General Samples: Guidelines & Protocols for Sample Preparation and Submission

Purity is the key to successful mass spectrometry of your sample. Salts, detergents, plasicizers, polymers, and other small-molecule contamination can significantly impact the ability to successfully ionize and analyze your sample. It is easy to inadvertently contaminate your sample.

The following are general guidelines for sample preparation and submission:

  • Use HPLC to purify and collect samples whenever possible. Be sure that there is no contamination from previous samples or background contamination from the HPLC system before collecting fractions of your samples. Collect fractions in clean microcentrifuge tubes (see below).
  • Always use “cleaned” or new glassware for preparations of solutions and reagents. Rinse common-lab glassware copiously with organic solvents and nanopure water before use; glassware may have traces of detergents or polymers adsorbed in the surface of the glass. Glassware that has been washed at one time with detergents will be contaminated. It’s a good idea to dedicate new glassware to certain solutions only, so that solutions don’t become inadvertently contaminated.
  • All polypropylene tubes and vials should be rinsed with acetonitrile, methanol, and nanopure water before making up solutions, storing, or collecting samples (including microcentrifuge tubes).
  • Use clean (rinsed with methanol and nanopure water) microcentrifuge tubes to submit samples, unless the solvent your sample is in is not compatible with polypropylene (e.g. acetone). (NalgeneTM has published tables of compatibility of polypropylene with different solvents).
  • Do not wrap sample vials with parafilm. Parafilm will contaminate samples that are in organic solvents in the microcentrifuge tubes.
  • Desalt samples prior to sample submission (e.g. HPLC, C18 sep-pack, C18 Ziptip, dialyses). Salts will have a significant impact on ionization, both for ESI or Maldi ionization. If needed, we can also desalt with Ziptip purification at the MS Facility.
  • Certain buffers (non-volatile buffers, such as HEPES) and purification methods, such as sephadex, will actually contaminate your sample. Polymers from certain types of sephadex preparations can inhibit or completely suppress ionization.
  • Always use HPLC-grade quality solvents and only nanopure water for sample preparations.
  • Do not prepare your samples in DI water or tap water (these are heavily contaminated); use only nanopure water.
  • Do not concentrate (e.g. lyophilization or speed-vac) your sample from large volumes of solvent or nanopure water (e.g. > 0.5 mL). This will concentrate non-volatile contaminants that are in all solvents (even HPLC-grade solvents and nanopure water).
  • Do not use glass vials with screw-on caps that can contaminate your sample. Paper or foil-lined caps will contaminate your sample when the solvent contacts the cap. Preferrably, use cleaned polypropylene microcentrifuge tubes (when the solvent is compatible with polypropylene).
  • Be careful not to touch microcentrifuge tubes or caps with fingers, and don’t use gloves coated with powder on the inside. This will contaminate samples. Gloves should be rinsed with nanopure water.
Nucleic Acids

Analysis of nucleic acids (i.e. DNA, RNA, and PCR products) requires that these samples be extremely pure (free from salts and small-molecule contaminants). Desalting nucleic acids is required. In most cases, this can be done by precipititation. However, it is highly recommended that the MS Facility do this desalting and purification for best results.

Proteomics/Protein/Peptide Samples: Guidelines & Protocols for Preparation and Submission

It is highly recommended that you discuss your specific proteomics project and the preparation of your protein samples with the MS Core Facility personnel prior to preparing samples and submitting them for mass spectrometry analysis!

Basic Guidelines for Protein Sample Handling and Preparation

High purity preparation of a protein sample (or protein complex) is the key to successful mass spectrometry. It is very important to prevent or minimize contamination of your sample. Problem contaminants include keratins, plasticizers, polymers, detergents, and non-volatile buffers. (Also refer to General Sample Guidelines)

  • Avoid contamination with keratins. It is extremely easy to contaminate your samples with keratins, which are ubiquitous and come from skin cells, hair, fingerprints, dust, lint, lint from Kim-wipes, etc. The smallest speck of dust or lint will probably overwhelm your sample with keratin proteins. Great care is required to prevent keratins from showing up in MS analyses. All vials, tubes, flasks, gel apparatus, etc. should be rinsed with solvents and nanopure water prior to use and should be kept in a dust-free environment, covered or in a laminar flow hood. It is best to use new polypropylene tubes to prepare aqueous solutions (and you should rinse new tubes first with solvents such as acetonitrile, methanol, then nanopure water before making up solutions).

