Biochemistry

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.

Protein kinase B, more commonly known as Akt, is a family of three serine/threonine kinases (Akt1, Akt2, and Akt3) that play a central role in regulating processes such as proliferation, survival, metabolism, and migration through phosphorylation of downstream targets. Given its involvement in numerous cellular processes, aberrant Akt signaling is prevalent across multiple cancer types, underscoring the need for Akt kinase assays to assess activity, regulatory mechanisms, and the efficacy of targeted interventions. Most existing Akt kinase assays rely on expensive commercial kits, some of which employ pre-purified, constitutively active Akt expressed in insect cells, bypassing physiologic autoinhibition of Akt; therefore, they are not suitable for evaluating allosteric inhibitors or context-dependent regulation. Here, we describe a detailed, step-by-step protocol for a nonradioactive Akt kinase assay using epitope-tagged, recombinant Akt1 expressed in a mammalian cell line and isolated by immunoprecipitation. This method eliminates the need to co-express Akt with upstream regulatory kinases or to purify active enzyme from insect cells, a time-consuming and technically demanding process, particularly when analyzing multiple Akt mutants. Because Akt is assayed in a regulated, autoinhibited state, this protocol enables direct evaluation of allosteric inhibitors that cannot be assessed using active Akt purified from insect cells. We note, however, that Akt1 kinase activity in this assay is measured from epitope-tagged, transiently overexpressed protein, which could influence cellular signaling dynamics. Despite this limitation, the cellular context preserves key regulatory features of Akt1 autoinhibition and membrane-dependent activation that are absent in assays using purified, pre-activated kinase. Together, this protocol supports analysis of Akt kinase activity under diverse experimental conditions, including receptor stimulation, pharmacologic treatment, allosteric inhibitor exposure, and mutations, using an accessible, economical, and physiologically relevant approach.

Peroxisomal β-oxidation is a key step in jasmonic acid biosynthesis. Quantitative biochemical characterization of enzymes involved in the β-oxidation pathway is essential for validating their catalytic functions and comparing differences among genetic variants. Existing enzyme activity assays largely rely on chromatographic techniques to quantify substrate consumption or product formation, but these approaches are not well-suited for high-throughput or continuous kinetic measurements. Here, we describe a spectrophotometric assay based on a plate reader determining OsAIM1 enzymatic activity by monitoring the decrease in NADH absorbance at 340 nm. The method employs a 96-well plate reaction system, enabling real-time kinetic measurements and providing a standardized workflow for calculating reaction rates. Reaction components, protein concentration ranges, and data processing parameters were systematically optimized to ensure linearity, reproducibility, and quantitative accuracy. This assay is simple to perform, requires small reaction volumes, and offers relatively high throughput, making it suitable for functional characterization and kinetic analysis of NADH-dependent enzymes.

Transfer RNAs (tRNAs) are important regulators of translation and cellular function. Several high-throughput sequencing methods have been developed to quantitatively analyze tRNA isoacceptors in cells. However, the strong secondary structures and extensive post-transcriptional modification of most tRNA molecules present significant challenges for many reverse transcriptases, negatively impacting sequencing library preparation and causing quantification biases. Currently, the field utilizes processive next-generation reverse transcriptases (ngRTs), such as Induro (New England Biolabs) and UltraMarathonRT (RNAConnect), to address these issues. Despite being used in multiple protocols, these commercial products face little competition and remain costly. However, non-commercial alternatives, such as the original MarathonRT (MRT), are available from gene repositories. MRT is a next-generation reverse transcriptase derived from the Eubacterium rectale group II intron maturase, which can read through RNA secondary structures and chemical modifications. Here, we present a simplified expression and purification protocol for producing highly active MRT that is stable over 1 year. This cost-effective protocol yields a heterogeneous protein preparation with no discernible competing enzymatic activities; it mitigates previously reported precipitation issues, saving one day of laboratory work and eliminating two chromatography-based purification steps. Moreover, the use of the resulting protein preparation has been verified in the mim-tRNAseq pipeline, where it was shown to perform equally to the commercial alternatives Induro and UltraMarathonRT. In addition, we have developed a simple and cost-effective assay for measuring the enzymatic activity of MRT, allowing for batch comparison.