Do I need to schedule the drop-off of my samples?
How long do the analyses take?
How much does it cost?
How much starting material do I need to submit for protein identification?
Unfortunately, there is not a clear rule as it depends on the sample, and on the question that you want to address. As a rule of the thumb, we prefer to work on gel bands visible on Coomassie-stained gel but we can still identify proteins coming from silver staining. For in-solution digestion, we usually request 10-20 ug total protein (if possible). Please note that for any type of purification (ex. Phospho-enrichment) significantly higher amounts are required.
In which buffer can I submit my sample?
For samples submitted in solution, we prefer MS compatible buffers like ammonium bicarbonate or Tris buffers (50-100 mM). Detergents, organic solvents and salts can strongly impair the MS measurements. It is necessary to provide us with a detailed description of the solution composition of your samples upon submission (Organic solvent? Detergents? Buffer? pH?…). This step is critical because we have different protocols for the removal of some incompatible substances (Desalting, FASP etc). For samples requiring special pH, detergents or salts to overcome solubility issues it is often possible to replace them by MS compatible formulations. Classical detergents (SDS, NP40,…) can be replaced by acid labile (Waters RapiGest…) or MS tolerant detergents like n-octylglucopyranoside or deoxycholate. Do not hesitate to contact us if you are not sure.
What are the most common reasons for obtaining poor mass spectrometric data?
- Use of a staining method that is not compatible for mass spectrometry based-proteomics.
- The presence of ion suppressing reagents, such as polymers, detergents, and glycerol in the sample.
- Overloading with contaminating proteins, such as antibodies in affinity purifications
- Protein specific properties; e.g. heavily modified proteins, very few lysine and arginine residues in a particular protein.
Which are the common contaminants for mass spectrometry-based proteomics?
- Polyethyleneglycol (PEG) and detergents containing PEG (common pegylated detergents include; Triton X-100, Tween, NP-40…). If detergents are present in your sample they will dramatically reduce the quality of your data.
- Keratin is a common protein contaminant and it comes from dust, skin, hair, clothes, etc. Wear gloves all the time, use clean glassware and tubes and try to work in clean environment.
What are some important considerations for a gel-based identification?
- Leave blank lanes in between samples to avoid any spill over effect.
- Use MS-compatible gel staining.
- Wash the gels extensively after the end of the run to remove as much remaining SDS as possible.
- Store the gels in clean boxes (pre-rinsed with organic containing solution) to minimize keratin contaminations.
Which gel staining protocol should I use?
Coomassie staining (R250 or colloidal G250) is the preferred one. You can also use silver staining if required but avoid any fixing step employing glutaraldehyde or paraformaldehyde. Ask us if you need a protocol.
Please remember that gels need to be washed extensively after the end of the run to remove as much remaining SDS as possible. Also, they need to be stored in clean boxes (pre-rinsed with organic containing solution) to minimize keratin contaminations. Finally, blank lanes in between samples are important to avoid any spill over effect.
Should I cut the gel band myself?
We prefer to receive the intact gel and excise the bands ourselves. In case you would like to bring already excised bands please follow the guidelines below:
- Always wear gloves.
- Only cut gels on a clean glass plate using a clean scalpel.
- Precisely cut around the stained protein band
- Transfer protein band in separate 0.5mL eppendorftubes
Gel bands can be kept at -20°C before MS analysis (except silver-stained bands which should be freshly processed).
Can I submit a silver-stained gel?
Yes, but it has to be stained with a mass spectrometry compatible silver staining gel protocol (without any protein cross-linker such as Glutaraldehyde). Many commercial vendors like Life technologies, Sigma, Pierce biotechnology (now Thermofisher) sell mass-spec silver stain kits.
Do I need a protein database for the organism of interest?
Yes! The preferred method for mass spectrometry-based protein identification is database searching during which experimentally generated spectra of mass to charge ratios are compared to in-silico generated ones. Therefore, the protein database used for the search is one of the major influencing factors in discovering proteins present in the sample; if a protein is not in the database it will not be identified no matter how abundant it is.
Can you identify phosphorylation sites?
Yes! However several factors should be taken into consideration.
- First, phosphorylation is often substoichiometric, meaning that phosphopeptides can be very low-abundant, and they may be masked from their non-phosphorylated counterparts.
- Second, ionization of phosphopeptides is less efficient than for non-modified peptides, which also makes detection challenging.
- Third, fragmentation of phospho-peptides does not follow the same rules that apply to non-modified peptides making spectra hard to interpret.
- Finally, the phosphorylation site may be present in a peptide that is tricky to be detected due to its amino-acid composition.
If you are working with a specific protein and you have an idea regarding the expected modification site, in-silico digestion with multiple enzymes can pinpoint which enzyme would generate the most optimal peptide.
In case of complex samples, enrichment of phosphopeptides by techniques such as IMAC or TiO2 is strongly recommended.
You could not identify a protein in my gel band. Why?
There may be several reasons why this can happen. We take care to ensure that this is not due to technical issues from our side (ex. instrument performance). Other common reasons include:
- The starting material was not enough.
- The protein is not present in the database.
- The amino acid composition and other properties of the protein (ex size, hydrophobicity) hinder the MS-based identification
- The protein under investigation does not contain a sufficient number of cleavage sites for trypsin (thus no peptides will be generated) or it contains too many cleavage sites (thus the generated peptides are too small to be detected). In-silico digestion with multiple enzymes can pinpoint which enzyme would generate the optimal peptides.
- The staining procedure followed is not MS-compatible. Avoid any fixation agents or microwaving your gel.
I expected 1 protein in my gel band but you gave me a list of 50. How is this possible?
Even if you excised a single band from a gel, it is very likely that it contains several proteins of very similar size, some of which may be below the detection level of the staining method used. Please note that mass spectrometers exceed by far the sensitivity of coomassie and silver staining. Try to avoid overloading the gel as it will affect the resolution and separation of proteins. Also, consider leaving a blank lane in between samples to avoid protein diffusion and cross contamination.
I see a lot of keratins in my list of proteins. Where do they come from?
Most likely they were introduced during sample preparation. Keratins are the most common contaminants and come from dust, skin, hair, clothes, etc. Wear gloves all the time, use clean glassware and tubes and try to work in clean environment.
I have never done an LC-MS/MS experiment. Is previous knowledge required?
We provide expert advice and support for your project, from experimental design to data interpretation. In addition, we offer trainings in case you are interested in learning more about proteomics.
Which enzymes you frequently use for protein digestion?
Most common and efficient proteolytic enzyme used is Trypsin (MS Grade) but extra ones are also used at the Facility such as Lys C, Chymotrypsin, Glu C. Optional enzymes are also available upon specific request such as Asp N or Arg C.