Biochemistry

OtUBD is a high-affinity ubiquitin-binding domain (UBD) derived from a large protein produced by the microorganism Orientia tsutsugamushi. The following protocol describes a step-by-step process for the enrichment of ubiquitinated proteins from baker's yeast and mammalian cell lysates using OtUBD. The OtUBD affinity resin can strongly enrich both mono- and poly-ubiquitinated proteins from crude lysates. The protocol further describes the use of different buffer formulations to specifically enrich for proteins covalently modified by ubiquitin with or without proteins that associate with them. Combining different OtUBD-mediated enrichment protocols with liquid chromatography–tandem mass spectrometry (LC–MS/MS) helps distinguish the pool of covalently ubiquitinated proteins (the ubiquitinome) from ubiquitin- or ubiquitinated protein-interacting proteins (the ubiquitin interactome). The OtUBD tool described in the protocol has been used successfully with downstream applications such as immunoblotting and differential proteomics. It provides researchers with a versatile and economical tool for the study of ubiquitin biology.

The phototransduction cascade allows photoreceptors to detect light across a wide range of intensities without saturation, with cGMP serving as the second messenger and calcium feedback as the key regulatory mechanism. While experimental evidence suggests that cAMP may also play a role in modulating this cascade, such regulation would necessitate rapid changes in cAMP levels on a timescale of seconds. However, data on the dynamics of intracellular cAMP changes in photoreceptors remain scarce, primarily due to the limitations of conventional fluorescence-based methods in this specialized sensory system. To address this gap, we developed a methodology combining rapid cryofixation of retinal samples following light stimulation with the isolation of outer segment preparations. The rapid cryofixation setup comprises six computer-controlled sections, each with a high-speed stepper motor-driven lever that rapidly moves the specimen in a 180° arc within ~80 ms to press it against a liquid nitrogen-cooled copper cylinder for fixation. Using highly sensitive metabolomics techniques, we measured cAMP levels in these samples. This approach enables the investigation of rapid cAMP dynamics and its potential regulatory role in phototransduction, providing a foundation for understanding the interplay between cAMP and PKA signaling in photoreceptor function.

Rhamnogalacturonan-II (RG-II) is one of the least studied domains of pectin, primarily due to its low abundance, the lack of reliable antibodies, and the complexity of its structure. The present study builds upon existing protocols and procedures used to analyse RG-II in tissues where it is more abundant, combining and adapting them for the isolation of RG-II from Arabidopsis seed mucilage—a structure previously thought to lack RG-II. By applying these adapted methods, we first confirmed the presence of RG-II in seed mucilage and subsequently succeeded in isolating it from a tissue where it is typically present in low abundance, thereby enabling future studies on this previously overlooked component.

Phospholipids are major structural and regulatory elements of biological membranes and are involved in many different cellular and physiological processes. In this protocol, we provide an easy, cost-effective, and efficient method to obtain an overview of the phospholipid composition using high-performance thin layer chromatography (HPTLC). While the currently known phospholipid separation methods based on HPTLC display co-migration of certain lipid classes, the method we describe here allows the separation of all phospholipid classes, including anionic phospholipids in plant samples. This protocol combines elements of the classical Vitiello and Touchstone solvent systems to optimize phospholipid separation in a scaled pattern. Here, we provide a full characterization of this method, including statistical analyses of the retention factor of each phospholipid to show the robustness of the method and its efficiency in separating all phospholipid classes of a biological sample.

The Sox (SRY-related HMG-box) protein family plays a crucial role in cellular differentiation, development, and gene regulation, with the HMG (high-mobility group) domain responsible for DNA binding and transcriptional regulation. Proteins in the SOX gene family contain an HMG domain that shares 50% homology with the HMG domain of the sex-determining factor SRY gene. The SOX gene family comprises 30 proteins, which are classified into 10 groups (A–H). As a member of this family, hSox2 has been shown to be involved in various biological processes, but its specific function remains unclear. Previous studies have used eukaryotic expression systems, GST-tag purification, and bacterial inclusion body refolding techniques to produce Sox family proteins. However, these methods are often limited by issues such as low yield, incorrect folding, or inefficient purification, restricting their application in functional and structural studies. In this study, a prokaryotic expression system for the hSox2-HMG domain was constructed using the pET22b vector and Escherichia coli BL21(DE3) as the host strain. Protein expression was induced by IPTG, and initial purification was performed using Ni-NTA affinity chromatography, followed by ultrafiltration concentration and size exclusion chromatography to improve purity. By optimizing lysis and elution conditions, we successfully obtained hSox2-HMG protein with high expression levels and purity. This method provides a cost-effective and scalable strategy for hSox2-HMG production, ensuring high purity and correct folding of the protein. The optimized experimental protocol lays a foundation for structural and functional studies of hSox2-HMG.

