2012 12:24 pm

I calculate the loading amount by working backwards from the MS. I use Michrom Captraps which have a capacity of 3-5ug according to their literature. So if I cut the strip into 10 pieces and assuming the peptide is evenly distributed across the strip (which it won't be), I can load 50ug. I typically load 100ug. All of the IEF eluate goes through the OMIX tip and then I'll load 1/10th or so of that into my LC/MS/MS to see how concentrated and complex the fraction is. I'll adjust the loading volume from there.

Getting a decent protein ID is related to the dynamic range of the sample. If the dynamic range is high (like serum/plasma) and you're interested in low abundance proteins, you will have to load more so there is more peptide to detect and get decent sequence. But then you have to fractionate more or cut the strip into smaller pieces to try and keep the high abundance peptides away from everything else. Unfortunatley, each sample has to be taken separately and there is no universal solution (yet. Start reading Mann's latest papers).

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Postby iheartlungs » Tue Jun 19, 2012 1:46 am

I have used an offgel fractionator (Agilent) and it works very well - but obvs this is a more expensive system. We have had success with GE Healthcare strips and ampholytes instead of the agilent ones which cost a crazy amount!

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Postby Toxic » Tue Jun 19, 2012 11:40 pm

I had an OFFGEL in the lab for a while as the distributor was using me as a demo site. If you need a new IEF unit, it's a good buy. Two 10kV power supplies that can be independently controlled and have sophisticated programs for voltage and current, its a great focusing unit. The it has the bonus of the OFFGEL. But I'm reluctant to buy it for the fractionating alone. The main problem I had with it is how much sample was getting left in the strip after focusing. Once the protein or peptide reaches its pI, the technique relies on simple diffusion to get the protein into the liquid above the strip. There is no active force pulling the protein/peptide out. So I tried it a few times to separate protein mixtures and after extracting the liquid, I'd equilibrate the strip in SDS and run it on an SDS-PAGE gel to see what was in it. The answer is lots and there was probably more left in the strip than was in the liquid. The simple way around this is to cut up the strip and elute the protein out by further diffusion, but the instructions don't mention this. I prefer the MicroRotoFor and Zoom IEF runner for liquid IEF of proteins. I haven't tried it for peptides. I really should.

And I believe the strips Agilent supply are rebadged GE strips. I think a rep told me that.

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Postby Toxic » Thu Jun 21, 2012 3:47 pm

I found this while looking for something else: http://www.bio-rad.com/webroot/web/pdf/lsr/literature/Bulletin_6140A.pdf

Taking the grain of salt of it being company propaganda in mind, this is a good example of how to perform peptide IEF. If they left out the TFA from their desalting, its likely both techniques would have got to higher focusing voltages.


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