Rapid universal MS sample prep.


The patent-pending S-Trap™ brings universal, simple and inexpensive sample preparation to bottom-up proteomics. One easy-to-use spin column combines sample concentration, clean up and digestion. S-Trap™ sample processing does not require any specialized equipment nor training. From µg to mg scales, your samples are ready in about an hour.

S-Traps™ eliminate one of the greatest sources of variability – inconsistent lysis and protein solubilization – by using strong 5% SDS for all samples. This almost universal protein solvent is removed in the protein trap of the S-Trap™, which retains all undigested proteins but does not retain small molecules including the SDS used to extract and handle proteins.

With S-Traps™, you will achieve better reproducibility within and between labs: proteins are consistently extracted and solubilized, and time and complexity are significantly reduced. Use S-Trap™ micros for sub-microgram to 50 µg scales; S-Trap™ minis for 50 – 300 µg and S-Trap™ midis for larger scales (300 µg to multiple mg).


S-Trap™ sample processing begins with strong, 5% SDS lysis and solubilization. This solution dissolves even poorly-soluble proteins like membrane proteins which are often left behind in the pellet. Reduction and alkylation are also performed in 5% SDS precluding any precipitation at this step. Proteins are further denatured by acidification to pH 1 to ensure complete destruction of all enzymatic activity (e.g. of proteases and phosphatases) and to maximize sensitivity to proteolysis.

Samples are then diluted with S-Trap™ binding buffer and loaded on the S-Trap™ column where proteins are captured within the submicron pores of the three-dimensional trap. Proteins captured within the trap present exceptionally high surface area, allowing them to be washed free of even pernicious contaminants (PEG, glycerol, detergents, salts, etc.). Protein capture and clean up takes only minutes.

Upon addition of your choice of protease, physical confinement of the substrate and protease within the pores of the trap forces fast digestion: the protease is either digesting your substrate or reflected off the sidewalls straight back to the protein to digest.

  • S-Trap™ technology enables fast (1 hr), reproducible and universal sample processing for bottom-up proteomics. It is simple, can be performed in any lab with standard equipment and works with your choice of protease.
  • S-Trap™ processing combines strong SDS-based protein solubilization with sample cleanup and rapid reactor-type protein digestion. Contaminants such as PEG, detergents, salts and glycerol are fully removed.
  • S-Trap™ units are available to the multiple milligram scale. Large scale is particularly useful for those doing pre-enrichment for example in phosphopeptide or SISCAPA/immunoaffinity analysis.
  • Sample processing is unaffected by protein solubility: cytosolic and membrane proteins are identically solubilized and processed.
  • S-Trap™ processing is suitable for all sample types including cells, tissues, membranes and serum. With serum, 8 M urea and sample dilution is not needed.
  • S-Trap™ sample processing is fast: removal of SDS and contaminants takes only few minutes, and complete digestion is achieved in as little as 30 min.
  • S-Trap™ processing integrates sample digestion and peptide clean-up in the same device. It is also capable of processing sub microgram levels of protein with low losses.
  • S-Traps™ also filter samples (no clogging from on-bead IP digestions, etc.)

What buffer components will the S-Trap remove?

Almost everything! Urea (even 8M), salts, glycerol, PEG, other detergents, Ficoll, tween, triton, Lamelli loading buffer… Just make sure to add SDS to 2% – 5% final concentration and follow the S-Trap protocol. The one thing to avoid is 6 M guanidinium chloride: it is positively charged, SDS is negative and when you put the two together they form an insoluble precipitant. If you’ve protocols with 6 M GuHCl, swap it out for 5% SDS. As with everything, we always recommend testing your specific conditions in particular for important samples.

How do I match my protein concentrations when I solubilize everything in 5% SDS?

