Direct Tissue Profiling of Protein Complexes: Toward Native Mass Spectrometry Imaging

Native mass spectrometry seeks to probe noncovalent protein interactions in terms of protein quaternary structure, protein–protein and protein–ligand complexes. The ultimate goal is to link the understanding of protein interactions to the protein environment by visualizing the spatial distribution of noncovalent protein interactions within tissue. Previously, we have shown that noncovalently bound protein complexes can be directly probed via liquid extraction surface analysis from dried blood spot samples, where hemoglobin is highly abundant. Here, we show that the intact hemoglobin complex can be sampled directly from thin tissue sections of mouse liver and correlated to a visible vascular feature, paving the way for native mass spectrometry imaging

High Field Asymmetric Waveform Ion Mobility Spectrometry in Nontargeted Bottom-up Proteomics of Dried Blood Spots

Despite the great potential of dried blood spots (DBS) as a source of endogenous proteins for biomarker discovery, the literature relating to nontargeted bottom-up proteomics of DBS is sparse, primarily due to the inherent complexity and very high dynamic range associated with these samples. Here, we present proof-of-concept results in which we have coupled high field asymmetric waveform ion mobility spectrometry (FAIMS) with liquid chromatography–tandem mass spectrometry (LC–MS/MS) for nontargeted bottom-up proteomics of DBS with the aim of addressing these challenges. We, and others, have previously demonstrated the benefits of FAIMS more generally in proteomics including improved signal-to-noise and extended proteome coverage, and the aim of the current work was to extend those benefits specifically to DBS. The DBS samples were either extracted by the more traditional manual “punch and elute” approach or by an automated liquid surface extraction (LESA) approach prior to trypsin digestion. The resulting samples were analyzed by LC–MS/MS and LC–FAIMS–MS/MS analysis. The results show that the total number of proteins identified increased by ∼50% for the punch and elute samples and ∼45% for the LESA samples in the LC–FAIMS–MS/MS analysis. For both the punch and elute samples and the LESA samples, ∼30% of the total proteins identified were observed in both the LC–MS/MS and the LC–FAIMS–MS/MS data sets, illustrating the complementarity of the approaches. Overall, this work demonstrates the benefits of inclusion of FAIMS for nontargeted proteomics of DBS

Subcritical Water Processing of Proteins: An Alternative to Enzymatic Digestion?

Subcritical water is an emerging tool in the processing of bioorganic waste. Subcritical water is an environmentally benign solvent which has the potential to provide an alternative to traditional methods of protein hydrolysis without the inclusion of expensive acids or enzymes. To date, most studies on the subcritical water mediated hydrolysis of proteins have focused on the production of amino acids, rather than the intermediate peptides. Here, we investigate the specificity of subcritical water with respect to the production of peptides from three model proteins, hemoglobin, bovine serum albumin, and β-casein, and compare the results with enzymatic digestion of proteins by trypsin. In addition, the effect of subcritical water (SCW) treatment on two protein post-translational modifications, disulfide bonds and phosphorylation, was investigated. The results show that high protein sequence coverages (>80%) can be obtained following subcritical water hydrolysis. These are comparable to those obtained following treatment with tryspin. Under mild subcritical water conditions (160 °C), all proteins showed favored cleavage of the Asp-X bond. The results for β-casein revealed favored cleavage of the Glu-X bond at subcritical water temperatures of 160 and 207 °C. That was similarly observed for bovine serum albumin at a subcritical water temperature of 207 °C. Subcritical water treatment results in very limited cleavage of disulfide bonds. Reduction and alkylation of proteins either prior to or post subcritical water treatment improve reported protein sequence coverages. The results for phosphoprotein β-casein show that, under mild subcritical water conditions, phosphorylation may be retained on the peptide hydrolysis products

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Large-Scale Analysis of Peptide Sequence Variants: The Case for High-Field Asymmetric Waveform Ion Mobility Spectrometry

