The past 25 years have seen amazing advances in mass spectrometry-based proteomics technologies and their use in a broad range of biological applications. While mass spectrometry is a well-established and indispensable tool for proteomics, the Yates lab continues to strive to improve MS performance for proteomics applications and to innovate techniques that expand the scope of biological questions that can be addressed by MS. Some of the methods recently developed in the lab are described here.
Interference-free proteome quantification
Chemical labeling of peptides prior to shotgun proteomics allows relative quantification of proteins in biological samples independent of sample origin. Current strategies utilize isobaric labels that fragment into reporter ions. However, quantification of reporter ions results in distorted ratio measurements due to contaminating peptides that are co-selected in the same precursor isolation window. We have developed a strategy for quantitation of isobaric peptide fragment isotopologues in tandem mass spectra that reduces precursor interference. The method is based on the relative quantitation of isobaric isotopologues of dimethylated peptide fragments in tandem mass spectra following higher energy collisional dissociation (HCD). Our approach enables precise quantification of a proteome down to single spectra per protein and quantifies >90% of proteins in a MudPIT experiment.
In-line separation by capillary electrophoresis prior to analysis by top-down mass spectrometry for characterization of protein complexes
Intact protein analysis via top-down mass spectrometry (MS) allows comprehensive characterization of protein variants, splice isoforms, and combinatorial post-translational modifications (PTMs), but the analysis of complex mixtures of intact proteins on a proteomics scale presents a challenge. We have developed a sensitive top down strategy by applying capillary electrophoresis through a sheathless CE-electrospray ionization interface (CESI) coupled to an LTQ Velos Orbitrap Elite mass spectrometer, and have utilized it to analyze the Dam1 complex from Saccharomyces cerevisiae, as well as the proteome from Pyrococcus furiosus. In analysis of the Dam1 complex, we achieved a 100-fold increase in sensitivity compared to a reversed-phase liquid chromatography coupled MS analysis of recombinant Dam1 complex and were able to observe N-terminal processing forms of individual subunits of the Dam1 complex as well as their phosphorylation stoichiometry upon Mps1p kinase treatment. CESI-top-down MS analysis of Pyrococcus furiosus cell lysate identified 134 proteins and 291 proteoforms with a total sample consumption of 270 ng in 120 min of total analysis time. Truncations and various PTMs were detected, including acetylation, disulfide bonds, oxidation, glycosylation, and hypusine. This is the largest scale analysis of intact proteins by CE-top-down MS to date.
Pulse AHA Labeling in Mammals
The quantification of newly synthesized proteins (NSP) at set time points using mass spectrometry has the potential to identify important early regulatory or expression changes associated with disease states. NSPs can be enriched from proteomes by employing pulsed introduction of the non-canonical amino acid, azidohomoalanine (AHA). AHA is accepted by the endogenous methionine tRNA and inserted into proteins in vivo. AHA can be enriched using “click chemistry” by reacting the azide of AHA to a biotin-alkyne. Thus, AHA containing proteins or peptides can be enriched and efficiently separated from the whole proteome. To date, the analysis of NSP have been mostly restricted to cultured cells. We developed a strategy, PALM (Pulse AHA Labeling in Mammals), to incorporate AHA into the entire rodent proteome for quantitative tissue proteomic analysis of NSPs in animal models of disease at discrete time points. Our analysis showed that less than a week of an AHA diet is sufficient to incorporate AHA safely into the proteome of multiple tissues and allows for robust identification of thousands of NSP by mass spectrometry. To quantify NSPs from tissues, we devised two different methods by employing heavy stable isotopes incorporated into biotin-alkynes and AHA molecules.