Biochemistry

Progressive neurodegeneration linked to the accumulation of misfolded proteins is a hallmark of several neurodegenerative disorders, including Parkinson’s disease, Huntington’s disease, and Alzheimer’s disease. Dysfunction in the protein homeostasis machinery correlates with pathology. The chaperone protein DNAJB6 is expressed in neurons and oligodendrocytes and has been shown to play a key role in preventing amyloid aggregation by binding to amyloidogenic proteins and facilitating their refolding or degradation, in cooperation with other chaperones. Here, we describe a simple and feasible assay that enables high-throughput screening for DNAJB6 activity in a plate reader format. We use genetically engineered HEK293 cells that stably express DNAJB6 fused to either CFP or YFP. These cells can be plated into multi-well plates, and the fluorescence resonance energy transfer (FRET) signal can be measured for analysis of DNAJB6 dimerization, which is linked to DNAJB6 activity. The protocol can be used for drug screening and to identify compounds that increase DNAJB6 dimerization, and can serve as a starting point for finding new medicines that act through modulating DNAJB6 activity.

Ribonucleoprotein (RNP) condensates are membraneless organelles that exist alongside many RNA-driven processes, such as transcription and splicing. Despite their ubiquity, the biological necessity of forming a condensed phase remains unclear, particularly because the same RNP components exist both within these organelles and in the surrounding dilute phase. Most current methods for studying biochemical interaction dynamics within condensates rely on in vitro reconstitution of minimal factors or low-throughput single-molecule studies. However, RNP condensates are complex organelles containing tens to hundreds of proteins and hundreds to thousands of different RNAs. Here, we describe a scalable, high-throughput fluorescence microscopy–based approach to analyze protein–protein interaction networks, allowing for the rigorous assessment of dynamic, process-critical interactions within RNP condensates from live cells. This method takes advantage of fluorescence lifetime imaging (FLIM) and phasor plot analysis to automate segmentation of condensate-localized fluorescence signals. Using suitable FLIM–Förster resonant energy transfer (FLIM-FRET) fluorescent pairs fused to proteins of interest, protein–protein interactions can be actively monitored throughout various conditions via changes in fluorescence lifetime. Results from this assay yield valuable insight into the organization and assembly of essential factors for different condensate-associated processes to infer the functional consequences of RNP granule partitioning. Although this protocol is tailored for studying protein interactions within condensates, the design and execution framework can be adapted to investigate protein–protein interactions across a wide variety of compartments within different biological systems.

Small ubiquitin-like modification (SUMOylation) is a crucial post-translational modification that modulates protein stability, localization, and interaction dynamics. Despite the identification of thousands of putative small ubiquitin-like modifier (SUMO) substrates, functional validation remains challenging due to the low abundance and highly dynamic nature of SUMOylated proteins. Here, we present a protocol for detecting protein SUMOylation, integrating bioinformatic site prediction, and rapid substrate screening via in vivo tobacco transient expression and in vitro E. coli assay, followed by precise validation using transgenic Arabidopsis lines. However, detection of low-abundance SUMOylated proteins may require coupling with mass spectrometry, and the in vitro system does not fully recapitulate the complex regulatory network in vivo. This workflow provides a useful tool for studying SUMOylation in plants.

Ornithine decarboxylase (ODC) is a rate-limiting enzyme in polyamine biosynthesis that plays a critical role in cell proliferation and tumorigenesis. Reliable quantification of ODC activity is essential for mechanistic and therapeutic studies. Traditional assays often rely on radiolabeled substrates or discontinuous endpoint measurements. Here, we describe a non-radioactive, continuous spectrophotometric assay for measuring ODC activity in cell lysates using a commercially available liquid-stable CO₂ detection reagent. In this assay, CO₂ generated by ODC is captured as bicarbonate and utilized in a coupled enzymatic system containing phosphoenolpyruvate carboxylase (PEPC) and malate dehydrogenase (MDH), leading to oxidation of thio-NADH. The decrease in absorbance at 405 nm due to thio-NADH oxidation is monitored in real time and is proportional to ODC activity. The protocol is performed in a 96-well plate format, requires minimal reagent preparation, and is suitable for medium- to high-throughput applications.

