生物化学

Extracellular vesicle (EV) proteomics can be used to study intercellular communication and find biomarkers of non-invasive diseases. Traditional separation methods (ultracentrifugation, size exclusion chromatography) and data-dependent acquisition (DDA) mass spectrometry usually have the drawbacks of copurification of pollutants, poor reproducibility, and insufficient sampling of low-abundance peptides. This protocol describes a workflow for label-free extracellular vesicle proteomics, which combines size exclusion chromatography for vesicle separation, data-independent acquisition (DIA) for deep discovery, and parallel reaction monitoring (PRM) for targeted verification. Plasma-derived extracellular vesicles are treated by standardized lysis, digestion, and LC-MS/MS procedures, so that the protein group of extracellular vesicles can be identified and quantified robustly. The main advantages of this scheme are that it can achieve high quantitative reproducibility, unbiased detection of low-intensity peptides, and seamless transition from discovery to targeted verification, while also being suitable for liquid biopsy samples and various cardiovascular diseases.

The conventional kinesin-1 is a plus-end-directed microtubule-dependent motor protein with distinct motor head, stalk, and tail domains. Along with the motor head, which binds and walks along microtubules in an adenosine 5’-triphosphate (ATP) dependent manner, kinesin also contains a C-terminal microtubule binding tail. Motor-driven collective motility is well characterized using in vitro gliding assays, which show uninterrupted, smooth trajectories of transport. However, gliding assays driven by the full-length Drosophila kinesin-1 with both head and tail resulted in the emergence of spontaneous spatial microtubule patterns and stop-and-go motion. This was reproduced by an equimolar ratio of the active head and passive tail. Here, we describe the detailed protocol to reconstitute these microtubule gliding assays using multiple motor types: the full-length kinesin-1, the motor head or tail, mixtures of both head and tail, and a rigor mutant of the kinesin. We provide details of the approach taken to acquire the image time-series, to then quantify the spatial patterns that result from these motor combinations. Our approach provides a framework to systematically characterize the spatiotemporal effects of molecular motor-driven collective microtubule transport.

Spatial proteomics enables the mapping of protein distribution within tissues, which is crucial for understanding cellular functions in their native context. While spatial transcriptomics has seen rapid advancement, spatial proteomics faces challenges due to protein non-amplifiability and mass spectrometry sensitivity limitations. This protocol describes a sparse sampling strategy for spatial proteomics (S4P) that combines multi-angle tissue strip microdissection with deep learning–based image reconstruction. The method achieves whole-tissue slice coverage with significantly reduced sampling requirements, enabling mapping of over 9,000 proteins in mouse brain tissue at 525 μm resolution within 200 h of mass spectrometry time. Key advantages include reduced sample processing time, deep proteome coverage, and applicability to centimeter-sized tissue samples.

Protein S-nitrosylation is a critical post-translational modification that regulates diverse cellular functions and signaling pathways. Although various biochemical methods have been developed to detect S-nitrosylated proteins, many suffer from limited specificity and sensitivity. Here, we describe a robust protocol that combines a modified biotin-switch technique (BST) with streptavidin-based affinity enrichment and quantitative mass spectrometry to detect and profile nitrosylated proteins in cultured cells. The method involves blocking free thiols, selective reduction of nitrosothiols, biotin labeling, enrichment of biotinylated proteins, and identification by tandem mass tag (TMT)-based quantitative mass spectrometry. Additionally, site-directed mutagenesis is employed to generate “non-nitrosylable” mutants for functional validation of specific nitrosylation sites. This protocol provides high specificity, quantitative capability, and versatility for both targeted and global analysis of protein nitrosylation.

Immunopeptidomics enables the identification of peptides presented by major histocompatibility complex (MHC) molecules, offering insights into antigen presentation and immune recognition. Understanding these mechanisms in hypoxic conditions is crucial for deciphering immune responses within the tumor microenvironment. Current immunopeptidomics approaches do not capture hypoxia-induced changes in the repertoire of MHC-presented peptides. This protocol describes the isolation of MHC class I-bound peptides from in vitro hypoxia-treated cells, followed by liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis. It describes optimized steps for cell lysis, immunoaffinity purification, peptide elution, and MS-compatible preparation under controlled low-oxygen conditions. The method is compatible with various quantitative mass spectrometry approaches and can be adapted to different cell types. This workflow provides a reliable and reproducible approach to studying antigen presentation under hypoxic conditions, thereby enhancing physiological relevance and facilitating deeper immunological insights.

