Biochemistry

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.

Xenopus laevis oocytes are widely used as a heterologous expression system for investigating the function of membrane proteins due to robust expression of heterologous protein and a low endogenous transport background. Traditionally, transporter activity in oocytes has been assessed using electrophysiology or radiolabeled uptake assays, approaches that are constrained by the requirement for electrogenicity of the transport process, availability of radiolabeled compounds, and instrumentation. Here, we describe a fluorescence-based uptake assay that enables direct and rapid quantification of transporter activity using a fluorescence plate reader. The protocol uses the Arabidopsis thaliana sucrose transporter 1 (SUC1) and its fluorescent substrate esculin as a case for how to set up the assay. The workflow includes optimizing assay conditions, sample preparation, fluorescent measurements, and downstream data analysis using R. This method can readily be adapted to other transporter-substrate pairs, and it supports applications such as transporter inhibitor screening, mutational analysis, characterization of kinetic properties, or indirect substrate specificity testing through competition assays. Overall, this protocol provides a simple and scalable alternative to traditional techniques, eliminating the need for radiolabeled compounds or electrophysiology while enabling easy quantitative assessment of transporter activity.

Metamorphosis in class Insecta is defined as the drastic transition of an organism from a larval stage to a morphologically very different adult form. Insects such as Drosophila melanogaster undergo metamorphosis, which requires precise temporal control, exerted by ecdysteroids, the primary regulators of these developmental transitions. The active form of ecdysone is 20-hydroxyecdysone (20E), which acts in pulses that trigger the onset of different developmental stages. A massive pulse of 20E occurs between the prepupal and pupal stages, triggering pupariation. Precise quantification of 20E levels is critical for understanding how this hormonal signaling governs the developmental timing of different stages. This protocol is sensitive for detecting low levels of 20E and describes a methanol-based extraction method coupled with a competitive ELISA to quantify whole-body 20E levels in third-instar Drosophila larvae. Further, this protocol enables detection of differences in ecdysone titers among genotypes affecting ecdysone biosynthesis, particularly at the late third-instar larval stage. This protocol can also be utilized for disease-causing insects (Aedes aegypti), as well as economically relevant beneficial insects (Apis mellifera), to study the effects of ecdysteroid analogs during development.

Deubiquitinases (DUBs) are attractive therapeutic targets within the ubiquitin-proteasome system, in part because four of the five DUB subfamilies are cysteine proteases amenable to the development of potent, selective inhibitors, as recently demonstrated for USP7. Identifying DUBs that deubiquitylate and stabilize specific human oncogenic proteins is therefore a promising approach to discovering new mechanism-based targets for cancer therapy. Several complementary experimental strategies are typically required to identify bona fide DUB–substrate pairs. Here, we present an efficient, straightforward in vitro immune-complex protocol to validate USP17-mediated deubiquitylation of the transcriptional co-activator β-catenin. In this assay, both β-catenin and USP17 are immunopurified from transiently transfected 293T cells and then combined to assess USP17 enzymatic activity. The protocol describes the in vitro enzymatic assay performed on immunopurified complexes and the immunoblot-based readout. It can be readily adapted to other DUBs and substrates for mechanistic studies.

Palmitoylation is a crucial post-translational modification, and bioorthogonal chemistry based on azide-alkyne cycloaddition is typically used to verify protein palmitoylation. Traditional copper-catalyzed click chemistry (CuAAC) proceeds with fast kinetics and is widely used, but it requires a copper catalyst and suffers from copper-induced toxicity and nonspecific labeling. By contrast, strain-promoted click chemistry (SPAAC) has slower kinetics but is catalyst-free, offering high specificity, low cytotoxicity, and simple operation. However, SPAAC is mostly applied to live-cell labeling and imaging of known palmitoylated proteins in the field of palmitoylation, and its use in identifying novel palmitoylated proteins is still limited. Here, we present a SPAAC-based method for detecting endogenous protein palmitoylation. Compared with CuAAC, this method eliminates the need for copper catalysts and reducing agents, thereby simplifying the procedure and reducing reagent usage.