Use a BCA assay and not Bradford: BCA is not sensitive to detergents and Bradford is. Note that the original BCA assay is not compatible with reducing agents. There exist, however, reducing agent compatible BCA assays, e.g. BCA-RAC assay from Pierce. You can also use amine-reactive fluorogenic reagents such as CBQCA (3-(4-carboxybenzoyl)-2-quinoline­carbox­aldehyde) and also absorbance at 280 nm. We use the standard Pierce BCA assay and get outstanding results: apply 20 uL of sample in various dilutions (i.e. dilute e.g. 1:1, 1:3, 1:10, 1:30 into 5% SDS lysis buffer (without reducing reagents) along with a standard curve in the same buffer and plate in triplicate) into a 96 well plate avoiding the outer row and column. To all samples, then add 200 uL of working reagent with a multichannel pipette (go fast and accurately). Then place the plate on a 95 C block and wait until you get a nice deep purple color at the e.g. 2 mg/mL max protein concentration. Measure and plate this. You will find the curve to be extremely linear.

My (favorite) protein is impossible: I can’t seem to see it! What to do?

That’s a tricky – and addressable – question, especially if your protein exists in another source such as recombinant or through an overexpression system. You must address two basic questions: 1) is your protein getting into your digestion and being observed?; and 2) if it makes it in, is it there in sufficient quantity to see in the background of your other proteins and your up-front separation? If you have a protein source, start with a positive control: do an S-Trap digest of, by example, a cell pellet from induced over expression. Verify that you have protein (e.g. you have a large band after overexpression on an SDS PAGE gel, or you have it from a purified source), that you’re sure your machine is working (positive control of e.g. HeLa cell tryptic digest) and that your digestion is working (run an SDS-PAGE gel of the input and output of an S-Trap digest). If these all check out but you still can’t see your protein by mass spec proteomics, you need to consider possibilities. Biochemical reasons include that the protein is not the expected protein (this happens more frequently than people realize); that it has a limited number of lysines or arginies; or that the peptides are insoluble in buffer A of your LC system, or that they are not coming off your separation column, or of course some combination of these reasons. Bioinformatics search reasons include the protein being absent from the database, the wrong databases being searched, and wrong search parameters (including enzyme specificity, fixed PTMs, etc.). Once you have verified that you can see your protein by proteomics, the next question is one of dynamic range. If you have purified protein, add it back to your sample at 0.1% wt:wt to begin and determine if you can detect it within that background given the current LC separation. Titrate up and down as needed to establish a lower limit of detection (LLD). If this LLD is insufficient to detect your protein at the levels present, you must increase chromatographic separation and/or determine a way to preferentially extract your protein, leaving behind interfering signal. For difficult proteins at particularly low levels, Western blots with extended incubation times may be more practical.

I have a detergent-containing buffer. The detergent isn’t SDS. Can I still use S-Traps? What should I do?

Add SDS to your buffer to at least a 2% w/v final concentration. Then proceed as normal: add phosphoric acid, then binding buffer and everything will be great! Like with other buffer components, if you run into a buffer component which is “sticky” and appears to not be fully removed by three washes, simply wash more.

How should I best remove DNA?

There are two simple and easy ways to remove DNA. First, apply high-intensity sonication with a probe sonicator or most preferably with a Covaris Adaptive Focused Acoustics (AFA) unit. (If you are manually processing samples, make sure to keep the time, intensity and depth of probe contact constant between samples). Alternatively, resuspend your samples in 5% SDS containing 2 mM MgCl2. Then, add a small amount of benzonase by Roche (e.g. 0.5 μL of 250 units per μL into 25 μL). Your DNA will almost instantly be gone. (Benzonase maintains activity in 5% SDS with magnesium sufficiently long to remove destroy DNA which then washes through.)

Covaris AFA technology is a non-contact process which imparts strong acoustic forces in a tightly controlled, isothermal environment. AFA treatment generates extreme sheer forces which homogenize samples, sheer DNA and force proteins into solution, especially in the presence of 5% SDS. AFA is amenable to low input material especially with 5% SDS which coats tubes and tips.

Can I use other buffers than TEAB?

Yes. In all buffers, you can use tris in place of TEAB at the same pHes and concentrations. You can also use ammonium bicarbonate in the lysis and digestion buffer. However, you cannot use ammonium bicarbonate in the methanolic S-Trap binding buffer: its solubility is not great enough in 90% methanol. You can use ammonium bicarbonate (or anything else) in the digestion buffer without issue. Remember however: if doing iTRAQ or TMT, you must use TEAB, HEPES or another buffer without primary or secondary amines (cf. Good Buffers).

If I make my own buffers, what can I adjust the pH with?