Working in a laminar flow hood is highly recommended. If keratins appear in the mass spectrometry data of your protein samples, it may be necessary to toss out all solutions and reagents involved.

  • Avoid contamination or addition (even trace levels) of polymers, detergents, plasticizers, non-volatile buffers, Protein-A Sephadex beads, BSA.
  • It’s a good idea to use powder-free nitrile gloves. Rinse gloves with ethanol and nanopure water before touching any tubes or surfaces that will come into contact with your protein prep. Don’t use Latex gloves. Latex is manufactured with proteins, which can contaminate your protein samples.
  • Use of “low-binding” (but not silanized) polypropylene microcentrifuge tubes is recommended to prevent adsorptive losses of proteins and peptides and for sample storage, not glass.
  • Protein storage recommendations:
    • Proteins in coomassie-stained gels are relatively stable at 4C or -20C. Silver-stained protein bands must be destained at the MS Facility immediately after staining (see below). Proteins in solution may not be stable and may be susceptible to proteases. If the protein to be analyzed is in solution, it should be submitted to the MS Facility and analyzed as soon as possible after it is isolated and purified or should be kept frozen (i.e. -20C or -80C).
Use of Protease Inhibitors and other Reagents

Do not use protease inhibitors in solutions that will be submitted directly for protein digestion and protein ID. We will not be able to digest the proteins with trypsin (due to the protease inhibitor).

Protein solutions that will be run on gels can be treated with protease inhibitor, since the protease inhibitors will not impact the in-gel digestion and LC/MS/MS analysis.

There are several issues associated with use of protease inhibitors and phosphatase inhibitors during protein preparations. Some protease inhibitors will affect mass spectrometry analyses and results and should be used with caution. See examples of protease inhibitors below.