Pyruvate kinase M2 (PKM2) is a key glycolytic enzyme that catalyzes the conversion of phosphoenolpyruvate (PEP) to pyruvate, producing ATP in the final step of glycolysis. Unlike other isoforms, PKM2 is uniquely regulated, shifting between active tetramers and less active dimers to balance energy production with biosynthetic demands. This flexibility is exploited in cancer cells to support the Warburg effect and anabolic growth. Additionally, PKM2 can translocate to the nucleus and act as a transcriptional co-activator, influencing gene expression and tumor progression. To facilitate functional studies of PKM2, we present a robust and reproducible protocol for its expression, purification, and enzymatic characterization. PKM2 is expressed in E. coli and purified via Ni-NTA affinity and size-exclusion chromatography to ensure high purity and proper folding. Enzymatic activity is measured using a lactate dehydrogenase (LDH)-coupled assay that tracks NADH oxidation at 340 nm, allowing sensitive kinetic analysis under various conditions, including different PEP concentrations, pH levels, and presence of the allosteric activator fructose-1,6-bisphosphate (FBP). This non-radioactive, high-resolution method is suitable for analyzing PKM2 regulation, post-translational modifications, and mutant variants, as well as for screening potential therapeutic modulators, providing a valuable tool for cancer metabolism research.

Protein synthesis and degradation (i.e., turnover) forms an important part of protein homeostasis and has been implicated in many age-associated diseases. Different cellular locations, such as organelles and membraneless compartments, often contain individual protein quality control and degradation machineries. Conventional methods to assess protein turnover across subcellular compartments require targeted genetic manipulation or isolation of specific organelles. Here we describe a protocol for simultaneous proteome localization and turnover (SPLAT) analysis, which combines protein turnover measurements with unbiased subcellular spatial proteomics to measure compartment-specific protein turnover rates on a proteome-wide scale. This protocol utilizes dynamic stable isotope labeling of amino acids in cell culture (dynamic SILAC) to resolve the temporal information of protein turnover and multi-step differential ultracentrifugation to assign proteins to multiple subcellular localizations. We further incorporate 2D liquid chromatography fractionation to greatly increase analytical depth while multiplexing with tandem mass tags (TMT) to reduce acquisition time 10-fold. This protocol resolves the spatial and temporal distributions of proteins and can also reveal temporally distinct spatial localizations within a protein pool.

Counting protein molecules helps reveal the organization of components within cellular structures and the stoichiometries of protein complexes. Existing protein and peptide quantitation methods vary in their complexity. Here, we report a straightforward workflow to measure the absolute number of HaloTag-labeled myosin 10 (Myo10) molecules in U2OS cells. Myo10 is a motor protein that plays a prominent role in cellular protrusion formation. Various biochemical and biological properties of Myo10 are established, but it is not well-defined how many molecules of Myo10 pack into narrow cellular structures called filopodia. We present a workflow for using SDS-PAGE to calibrate Myo10 signal with a reference protein, segmenting epifluorescence microscopy images to map Myo10 intracellular distribution, and interpreting the results to derive biological and functional insights. Our protocol is simple to employ and not only applicable for Myo10 research but also easily adaptable for other biological systems that use HaloTag.

We have observed that some proinsulin molecules in pancreatic islets and beta cell lines have incomplete or improper intramolecular disulfide bonds. These misfolded monomers can form intermolecular disulfide-linked complexes. Accurately quantifying the fraction of proinsulin molecules contained in these complexes is challenging. By proinsulin immunoblotting after nonreducing SDS-PAGE, the signal for disulfide-linked complexes can exceed the total proinsulin signal detected after reducing SDS-PAGE (i.e., overestimating the abundance of misfolded species due to antibody affinity differences). However, after modification of the SDS-PAGE and electrotransfer protocol, we have been able to more accurately estimate the fraction of proinsulin monomers folded to the native state, as well as misfolded proinsulin monomers and disulfide-linked complexes. Our improved technique offers the ability to obtain a more precise assessment of proinsulin misfolding under different environmental conditions in beta cells and normal islets and in diabetes.

Cytosolic peptide: N-glycanase (PNGase/NGLY1 in mammals), an amidase classified under EC:3.5.1.52, is a highly conserved enzyme across eukaryotes that catalyzes the removal of N-glycans from glycoproteins, converting N-glycosylated asparagine residues into aspartic acid. This enzyme also plays a role in the quality control system for nascent glycoproteins. Despite the development of non-radioisotope-based assay systems such as those using S-alkylated RNase or fluorescent-labeled glycopeptides as substrates, these methods are incompatible with crude enzyme sources, primarily due to the degradation of reaction products by contaminating endogenous proteases. We previously developed an assay system using a 5-carboxyfluorescein-labeled glycosylated cyclo-heptapeptide (5FAM-GCP), a substrate remarkably resistant to endogenous peptidase activity. This system enables the accurate measurement of endogenous NGLY1 activity in various samples, including cell lines, tissues, peripheral blood mononuclear cells, and NGLY1-deficient patient-derived cells, without the interference of proteolytic degradation. We recently advanced this approach by producing a novel fluorescence resonance energy transfer (FRET)-based GCP probe (fGCP) and demonstrated its ability to detect endogenous NGLY1 activity across diverse enzyme sources via fluorescence on multiarray plates. This innovative and straightforward assay now offers reliable disease diagnostics and also allows the measurement of endogenous PNGase/NGLY1 activities across various organisms.