UFMylation is an evolutionarily conserved ubiquitin-like modification that covalently conjugates UFM1 to lysine residues of substrates via a sequential E1-E2-E3 enzymatic cascade. UFMylation plays a pivotal role in maintaining cellular homeostasis, and its dysregulation is closely linked to multiple major diseases, including malignant tumors, hematopoietic defects, neurodegenerative disorders, and congenital developmental defects, highlighting its important biological significance. However, few substrates of UFMylation have been reported to date, limiting our deep understanding of the mechanistic functions of this modification. This major bottleneck stems from two major technical limitations: the overwhelming abundance of ribosomal protein L26 (RPL26)-UFM1 conjugates masks signals from low-abundance substrates, and conventional methods rely on cumbersome cotransfection of multiple pathway components with poor efficiency and specificity in UFMylated peptides enrichment. To address these challenges, we have developed an effective and specific experimental protocol for UFMylation detection and large-scale substrate identification. This protocol employs CRISPR-Cas9-mediated gene editing to generate UFSP1/UFSP2 double-knockout (UFSP1^KO/UFSP2^KO, DKO) HEK293T cells, which completely abrogate de-UFMylation and thus significantly elevate global protein UFMylation levels upon exogenous introduction of mature UFM1-ΔC2. In addition, exogenous co-expression of the E3 ligase core components UFL1 and DDRGK1 can further improve the sensitivity of substrate detection. This protocol enables large-scale identification of UFMylation substrates with modification sites via high-efficiency enrichment with the K-ε-VG antibody and LC-MS/MS analysis.

This protocol describes an optimized strategy for the efficient generation of peptide-loaded major histocompatibility complex (MHC) class II (pMHC) tetramers, which are essential tools for detecting and characterizing antigen-specific T cells in immunological research. Traditional methods require separate expression of MHC proteins followed by in vitro biotinylation—a multi-step process that is time-consuming and prone to protein loss. Here, we present an integrated approach based on co-expression of MHC monomers and BirA biotin ligase in Expi293F T cells, enabling site-specific biotinylation in vivo during protein synthesis. At the same time, the incorporation of a thrombin-cleavable class II–associated invariant chain peptide (CLIP) peptide into the MHC construct allows flexible loading of any antigenic peptide of interest without the need for re-cloning or re-expression of the MHC molecule. Pre-biotinylated MHC molecules are subsequently purified, loaded with antigenic peptides, and assembled into fluorescent tetramers via streptavidin conjugation. This streamlined workflow significantly reduces handling steps, improves protein yield, and enhances reproducibility. The resulting tetramers are suitable for sensitive detection and isolation of antigen-specific T cells by flow cytometry, supporting applications in T-cell immunogenicity studies, vaccine development, and autoimmune disease research.

Seahorse metabolic assays are now widely utilized across numerous fields for performing functional assessments of glycolysis and mitochondrial function in adherent or suspension cell culture samples. Seahorse assays measure extracellular acidification rate (ECAR) and oxygen consumption rate (OCR) as a means of assessing glycolysis and mitochondrial function, respectively. Currently, the vast majority of Seahorse metabolic assays are performed using in vitro samples due to the current established standardized method. However, a uniform approach to assess real-time functional measurements of glycolysis and mitochondrial function in ex vivo tissue samples remains elusive. In particular, this protocol was designed to assess glycolysis in ex vivo murine intestinal samples through ECAR measurements using the Agilent Seahorse XFe24 platform with corresponding Islet Capture microplates and screens. This protocol was developed to provide functional measurements of glycolytic metabolism in murine intestinal tissue samples. This protocol details a method to assess glycolysis in tissue samples and represents the next stage of ex vivo metabolic methods to complement existing standardized in vitro approaches. While this protocol was developed to assess ECAR in ex vivo murine intestinal samples, the same approach can be applied to assessing mitochondrial respiration through measurements of OCR in other tissue types. Overall, this protocol expands the purview of Seahorse metabolic assays through the inclusion of tissue samples and provides the framework to interrogate organ-level metabolism in the context of systemic nutrient metabolism and physiology.