Large scale analysis of proteins by mass spectrometry is becoming increasingly routine; however, the presence of peptide isomers remains a significant challenge for both identification and quantitation in proteomics. Classes of isomers include sequence inversions, structural isomers, and localization variants. In many cases, liquid chromatography is inadequate for separation of peptide isomers. The resulting tandem mass spectra are composite, containing fragments from multiple precursor ions. The benefits of high-field asymmetric waveform ion mobility spectrometry (FAIMS) for proteomics have been demonstrated by a number of groups, but previously work has focused on extending proteome coverage generally. Here, we present a systematic study of the benefits of FAIMS for a key challenge in proteomics, that of peptide isomers. We have applied FAIMS to the analysis of a phosphopeptide library comprising the sequences GPSGXVpSXAQLX­(K/R) and SXPFKXpSPLXFG­(K/R), where X = ADEFGLSTVY. The library has defined limits enabling us to make valid conclusions regarding FAIMS performance. The library contains numerous sequence inversions and structural isomers. In addition, there are large numbers of theoretical localization variants, allowing false localization rates to be determined. The FAIMS approach is compared with reversed-phase liquid chromatography and strong cation exchange chromatography. The FAIMS approach identified 35% of the peptide library, whereas LC–MS/MS alone identified 8% and LC–MS/MS with strong cation exchange chromatography prefractionation identified 17.3% of the library

High Field Asymmetric Waveform Ion Mobility Spectrometry in Nontargeted Bottom-up Proteomics of Dried Blood Spots

Despite the great potential of dried blood spots (DBS) as a source of endogenous proteins for biomarker discovery, the literature relating to nontargeted bottom-up proteomics of DBS is sparse, primarily due to the inherent complexity and very high dynamic range associated with these samples. Here, we present proof-of-concept results in which we have coupled high field asymmetric waveform ion mobility spectrometry (FAIMS) with liquid chromatography–tandem mass spectrometry (LC–MS/MS) for nontargeted bottom-up proteomics of DBS with the aim of addressing these challenges. We, and others, have previously demonstrated the benefits of FAIMS more generally in proteomics including improved signal-to-noise and extended proteome coverage, and the aim of the current work was to extend those benefits specifically to DBS. The DBS samples were either extracted by the more traditional manual “punch and elute” approach or by an automated liquid surface extraction (LESA) approach prior to trypsin digestion. The resulting samples were analyzed by LC–MS/MS and LC–FAIMS–MS/MS analysis. The results show that the total number of proteins identified increased by ∼50% for the punch and elute samples and ∼45% for the LESA samples in the LC–FAIMS–MS/MS analysis. For both the punch and elute samples and the LESA samples, ∼30% of the total proteins identified were observed in both the LC–MS/MS and the LC–FAIMS–MS/MS data sets, illustrating the complementarity of the approaches. Overall, this work demonstrates the benefits of inclusion of FAIMS for nontargeted proteomics of DBS

Large-Scale Analysis of Peptide Sequence Variants: The Case for High-Field Asymmetric Waveform Ion Mobility Spectrometry

Large scale analysis of proteins by mass spectrometry is becoming increasingly routine; however, the presence of peptide isomers remains a significant challenge for both identification and quantitation in proteomics. Classes of isomers include sequence inversions, structural isomers, and localization variants. In many cases, liquid chromatography is inadequate for separation of peptide isomers. The resulting tandem mass spectra are composite, containing fragments from multiple precursor ions. The benefits of high-field asymmetric waveform ion mobility spectrometry (FAIMS) for proteomics have been demonstrated by a number of groups, but previously work has focused on extending proteome coverage generally. Here, we present a systematic study of the benefits of FAIMS for a key challenge in proteomics, that of peptide isomers. We have applied FAIMS to the analysis of a phosphopeptide library comprising the sequences GPSGXVpSXAQLX­(K/R) and SXPFKXpSPLXFG­(K/R), where X = ADEFGLSTVY. The library has defined limits enabling us to make valid conclusions regarding FAIMS performance. The library contains numerous sequence inversions and structural isomers. In addition, there are large numbers of theoretical localization variants, allowing false localization rates to be determined. The FAIMS approach is compared with reversed-phase liquid chromatography and strong cation exchange chromatography. The FAIMS approach identified 35% of the peptide library, whereas LC–MS/MS alone identified 8% and LC–MS/MS with strong cation exchange chromatography prefractionation identified 17.3% of the library

