Molecular Biology

Enzyme-catalyzed proximity labeling is a potent technique for the discernment of subtle molecular interactions and subcellular localization, furnishing contextual insights into the protein of interest within cells. Although ascorbate peroxidase2 (APEX2) has proven effective in this approach when overexpressed, its compatibility with endogenous proteins remains untested. We improved this technique for studying native protein–protein interactions in live Drosophila ovary tissue. Through CRISPR/Cas9 genome editing, APEX2 was fused with the endogenous dysfusion gene. By pre-treating the tissue with Triton X-100 to enhance biotin-phenol penetration, we successfully identified multiple proteins interacting with the native Dysfusion proteins that reside on the inner nuclear membrane. Our protocol offers a comprehensive workflow for delineating the interactome networks of ovarian components in Drosophila, aiding future studies on endogenous protein–protein interactions in various tissues of other animals.

Candida albicans is the most common human fungal pathogen, able to reside in a broad range of niches within the human body. Even though C. albicans systemic infection is associated with high mortality, the fungus has historically received relatively little attention, resulting in a lack of optimized molecular and fluorescent tools. Over the last decade, some extra focus has been put on the optimization of fluorescent proteins (FPs) of C. albicans. However, as the FPs are GFP-type, they require an aerobic environment and a relatively long period to fully mature. Recently, we have shown the application of a novel type of fluorogen-based FP, with an improved version of fluorescence activating and absorption shifting tag (iFAST), in C. albicans. Due to the dynamic relation between iFAST and its fluorogens, the system has the advantage of being reversible in terms of fluorescence. Furthermore, the combination of iFAST with different fluorogens results in different spectral and cellular properties, allowing customization of the system.

Progress in bioinformatics has facilitated the identification of a large number of putative glycosyltransferases (GTs) associated with many physiological processes. However, many of these GTs remain with unknown biochemical function due to numerous technical limitations. One of these limitations is the lack of innovative tools for large-scale screening of enzyme activity in vitro and testing protein–protein interactions (PPIs) between GT partners. Currently, testing the enzyme activity of a protein requires its production in a heterologous expression system and purification before enzyme assays, a process that is time-consuming and not amenable to high-throughput screening. To overcome this, we developed a platform called in vitro GT-array (i-GT-ray). In this platform, 96-well microplates are coated with plasmid DNA encoding for tagged GTs and a capture antibody. Tagged GTs are produced from plasmid DNA via a cell-free in vitro transcription/translation (IVTT) system and captured through the anti-tag capture antibody directly on microplates. After washing to remove IVTT components, the captured enzymes can be considered purified, and their activity can be tested directly on microplates. The whole process can be performed in less than two days, compared to several weeks for currently available screening methods. The i-GT-ray platform has also been adapted to investigate PPIs between GTs. Here, we provide a practical user guide for the preparation of GT-arrays coated with plasmid DNA and a capture antibody that can be used for monitoring enzyme activity and PPIs of GTs in a high-throughput manner.

Calcium channels at synaptic boutons are critical for synaptic function, but their number and distribution are poorly understood. This gap in knowledge is primarily due to the resolution limits of fluorescence microscopy. In the last decade, the diffraction limit of light was surpassed, and fluorescent molecules can now be localized with nanometer precision. Concurrently, new gene editing strategies allowed direct tagging of the endogenous calcium channel genes—expressed in the correct cells and at physiological levels. Further, the repurposing of self-labeling enzymes to attach fluorescent dyes to proteins improved photon yields enabling efficient localization of single molecules. Here, we describe tagging strategies, localization microscopy, and data analysis for calcium channel localization. In this case, we are imaging calcium channels fused with SNAP or HALO tags in live anesthetized C. elegans nematodes, but the analysis is relevant for any super-resolution preparations. We describe how to process images into localizations and protein clusters into confined nanodomains. Finally, we discuss strategies for estimating the number of calcium channels present at synaptic boutons.

Bottom-up proteomics utilizes sample preparation techniques to enzymatically digest proteins, thereby generating identifiable and quantifiable peptides. Proteomics integrates with other omics methodologies, such as genomics and transcriptomics, to elucidate biomarkers associated with diseases and responses to drug or biologics treatment. The methodologies employed for preparing proteomic samples for mass spectrometry analysis exhibit variability across several factors, including the composition of lysis buffer detergents, homogenization techniques, protein extraction and precipitation methodologies, alkylation strategies, and the selection of digestion enzymes. The general workflow for bottom-up proteomics consists of sample preparation, mass spectrometric data acquisition (LC-MS/MS analysis), and subsequent downstream data analysis including protein quantification and differential expression analysis. Sample preparation poses a persistent challenge due to issues such as low reproducibility and inherent procedure complexities. Herein, we have developed a validated chloroform/methanol sample preparation protocol to obtain reproducible peptide mixtures from both rodent tissue and human cell line samples for bottom-up proteomics analysis. The protocol we established may facilitate the standardization of bottom-up proteomics workflows, thereby enhancing the acquisition of reliable biologically and/or clinically relevant proteomic data.