For the lysis buffer, we recommend using phosphoric acid to adjust the pH as the samples will be further denatured with that acid. For the methanolic S-Trap binding/wash buffer, we recommend phosphoric acid. For the digestion buffer, anything compatible with your protease can be used including whatever acid or base is needed to correctly set the pH. Note that typical digestion buffers of ammonium bicarbonate and TEAB do not require pH adjustment for trypsin. However buffers made with tris or HEPES free base (or acid) will need pHing.

How essential is the correct pH value?

For the lysis buffer, not important at all. For the binding buffer, important. Note that TEAB is volatile and can change pH if left to sit or stored with a large air headspace. We recommend adjusting the pH of a 1 M solution with 85% phosphoric acid, aliquoting this solution and keeping it frozen.

Will a 1-hour 47 C digestion give the same results (numbers of identified peptides, missed cleavages, etc.) as an overnight in-solution digestion?

That actually depends on your instrument and how it’s run. On a relatively fast 1D run of a complex sample, especially with a machine that is a year or two old, you will probably not see a difference. On a fast and new instruments such as new Orbitrap class machines, by peptide counting, you will see an increase in missed cleavages from ~10% – 15% for an overnight tryptic digest to ~20% – 30% for a 1-hr digest. However ion current tells a different story: if we set to 100% the total amount of ion current in peptides with no missed cleavages generated in an overnight tryptic digest, the corresponding amount of ion current for fully tryptic peptides in a 1-hr digestion is in our experience always >90% with >95% being quite common. This discrepancy follows from the fact that new machines are simply so fast and so good at picking up low level missed cleavages i.e. little peaks. A 1-hr digestion time is thus “95% of the way there” and matches the duty cycle of typical runs (around 1 hr) to the duty cycle of sample preparation. However, you can and should optimize digestion for your samples and workflows. We have people doing everything from 10 min digestions (at 1:1 wt:wt substrate:trypsin to limit deamidation) to overnight digestions (use a water bath to keep columns from drying out, or place a beaker of water in an incubator).

Why 47 C? Does 47 C work for other enzymes?

47 C for 1 hr is for trypsin only and is an intermediate between the S-Trap and digestion buffer warming up, trypsin working very rapidly and then starting to thermally denature to the end of the 1 hr. It was empirically determined for trypsin. Other enzymes require different temperatures and times.

Can I use other enzymes than trypsin in the S-Trap? Are the conditions the same?

Yes you can use other enzymes. No the conditions are not the same as for trypsin.

What happens if the columns dry out?

Just rehydrate them with digestion buffer, let them sit for 15 min at 37 C and follow the standard elution procedure. The protein trap is designed to have no affinity for digested peptides; they just need to be solubilized.

How do I remove bubbles from the trap?

The easiest way is to “flick” the tubes. Alternatively you can pipette the digestion solution up and down. Just be sure that all the solution is on the trap and not on the side of the spin column where it will dry out.

What is the best way to lyse my cells?

If you want to simultaneously lyse your cells and instantly stop all enzymatic activity, “freezing” the sate of biology, heat 1X lysis/solubilization buffer (5% SDS, 50 mM TEAB pH 7.55) to 95 C and add TCEP to 5 mM. Squirt this on your cells and sonicate (better: process with AFA) to sheer the DNA. Let cool to room temperature, add MMTS to 15 mM, let stand for 30 min (or 10 min at 37 C) and you’re ready to process. Note that this approach is also one of the best ways to prevent loss of PTMs by endogenous enzymatic activity.

Do S-Traps trap low molecular weight proteins?

Yes, especially if they are in a complex mixture with many other proteins which tend to carry them into the protein trap. However proteins have extraordinarily diverse properties and if you’re working with a purified protein, there is a chance that it has unusual properties like methanol solubility (see below).

Is the S-Trap compatible with all proteins?

Yes with the single exception of alcohol soluble proteins, which we have observed only in plant seeds samples. (You can verify that your sample has no methanol soluble proteins by speed-vacing the flow-through and washes of the S-Trap and running that fraction on an SDS-PAGE gel; we have not observed this with anything outside of seeds.) If you are studying plant seeds, remember that prolamin storage proteins are alcohol soluble. These include the gliadins of wheat, the avenins of oats, the zeins of corn, the hordeins of barley and secalins of rye. In addition, while glutelin proteins are typically oligomerized via interchain difulfide bonds and thus insoluble, they become alcohol soluble when reduced.