Phosphatase and tensin homolog-induced kinase 1 (PINK1) is a serine/threonine kinase that plays a key role in mitophagy initiation. Loss-of-function autosomal recessive mutations in PINK1 cause early onset Parkinson’s disease (EOPD). Current approaches for studying PINK1 function depend on bulk techniques that can only provide snapshots of activity and could miss the dynamics and cell-to-cell heterogeneity of PINK1 activity or provide an indirect readout of PINK1 activity. Here, we present a protocol using our newly developed phase separation–based PINK1 biosensor (PINK1-SPARK) to observe real-time activity of endogenous PINK1 in single cells. Following transfection of live cells with PINK1-SPARK, cells are treated with mitochondrial depolarizing agents and visualized using widefield or confocal fluorescence microscopy, either following the same cells over time for time-lapse imaging of PINK1 activity or end-point measurements. Thus, PINK1-SPARK is a new tool that enables the measurement of PINK1 activity in single live cells, allowing for further elucidation of the role of PINK1 in mitophagy and cell function.

Glycogen is a highly conserved macromolecule across species, and its visualization provides critical insights into both physiological processes and disease states. Existing approaches for glycogen imaging in Caenorhabditis elegans rely primarily on traditional microscopy slides, which introduce variability in image acquisition and downstream data analysis, limit throughput, and require substantial hands-on time and technical expertise.

Here, we present a standardized, cost-effective, and high-throughput imaging method that enables efficient visualization and quantification of glycogen in C. elegans. Our approach utilizes a custom-designed three-dimensional pad containing two to four chambers, allowing control and experimental samples to be processed simultaneously under identical conditions. Worms are exposed to iodine crystals, ensuring uniform staining while minimizing reagent use and handling variability. Imaging is performed using a simple binocular microscope, and analysis is conducted in Fiji, making the workflow accessible to laboratories with minimal specialized equipment or training.

This method also reduces technical variability, shortens turnaround time, and requires only basic reagents and expertise, making it well-suited for both research and teaching laboratories. Importantly, the platform is readily adaptable to other nematode species and scalable for large-scale genetic or pharmacological screening applications. Together, this workflow minimizes technical variability and provides a robust platform for comparative glycogen analysis in C. elegans.

Apolipoprotein B–containing lipoproteins (ApoB-LPs) transport lipids throughout the circulation and are closely associated with cardiovascular disease in humans. Many aspects of ApoB-LP biology remain elusive, often due to their indirect characterization through the measurement of plasma triglycerides and cholesterol. The conventional approach provides limited information on ApoB-LPs number and size distribution, essential features that influence cardiovascular disease risk. Additionally, drug studies have historically been limited to the use of mammalian research models, which are not suited for high-throughput experiments. Therefore, we generated a reporter system (LipoGlo) utilizing a luciferase enzyme (NanoLuc) fused to the C-terminus of the zebrafish (Danio rerio) ApoBb.1 protein. In metazoans, ranging from insects to humans, each ApoB-LP contains a single ApoB molecule, such that the luminescence emitted from these transgenic fish is proportional to the total number of ApoB-LPs. The LipoGlo zebrafish reporter generates a quantitative chemiluminescent signal that can be used in plate-based assays to measure lipoprotein quantities, a gel-based assay that can measure lipoprotein size distribution, and chemiluminescent microscopy that can, for the first time, visualize lipoprotein localization in a larval zebrafish. LipoGlo, combined with the amenability of zebrafish to genetic approaches, facilitates the rapid assessment of any gene or drug’s role in ApoB-LP molecular and cell biology. This protocol describes three optimized LipoGlo assays that facilitate ApoB-LP characterization with 100× less starting material than prior assays routinely used for mammalian lipoprotein analysis.