Proteomics: Frequently Asked Questions

FAQs

Methodologies

ESI, and LC-MS mass spectrometry

Sample Submission

How much sample should I submit?

For SDS PAGE gel bands, we recommend providing enough sample that your band of interest can be seen by Coomassie stain.

For immunoprecipitations/affinity tag pull-downs, your chances of LC-MS identification are better if you are able to run an elution on a gel and see a band corresponding to your protein. If you can only see your protein by western blot, LC-MS may or may not be sensitive enough.

For purified proteins, a few micrograms is typically enough. For more complex liquid samples like cell lysate, we recommend > 20 ug. For solid samples, consult us ahead of time.

Can I submit protein/peptide samples that I have already digested?

Yes, you can pre-digest your samples. If your digestion buffer is not MS compatible, it will need to be cleaned up by either you or us. If you digest your own samples, we are not responsible for the digestion efficiency.

Can I submit beads from an immunoprecipitation for protein identification?

Yes, we accept immunoprecipitation beads as well as IP elutions. If you used any detergents in the binding and/or wash steps, you must tell us what you used and what the concentration was. We reserve the right to wash your beads several times after receiving them to remove residual detergent.

What buffer should I use?

Common buffers like HEPES, PBS, Tris, and ammonium bicarbonate are all acceptable for shotgun analysis. Water and organic solvents like methanol and acetonitrile are also acceptable. For native intact protein analysis, use a volatile buffer like ammonium acetate, ammonium bicarbonate, or ammonium formate.

Are there any detergents I can use during sample preparation? Can the facility remove detergents for me?

Nearly all detergents for biological applications are incompatible with mass spectrometry. This includes Triton, Tween, NP-40, and Brij. For a charge, we can remove some common detergents at low concentrations from samples. However, we may refuse samples with certain detergents altogether.

If you’re looking for a detergent that is MS-compatible, you can check out Waters RapiGest or sodium deoxycholate (SDC), which can be acid precipitated and centrifuged out of samples. These are just suggestions—we can’t guarantee that these will work for your application.

Is RIPA buffer or Laemmli SDS buffer okay?

Laemmli buffer is never acceptable. If you use RIPA buffer, cleanup steps are required and will be billed accordingly.

Do you perform RIME and BioID analyses?

Yes, we can analyze RIME and BioID samples.

Can you do crosslink analysis?

Yes, depending on the crosslink, its specificity, and the complexity of your system. In general, we don’t support untargeted/nonspecific crosslinks on large systems.

Do you do isobaric labeling for quantitation?

Yes, we can use either iTRAQ or TMT isobaric labels for relative quantitation. Consult us beforehand to discuss sample requirements, reagent costs, etc.

What information do I need when submitting samples?

You will need to provide a CFOP or other payment information along with your email, phone number, and PI.

Be prepared to tell us about your samples: protein amount, buffer, etc. For shotgun proteomic analysis, specify which organism(s) to use in the data search. If you need us to analyze a custom database or post-translational modification, provide that information by email. If you need quantitation or analysis through a platform other than Mascot, tell us that as well.

For a better idea, see the Sample Submission Form.

How do I submit a custom database for searching? Is there an extra cost?

Let us know when you submit your samples that you’d like for us to search a custom database. If it’s a particular organism strain, give us the specific taxon identifier. If it’s a unique sequence (e.g. a protein with a set of point mutations or a tag), email us your database in FASTA format. There is an extra cost when you submit a custom database for the first time. See our price list for more details.

How do I submit a custom PTM or chemical derivative for searching? Is there an extra cost?

Let us know when you submit your samples that you are looking for a special modification. If it is not in our database, there will be a charge to add it the first time. Please email us the expected mass, the exact chemical formula, and which amino acid(s) may be modified.

If your modification is in Unimod, there is a good chance it’s already in our database. Provide the Unimod accession number to be sure: https://www.unimod.org/modifications_list.php

Do you take samples from outside the University of Illinois?

Yes, we take outside samples. Please fill out an off-campus user form and arrange payment information ahead of time. We can only accept packages Monday through Friday, and we recommend that you let us know when you ship your samples. Please use the address below.

Proteomics Core Facility
Carver Biotechnology Center
315 Noyes Laboratory, MC 712
600 S. Mathews Ave
Urbana, IL 61801

Data Analysis

How will I get my protein identifications?

Protein identification by LC-MS includes one Mascot analysis. After the analysis is complete, we will send you a unique link with log in information to access your data. Your data is kept on our in-house server and is available for one year after project completion.

Additional data processing will be charged according to an hourly rate.

Can I get the raw data?

Yes, if you want to analyze your raw data, we can upload it to University of Illinois Box.

What types of software do you use?

The default analysis included with our LC-MS protein identification service is Mascot. We can also analyze data by ProteomeDiscoverer and MaxQuant.

Can you do protein quantitation?

Basic protein quantitation based on spectral counting is included by default with Mascot analysis. Each protein is assigned an emPAI score; larger scores indicate higher abundance while smaller scores indicate lower abundance. This type of quantitation is excellent for determining the relative amounts of proteins within a sample.

For comparisons of relative protein abundances across many samples, we recommend either extracted ion chromatogram (XIC) label-free quantitation or isotopic label-based quantitation. XIC label-free quantitation can be done in Mascot and MaxQuant. Isotopic label-based quantitation (e.g. iTRAQ or TMT) is typically done in MaxQuant or ProteomeDiscoverer.

If you need additional quantitation beyond the Mascot default, tell us when you submit your samples. For iTRAQ and TMT quantitation, consult with us ahead of time.

What do you mean by spectral counting, emPAI scores, and extracted ion chromatograms?

Spectral counting is a label-free quantitation strategy that simply adds up the number of spectra for each identified peptide; those peptide numbers are then combined according to their corresponding proteins.

An extension of this strategy is the emPAI (exponentially modified Protein Abundance Index) score, which is calculated using the number of observed peptides as well as the number of observable peptides. The emPAI score is similar to spectral counting in concept, but it is a more accurate representation of relative abundances because it takes into account protein size and sample complexity. See the original paper for more detail about emPAI: https://doi.org/10.1074/mcp.M500061-MCP200

Extracted ion chromatograms (XIC) are generated by recording the intensity of the signal from a particular m/z value (i.e. a peptide) as a function of retention time. The area under the resulting curve can then be integrated and correlated to peptide abundance.

Should I pursue label-free quantitation or isotopic label-based quantitation?

There is no one-size-fits-all answer to this question. In general, label-free quantitation can be used for any number of samples and sometimes permits deeper coverage of the proteome. However, it is more dependent upon instrument conditions, so all samples for quantitation analysis should be run sequentially.

Isotopic label-based quantitation (including SILAC or reporter based techniques like iTRAQ or TMT) requires much less instrument time and makes it easy to directly compare samples since they are run at the same time. However, study design is limited by the number of available isotope labels.

Neither strategy is better per se, so you should carefully consider which one will work best for your particular experiment.