Proteomics strategies

We apply diverse strategies to answer the needs of your research projects. If you cannot find the appropriate strategy for your specific research project, contact us to discuss your requirements. 

Quantitative proteomics

The field of proteomics has shifted from the use of just descriptive strategies, in which catalogues of proteins and PTM’s were generated, to the development and use of quantitative approaches where temporal aspects of protein function can be assayed.

The inherent dynamics of proteins, protein modifications and protein complexes require sensitive and comprehensive quantitative methods for their study.

Large-scale protein identification and quantification can be performed using a number of different strategies. Firstly, in order to simplify highly complex samples such as whole cell extracts, an extra dimension of separation must be performed prior to LC-MS/MS analysis. Proteins can be separated by gel electrophoresis and the entire lane excised to generate fractions of increasing protein molecular weight.

In gel digestion is then performed and extracted peptides are analysed by LC-MS/MS. Complex samples can also be digested into peptides and fractionated by ion-exchange chromatography or peptide isoelectric focussing (OFFGEL) and then each fraction analysed by LC-MS/MS.

In all of these approaches the second dimension of peptide separation is online reversed phase chromatography; long analytical columns (50 cm) are typically used to increase peak capacity and depth of analysis.

Proteins can be quantified using a number of methods but it is recommended that samples are SILAC labelled if possible as this increases accuracy of quantification and has the added benefit of reducing instrument time as samples are pooled upstream in the sample preparation workflow. Ultimately, the choice of quantification method is often dependent on the type of biological system being studied but we use a number of different approaches to broaden the utility of proteome profiling.

We generally advise that 4-5 biological replicates are analysed for proteome profiling experiments to maximise coverage and allow meaningful statistics to be used, however in cases where is this not possible a minimum of 3 biological replicates can be used.

Stable isotopic labelling (SILAC, TMT)

Stable isotope labelling by amino acids in cell culture (SILAC) is an approach developed by the Mann lab that uses isotopically labelled (13C6, 15N2 etc.) amino acids (usually lysine (K) and arginine (R) as they are always present in tryptic peptides) that are substituted for their naturally occurring forms in cell culture media.

Cells can be labelled to near completion with these amino acids, allowing pooling of differentially labelled samples (whole cell lysates, organelle proteomes, protein complexes etc.) for analysis by mass spectrometry. Differentially labelled peptides co-elute during liquid chromatography, allowing quantification of light and heavy peptide pairs by their relative intensity as they elute in time in an LC-MS/MS experiment.

The main advantage of SILAC quantification is that samples can be pooled very early in proteomic workflows, thereby reducing experimental variation as well as reducing MS acquisition time. In duplex experiments, control cells can be unlabelled and experimental cells can be labelled with heavy K/R amino acids so that after drug treatment or RNAi for example, both cell populations can be pooled and processed for MS analysis. Tri-plex experiments can also be performed by labelling cells with medium and heavy amino acids thereby expanding the number of samples that can be pooled and analysed together. We use MaxQuant and Proteome Discoverer to process SILAC labelled data and we can achieve highly sensitive protein identification and quantification. Quantification is performed by integrating the area under the curve for precursors (intact peptides) as they elute chromatographically.

In cases where SILAC labelling is not possible (e.g. human tissue, cells that cannot be cultured etc.) or cost effective (higher model organisms), alternative approaches such as chemical labelling are used. Peptides from digested proteins (solution digest/in gel digestion) can be differentially labelled using a combination of light, medium and heavy forms of formaldehyde and cyanoborohydride (Heck lab). Labelled peptides are pooled and analysed by LC-MS/MS analysis in which a 4 Da mass difference is observed between differentially labelled peptides. Quantitative information is collected at the MS level at high resolution allowing accurate and sensitive quantification.

Quantification can also be performed by labelling peptide samples with tandem mass tag (TMT) reagents which barcode samples and permit reporter ion quantification. The reagents add the same overall mass to peptides from each of the samples to be quantified; labelled peptides from pooled samples elute at the same retention time and will have the same precursor mass in MS1 spectra but upon MS2 fragmentation the barcode becomes decoded and quantification is performed on signature low mass fragment ions.

