生物化学

Liquid–liquid phase separation (LLPS) underlies the spatial organization of the nucleolus, a membraneless organelle responsible for ribosomal RNA (rRNA) transcription and ribosome subunit assembly. One of the key proteins involved in the formation of the fibrillar center of the nucleolus is the treacle, an intrinsically disordered protein that contains low-complexity repeats enriched in charged amino acid residues. In this work, we present a detailed protocol for the bacterial expression and purification of a recombinant fragment of treacle comprising two tandem low-complexity repeat (LCR) modules, with a total length of 136 amino acids. This fragment is intended for subsequent in vitro investigation of its ability to undergo LLPS. The described method enables the production of a soluble, biochemically pure protein preparation suitable for studying the mechanisms of spontaneous condensate formation in a cell-free system. This approach allows for the controlled modeling and quantitative evaluation of the contribution of low-complexity sequences to the phase behavior of treacle, independently of its interactions with cellular partners in vivo.

Candida albicans is the pathogenic fungus that most frequently causes infections in humans. It is part of the microbiota commonly found in the skin, gastrointestinal tract, and vaginal mucosa. However, certain conditions, including immunosuppression, excessive use of antibiotics, hormonal changes, the use of medical devices in patients, and individual nutritional status, promote the development of opportunistic infections caused by this fungus. One of the main fungal structures interacting with the host is the cell wall, which is principally composed of chitin, glucan, and proteins. The cell wall plays key functions for the cell, such as osmotic protection; it is also responsible for cellular shape and acts as a signaling hub in response to environmental changes. Cell wall proteins participate in diverse cellular functions, such as attachment to surfaces and cell wall structure; some possess catalytic or transport activities. In this protocol, we show the methodology for isolating cell wall proteins covalently linked or not to cell wall components that can be previously labeled with [¹⁴C]-L-lysine by the action of the fungal transglutaminase localized in the cell wall. We use an extraction method by mechanical cell disruption and washing with 2 M NaCl, whose ionic strength eliminates contaminating proteins from other organelles, through subsequent serial treatments with SDS, chitinase, and zymolyase.

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.

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.

The CRISPR-Cas system of Thermus thermophilus has emerged as a potent biotechnological tool, particularly its Cas6 endonuclease, which plays a crucial role in CRISPR RNA (crRNA) maturation. This protocol details a robust and reproducible method for the high-level expression and purification of recombinant T. thermophilus Cas6 proteins (Cas6-1 and Cas6-2) in E. coli. We describe a streamlined approach encompassing plasmid construction using seamless assembly, optimized bacterial heterologous expression, and multi-step purification leveraging affinity and size-exclusion chromatography. The protocol outlines the generation of both His-tagged and GST-tagged Cas6 variants, enabling flexibility in downstream applications. Key steps, including primer design, PCR optimization, competent cell transformation, and chromatography strategies, are meticulously detailed with critical parameters and troubleshooting guidance to ensure experimental success and high yields of highly pure and active T. thermophilus Cas6 proteins. This protocol is useful for researchers requiring purified T. thermophilus Cas6 for structural studies, biochemical characterization, and the development of CRISPR-based biotechnological tools.

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.

Protein purification is essential for drug development, antibody production, and structural biology. We propose a cost-effective chromatography method using elastin-like polypeptide (ELP) as an aggregating core. In this approach, a chilled (target protein)-ELP fusion is loaded onto an immobilized metal affinity chromatography (IMAC) column equilibrated with low-salt buffer. Impurities are removed with warm high-salt buffer washes. Warming the column above the ELP’s transition temperature (T_m) triggers ELP aggregation, physically trapping the target protein between beads. Subsequent stringent washing (high salt/imidazole) eliminates residual contaminants. Finally, cooling with cold low-salt buffer reverses aggregation, eluting the purified target protein. This method eliminates the need for advanced chromatography systems while achieving high purity through dual mechanisms: (1) IMAC affinity and (2) temperature-dependent physical capture. The ELP’s reversible phase transition offers a simplified yet efficient purification platform, particularly valuable for lab-scale production of challenging proteins.

The voltage-gated proton channel (H_v1) is a membrane protein that dissipates acute cell proton accumulations. To understand the molecular mechanisms explaining H_v1 function, methods for purifying the protein are needed. Previously, methods were developed for expressing and purifying human H_v1 (hH_v1) in yeast and later in bacteria. However, these methodologies produced low protein yields and had high production costs, considerably limiting their usefulness. The protocol described in this work was developed to overcome those limitations. hH_v1 is overexpressed in bacteria, solubilized with the detergent Anzergent 3–12, and purified by immobilized metal affinity chromatography (IMAC) and size-exclusion chromatography (SEC). Our protocol produced higher protein yields at lower costs than previously published methodologies.

De novo synthesis of purine nucleotide is essential for the production of genetic materials and cellular chemical energy. PRPP amidotransferase (PPAT) is the rate-limiting enzyme in de novo purine synthesis, thereby playing a crucial regulatory role in this pathway. Recent studies suggest that metabolic enzymes, including PPAT, form condensates through phase separation to regulate cellular metabolism in response to environmental changes. However, due to the lack of methods for purifying eukaryotic PPAT, the biophysical properties of the enzyme have remained unknown. Here, I describe a protocol for purifying budding yeast PPAT tagged with green fluorescent protein from yeast cells, as well as an in vitro assay to examine condensation of the fluorescent PPAT by microscopy. These techniques enabled us to elucidate the mechanism controlling PPAT condensation and may also be applicable to the purification and condensation assay of other enzymes.