Category Archives: In-solution digestion


Which reagent is better for reducing disulfides, DTT or TCEP? The two reagents are quite different in their reactivity, stability towards oxidation, reaction mechanisms, and other categories.


DTT is a thiol-containing reagent, and this must be considered in applications involving thiol labeling. TCEP is charged in solution and should not be used in isoelectric focusing. Aqueous solutions of TCEP are quite acidic (pH 2-3).

TCEP HCl is odorless, air-stable crystalline solid, soluble in water at a > 1 M concentrations. It reduces disulfides at room temperature in < 5 min in dilute solutions (5 -50 mM). There is no need to remove TCEP prior to the use of sulfhydryl-reactive labels or crosslinkers. TCEP is selective toward disulfides, and is reactive at a broad pH range.

For those who love Chemistry:

TCEP mechanism

Reduction of disulfide with TCEP. First step is rate-determining, kinetic rather than thermodynamic control.



Reduction of disulfide with DTT. Formation of stable cyclic disulfide drives the reaction.

You can learn more about DTT and TCEP from this article: “A Comparison between the Sulfhydryl Reductants Tris(2-carboxyethyl)phosphine and Dithiothreitol for Use in Protein Biochemistry”, Elise Burmeister Getz et al.  Analytical Biochemistry 273, p. 73–80 (1999)

In-solution digestion of proteins

Purified proteins or protein mixtures can be digested in solution if an additional separation step is undesirable or unnecessary.

Proteins in solution are usually denatured by boiling or using denaturing buffers. During this step, the disulfide bonds must be reduced, and the sulfhydryl groups must be alkylated to prevent the disulfides from re-forming. The protein samples are then incubated with trypsin for several hours, and the resulting peptides can be analyzed by MS.

Denaturing buffers contain chaotropic agents, salts, and detergents at concentrations that inactivate trypsin. Before adding trypsin, you should desalt your protein sample and remove detergents. There are a number of the detergent removal and desalting options: detergent removal spin columns, size-exclusion and MW-cut-off spin columns, ion-exchange membranes and resins, etc.  Gel-assisted proteolysis is another option, but then it is not really an ‘in-solution’ procedure, although it does not involve electrophoresis. I thought I should mention it here in case it could be of interest to you.

You will need

Digestion buffer: 16 mg/mL ammonium bicarbonate in water

Reducing reagent: 30 mg/mL TCEP (~100 mM, Sigma C4706)  or 15 mg/mL DTT (Sigma D0632 ) in digestion buffer. NOTE: The 30 mg/mL TCEP stock solution must be prepared in 16 mg/mL (~200 mM) ammonium bicarbonate to bring up its pH. The final concentration of TCEP in the digestion mixture should be 5-10 mM.

Alkylating reagent: 18 mg/mL iodoacetamide (Sigma I1149) prepared fresh in the digestion buffer

Proteomics grade trypsin (e.g. Sigma T6567-5x20UG or Thermo Pierce 90057, 5 vials x 20 ug lyophilized powder). Trypsin, 20 ug can be dissolved in 20 uL of 1 mM HCl or 50 mM acetic acid, pH ~ 3, aliquoted and stored at -20C (stock solution).

To prepare activated (or working) trypsin solution, dilute trypsin stock solution with digestion buffer 10-fold to 0.1 ug/uL concentration.


Volumes are approximate, it is a sample procedure after all. Trypsin should not exceed 5% of the total protein, provided the protein concentration range is known.

Combine 15 uL digestion buffer, 3 uL reducing reagent, and up to 12 uL sample solution containing 0.025 – 10 ug protein (total volume 30 uL)

Denature/reduce at 50-60 C (TCEP) or in a boiling water bath (DTT) for 5 – 10 min, cool to r.t., spin down to collect the sample

Add 3 uL alkylating reagent and incubate in the dark at r.t. for 20 min

If protein sample contains detergents, salts, or chaotropic agents, perform buffer exchange after the alkylation using a 3,000 MWCO centrifugal filter. It will be impossible to remove detergents after the digestion; and most detergents are not compatible with LC MS analysis. You can find a list of MS-compatible detergents here.

Add 1-5 uL activated trypsin and incubate at 37 C for 3 hrs. Optional: add 1-5 uL of fresh activated trypsin and incubate for an additional 2 hrs at 37 C or overnight.

Once the incubation is complete, the peptides can be submitted for analysis or stored at -20 C.

Helpful tips

Minimum sample amount required for MS analysis is in the fmol/uL range (ng/uL). Solutions of peptides at very low concentrations (e.g. less than 100 fmol/uL) should not be stored for more than 1-2 days.

Always run a control along with your sample. It could be a 1 mg/mL solution of bovine serum albumin or other standard protein that you have in your lab prepared in the same buffer as your sample and taken through the entire procedure. We don’t charge for analyzing your controls.

TCEP is a great reducing reagent because it does not contain -SH groups and thus does not consume iodoacetamide during alkylation, unlike DTT. TCEP solutions in water are acidic. Depending on your buffer composition, you might observe your sample coming out of the sample tube as a soapy foam the moment you add TCEP. It is pretty much impossible to put that foam back into the tube, don’t ask me how I know. So, prepare your TCEP solutions in ammonium bicarbonate buffer to get a pH close to 8. Another note about TCEP: it should never be stored in phosphate buffers because it quickly decomposes in the presence of phosphate.

Have I missed anything? Let me know!

Ammonium bicarbonate or triethylammonium bicarbonate?

