Mass Spectrometery Sample Preparation Tips
By far the most common contaminant in proteomics is keratin (unless you happen to be working with skin cells). This protein comes from skin, hair, and dust. It's everywhere! It's on everything! It will be in your sample, but we can minimize it! Trypsinized Keratin generates peptides of the ideal size. The peptides ionized readily and generate beautiful spectra. The cleaner your sample, the more likely it is that your protein of interest (along with any PTMs that you may be interested in!) will be detected. So, always wear gloves! Never touch your tubes with bare hands. Soap and water are your friends. Always work on freshly cleaned bench surfaces Wash everything that will encounter your protein(s): e.g., gel boxes, glass plates, Coomassie/silver staining containers and those fresh gloves that you put on. Use a fresh box of pipette tips, a fresh bag/box of Eppendorf tubes, fresh blades, etc. Consider a small separate stock of "keratin-free" chemicals – you never know how many lab mates have had their bare hands stuck in that bottle! Wear a clean lab coat, particularly when wearing any woolen clothing. Keratin-laden dust particles adhere to Eppendorf tube racks and other plastic holders, particularly during dry winter months. We strongly advise daily rinsing of dust out of actively used Eppendorf tube racks.
Mass Spec-incompatible compounds
Some chemical compounds can also cause problems during MS detection.
- PEG and detergents: design your isolation procedure so that you can get rid of SDS, Triton X-100, NP-40, Tween, CHAPS, etc., before MS. These compound and related polymers are observed very well by MS and will likely obliterate the signal for your protein.
- DMSO, DMF, glycerol: to be avoided. These compounds are highly viscous, can be observed in spectra, and/or can interact with tubing and other machine parts.
- Acids/plastic polymers: while acetic acid is often used in MS protocols, it is imperative that you don't contaminate your stock glacial acetic acid bottle with plastic. Don't put plastic pipettes or pipette tips into the stock bottle; every time you do this, a little bit of plastic is released into the solution. Plastic polymers can thus easily build up to a point where we will begin to see them in your sample. Instead, measure out what you need with a clean glass graduated cylinder. You can also pour from the stock bottle into a separate container, and pipette from this – but dump this out after removing what you need.
Avoid salts and non-volatile buffers in your final purification steps, if possible. In most cases, we will simply dry your sample down in our speed-vac and re-suspend it in 50mM ammonium bicarbonate pH 8.3 for an overnight tryptic digestion. If your sample contains non-volatile salts or buffers, they will still be there. Besides interfering with the protease, these compounds can also suppress ionization of your peptides, affect chromatography, and form adducts that interfere with peptide detection. Consider using the following volatile buffers in your final purification step(s).
|Triethylammonium phosphate||pH 3|
|Ammonium formate||pH 3-5|
|Pyridinium formate||pH 3-6|
|Ammonium acetate (-pH with acetic acid)||pH 4-6|
|Pyridinium acetate||pH 4-6|
|Trimethylammonium formate||pH 5-6|
|Triethylammonium acetate||pH 6-7|
|Ammonium bicarbonate||pH 8|
|Ammonium carbonate||pH 8-10|
Some acids and bases are also volatile. These are great, e.g. for protein elution from a solid support/column.
|Volatile Acids||Volatile Bases|
|Carbonic acid||Ammonium hydroxide|
The following are the key publications that we draw upon for gel and solution based peptide preparations, by the some of the originators of the techniques:
- In-gel digestion procedure for preparing peptides for mass spectrometry from SDS-PAGE gel bands
- FASP methodology for preparing peptides from biological samples
The following schematic depicts the process of SDS-PAGE gel band excision and dicing:
While we are glad to conduct gel band excising, those groups comfortable with this procedure may accelerate analytical times by one or more days by taking on this task in their own lab.
Sample Submission Guidelines
For submission of samples in solution for intact molecular weight determination please make sure that the solutions are free of detergents (e.g., Triton X, NP-40, TWEEN, SDS, etc.), PEG or other polymers, and solvents such as DMSO, DMF, or NMP. Even a small amount of these substances interferes with intact protein mass spectrometry and may be very difficult for us to remove. You may need to perform an additional final dialysis or buffer exchange step prior to submission to transfer your proteins from their storage solutions into a solution compatible with mass spectrometry. Low percentages of acetonitrile or methanol may be added to solutions if that assists in solubility. If a mild detergent appears to be necessary to maintain your protein in solution, you may consider using an acid-labile detergent designed for use with mass spectrometry, such as Rapigest SF (Waters) or ProteaseMax (Promega). These can be left intact until the final moments prior to analysis, then decomposed into inert species.
For membrane protein or hydrophobic protein-containing samples, the S-trap protocol is used. See https://protifi.com/products/s-trap-micro-kit. A sample amount of 1-100ug can be submitted for S-trap micro. We can provide 1M TEAB pH 8.5 buffer, if needed. Please contact Henry Shwe [email protected] for details.
We accept both commercial precast SDS-PAGE gels and homemade gels. The acrylamide percentage of the gels should be kept ≤12% if possible, to facilitate diffusion of endoproteinases into the gel matrix for in-gel digestion. For most applications, we prefer 4-12% gradient gels or 10% fixed concentration gels, in either the Bis-Tris (BioRad or Life Technologies) or TGX (BioRad) formulations. Be sure that the gel buffers and other reagents are paired appropriately with the formulation type. If handled properly, commercial gels will provide much lower levels of keratin peptides in the final analysis.
We suggest using a fresh colloidal Coomassie stain, such as GelCode Blue (ThermoFisher/Pierce), which is safe and highly compatible with mass spectrometry sample prep methodologies. Maximum staining is achieved by thorough washing (4-5 X) in distilled water and soaking twice, each for several hours, in fresh Coomassie stain. Alternatively, non-fixative fluorescent or silver stains are also acceptable. If you would like protocols/recipes, please get in touch with us.
Excise the band of interest including the least amount of extraneous gel material possible. Gel slices should be submitted to us in labeled Eppendorf tubes. You may accelerate our processing by supplying the slices cut into 1 mm cubes. Please see our excision schematic (also referenced in the Protocols section above).