Label free

Peptide/protein intensities obtained from software such as MaxQuant can be used for label free quantitative analyses although the accuracy of the quantification is not as good as SILAC quantification. The match between runs feature of MaxQuant helps to transfer identifications across replicate experiments reducing the missing value problem. Overall, this approach is best suited for quantification of enrichments in biochemical purifications in which fold changes to be quantified are reasonably large.

Absolute protein quantification

Selected reaction monitoring (SRM)

The gold standard of MS-based absolute protein quantification is the use of spiked-in isotopically labelled peptides in conjunction with selected reaction monitoring (SRM), also called multiple reaction monitoring (MRM) on triple quadrupole instruments.

The unrivalled specificity and sensitivity of this approach is best utilised when a small number of proteins are being assayed in a MS-based “western blotting” approach. This is ideally suited to quantifying a limited set of proteins in a medium to large number of samples.

Post-translational modification analysis

Phosphorylation

Phosphoproteomics, the large scale analysis of the phosphorylated portion of the proteome is maturing as a technology and routinely allows the identification of in excess of 10,000 phosphorylation sites from mammalian cell lines in less than 24 hrs acquisition time.

The predominant strategy for phosphoproteomics involves sample digestion into peptides, offline desalting of peptides, fractionation (usually SCX), phosphopeptide enrichment (IMAC/TiO2) and LC-MS/MS analysis using long analytical columns and long analytical gradients. In proteome profiling experiments, the ultimate goal is to quantify proteins and it is not necessary to identify and quantify the same set of peptides in all replicate experiments.

However, in phosphoproteomics a phosphorylation site must be identified and quantified in at least 3 replicates in order to get meaningful data. T

he selection of peptides for sequencing can be a stochastic process, especially in highly complex samples and as a result we recommend that 5 biological samples are analysed in order to increase the chances of phosphopeptides being sequenced, identified and quantified in enough replicates for confident differential analysis.

Other post-translational modifications

Generally, post-translational modifications are identified by digesting proteins into peptides and using an enrichment step like that described for protein phosphorylation. Increasingly, PTM-specific antibodies that allow immunoaffinity purification of modified peptides are becoming very useful in cases there modified peptides cannot be readily isolated by their physiochemical properties.

This approach allows specific purification of tyrosine phosphorylated peptides which are generally a minor component of the phosphoproteome captured using IMAC or TiO2.

Ubiquitination sites can also be targeted using a similar approach; samples are digested with trypsin and this not only digests ubiquitin modified proteins themselves but cleaves the lysine attached ubiquitin chain leaving a remnant of K-GG (two glycine residues attached to the modified lysine residue). Antibodies that specifically recognise this remnant motif can be used to enrich thousands of peptides harbouring ubiquitin sites.

We have experience with the enrichment and identification of other modifications such as palmitoylation in which the modification itself is selectively released from proteins and previously modified sites are biotin labelled allowing enrichment of previously modified proteins and peptides.

Protein-protein interaction study

Affinity-purification mass spectrometry

This approach is incredibly useful for unbiased identification of components of multiprotein complexes. A key aspect of this method is the specificity of both bait protein capture and release of purified protein complexes. Traditional immunoprecipitation experiments in which interacting proteins are identified by western blotting can tolerate samples that are of lower purity when the necessary controls are included, as candidate interacting proteins are targeted by blotting. However, in AP-MS, all proteins that are present in the enriched sample can theoretically be identified and therefore the relative enrichment of the bait and associated proteins over the background is very important.

Samples with high background (non-specifically interacting proteins) will result in much of the sequencing time being devoted to irrelevant proteins and therefore a reduced ability to identify proteins interacting with the bait. Bait specific antibodies can be used for AP-MS as along as the antibody specificity is good and complexes can be cleanly eluted from immobilised antibodies.

Generally, the method of choice is to express a version of the bait protein that contain a generic affinity tag (e.g. FLAG, HA, GFP etc.) that can be used for capture by high quality tag specific antibodies.

Tags such as FLAG are particularly good as proteins complexes can be competitively eluted by incubation with FLAG peptide resulting in a clean complex with greatly reduced contamination from antibody chains and non-specifically bound proteins commonly found in pH elutions or by boiling resin in SDS buffers. TAP (tandem affinity) tags offer two rounds of enrichment and are particularly useful for generating highly enriched protein complexes but can result in the loss of less stably interacting proteins that are better identified by faster single step affinity enrichments.