Top-Down Mass Analysis of Protein Tyrosine Nitration: Comparison of Electron Capture Dissociation with “Slow-Heating” Tandem Mass Spectrometry Methods

Tyrosine nitration in proteins is an important post-translational modification (PTM) linked to various pathological conditions. When multiple potential sites of nitration exist, tandem mass spectrometry (MS/MS) methods provide unique tools to locate the nitro-tyrosine(s) precisely. Electron capture dissociation (ECD) is a powerful MS/MS method, different in its mechanisms to the “slow-heating” threshold fragmentation methods, such as collision-induced dissociation (CID) and infrared multiphoton dissociation (IRMPD). Generally, ECD provides more homogeneous cleavage of the protein backbone and preserves labile PTMs. However recent studies in our laboratory demonstrated that ECD of doubly charged nitrated peptides is inhibited by the large electron affinity of the nitro group, while CID efficiency remains unaffected by nitration. Here, we have investigated the efficiency of ECD versus CID and IRMPD for top-down MS/MS analysis of multiply charged intact nitrated protein ions of myoglobin, lysozyme, and cytochrome c in a commercial Fourier transform ion cyclotron resonance (FT-ICR) mass spectrometer. CID and IRMPD produced more cleavages in the vicinity of the sites of nitration than ECD. However the total number of ECD fragments was greater than those from CID or IRMPD, and many ECD fragments contained the site(s) of nitration. We conclude that ECD can be used in the top-down analysis of nitrated proteins, but precise localization of the sites of nitration may require either of the “slow-heating” methods

Tissue Washing Improves Native Ambient Mass Spectrometry Detection of Membrane Proteins Directly from Tissue

Native ambient mass spectrometry enables the in situ analysis of proteins and their complexes directly from tissue, providing both structural and spatial information. Until recently, the approach was applied exclusively to the analysis of soluble proteins; however, there is a drive for new techniques that enable analysis of membrane proteins. Here we demonstrate native ambient mass spectrometry of membrane proteins, including β-barrel and α-helical (single and multipass) integral membrane proteins and membrane-associated proteins incorporating lipid anchors, by integration of a simple washing protocol to remove soluble proteins. Mass spectrometry imaging revealed that washing did not disrupt the spatial distributions of the membrane and membrane-associated proteins. Some delocalization of the remaining soluble proteins was observed

Higher Energy Collision Dissociation (HCD) Product Ion-Triggered Electron Transfer Dissociation (ETD) Mass Spectrometry for the Analysis of <i>N</i>‑Linked Glycoproteins

Large scale mass spectrometry analysis of <i>N</i>-linked glycopeptides is complicated by the inherent complexity of the glycan structures. Here, we evaluate a mass spectrometry approach for the targeted analysis of <i>N-</i>linked glycopeptides in complex mixtures that does not require prior knowledge of the glycan structures or pre-enrichment of the glycopeptides. Despite the complexity of <i>N</i>-glycans, the core of the glycan remains constant, comprising two <i>N</i>-acetylglucosamine and three mannose units. Collision-induced dissociation (CID) mass spectrometry of <i>N</i>-glycopeptides results in the formation of the <i>N</i>-acetylglucosamine (GlcNAc) oxonium ion and a [mannose+GlcNAc] fragment (in addition to other fragments resulting from cleavage within the glycan). In ion-trap CID, those ions are not detected due to the low <i>m</i>/<i>z</i> cutoff; however, they are detected following the beam-type CID known as higher energy collision dissociation (HCD) on the orbitrap mass spectrometer. The presence of these product ions following HCD can be used as triggers for subsequent electron transfer dissociation (ETD) mass spectrometry analysis of the precursor ion. The ETD mass spectrum provides peptide sequence information, which is unobtainable from HCD. A Lys-C digest of ribonuclease B and trypsin digest of immunoglobulin G were separated by ZIC-HILIC liquid chromatography and analyzed by HCD product ion-triggered ETD. The data were analyzed both manually and by search against protein databases by commonly used algorithms. The results show that the product ion-triggered approach shows promise for the field of glycoproteomics and highlight the requirement for more sophisticated data mining tools