What should I do with lipid rich samples?

The SDS lysis/solubilization buffer plus methanolic S-Trap washes are usually sufficient to take care of lipids from common samples like cell culture. However, like proteins, lipids have very disparate physiochemical properties and not all are methanol soluble. If you are working with a particularly lipid rich tissue, such as brain or adipose tissue, you will probably want to do additional lipid cleanup.

There are two main approaches: remove the lipids before protein processing, or remove the lipids when the proteins (and potentially methanol insoluble proteins) are on the trap. The best way to remove lipids before S-Trap processing is to use a cryogenic bead beater at liquid nitrogen temperature where the sample and an organic such as DCM, chloroform or ether and pulverized together. (Note at –196 ºC, these solvents are solid. Typically a 5 – 10-fold volume excess of organic solvent over tissue is used.) This sample is then warmed to 4 ºC, vortexed, the organic removed (filtration or centrifugation; be cautious with the density of chloroform and DCM as proteins float) and the samples dissolved in 1X 5% SDS with harsh sonication (probe) or Covaris AFA. Proteins tend to be denatured by organic lipid extractions, and require extra encouragement to go into solution.

To remove lipids on the trap, follow the S-Trap protocol up to the binding and first wash step. Then, rinse the proteins with an appropriate solvent (see below table). We recommend 2:1 chloroform/methanol to begin. Do the next two washes with the 90% methanol S-Trap binding/wash buffer and proceed with the protocol.

Solvent mixture(s); all v/v Citation
10:3:2.5 MTBE/methanol/water; 60:30:4.5 chloroform/methanol/water Cai, T., Shu, Q., Liu, P., Niu, L., Guo, X., Ding, X., … & Wu, P. (2016). Characterization and relative quantification of phospholipids based on methylation and stable isotopic labeling. Journal of Lipid Research, 57(3), 388-397.
30:25:41.5:3.5 chloroform/isopropanol/ methanol/water Shiva, S., Enninful, R., Roth, M. R., Tamura, P., Jagadish, K., & Welti, R. (2018). An efficient modified method for plant leaf lipid extraction results in improved recovery of phosphatidic acid. Plant methods, 14(1), 14.
10:3:2.5 MTBE/MeOH/water Matyash V, Liebisch G, Kurzchalia TV, Shevchenko A, Schwudke D (2008) Lipid extraction by methyl-tert-butyl ether for high-throughput lipidomics. J Lipid Res 49: 1137-1146.
2:1 chloroform/methanol Folch J, Lees M, Stanley GHS (1957) A simple method for the isolation and purification of total lipides from animal tissues. The Journal of Biological Chemistry 226: 497-509.

Knittelfelder, O. L., Weberhofer, B. P., Eichmann, T. O., Kohlwein, S. D., & Rechberger, G. N. (2014). A versatile ultra-high performance LC-MS method for lipid profiling. Journal of Chromatography B, 951, 119-128.

4:1 methanol/chloroform Dawson, G. (2015). Measuring brain lipids. Biochimica et Biophysica Acta (BBA)-Molecular and Cell Biology of Lipids, 1851(8), 1026-1039.
1:2 chloroform/methanol Bligh EG, Dyer WJ (1959) A rapid method of total lipid extraction and purification. Can J Biochem Physiol 37: 911-917.
1:1 chloroform/methanol; others Reis, A., Rudnitskaya, A., Blackburn, G. J., Fauzi, N. M., Pitt, A. R., & Spickett, C. M. (2013). A comparison of five lipid extraction solvent systems for lipidomic studies of human LDL. Journal of lipid research, jlr-M034330.
3:1 butanol/methanol Löfgren, L., Forsberg, G. B., & Ståhlman, M. (2016). The BUME method: a new rapid and simple chloroform-free method for total lipid extraction of animal tissue. Scientific reports, 6, 27688.
Many Christie, W. W., & Han, X. (2010). Lipid Analysis-Isolation, Separation, Identification and Lipidomic Analysis, 446 pages.

My sample is not going into solution. What should I do?

