Biochemistry

Peptergent is a novel class of amphipathic peptides that enables detergent-free extraction of membrane proteins (MPs) from lipid bilayers. This reagent self-assembles around hydrophobic transmembrane regions, forming stable, water-soluble complexes that can be isolated directly from biological membranes. Peptergent therefore bypasses the limitations imposed by traditional detergents, which often destabilize protein assemblies. Since detergents are completely avoided, MPs are directly amenable to structural and mass spectrometry (MS) analysis, thereby addressing their persistent underrepresentation in proteomic datasets and improving their accessibility in drug-screening strategies. We present here a streamlined protocol for MPs extraction with the Peptergent PDET-1, followed by exchange into His-tagged Peptidiscs for Ni-NTA-based affinity purification. The method encompasses membrane isolation, peptide preparation, protein extraction, clarification, and MPs exchange from Peptergents to Peptidiscs. This workflow yields an enriched membrane proteome compatible with downstream LC-MS/MS analysis for improved identification of multi-pass MPs.

Bottom-up proteomics workflows encompass several key stages, including sample preparation, data acquisition, and data analysis. Of these, sample preparation is the initial and critical stage, as it significantly influences the depth, reproducibility, and reliability of subsequent mass spectrometry–based analyses. While several main digestion strategies exist, including in-gel, in-solution, and filter-aided methods, each presents distinct trade-offs in terms of throughput, contamination removal, and applicability to complex biological matrices. The Suspension Trapping (S-Trap) method offers a compelling alternative by efficiently capturing and digesting proteins while removing interferents like sodium dodecyl sulfate (SDS), which can compromise downstream LC–MS/MS performance. This protocol details a S-Trap workflow optimized for biofluid proteomics, specifically plasma, serum, and cerebrospinal fluid (CSF). We describe two complementary formats: a manual tube-based procedure for individual or small-batch samples and a 96-well-plate-based system enabling high-throughput processing. The protocol integrates optional high-abundance protein depletion to enhance coverage of low-abundance analytes and includes steps for reduction, alkylation, digestion, and peptide elution for low total protein content samples, such as plasma, serum, and cerebrospinal fluid. By providing a detailed protocol, this work aims to improve the consistency and accessibility of S-Trap-based sample preparation, facilitating robust and reproducible discoveries in bottom-up proteomics.

Extracellular vesicles (EVs) are nanoscale particles secreted by all cells and present in all biological fluids, where they carry molecular cargo reflective of health and disease states. Their diagnostic potential is often obscured by the high abundance of non-EV proteins and lipoproteins (e.g., albumin, apolipoproteins) that complicate proteomic analysis of primary biofluids, such as ascites fluid. Conventional isolation strategies face a persistent trade-off between EV purity and yield. To overcome this, a magnetic bead-based protocol (Mag-Net) to enrich EVs according to electrochemical surface charge using strong anion-exchange chemistry (SAX) was adapted for proteomics. Our workflow is specifically adapted to ascites fluid from human or murine sources. This approach effectively separates EVs from high-abundance proteins and lipoproteins, enabling proteomic profiling from as little as 2 μL of ascites fluid. Demonstrated in both murine and human ovarian cancer models, Mag-Net offers a reproducible, scalable, and automation-ready solution for EV isolation from various biofluids.

The CRISPR/Cas9 system is a cornerstone technology in genome editing. Delivery of pre-assembled Cas9 ribonucleoprotein (RNP) complexes exhibits distinct advantages, including reduced off-target effects and lower immunogenicity. Conventional methods for purifying Cas9 protein typically involve multi-step chromatography and the cleavage of fusion tag, which are time-consuming and result in diminished yields. In this study, we present a simplified, one-step purification strategy for functional Streptococcus pyogenes Cas9 (SpCas9) using the ubiquitin (Ub) fusion system in Escherichia coli. The N-terminal Ub fusion not only improves protein solubility but also facilitates high-yield production of the His-Ub-Cas9 fusion protein. Importantly, the Ub tag does not require proteolytic removal during purification, allowing direct one-step purification of the fusion protein via nickel-affinity chromatography. The purified His-Ub-Cas9 retains robust DNA cleavage activity in vivo, as validated in zebrafish embryos. This protocol greatly simplifies the production of functional Cas9 protein, facilitating its broad application in genome editing.

This article presents an efficient protocol for refolding recombinant proteins that are prone to aggregation and form inclusion bodies during expression in Escherichia coli. As a model system, the homolog of CRISPR-associated effector protein CasV-M was investigated. The key element of the developed approach is refolding directly on a metal-affinity Ni-TED (N,N,N´-tris(carboxymethyl)ethylendiamine) resin using a dual-gradient system: a stepwise reduction in the concentration of the chaotropic agent combined with a simultaneous increase in the concentration of a mild nonionic detergent. This combination ensures spatial separation of protein molecules, minimizes aggregation, and promotes the recovery of the native conformation. The resulting method appears to be an alternative to conventional refolding strategies, with potential improvements in the reproducibility and yield of soluble protein compared to dialysis or dilution. The proposed approach can be extended to a broad range of aggregation-prone proteins and is considered a promising strategy for obtaining otherwise insoluble recombinant proteins.

The cellular secretome is a rich source of biomarkers and extracellular signaling molecules, but proteomic profiling remains challenging, especially when processing culture volumes greater than 5 mL. Low protein abundance, high serum contamination, and sample loss during preparation limit reproducibility and sensitivity in mass spectrometry–based workflows. Here, we present an optimized and scalable protocol that integrates (i) 50 kDa molecular weight cutoff ultrafiltration, (ii) spin column depletion of abundant serum proteins, and (iii) acetone/TCA precipitation for protein recovery. This workflow enables balanced recovery of both low- and high-molecular-weight proteins while reducing background from serum albumin, thereby improving sensitivity, reproducibility, and dynamic range for LC–MS/MS analysis. Validated in human mesenchymal stromal cell cultures, the protocol is broadly applicable across diverse cell types and experimental designs, making it well-suited for biomarker discovery and extracellular proteomics.

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.