  • AEBSF – probably the most commonly used protease inhibitors contain AEBSF (4-(2-aminoethyl)-benzenesulfonylfluoride. This reagent is found, for example, in Roche Complete Mini tablets and Roche Pefabloc. It is not recommended using AEBSF as a protease inhibitor for most protein preparations, but depends on the protein information being requested and how the protein is to be analyzed (you can discuss with MS Facility personnel).
    • AEBSF is a serine-protease inhibitor and, as such, will affect the ability for us to digest the protein sample in solution using trypsin, chymotrypsin, and other enzymes, and the subsequent affect the ability for us to obtain good MS results on the proteins in the sample.
    • AEBSF functions by modifying or derivatizing proteins, preferentially on Tyr, and to a lesser extent on Lys, and His residues, and the amino-terminus. These modifications result in a net mass addition of 183 Da per AEBS-group and affect the outcome of peptide identification, sequence coverage, and protein ID. The extent of derivitization depends on the time and conditions of exposure to the AEBSF reagent. AEBSF can be mostly removed from solutions by Ziptip purification (or by running the protein on a gel), prior to protein digestion and LC/MS/MS analysis, however, any AEBS derivatizations will remain intact and can potentially impact protein digestions and identification of peptides and proteins in database searches (by shifting the mass of the derivatized peptides).
  • PMSF –protease cocktails with PMSF apparently do not derivatize proteins, or there is minimal derivatization. If a protease inhibitor is an essential part of your protein isolation, we recommend using PMSF. For example, the following cocktail has been successfully used with protein ID and identification of phosphorylations:
    • Cocktail: PMSF (100 mM), phenanthroline (1 mg/ml), benzamidine HCl (100 mM), pepstatin A (1mg/ml). This stock is used at 1/500-1/1000 dilution in all the buffers, then removed by dialysis (into ammonium bicarbonate) just immediately prior to submitting the protein for mass spec analysis. No phosphatase inhibitors were used.
  • Other Protease Inhibitors –there are many other protease cocktails and inhibitors, such as NaF, TLCK and TPCK. Before using these reagents you should investigate the functionality of the inhibitor and discuss the potential impact on protein ID and identification of post-translational modification with the MS Facility personnel prior to using. TLCK and TPCK are also serine-protease inhibitors which will inhibit the activity of trypsin, chymotrypsin, and other enzymes.
  • EDTA – EDTA and phosphate buffers should not be used in protein solutions that will be analyzed directly for phosphate modifications with IMAC purification.
  • Derivatization Reagents –be aware that cocktails or commercial kits for protein preparations sometimes contain reagents that derivatize proteins. For instance, NEM or alkylation reagents are sometimes part of protein protocols. These reagents derivatize cysteine and other residues and can impact protein ID by altering digestion efficiency and changing the mass and ionization of peptides, and thereby affecting the identification of peptides and proteins in nanoLC/MS/MS analysis and database searches.
  • Reducing Agents –proteins preparations in solution are commonly stored with reducing agents DTT (dithiothreitol) and BME (beta-mercapto ethanol). These reducing agents degrade after a short time (i.e. within a few days, even stored cold or frozen) and will oxidize and derivatize cysteine residues, such that proteins become modified (mostly at cysteines). These unwanted derivatizations impact analysis of proteins, but can be removed prior to gel electrophoresis and/or mass spectrometry analysis of the sample, with addition of freshly-made (same day) reducing reagent immediately before analysis of the sample (e.g. 20 mM DTT for 30 min).
  • Affinity purifications –reagents used to displace proteins from affinity columns can greatly impact the mass spectrometry analysis of proteins. (Please discuss details of your affinity purification method with the MS Facility personnel).
    • For instance, high concentrations of flag-tag peptide from flag-tag affinity isolations will affect the analysis of proteins in solution, affect chromatography during nanoLC/MS/MS, and analysis of phosphorylations (with IMAC purification). The flag-tag peptide will have to be removed if proteins will be analyzed from solution. This is not an issue for proteins analyzed from a gel.
    • High concentrations of glutathione from GST affinity purifications will derivatize your proteins in a short time in solution. Cysteine residues become derivatized with glutathione (a net mass change of 305 Da per derivative), which will greatly impact the analysis of the protein. These derivatives can be removed, however, with exposure to fresh reducing reagent (i.e. DTT at 20 mM for 30 min) prior to analysis.
  • Denaturants –if denaturants are necessary for isolation of your proteins, guanidine HCl is preferred over urea. Urea can modify proteins by forming a carbamoylate derivative of lysine and other residues. This can happen from cyanates formed from urea at temperatures above 30C, from degraded or old urea, or from exposure to urea for an extend time.
Protein Purification Methods and Buffers

Protease inhibitors, detergents, PEG and other polymers, plasticizers, polymer from Sepharose beads, silica, buffers, salts, and other contaminants greatly impact the ability to analyze proteins by mass spectrometry, particularly analysis of proteins in solution.

Purification is essential, and proteins can be purified many different ways. However, there are several issues associated with methods of purification which affect the recovery and ability to successfully analyze proteins by mass spectrometry, depending how the protein will be analyzed (i.e. from a gel, or from a solution).