For particularly difficult samples, be sure to start with samples that are as finely pulverized as possible. Especially for tough samples like tissues, either find and use a bead beater under liquid nitrogen, or use a Covaris cryoPREP to pulverize it. Alternatively for larger scales and absolute maximum pulverization, dice your sample (preferably frozen), lyophilize it and use a laboratory-scale jet mill. Note the losses here will be higher.

If you are sample limited and/or want to improve your recovery, use a Covaris AFA unit which is well suited to small sample amounts, or a cryogenic bead beater with frozen 5% SDS. For AFA, place the sample in 5% SDS and immediately AFA treat until the sample is fully disaggregated. Samples are kept isothermal typically at 4 C. For cryogenic bead beating, freeze 5% along with the sample, make sure the bead(s) are able to move, and pulverize until a free-flowing powder results. In both cases, 5% SDS will coat the surfaces of tubes, tips and beads, limiting loss. Surfaces can be washed if desired with additional buffer.

If a Covaris AFA unit or cryogenic bead beater is not available, probe sonication is an option. Be careful, however, to not explode your sample out the top of the tube. Alternatively or additionally, incubate samples with agitation at elevated temperatures (e.g. 55 C for 2 hrs on a heating block or overnight at 37 C on an end-over-end rotator). Note that elevated temperatures can cause sample degradation, even with enzymes like proteases and phosphatases inhibited by 5% SDS.

For tissues and other samples which may be crosslinked through disulfide bonds, we recommend including 5 – 10 mM TCEP during the solubilization step. (Note however that this must be removed before a standard BCA assay or use a reducing reagent compatible BCA assay.)

With extensive solubilization, it may be useful 1) to sparge your solutions with nitrogen or argon and to keep the protein sample under the same atmosphere (to reduce oxidation) and 2) to do an initial extraction (remove the supernatant and keep it) followed by a harsh extraction. Sequential extraction can be useful to 1) preserve the chemical integrity of the proteins which more easily go into solution and 2) to provide different classes of proteins. Frequently, the protein of interest is more concentrated in one of the fractions.

Do I need to clean my peptides before running them?

We generally recommend concentrating the peptides using desalting columns, however, C18 columns and tips have much lower recoveries than most people are aware of: around 50% – 60% in typical use. If you are working with low-level and/or hydrophobic samples, the serial loss over desalting steps, a LC trap column and then an analytical column can result in seeing nothing or almost nothing. We and customers have observed this in particular with exosome samples: the proteins are typically hydrophobic and membrane bound and often people have only a microgram or two. If you are doing a large scale digestion for PTM enrichment, you will want to do desalting so that you can dry your peptides down and solubilize them in the binding buffer for your PTM enrichment.

I dried down my peptides and have a larger pellet than I think I should have. What is going on? Or: my LC system is working great but the peptide chromatography doesn’t look right. What’s happening?

Both of these are signs that the protein must be washed more to further remove SDS and/or any buffer components: do further washes. SDS is moderately hydrophobic and will come off C18 columns. Before it does however, it alters the surface to make it more of a cation exchange.

Can I use the S-Trap to prepare samples (for gel analysis, etc.)?

Yes, it is completely possible to use S-Traps to clean up sample especially for SDS-PAGE gels. In this case, concentrate and clean your proteins in exactly the same way but do not add trypsin. Rather, add 1X SDS-PAGE buffer to the trap (the volume depends on the size of the wells in your gel; at least 20 uL however), heat the trap to solubilize the proteins (5 min at 95 C) and spin out your eluted proteins. If you have a lot of protein, this single elution is fine and recovery is typically around 90%. If you have only a small amount of protein, then do three elutions with 0.5X Laemmli loading buffer, put it on a speed vac to concentrate it (the SDS will be a chunk then) and resuspend back to the right volume. It is a good idea to both reduce and alkylate proteins before purifying them with an S-Trap: first you don’t have to worry about disulfide bond formation causing a ladder in your gel and because you can then go straight to Gel-LC if you want to do MS analysis.

I’m doing iTRAQ or TMT. Do S-Traps work for that?

Yes. The standard protocol is designed for use with isobaric amine labels (i.e. iTRAQ and TMT).