Regulated IRE1-dependent decay (RIDD) is a critical cellular mechanism mediated by the endoplasmic reticulum (ER) stress sensor IRE1α, which cleaves a variety of RNA targets to regulate ER homeostasis. Current in vitro assays to study IRE1α activity largely rely on synthetic or in vitro transcribed RNA substrates, which may not fully replicate the physiological complexities of native RNA molecules. Here, we present a comprehensive protocol to assess IRE1α-dependent RNA cleavage activity using total RNA isolated directly from mouse tissues. This protocol provides a step-by-step guide for tissue collection, RNA isolation, an ex vivo RIDD assay, cDNA synthesis, and subsequent RT-PCR analysis of target mRNA cleavage products. Key reagents include active IRE1α protein, the RIDD-specific inhibitor 4μ8C, and target-specific primers for RIDD-regulated genes such asBloc1s1 and Col6a1. Quantitative assessment is achieved using agarose gel electrophoresis and imaging software. This methodology enables the study of IRE1α's RNA cleavage activity under conditions that closely mimic in vivo environments, providing a more physiologically relevant approach to understanding the role of RIDD in cellular and tissue-specific contexts.

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.

Zinc-finger (ZF) arrays are compact, sequence-specific polynucleotide-binding domains, which have been used to target the delivery of diverse effector domains, enabling applications such as gene identification, localization, regulation, and editing. To facilitate in vitro applications of ZF arrays, we have developed a general method for their expression and purification. Here, we describe a protocol involving two chromatographic steps that yields homogeneous and functional ZF arrays in milligram quantities.

Oxidative protein damage is important in various biological processes and age-related diseases. Protein carbonylation is the predominant and most frequently studied form of protein oxidation. It is most frequently detected following its derivatization with 2,4-dinitrophenylhydrazine (DNPH) hapten, followed by its detection with an anti-DNP antibody. However, when used to detect protein carbonylation by western blotting, this method suffers from diminished sensitivity, distortion of protein migration patterns, and unsatisfactory representation of low-abundance proteins. This is due to the poor solubility of DNPH in typical buffer solutions, the acidic protein precipitation due to the use of strong acid for its dissolution, the instability in solution, and the distorted protein migration patterns introduced by an additional salt content generated by the required pH adjustment prior to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). To address the DNPH method limitations, a new Oxime blot technique was developed. This method is based on forming the stable oxime bonds between the protein carbonyl groups and biotin-aminooxy probe in the presence of a p-phenylenediamine (pPDA) catalyst at neutral pH conditions. The derivatization reaction reaches a plateau within 3 h. It ensures efficient and complete derivatization of carbonylated proteins, which are separated by SDS-PAGE without additional manipulation and detected with avidin-HRP and enhanced chemiluminescence (ECL) in western blotting. The Oxime blot protocol allows researchers to reliably and sensitively detect carbonylated proteins and provides a valuable tool for studying oxidative stress in diverse biological settings.

Studying G protein-coupled receptor (GPCR) activation of heterotrimeric G proteins is crucial for understanding diverse physiological processes and developing novel therapeutics. Traditional methods to assay GPCR activation of G proteins, including assays of second messengers and biosensors, involve complex or indirect procedures. However, second messengers like cAMP and calcium are not direct readouts of GPCR activity due to signaling crosstalk, while biosensors can have undesired consequences due to structural alteration caused by fluorescent protein insertion. Here, we present a streamlined protocol employing GST-tagged bait proteins and epitope-embedded Gα subunits to achieve direct monitoring of Gα activity within cells. This method involves purification of GST-tagged bait constructs from bacteria and subsequent direct interaction studies with GluGlu-tagged Gα proteins expressed in any human cells of interest by including GST-tagged bait proteins in the cell lysis buffer. The approach enables sensitive detection of activated Gα within cells following extracellular stimulation. Advantages of this protocol include high sensitivity, enhanced monitoring of GPCR signaling dynamics under physiologically relevant conditions with minimum alteration in Gα, and the ability to distinguish between highly homologous isoforms within the same Gα family.