We provide a full proteomic infrastructure for the identification and characterization of proteins that include various platforms for protein and peptide separation, and state-of-the-art mass spectrometry for MS and LC-MS/MS experiments.
This is a list of our most frequently asked questions:
This is the most frequent question we get, and the hardest one to answer. Theoretically MS can identify at zeptomoles level, but correct answer is that this depends on the sample, and on the question that you want to address. If you want to identify a single protein from gel, an amount that produces a colloidal coomassie-stainable band will almost always give you a protein identification this should be in the range of 10-20 ng. In general, always send us the maximum amount you can get. We process also silver stained faint band if this is the maximum amount you can get, but the results are not ensured. For identification of proteins in complex mixtures (e.g. separated over a gel lane), the amount of protein should be upwards from 10 μg.
To determine the weight of intact proteins, you should send at least 100 μl at a minimum concentration of 10pmo/μl. Usually this is sufficient to get a molecular weight determined under denaturing conditions.
We are better able to obtain an unambiguous protein identification if we know the species of origin and any relevant details about how the sample was prepared (any known or suspected post-translational modifications, was the sample from a 2D gel, an SDS-PAGE gel, was the sample artificially modified by alkylation etc.).
Gels can be sent well-sealed in plastic foils, gel bands in closed eppendorf tubes. There is no need to ship them on ice. For MW determination of intact proteins, the buffer composition is not highly criticial, as long as it does not contain detergents or more than 5% glycerol.
Paolo Soffientini and Angela Cattaneo
Via Adamello 16
20139 Milan, Italy
We do not sequence proteins, we identify them. Protein identification by mass spectrometry is probabilistic, meaning that the best match is sought between an experimental spectrum and a theoretical spectrum for a peptide in the database. The score assigned to this match, and therefore the probability for the match to be right, depends on a number of parameters, such as spectral quality, mass accuracy, the size of the database, and the algorithm used for database searching. In addition, the more peptides are assigned to a given protein, the higher the protein score will be. A protein is considered identified if the software can correctly assign 2 peptides to this protein.
We identify proteins by matching spectra to protein sequences in a database. Thus, in principle, if a protein is not in the database it will not be identified. But there are exception, Please contact us for further information about this.
We are set up to do this, but detection of phosphopeptides and site-localisation of the modification is never straightforward. First, phosphorylation often is substoichiometric, meaning that phosphopeptides can be very low-abundant, and that they may even go unnoticed among non-phosphorylated peptides from the same or other proteins. Second, ionisation of phosphopeptides is less efficient than for 'normal' peptides, which also does not help to detect them. Third, fragmentation of phosphopeptides does not obey the same rules that apply to normal peptides, which sometimes makes spectra hard to interpret. Finally, the phosphorylation site may be present in a peptide that is too large or too small to be detected and fragmented efficiently. If you know your protein, and the expected modification site, the choice of a different protease may be a good alternative for mapping the desired domain. We can enrich for phophopeptides by TiO2 techniques, especially for complex samples, but all the problem pointed before remain.
We can identify some PTMs (e.g. acetylation, methylation, ubiquitination, glycosylation sites and others). It will be helpful if you tell us which one(s) you expect to occur so we can include it in our search and then look for this specifically. Please contact us for further information about this.
If you excised a single band from a gel, it is not unlikely that this contains several proteins of (almost) the same size, some of which may be below the detection level of the staining method used. Mass spectrometers by far exceed the sensitivity of coomassie and silver staining.
Another reason of multiple proteins in a band is that proteins, expecially the most abundant ones, can smear all over the lane be degraded, so it is common that you find proteins of different molecular weight in your bands.
Most likely they were introduced during sample preparation, or (in case you used gels) during staining or cutting. Dust in the lab is the most likely source, so make sure you work in a clean area. We can provide you a list of tricks to avoid contaminations.
For most users, an Excel-list of identified proteins is usually sufficient, with an accession number to an appropriate database (e.g. Uniprot). However, we can provide much more than that, including sequences of identified peptides, peptide scores, sequence coverage, position of the peptide in the protein, annotated spectra for all peptides, etc. The latter may be required by some journals if you report identification of PTMs. Everything is however present in the Scaffold file that we will send you for results. In addition, we provide details on protein and peptide quantification, either using label-free or stable isotope-labeling approaches.
For protein identification and PTM analysis, you will be provided with a Scaffold file that resume the majority of the information. We are happy to help you in learn how to use it. For protein quantitation via MaxQuant (SILAC and Label Free) you will receive an excel file with significant up/down regulated proteins, all proteins quantified and a volcano-plot or Christmas three representation of your results. For Targeted proteomics you will receive an excel file with peptides/protein quantification. You will receive a detailed email with results of the analysis and a powerpoint to graphically explain how we process your samples. Please contact us for further information about this.
We aim to return data within 1 week of sample receipt for protein identification and in maximum 3 weeks for protein quantification and PTM analysis.
Please contact us for costs, as these will vary from project to projects depending on instrumentation used and analysis requirements.
This can happen only in very specific circumstances, sorry. In order to provide a high quality facility, it is essential that we limit access to the instrumentation to a handful of expert users.