STrap method

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zougman
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STrap method

Postby zougman » Mon Mar 10, 2014 11:45 pm

To my proteomics colleagues:

I have just come up with a new bottom-up proteomics sample prep method. The method (called STrap - Suspension Trapping) is based on a novel simple concept utilizing an old-fashioned SDS extraction of the protein material (thus resulting in the unbiased proteome coverage), provides very fast reactor-type in situ sample processing and peptide clean-up, works with low µg sample loads and all types of proteins. I am 100% confident it is going to save a significant amount of the sample processing efforts in your research. Please try it - I am happy to answer any questions. Alexandre :)

http://onlinelibrary.wiley.com/doi/10.1002/pmic.201300553/abstract

Suspension trapping (STrap) sample preparation method for bottom-up proteomics analysis. Alexandre Zougman, Peter J. Selby, and Rosamonde E. Banks. DOI: 10.1002/pmic.201300553

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Postby Omics » Thu Mar 13, 2014 9:58 am

Theoretically, both the proteins and trypsin are to be immobilized on the column. Will the digestion efficiency be affected?

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Postby zougman » Thu Mar 13, 2014 10:32 am

Dear Omics,

The trapping depth filter part of the STrap tip is made of pure quartz (SiO2) - thus, it is unlikely that any significant protein immobilization to the depth filter matrix is taking place under the digest conditions (50 mM AmBic).

Sincerely,

Alexandre

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digestion time

Postby Joe » Mon Mar 17, 2014 5:42 am

Dear Alexandre,

I am really interested in the STrap method and will definitely try it. I just wonder about your short digestion times. Is the protocol optimized for maximum peptide yield or was your aim to prepare your samples really fast. What happens if I would increase the digestion time e.g. trypsin @37°C overnight? Can I expect an increase of the peptide yield?

thanks

Joe

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Postby zougman » Mon Mar 17, 2014 8:57 am

Dear Joe,

The method is already optimized - according to my experience there is no difference in the STrap-tip method performance digesting for 1 hour at 47C versus 4 hours at 37C using Promega trypsin. When employing other (non-thermostable) enzymes, such as WAKO Lys-C, you do have to abide by the conventional temperatures and incubation times.

I am happy to provide some STrap tips to try, if needed.

Sincerely,

Alexandre

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Postby leopardyu » Mon Apr 28, 2014 11:02 am

Dear Alexandre,

I want to thank you again for your previous answers to my questions. I'm gonna post my following questions here, so other people who is interested in this method may also benefit from your answers.

I have never worked with quartz fiber before, so I'm still not so clear with the mechanism underlying STrap method. I have more than one questions here.

1. How the "particulate matter” is generated?
It seems when you add the SDS-containing lysate into methanol (90% in 100 mM Tris/HCl, pH 7.1), then you’ll get the protein particulate suspension, is that right? But why is that? Is it because the proteins are not soluble in methanol?

2. What if you add the SDS-containing lysate into the tip and wash with 8M urea directly? Are the SDS detergents going to be depleted in this case?

3. What’s the purpose of acidifying protein samples with phosphoric acid? Otherwise you will not get particulate matter?

4. If methanol can solubilize SDS (up to 2%), can we replace 8M urea in the traditional FASP method for SDS depletion? If so, we don’t have to deal with high concentration urea buffer then.

Thanks in advance for your kind reply.

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Postby leopardyu » Mon Apr 28, 2014 11:03 am

Dear Alexandre,

I want to thank you again for your previous answers to my questions. I'm gonna post my following questions here, so other people who is interested in this method may also benefit from your answers.

I have never worked with quartz fiber before, so I'm still not so clear with the mechanism underlying STrap method. I have more than one questions here.

1. How the "particulate matterâ€￾ is generated?
It seems when you add the SDS-containing lysate into methanol (90% in 100 mM Tris/HCl, pH 7.1), then you’ll get the protein particulate suspension, is that right? But why is that? Is it because the proteins are not soluble in methanol?

2. What if you add the SDS-containing lysate into the tip and wash with 8M urea directly? Are the SDS detergents going to be depleted in this case?

3. What’s the purpose of acidifying protein samples with phosphoric acid? Otherwise you will not get particulate matter?

4. If methanol can solubilize SDS (up to 2%), can we replace 8M urea in the traditional FASP method for SDS depletion? If so, we don’t have to deal with high concentration urea buffer then.

Thanks in advance for your kind reply.

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Postby zougman » Tue Apr 29, 2014 5:23 am

Dear leopardyu,

Thanks for the questions. The basic principle underlying the STrap methodology is creation of the fine particles from the SDS-solubilized proteins and trapping the particles in the depth filter media. The quartz depth filters were chosen because of their pure composition (SiO2), low metal content, low peptide background binding under the digest conditions, as well as commercial availability (you may also try to use glass depth filters, for example). While the particles are indeed retained throughout the depth filter dimensions, SDS and other contaminants are removed in the flow through, and then a protease is introduced which processes the fine protein particulate into the peptides. This is, in my opinion, as close as one can get to the high-throughput robust unbiased comprehensive etc. proteome profiling.

1. How the "particulate matter” is generated?
It seems when you add the SDS-containing lysate into methanol (90% in 100 mM Tris/HCl, pH 7.1), then you’ll get the protein particulate suspension, is that right? But why is that? Is it because the proteins are not soluble in methanol?

When you add the SDS-protein solution into the methanolic buffer the SDS-micelles are broken, the proteins are exposed to the alcohol medium which has a low dielectric constant and thus the proteins start precipitating out.