  • Ziptip –reverse-phase purification using C18 or C4 Ziptip (Millipore) is generally a good way to remove salts and other small-molecule contaminants prior to intact protein analysis or digestion of proteins in solution and nanoLC/MS/MS. For a final purification step of proteins in solution, it is recommended that this be performed at the MS Facility, just prior to analysis. Ziptip purification, however, is not effective in removing detergents, polymers, and other large molecules, which also display an affinity for reverse-phase material.
  • Sepharose Beads – purification with some Sepharose beads (e.g. Protein-A Sepharose) can introduce a polymer contaminant in solution that will significantly impair chromatography and the ability to analyze protein digests and identify proteins. Please discuss with the MS Facility before using Sepharose-bead slurry purifications.
  • HPLC – purification by reverse-phase HPLC is an excellent method of protein purification when possible.
  • Microcon purification – microcon size-exclusion membranes can be a good method of removing small molecule impurities, small polymers, and some detergents (when not tightly bound to the protein). The membrane should be washed several times with nanopure water before recovering proteins back off the membrane. However, some proteins, particularly large proteins, can bind to the membrane and not be recoverable. Proteins recovery can be improved by eluting with one or two 50ul aliquots of 30% to 50% acetonitrile in nanopure water with 1% formic acid. Low amounts of protein (low picomole or femtomole levels) may be lost with Microcon purification.
  • Buffers — HEPES is a nonvolatile buffer that is not compatible with mass spectrometry of proteins in solution. However, HEPES can be mostly removed with Ziptip purification prior to analysis, but using HEPES buffer is not recommended.
    • Recommended buffers: 50mM Tris, 50 mM ammonium bicarbonate, or other volatile ammonium salts.
    • Phosphate buffers cannot be used in protein solutions where identification of phosphorylated peptides will be performed using IMAC purification.
  • Protein Precipitation – generally, protein precipitations with TCA require relatively high amounts of protein for the precipitation to be successful with good recoveries. Addition of carrier molecules, such as deoxycholate, is not recommended. Deoxycholate, a detergent, acts similarly to triton in its affinity for BSA and other proteins, and trace levels of triton can have a serious impact on ionization of proteins and peptides. Therefore, it is likely that DOC will significantly inhibit or interfere to some degree with MS analysis, particularly with analysis of intact proteins.
  • Use of Detergents – even trace levels of detergents (<0.01%), such as Triton X-100 or SDS, will have a serious impact or completely inhibit ionization of intact proteins. Analysis of protein digests from solutions containing detergents will also be adversely affected, impacting the ability to identify peptides and affecting total protein sequence coverage and protein ID. The extent of impact will depend on the amount of detergent and the affinity of the detergent for the particular protein and peptides. Ziptip or reverse-phase purification will not affectively remove detergents, which bind to reverse-phase media.
    • When detergents must be used for isolation of proteins, it is best that the protein is loaded on a SDS gel and MS analyses are performed from an in-gel digest of the band or spot. Another option is to perform an acetone extraction/purification of the protein isolate to remove detergents prior to protein digestion or intact protein analysis, but this may not adequately or completely remove the detergent.
    • Mass spectrometry compatible detergents, such as ProteaseMax (Promega) can be used.
Preparation of Proteins in Solution
  • Protein solutions containing very low levels of protein (i.e. femtomole or picomole) should be prepared and concentrated with techniques that result in absolute minimum volume (i.e. uL volumes). When possible, protein samples should be prepared in concentrations greater than 10 femtomole/uL.
    Do not concentrate the protein from mL volumes using lyophilization or speed-vac, this only concentrates nonvolatile contaminants.
  • Protease inhibitors, detergents, PEG and other polymers, plasticizers, polymer from Sepharose beads, silica, buffers, salts, and other contaminants greatly impact the ability to analyze proteins in solution by mass spectrometry. (See Protein Purification Methods, above).
  • In most cases, Ziptip purification will be required of protein samples in solution prior to any type of mass spectrometry analysis. Samples that contain detergents and/or polymers may not be accepted for analysis; you may be asked to run the sample on a gel instead, for subsequent in-gel digest and MS analysis.
  • Phosphate buffers cannot be used in protein solutions where identification of phosphorylated peptides will be performed using IMAC purification.EDTA, flag-tag peptide, and other acidic compounds, should not be used in protein solutions where identification of phosphorylated peptides will be performed using IMAC purification.
    • For protein ID’s from IP pull-down experiments, the best strategy for mass spec analysis (i.e. analysis of gel bands vs. solution) will depend on the complexity of proteins and relative protein concentrations in the sample (e.g. is SCX fractionation of the protein digest required?). Before deciding to analyze the proteins from a solution or from a gel, we will need to know exactly what all the constituents are in the solution (e.g. any detergents, polymers, buffers, tags and elution reagents, etc.). Running an aliquot on a gel first is recommended to get an idea of the protein complexity.
    • Amount of Protein Required for Protein ID from a Solution Digest: The amount of protein needed in the sample will depend on the purity of the sample, MW of the protein(s), complexity of the proteins, and what information or type of analysis is being requested. From a highly-purified protein preparation, protein ID can potentially be performed from perhaps 10 to 100 femtomoles of protein digest, but is depends on the specific proteins. Identification of post-translational modifications will usually require more protein, sometimes as much as picomole quantities of purified protein.
    • The following components are generally acceptable for submission of a protein sample in solution (but may require Ziptip purification at the MS Facility). For compatibility of other buffers and reagents, talk to the MS Facility personnel.
      • Tris or ammonium buffers (<100mM
      • Glycerol (<10%)
      • DTT or BME (<50mM) – must be added fresh within 2 days
      • Protease inhibitor (may be partially or entirely removed with Ziptip or dialysis prior to analysis)
      • Guanidine HCl or urea (<1 M)
      • Salts (e.g. NaCl, KCl, ammonium salts) (<500 mM)
      • EDTA
      • Imidizole
Preparation and Submission of Proteins in Gels
  • Thin gels should be used. 0.75mm is preferred, but 0.5mm or 1mm is acceptable.
  • SDS gels should be cast at least 24 hrs prior to use. If the gel is not fully polymerized, the in-gel digest, peptide extraction, and nanoLC/MS/MS analysis will not be successful.Tris, tricine, gradient gels, 2D gels, and commercial precast gels can be used. However, SDS gels with 8 to 12% acrylamide are preferred.
  • Stains – Coomassie, Colloidal Blue, and similar blue stains are preferred and will generally result in the best peptide recoveries and protein ID results. However, gels will be accepted with the following stains:
    • Coomassie
    • Colloidal blue
    • Silver*
    • Sypro Ruby
*Special Instructions for Silver Staining