Membrane proteins play critical roles in cell physiology and pathology. The conventional way to study membrane proteins at protein levels is to use optimal detergents to extract proteins from membranes. Identification of the optimal detergent is tedious, and in some cases, the protein functions are compromised. While this detergent-based approach has produced meaningful results in membrane protein research, a lipid environment should be more suitable to recapture the protein’s native folding and functions. This protocol describes how to prepare amphipathic membrane scaffold-proteins (MSPs)-based nanodiscs of a cation-coupled melibiose symporter of Salmonella enterica serovar Typhimurium (MelB_St), a member of the major facilitator superfamily. MSPs generate nano-assemblies containing membrane proteins surrounded by a patch of native lipids to better preserve their native conformations and functions. This protocol requires purified membrane protein in detergents, purified MSPs in solution, and detergent-destabilized phospholipids. The mixture of all three components at specific ratios is incubated in the presence of Bio-Beads SM-2 resins, which absorb all detergent molecules, allowing the membrane protein to associate with lipids surrounded by the MSPs. By reconstituting the purified membrane proteins back into their native-like lipid environment, these nanodisc-like particles can be directly used in cryo-EM single-particle analysis for structure determination and other biophysical analyses. It is noted that nanodiscs may potentially limit the dynamics of membrane proteins due to suboptimal nanodisc size compared to the native lipid bilayer.

The cellular thermal shift assay (CETSA) and isothermal dose-response fingerprint assay (ITDRF _CETSA) have been introduced as powerful tools for investigating target engagement by measuring ligand-triggered thermodynamic stabilization of cellular target proteins. Yet, these techniques have rarely been used to evaluate the thermal stability of RNA-binding proteins (RBPs) when exposed to ligands. Here, we present an adjusted approach using CETSA and ITDRF_CETSA to determine the interaction between enasidenib and RBM45. Our assay is sensitive and time-efficient and can potentially be adapted for studying the interactions of RBM45 protein with other potential candidates.

The cell–cell adhesion molecule E-cadherin has been intensively studied due to its prevalence in tissue function and its spatiotemporal regulation during epithelial-to-mesenchymal cell transition. Nonetheless, regulating and studying the dynamics of it has proven challenging. We developed a photoswitchable version of E-cadherin, named opto-E-cadherin, which can be toggled OFF with blue light illumination and back ON in the dark. Herein, we describe easy-to-use methods to test and characterise opto-E-cadherin cell clones for downstream experiments.

The assessment of peptide–protein interactions is a pivotal aspect of studying the functionality and mechanisms of various bioactive peptides. In this context, it is essential to employ methods that meet specific criteria, including sensitivity, biocompatibility, versatility, simplicity, and the ability to offer real-time monitoring. In cellular contexts, only a few proteins naturally possess inherent fluorescence, specifically those containing aromatic amino acids, particularly tryptophan. Nonetheless, by covalently attaching fluorescent markers, almost all proteins can be modified for monitoring purposes. Among the early extrinsic fluorescent probes designed for this task, dansyl chloride (DNSC) is a notable option due to its versatile nature and reliable performance. DNSC has been the primary choice as a fluorogenic derivatizing reagent for analyzing amino acids in proteins and peptides for an extended period of time. In our work, we have effectively utilized the distinctive properties of dansylated-calmodulin (D-CaM) for monitoring the interaction dynamics between proteins and peptides, particularly in the context of their association with calmodulin (CaM), a calcium-dependent regulatory protein. This technique not only enables us to scrutinize the affinity of diverse ligands but also sheds light on the intricate role played by calcium in these interactions.

Key features

• Dynamic fluorescence and real-time monitoring: dansyl-modified CaM enables sensitive, real-time fluorescence, providing valuable insights into the dynamics of molecular interactions and ligand binding.

• Selective interaction and stable fluorescent adducts: DNSC selectively interacts with primary amino groups, ensuring specific detection and forming stable fluorescent sulfonamide adducts.

• Versatility in research and ease of identification: D-CaM is a versatile tool in biological research, facilitating identification, precise quantification, and drug assessment for therapeutic development.

• Sensitivity to surrounding alterations: D-CaM exhibits sensitivity to its surroundings, particularly ligand-induced changes, offering subtle insights into molecular interactions and environmental influences.

Graphical overview

Fluorescence emission profiles of dansylated-calmodulin (D-CaM) in different states. Fluorescence emission spectra of D-CaM upon excitation at 320 nm are depicted. Conditions include apo-D-CaM (gray), holo-D-CaM (red), apo-D-CaM bound to peptide (blue), and holo-D-CaM bound to peptide (purple). Corresponding structural representations of D-CaM next to each condition are superimposed on the respective spectra along with the hydrophobicity of the dansyl environment, which increases upon binding of peptide or Ca²⁺ to D-CaM. Upon peptide binding to D-CaM, there is an enhancement in the fluorescent intensity of the spectra; upon Ca²⁺ binding, there is an enhancement of the intensity and a leftward shift of the spectra.

Camelina sativa, a Brassicaceae family crop, is used for fodder, human food, and biofuels. Its relatively high resistance to abiotic and biotic stresses, as well as being a climate-resilient oilseed crop, has contributed to its popularity. Camelina's seed yield and oil contents have been improved using various technologies like RNAi and CRISPR/Cas9 genome editing. A stable transformation system for protein localization and other cell autonomous investigations, on the other hand, is tedious and time consuming. This study describes a transient gene expression protocol for Camelina sativa cultivar DH55 leaves using Agrobacterium strain C58C1. The method is suitable for subcellular protein localization and colocalization studies and can be used with both constitutive and chemically induced genes. We report the subcellular localization of the N-terminal ER membrane signal anchor region (1–32 aa) of the At3G28580 gene-encoded protein from Arabidopsis in intact leaves and the expression and localization of other known organelle markers. This method offers a fast and convenient way to study proteins in the commercially important Camelina crop system.

Key features

• This method is based on the approach of Zhang et al. [1] and has been optimized for bioenergy crop Camelina species.

• A constitutive and inducible transient gene expression in the hexaploid species Camelina sativa cultivar DH55.

• Requires only 16–18 days to complete with high efficacy.

Graphical overview

Agrobacterium-mediated transient gene expression optimized for Camelina sativa