This was a question from one of my blog’s secret readers. Actually, most of the time I feel like I am talking to myself: “Hey Tania, how do you prepare a protein sample for proteolysis?” “Well, Tania, let me show you in a step-by-step tutorial.” No comments, no questions, no pointing out typos, no “thank you, Tania, but there’s a better way to do this”?

Oh well, back to ammonium bicarbonate. This is a volatile salt which breaks down to ammonia, carbon dioxide, and water. Volatile salts are the only salts compatible with MS. Aqueous solutions of ammonium bicarbonate (0.01 – 0.1 M) have pH around 8, the optimal pH for trypsin activity. Ammonium bicarbonate competes with basic amino acids for Coomassie dye, which makes it a great de-staining reagent for the in-gel digestion procedure. All this goodness comes at a very reasonable price – what not to like? Another ammonium salt, triethylammonium bicarbonate (TEAB), is more volatile than ammonium bicarbonate; it is also more expensive. TEAB is a buffer of choice for LC-MS applications: TMT (iTRAQ) amine-reactive labeling, ion-exchange chromatography, protein solubilization (when neutral and acidic pH is undesirable), in-gel digestion, etc.


Detergent removal 2: Affinity resin

To continue our detergent theme, here’s an affinity-based detergent removal method that is faster than the gel-assisted proteolysis but removes only the detergents, leaving the salts and chaotropic agents for you to deal with later. Unlike the gel-assisted method, it works for both proteins and peptides. The gel-assisted method is best suited for proteins, because the small, more soluble peptides are likely to elute out of the gel matrix during the washing.

I use Pierce detergent removal spin columns (0.125 mL format) in my lab, but there are other options available such as Bio-Beads or HyperD, each with its own pros and cons in the business of detergent removal. Pierce also sells a so-called HiPPR detergent removal resin (high protein and peptide recovery) for low-protein-concentration samples. The initial % detergent in such samples must also be low (ca. 1%).

The Pierce resin removes common ionic, nonionic, and zwitterionic detergents from protein and peptide solutions. This oligosaccharide-based affinity resin has a small hydrophobic cavity which creates a microenvironment for a detergent’s nonpolar moiety to enter and form an inclusion complex.

The workflow is simple: (1) centrifuge the column to remove storage buffer, (2) wash 3 times with your favorite buffer (pH 5-10), discarding the buffer each time, (3) add protein or peptide solution and let the resin do its magic for 2-5 min at room temperature, (4) centrifuge to collect your >95% detergent-free sample, i.e. don’t discard the flow-through this time!

Antharavally and co-workers from Thermo Scientific Pierce Protein Research published a study examining the detergent removal efficiency and protein recovery using their resin under several conditions (doi:10.1016/j.ab.2011.05.013). A table from this reference gives you some idea of the detergent concentrations removable with the Pierce resin.

Detergent removal efficiency (640x243)

Samples (0.1 ml containing 0.100 mg BSA + detergent at maximum concentration) were processed through 0.5 ml of Pierce detergent removal resin, and the residual detergent was measured as described in Materials and Methods. Protein concentration was determined by BCA protein assay (Pierce).

Detergent removal 1: Gel-assisted proteolysis

As promised, here’s a straightforward way to remove detergents, urea, and other LC-MS incompatible nasties from small-volume samples. The literature calls it ‘gel-assisted’ proteolysis. The idea is to entrap the protein solution in a polyacrylamide gel matrix, wash out detergents, salts, and chaotropic agents, and perform in-gel digestion. This technique works great for membrane proteins which are notoriously difficult to dissolve, and it is quite useful for any protein sample clean-up.


For my little demo, I used a 1 mg/mL BSA solution in 2% SDS. The disulfides were reduced with TCEP and alkylated with IAA, after which the protein solution was very thoroughly mixed with a 30% T acrylamide monomer solution. I quickly added 10% APS and TEMED and immediately vortexed and centrifuged this mixture so that the liquid is collected at the bottom of the tube. The polymerization time is very short, a minute or two! I left it to completely polymerize for another 20 min.


Using scalpel I removed the gel plug from the tube and diced it into small pieces. After 6 washes with 8 mg/mL ammonium bicarbonate in 50% acetonitrile, I dried the gel pieces in neat acetonitrile, removed the acetonitrile and added trypsin (see the in-gel digestion tutorial for details).



Fast forward to the MS analysis: Since the original BSA solution was very concentrated, I dissolved the peptides in 540 uL of mobile phase and injected 1 uL of this solution (55 ng total protein on column or approximately 1 pmol).








Second example shows an ion chromatogram from a 10-uL IP eluent containing 2% detergent which I cleaned up and digested using this technique.





















As always, let me know if you have questions!


doi: 10.1074/mcp.M500138-MCP200

doi: 10.1074/mcp.M800068-MCP200



LC-MS compatible detergents

They do exist! They are compatible with the reverse-phase chromatography! They solubilize stubborn proteins and improve proteolysis! Here are their names:

Acid-labile surfactants are hydrolyzed at low pH, and the hydrolysis products are compatible with reversed-phase separations and MS. These include RapiGest SF and PPS Silent Surfactant.

Invitrosol is a homogeneous surfactant whose elution profile does not overlap with the proteolytic peptides elution profiles.

ProteaseMAX is a surfactant that degrades during proteolysis, and its degradation products do not interfere with LC-MS.

All other detergents are not compatible with LC-MS and must be removed from the samples prior to analysis. These detergents include SDS (sodium dodecyl sulfate) and LDS (Lithium dodecyl sulfate), NP-40, Triton, Octyl glucoside and octyl thioglucoside, CHAPS, sodium deoxycholate, lauryl maltoside, Brij-35, etc. There are several ways to remove detergents, which is a topic for another blog post!