For silver staining, we recommend that you only use mass spectrometry-compatible silver stain (such as SilverQuest from Invitrogen).
Don’t over-stain the gel; stain for the least amount of time possible until you can just see your bands of interest.

For submission of gel bands with silver staining, you must contact the MS Facility a few days in advance of submitting the gel and arrange a time to submit the gel. The gel must be delivered to the MS Facility as soon as possible (within an hour is best) after staining, in order for us to successfully destain and analyze proteins from the gel. Silver-stained protein gels that have been stored for more than 1 day will not be accepted for protein ID.

Sypro Ruby-stained gels can be submitted, but these gels are difficult to work with, since the gel must be handled under UV light to see and excise bands.

After staining, gels may be stored at 4C in nanopure water or 1% acetic acid in nanopure water (except in the case of silver-stained gels, see above).
In general, chemical crosslinkers and strong oxidizing or reducing agents should be avoided.

  • Obtain an image of your gel and bring the image with you when you submit the gel for analysis.
  • It is preferred that gel bands be excised at the MS Facility. Bring your gel (and image of the gel) to the MS Facility and discuss which bands/ spots you would like analyzed.

We don’t recommend it, but if you prefer to excise your own gel bands, follow these guidelines:

First, obtain an image of your gel. Be careful not to allow the gel to contact any contaminated surface.

Avoid contamination with keratins and other proteins. Use extremely clean surfaces (e.g. glass cleaned with ethanol or isopropanol) and clean, new razor blades or scalpels (rinsed with ethanol or isopropanol first). Ideally, this should be done in a laminar-flow hood to minimize keratin contamination. Wear clean nitrile gloves (rinsed with ethanol and nanopure water).

Cut the band out as close as possible; be very selective and cut out only the center of the band; don’t include any excess gel material. The best gel bands are very narrow (<1mm); the smaller, the better. Large gel pieces will not be accepted.

Place the gel band in a clean, labeled 1.5mL microcentrifuge tube. (The microcentrifuge tube should be rinsed in advance with methanol or acetonitrile, then nanopure water). Coomassie, Colloidal Blue, or Sypro Ruby-stained bands can be stored at 4C or frozen until the band is submitted for analysis.