I’ve got an immunoprecipitation (IP)/I need to do an IP. Now what?

There are a few considerations with IPs including how dirty or clean they are, how much protein has been immunoprecipitated (a function both of how much protein is present and the amount and affinity of antibody, as well as binding conditions) and the signal to noise ratio of capture antibody to target antigen. To improve the cleanliness of IPs, run experiments of increasing wash stringencies monitored by Western and silver stained gel: you should increase stringency so that the background is clear while maintaining clear differences between a control lane (ideally an isotype-controlled antibody without affinity to your target antigen) and the experimental lane. As the S-Trap removes all such buffer component, you can use salts, chaotropes and detergents including polymeric detergents such as tween or triton. This optimization is typically the key to getting reliable IP results and additionally improves the signal to noise for small amounts of captured target protein. To improve the relative amount of captured antigen to capture antibody, you can covalently anchor your antibody (e.g. use BS^3 on protein A/G after Ab binding, or put your antibody on covalently with for example epoxy or CNBr derivatized beads). You can elute the IPed proteins either with 5% SDS (and heating to 95 C as desired; often this helps), which will strip everything especially if reduction and alkylation is done on beads. Alternatively, you can try to use other detergents or conditions which elute the protein(s) of interest and leave behind the “garbage.” Cf. Impact of Detergents on Membrane Protein Complex Isolation. http://www.ncbi.nlm.nih.gov/pubmed/29110486. If you didn’t elute in SDS, add it to 5% and the SDS will carry the (for example) nanograms of IPed material which you will then efficiently retain with the high recovery protocol. The high recovery protocol uses trypsin itself as both a carrier and a proteolytic enzyme for three reasons: first, without a carrier, the small amount of antigen will be poorly retained; second adding trypsin does not further complicate your spectrum (often tryptic autodigestion products are already in exclude lists); and third, at those very low levels, in an [ES] reaction the substrate is at a very small concentration (and thus rate limiting, rather than the protease), so we must significantly increase the concentration of trypsin for efficient proteolytic digestion.

Someone brought me an immunoprecipitation (IP) in blue Laemmli loading buffer. Can S-Traps help?

Yes. Reduce and alkylate as normal, acidify as normal and proceed as normal. You can also use the high recovery protocol if you believe the amount of protein is low. Note that you must use very high concentrations of alkylating reagents if proteins were eluted in 5% beta-mercaptoethanol (BME): neat BME is 14.21 M making 5% BME 710 mM! Avoid this if possible.

What is the reproducibility of S-Trap sample processing?

Typically  less than 10% CVs for replicate digestions.

I’m analyzing urine. What should I do?

First concentrate the urine either by lyophilization or a centrifugal filter to around 1 – 2 mg/mL then follow the standard S-Trap procedure including the addition of SDS to 5% to begin, acidification and addition of binding buffer. Urine is usually around .08 mg/mL so typically a 10 – 20x concentration is necessary. Conditions of pathology may however significantly alter the protein concentration; determine protein content before processing.

I’m analyzing CSF. What should I do?

If your CSF samples are already aliquoted and frozen, lyophilize them. Bring them up in 8 M urea, 5% SDS, 50 mM TEAB pH 7.4 to approximately 1 – 2 mg/mL. (CSF is normally around 0.2 – 0.4 mg/mL so a 5 – 10x reduction in volume is typical.) Sonicate the samples (keeping the time of sonication identical), reduce and alkylate and process as per the standard protocol. Speed-vacing in the liquid state is to be avoided as it may introduce changes. If you have to aliquot the CSF samples, minimize freeze thaw.

I’m analyzing serum/plasma. What should I do?

Dilute the serum or plasma into 1X 5% SDS lysis buffer to a final concentration of 2 mg/mL. Serum and plasma are typically around 80 mg/mL so a typical dilution factor is 40x. Reduce and alkylate the proteins in this SDS solution, then proceed with the standard protocol including acidification, dilution with the binding buffer, application to the S-Trap, and cleaning and digestion in the trap.

Do S-Traps trap peptides?


I’m analyzing tissue. What should I do?

Please see the S-Trap protocol for mammalian samples.

Can I use another acid than phosphoric acid?

Yes, lactic acid at 3% or citric acid at 4% final concentration.

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