Memory is a fundamental process, regulated by protein–protein interactions within neuronal proteome networks. Learning-dependent changes in specific brain regions important for memory have been detected by mass spectrometry, by comparing proteins from animals trained to learn with mock-trained controls. Detection through this method relies on relative protein abundance; brain dissection is readily available for macroscopic animals to spatially control protein identification by mass spectrometry. In the nematode C. elegans, however, such spatial control is limited due to its microscopic size, hindering its utilization in proteomics. A protocol to address this limitation would strengthen an already excellent model to study memory, given that many proteins for learning are evolutionarily conserved in the worm and single-cell expression is uniquely defined across all 302 neurons. We modified existing protocols to enable (i) proximity labeling detection of neuronal proteins in C. elegans and (ii) high-throughput enrichment of these proteins from >3,000 whole worm bodies simultaneously, to assess trained vs. mock-trained proteomes. This involved the biotin ligase enzyme TurboID, which promiscuously labels nearby proteins with its substrate biotin. Enzyme expression was transgenically restricted to the nervous system, and biotin supplementation was limited to the training (or mock training) period in a classical (gustatory) conditioning paradigm. Labeled proteins were enriched by pull-down using streptavidin, which has a high binding affinity to biotin, and then processed for mass spectrometry runs and qualitative data analysis. This protocol is uniquely advantageous in that it minimizes proteins present before a temporal window of interest (training/mock training), improving the detection of lowly abundant proteins from a specific tissue in the worm (neurons). We have demonstrated that the protocol can sufficiently detect novel learning regulators, thus providing a useful framework to interrogate proteomes in microscopic brains.

The transient receptor mucolipin subtype 1 (TRPML1) is a ubiquitously expressed ion channel involved in lysosomal homeostasis. Recent pharmaceutical interest in developing agonist ligands has emerged due to beneficial effects in neurodegenerative diseases. The major high-throughput screening techniques to investigate this ion channel involve fluorescent calcium imaging and electrophysiology. Despite their high capacity for screening compounds, it is well known that both methods face hurdles, such as the need for expensive, specialized equipment. Here, we present a novel technique to screen for ligands of TRPML1 using a bioluminescence resonance energy transfer (BRET) assay. This assay consists of a target engagement assay in live cells, which permits the determination of binding constants between ligands and the target of interest in equilibrium or time-dependently. We employ a full-length TRPML1 C-terminally tagged with the small bioluminescent protein nanoluciferase. This ensures the correct localization of the ion channel in the lysosomal membrane and an optimal placement of the luciferase in the cytoplasm. We also developed a cell- and lysosome-permeable fluorescent BRET tracer that gives a BRET signal only when bound to the ion channel. This new protocol allows researchers worldwide to screen compounds that would interact with TRPML1 by using any plate reader with luminescent and fluorescence filters.

ADP ribosylation (ADPr) is a crucial post-translational modification that plays a vital role in DNA damage repair. Catalyzed by ADP ribose polymerases using NAD⁺ as a substrate, ADPr activates DNA repair pathways rapidly, thereby maintaining genomic integrity. The involvement of ADP ribose hydrolases in this process is significant, as they hydrolyze PAR chains, facilitating the release of ADPr-modified proteins from DNA or other proteins, which is essential for subsequent DNA repair steps. This protocol outlines a high-throughput screening method for identifying inhibitors of ADP ribose hydrolases, utilizing His-Tb-conjugated and ADPr-modified His-ADP ribose polymerase as the signal donor, and GST-d2-conjugated GST-XRCC1 as the signal receptor. The detection of time-resolved fluorescence signals enables efficient evaluation of compounds with potential therapeutic activity against cancer.

Nucleases are key tools in molecular biology, enabling controlled nucleic acid digestion for applications such as ribosome profiling. Micrococcal nuclease (MNase) from Staphylococcus aureus is widely used as a tool in molecular biology and biochemistry, but its reduced activity under high-salt conditions necessitates higher enzyme input to achieve efficient digestion, increasing costs in studies of halophilic organisms. Here, we present an optimized protocol for the heterologous expression and purification of the recombinant staphylococcal MNase. The procedure enables reproducible production of a highly active, stable enzyme and incorporates an enzymatic activity assay to standardize batches to minimize variability. The resulting MNase exhibits robust activity in high-salt environments and remains stable during storage, providing a cost-effective and reliable alternative to commercial nucleases for ribosome profiling and related applications.

Chromatin modifications regulate genome function by recruiting proteins that control transcription, genome organization, and DNA repair. Identifying the proteins associated with specific chromatin modifications is therefore essential for understanding how these regulatory processes operate. Traditional approaches, including chromatin immunoprecipitation and affinity purification coupled to mass spectrometry, have uncovered many chromatin-associated proteins. However, they often rely on crosslinking and chromatin fragmentation, which can disrupt native chromatin architecture and limit the detection of transient interactions. Here, we describe a proximity-labeling protocol for identifying the chromatin-dependent protein interactome associated with specific chromatin marks, termed ChromID. ChromID uses engineered chromatin readers (eCRs) fused to a promiscuous biotin ligase, which labels proteins in the immediate vicinity of the targeted chromatin mark. The protocol includes in vivo biotin labeling, nuclear extract preparation, streptavidin-based enrichment, and tryptic digestion for downstream LC-MS/MS analysis. The protocol has been validated across multiple cell types and chromatin contexts and can be extended to other chromatin-associated proteins, providing a versatile approach to profile chromatin-associated proteomes within their native cellular environment.