生物化学

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.

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.

This manuscript details protocols for the ZnCl₂ precipitation-assisted sample preparation (ZASP) for proteomic analysis. By inducing protein precipitation with ZASP precipitation buffer (ZPB, final concentration of ZnCl ₂ at 100 mM and 50% methanol), ZASP depletes harsh detergents and impurities, such as sodium dodecyl sulfate (SDS), Triton X-100, and urea, at high concentrations in solution from protein solutions prior to trypsin digestion. It is a practical, robust, and cost-effective approach for proteomic sample preparation. It has been observed that 90.2% of the proteins can be recovered from lysates by incubating with an equal volume of ZPB at room temperature for 10 min. In 1 h of data-dependent acquisition (DDA) analysis on an Exploris 480, 4,037 proteins and 25,626 peptides were quantified from 1 μg of mouse small intestine proteins, reaching a peak of 4,500 proteins and up to 30,000 peptides with 5 μg of input. Additionally, ZASP outperformed other common sample preparation methods such as sodium deoxycholate (SDC)-based in-solution digestion, acetone precipitation, filter-aided sample preparation (FASP), and single-pot, solid-phase-enhanced sample preparation (SP3). It demonstrated superior performance in protein (4,456 proteins) and peptide identification (29,871 peptides), lower missing cleavage rates (16.3%), and high reproducibility (Pearson correlation coefficients of 0.96 between replicates) with similar protein distributions and cellular localization patterns. Significantly, the cost of ZASP per sample with 100 μg of protein as input is lower than 30 RMB, including the expense of trypsin.

Cathepsin L (CTSL), a lysosomal cysteine protease belonging to the papain-like protease family, is primarily involved in intracellular protein degradation, antigen processing, and extracellular matrix remodeling. It plays critical roles in pathological conditions, including cancer metastasis, neurodegenerative disorders, and viral infection, due to dysregulated activity or overexpression. Thus, inhibitors targeting CTSL are under investigation for therapeutic applications. Current approaches for identifying CTSL inhibitors predominantly rely on fluorescence-labeled substrates, fluorescence resonance energy transfer (FRET), and cell-based screening assays. Here, we applied the principle of fluorescence polarization (FP) to the detection of substrate cleavage activity by CTSL through changes in millipolarization unit (mp) values and established a cost-effective, quantitative, reagent- and time-saving inhibitor high-throughput screening (HTS) assay. We also provide detailed steps for the expression and purification of highly active CTSL from eukaryotic cells, which lays a solid foundation for the FP-based assay. A key advantage of this assay lies in its reduced susceptibility to fluorescence interference, as the fluorescein isothiocyanate (FITC) fluorophore exhibits high quantum efficiency with an emission peak at 535 nm—a wavelength range distinct from most naturally occurring fluorescent molecules. The assay’s adaptability to reaction time, temperature, and dimethyl sulfoxide (DMSO) concentration minimizes false-positive or false-negative results caused by minor experimental inconsistencies, streamlining the screening process. Furthermore, the protocol requires fewer operational steps, reduced incubation time, and lower quantities of CTSL and substrates compared to conventional methods. This rapid, cost-effective, and scalable approach aligns well with the demands of HTS platforms.

Accurate measurement of protein translation rates is crucial for understanding cellular processes and disease mechanisms. However, existing methods for quantifying translation rates in yeast cells are limited. Here, we present a streamlined protocol for measuring protein translation rates in Saccharomyces cerevisiae using the methionine analog L-azidohomoalanine (AHA), which is the L isoform of this synthetic amino acid, and fluorophore-labeled alkyne dye-based Click chemistry. Our method involves incorporating AHA into newly synthesized proteins, followed by detection using confocal microscopy, flow cytometry, and SDS-PAGE. We validated our protocol by measuring translation rates under various stress conditions, including heat stress, endoplasmic reticulum (ER) stress induced by tunicamycin, and translation inhibition by cycloheximide. Confocal microscopy revealed differential AHA incorporation and fluorescence intensity across conditions. Flow cytometry quantitatively confirmed significant increases in translation rates under heat stress and decreases under ER stress compared to unstressed conditions at 6 and 24 h post-treatment. Imaging of gels under fluorescence detectors following SDS-PAGE further visualized newly synthesized proteins, with no detectable translation after cycloheximide treatment. Our protocol offers enhanced precision and selectivity compared to existing methods for mammalian cells and represents the first standardized approach for measuring translation rates in yeast. Despite limitations in required specialized equipment and expertise, this method holds promise for diverse applications in biotechnology and biomedical research, enabling investigations into protein synthesis regulation in yeast systems.