  • A general guideline for the amount of protein required for a protein ID is that if you can see the band (even a very faint band), the mass spectrometry analysis is likely to be successful for identifying the protein(s) in the band or spot. For identification of post-translational modifications from a gel band, picomole amounts may be necessary.
  • It is highly recommended that a control band or appropriate blank part of the gel is included with sample submission. Quite often, keratins or other proteins are present throughout the gel, although they may not appear as a distinct band.
Intact Protein Analysis
  • Detergents, polymers, and salts – purity of the intact protein sample is critical for a successful analysis of the molecular weight of a protein. It is critical that the sample does not contain any detergents or polymers (even at trace levels). Avoid using Triton, Tween, SDS, or PEG during any part of the protein preparation. It will be very difficult or impossible to remove them. Avoid using glassware that has been exposed to detergents or polymers, since this will introduce trace contamination into your protein sample. Electrospray ionization of intact proteins will not be successful with even trace contaminants present, and they cannot be removed with Ziptip purification.
    • The protein sample must be free of salt and other small-molecule contaminants. These can usually be adequately removed with reverse-phase purification or Ziptip. Ziptip purification is best performed at the MS Facility, just prior to analysis of the protein.
  • Amount of protein required – the amount of protein required will depend on the purity of the sample, relative hydrophobicity of the proteins, complexity or number of proteins, and molecular weight of the proteins. Generally, a minimum concentration of picomoles/ul is required. More is better, if possible, but purity is far more important than the amount of protein. Analysis of complex protein mixtures is likely not to show all the proteins present in the mixture, as a result of ion suppression effects during electrospray ionization of proteins with different hydrophobicities and relative concentrations in the mixture (protein samples are usually analyzed by ESI/MS with sample introduction by infusion).
    • If the protein sample is extremely pure and in nanopure water, the analysis of relatively small, hydrophilic proteins (e.g. <30k Da) can be successfully done with concentrations as low as 10 femtomoles/ul, but this is not typical.
  • Size of proteins that can be analyzed – proteins as large as 150k Da can be successfully analyzed if the protein is homogeneous (i.e. the molecular species is a single molecule, not a population of molecules with small molecular weight differences, such as from variants in amino-acid sequence or substitution, methylation, formylation, or sulfur oxidation). The protein purity must be very high.
  • Acceptable components in solution for sample submission – ideally, submit the purified protein in a solution of nanopure water only, or nanopure water and acetonitrile (1% formic acid can be added). HPLC-purified protein is ideal, when possible. However, the following components are acceptable for sample submission for intact protein analysis from a solution (Ziptip purification at the MS Facility will be required prior to analysis):
    • tris or ammonium buffers (<100mM)
    • glycerol (<10%)
    • DTT or BME (<50mM) – must be added fresh within 2 days
    • guanidine HCl or urea (<500mM)
    • salts (e.g. NaCl, KCl, ammonium salts) (<200mM)
    • EDTA (<50mM)
    • Imidizole (<50mM)
  • Reducing agents – if the protein has been exposed to DTT, BME, or other reducing agents for longer than 1 or 2 days, it is likely that cysteine residues in the protein have been derivatized from oxidized or degraded forms of the reducing agent. Derivatives from DTT or BME are recognized by complex mass additions of 32 Da or 76 Da in the molecular mass spectra. These modifications can be removed by addition of freshly-made 20mM DTT for 30 min prior to analysis (disulfide linkages will also be reduced).
Guidelines for Identification of Phosphate Modifications
  • Proteins being investigated for phosphorylations should be analyzed as soon as possible after isolation and purification, to minimize possible loss of labile phosphate modifications. If the protein must be stored, -80C is recommended.
  • Selective purification of phosphopeptides can be performed at the MS Facility using IMAC (immobilized metal affinity chromatography) or TiO2 columns. Roughly 5 or more pmoles of protein is required for IMAC purification.
  • TiO2 is a new technique that is perhaps 10 to 100 times more sensitive for selective purification of phosphopeptides, compared to IMAC. (The TiO2 purification technique will be available soon at the MS Facility).
  • Phosphopeptides and the exact site of modification can be identified from protein digests (i.e. in-gel or solution), nanoLC/MS/MS, and database searches (specifically for phosphate modification).
  • Phosphorylations can be identified from analysis of intact proteins. Although the site of modification will not characterized from an intact protein analysis, the relative extent and number of phosphorylations can be determined.
  • Phosphate buffers cannot be used in protein solutions where identification of phosphorylated peptides will be performed using IMAC purification.
  • EDTA, flag-tag peptide, and other acidic compounds, should not be used in protein solutions where identification of phosphorylated peptides will be performed using IMAC purification. Unbound flag tag peptide in solution must be removed before IMAC purification.
  • For identification of phosphorylations and other post-translational modifications, it is recommended that the protein be analyzed from solution (if the solution is suitable), since in principle, all peptides products from digestion of the protein will be injected during nanoLC/MS/MS analysis.
  • Protein digests with chymotrypsin may result in loss of phosphorylation (there may be evidence of some phosphatase activity in chymotrypsin).
Enzyme Digestion of Protein Samples