2. What if you add the SDS-containing lysate into the tip and wash with 8M urea directly? Are the SDS detergents going to be depleted in this case?

If you do such a thing you will loose your proteins in the flow through because urea retains the proteins in solution and thus the proteins are going to simply pass through the filter pores.

3. What’s the purpose of acidifying protein samples with phosphoric acid? Otherwise you will not get particulate matter?

The acidification helps to obtain the fine protein particles which fit the quartz depth filter particle retention characteristics. This was found by trial and error.

4. If methanol can solubilize SDS (up to 2%), can we replace 8M urea in the traditional FASP method for SDS depletion? If so, we don’t have to deal with high concentration urea buffer then.

Providing your centrifugal filter unit withstands the action of methanol, the particulate matter created in such an exercise is going to block the filter membrane and, basically, that is it :(

Just to add, when working with the STrap tip please do not overload it - it will result in the increased backpressure and reduced performance.
Best,

Alexandre :) .

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Postby leopardyu » Tue Apr 29, 2014 6:54 am

Dear Alexandre,

Thanks for your answers! They all make sense to me. We'll definitely try it out in our lab.

Thanks,

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Postby zougman » Wed May 21, 2014 9:38 am

Hi all,

Lately, I have been getting enquiries regarding the basic STrap tip stack composition used in my lab. What we use currently for routine applications (done with gauge 14 needle, from top to bottom):

6 QMA plugs (Whatman) + 7 MK360 plugs (Munktell) + 3 C18 plugs (Empore)

Alexandre

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source of MK360 Munktell filters?

Postby Lengqvist » Thu May 29, 2014 6:22 am

Dear Alexandre

I'm having some trouble locating a distributor for the Munktell MK360 filters...

Perhaps you could share where you get them?

Thanks!

Best regards

Johan

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Postby zougman » Thu May 29, 2014 7:35 am

Dear Johan,

In the UK I buy it from http://www.gilsonuk.com/search/Catalogue/mk360
I guess you can always contact Munktell http://munktell.com/en/Munktell/Contact-us/ to find out the local distributors for the product

Sincerely,

Alexandre

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Postby chengh1 » Tue Jul 01, 2014 8:18 pm

Hi Alexandre:

Thanks for developing a new methodology and post it for everyone. We might want to have a try STrap, but I come to one question, if I am going to start with like 1mg or maybe 10mg of protein, do you think it is feasible?

Cheers
Cheng

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Postby zougman » Fri Jul 04, 2014 1:38 am

Dear chengh1,

The STrap technology was designed, first of all, to overcome the challenges one faces when processing small protein amounts. Thus the protein load capacity of the STrap tip is up to ca. 50-60 ug . In the supplementary material to the STrap paper you can find the descriptionof the STrap tube unit which allows for processing of larger protein amounts (ca. 250 ug). I have not had the chance yet to try processing mg protein loads. However, I believe that by adhering to the STrap method principles and designing bigger processing units you can do so. Assuming that such large peptide quantities are needed for some PTM enrichment studies (e.g. phosphopeptides), you can even facilitate the digestion process by applying larger enzyme/protein ratios as, in this case, the enzyme autodigestion products are not going to interfere with the consequent peptide identifications. Of course, ideally, it could be much easier if STrap units of different load capacities were made commercially available…

Sincerely,

Alexandre

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Postby chengh1 » Sun Jul 27, 2014 7:22 pm

Hi Alexandre:

Thanks a lot for the information.

Cheers
Cheng

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STrap and cysteine peptide capture (C-STrap)

Postby zougman » Sat Sep 26, 2015 12:14 am

As the STrap depth filter surface is made of silica we can modify it using silane-based chemistries. Therefore, during the STrap-based digest, peptides with specific features could be targeted in-situ for enrichment. In the example presented, the STrap tip is functionalized with SPDP, enabling reversible capture of the cysteinyl peptides while unbound peptides are collected as a separate fraction.

http://journals.plos.org/plosone/article?id=10.1371/journal.pone.0138775

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Postby Infinity » Tue Oct 06, 2015 11:30 am

Hi, I'm mostly interested in processing large amounts of proteins (starting from 1 mg) for subsequent phosphopeptide enrichment. So I was wondering if I can use similar approach w/o column format. Let's say I lyse my cells in SDS buffer, treat with DTT/IAA, acidify with phosphoric acid and mix with excess of MeOH. Subsequently I will spin down protein pellet and wash it several times with MeOH and finally resolubilize it in ammonium bicarbonate and digest with trypsin.
Do you think that might work?

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Postby zougman » Wed Oct 07, 2015 7:03 am

Hi Infinity,

Thanks for the question. You are basically suggesting to use a kind of a precipitation approach for protein digestion. This approach is indeed milder than the one with TCA or acetone, and may result in partial digestion of proteins. However, the results arenot going to be perfect. The main idea of STrap is to minimize particulate aggregation aiming at separation of the trapped protein particles in the trap's space, making it easier for an enzyme to digest them - the aggregation happens as soon as the particulate matter hits the bottom of the tube. I am aware of the need to process larger protein quantities and hope to make simple larger STrap processing units available in the foreseeable future. In fact, for such studies you do not need the underlying C18 membrane as in the case of the basic STrap - all you need is a sole depth filter trap unit of the right capacity which makes sample processing even faster and easier without centrifugation.

AZ

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Re: STrap method

Postby zougman » Sat Oct 08, 2016 3:50 am

It has taken some time but now various S-Trap units for processing protein loads in the sub-microgram to 10 mg range are commercially available.
www.protifi.com


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