In-gel Protein Digest**

  • 1D gel bands or 2D gel spots are preferably excised and destained by personnel at the MS Facility.
    In-gel digests can be performed with trypsin or chymotrypsin enzymes only.
  • The following gel stains are acceptable. However, coomassie, Colloidal Blue (Invitrogen) and other blue-type stains are preferred for protein ID from in-gel analysis.
  • (See Sample Submission and Guidelines and Protocols for recommendations prior to preparing, running, and staining gels).
    • Coomassie
    • Colloidal Blue
    • Silver (i.e. must be mass spec compatible, see special submission requirements described above)
    • Sypro-ruby

**We do not offer gel electrophoresis or gel staining services; the user must provide the gel or gel band/spot to the MS Facility.

Protein Solution Digests

Digests of proteins in solution can be performed with the following enzymes. However, trypsin and chymotrypsin are preferred and will be the most specific and informative in most cases.

  • Trypsin
  • Chymotrypsin
  • Thermolysin
  • Pepsin
  • For other proteases (e.g. Asp-N, Glu-C, O-glycosylase, N-glycosylase) or other reagents, such as CNBr, please inquire.
Identification of Protein Post-translational Modifications and Derivatives

Identification and mapping of post-translational modifications can be difficult, particularly when the modification is at relatively very low levels. Often, several mass spectrometry experiments may be required and may involve different sample preparations and mass spectrometry strategies.

For example, if the type and extent of modifications are unknown, a combination of analysis strategies are needed. When feasible, ESI/MS analysis of the intact protein can be highly informative in determining whether the protein has any modifications, what those modifications may be (such as phosphorylation), at what approximate levels, and whether the protein amino-acid sequence is different than anticipated, truncated, or degraded from protease activity.

Analysis of protein digests is used to identify and map sites of modification. When searching for possible modifications, it is recommended analyzing the protein using two or more different enzyme-digest experiments, with subsequent nanoLC/MS/MS analyses and/or Maldi/MS/MS analyses, and multiple database searches for possible modifications. Accurate mass measurements with FTMS (LTQ-FT instrument), along with MS/MS sequence, provide a high-degree of confidence in the assignment and identification of modified peptides.

Phosphorylations can be targeted using IMAC (immobilized metal affinity chromatography) or TiO2 columns to selectively isolate and purify phosphopeptides from the digest. These procedures can sometimes greatly improve the ability to find phosphorylations.

If a specific site or region of the protein is anticipated to be modified (such as phosphorylation), a custom method for the mass spectrometry analysis can be setup (i.e. create an inclusion list of ions) to specifically target one or more putative modified peptides of interest, thereby increasing the capability of the instrument to identify those peptides or modified peptides.

Proteins with labile modifications (such as GlcNAc or phosphorylations) can be identified using ECD (electron-capture dissociation) fragmentation of peptides for MS/MS sequencing. The ECD technique (in the FTMS instrument) generally leaves labile modifications intact during fragmentation of peptides, such that the position of the modification can be clearly determined. Other lower-energy fragmentation methods, such as CID (collision-induced dissociation) or IRMPD (infrared multiphoton dissociation) can result in dissociation of labile modifications, but often the modification and site of modification can still be identified by virtue of mass-loss differences observed in the MS/MS sequence spectra. Modifications can potentially be located by “top-down” analysis of intact proteins using ECD fragmentation of the intact protein in the FTMS.

Database searches using MascotTM software and other search engines are used to process the MS/MS spectra of peptides to locate the sites of modifications.

Many types of protein modifications can potentially be characterized, such as:

  • Phosphorylation
  • Sulfation
  • Oxidation
  • Acetylation
  • Methylation
  • Ubiquitination
  • Sumoylation
  • GlcNAc and other glycosylations
  • Disulfide linkages
  • NEM and other derivatives
  • AEBSF modification from exposure to protease inhibitors