分子生物学

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.

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.

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 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.

Understanding protein–protein interactions is crucial for unravelling subcellular protein distribution, contributing to our understanding of cellular organisation. Moreover, interaction studies can reveal insights into the mechanisms that cover protein trafficking within cells. Although various techniques such as Förster resonance energy transfer (FRET), co-immunoprecipitation, and fluorescence microscopy are commonly employed to detect protein interactions, their limitations have led to more advanced techniques such as the in situ proximity ligation assay (PLA) for spatial co-localisation analysis. The PLA technique, specifically employed in fixed cells and tissues, utilises species-specific secondary PLA probes linked to DNA oligonucleotides. When proteins are within 40 nm of each other, the DNA oligonucleotides on the probes interact, facilitating circular DNA formation through ligation. Rolling-circle amplification then produces DNA circles linked to the PLA probe. Fluorescently labelled oligonucleotides hybridise to the circles, generating detectable signals for precise co-localisation analysis. We employed PLA to examine the co-localisation of dynein with the Kv7.4 channel protein in isolated vascular smooth muscle cells from rat mesenteric arteries. This method enabled us to investigate whether Kv7.4 channels interact with dynein, thereby providing evidence of their retrograde transport by the microtubule network. Our findings illustrate that PLA is a valuable tool for studying potential novel protein interactions with dynein, and the quantifiable approach offers insights into whether these interactions are changed in disease.

The identification and characterization of the ubiquitin E-ligase complexes involved in specific proteins’ degradation via the ubiquitin-proteasome system (UPS) can be challenging and require biochemical purification processes and in vitro reconstitution assays. Likewise, evaluating the effect of parallel phosphorylation and ubiquitination events occurring in vivo at dual phospho/ubiquitin-regulated motifs (called Phospho-Degrons or pDegrons) driving UPS degradation of the targeted protein has remained elusive. Indeed, the functional study of such E1-E2-E3 complexes acting on a protein-specific level requires previously or otherwise acquired knowledge of the nature of such degradation complex components. Furthermore, the molecular basis of the interaction between an E3 ligase and its pDegron binding motif on a target protein would require individually optimized in vitro kinase and ubiquitination assays. Here, we describe a novel enzymatically enhanced pull-down method to functionally streamline the discovery and functional validation of the ubiquitin E-ligase components interacting with a phospho-degron containing protein domain and/or sub-domain. The protocol combines key features of a protein kinase and ubiquitination in vitro assay by including them in a pull-down step exerted by a known or putative pDegron-tagged peptide using the cell extracts as a source of enzymatically active post-translational modification (PTM) modifying/binding native proteins. The same method allows studying specific stimuli or treatments towards the recruitment of the molecular degradation complex at the target protein’s phospho-degron site, reflecting in vivo–initiated events further enhanced through the assay design. In order to take full advantage of the method over traditional protein–protein interaction methods, we propose to use this PTM-enhanced (PTMe) pull down both towards the degradation complex discovery/ID phase as well as for the functional pDegron recruitment validation phase, which is the one described in the present protocol both graphically and in a stepwise fashion for reproduceable results.

Key features

• Suitable to study UPS-regulated (a) cytosolic and/or nuclear proteins, (b) intracellular region of transmembrane proteins, and (c) protein sub-domains bearing a known/putative pDegron motif.

• Requires a biotin-tagged recombinant version of the target protein and/or sub-domain.

• Allows the qualitative and quantitative analysis of endogenous ubiquitin (Ub) E-ligases recruitment to a known or putative pDegron bearing protein/sub-domain.

• Allows simultaneous testing of various treatments and/or conditions affecting the phosphorylative and/or ubiquitylation status of the studied pDegron bearing protein/sub-domain and the recruited factors.

Graphical overview

The centrosome governs many pan-cellular processes including cell division, migration, and cilium formation. However, very little is known about its cell type-specific protein composition and the sub-organellar domains where these protein interactions take place. Here, we outline a protocol for the spatial interrogation of the centrosome proteome in human cells, such as those differentiated from induced pluripotent stem cells (iPSCs), through co-immunoprecipitation of protein complexes around selected baits that are known to reside at different structural parts of the centrosome, followed by mass spectrometry. The protocol describes expansion and differentiation of human iPSCs to dorsal forebrain neural progenitors and cortical projection neurons, harvesting and lysis of cells for protein isolation, co-immunoprecipitation with antibodies against selected bait proteins, preparation for mass spectrometry, processing the mass spectrometry output files using MaxQuant software, and statistical analysis using Perseus software to identify the enriched proteins by each bait. Given the large number of cells needed for the isolation of centrosome proteins, this protocol can be scaled up or down by modifying the number of bait proteins and can also be carried out in batches. It can potentially be adapted for other cell types, organelles, and species as well.

Graphical overview

An overview of the protocol for analyzing the spatial protein composition of the centrosome in human induced pluripotent stem cell (iPSC)-derived neural cells. ① Human iPSCs are expanded, which serve as the starting cell population for the neural induction (Sections A, B, and C in Procedure). ② Neurons are induced and differentiated for 40 days (Section D in Procedure), in at least four biological replicates. ③ Total protein is isolated either at 15th or 40th day of differentiation, for neural stem cells and neurons, respectively (Sections E and F in Procedure). ④ Selected bait proteins are immunoprecipitated using the respective antibodies (Sections G and H in Procedure). ⑤ Co-immunoprecipitated samples are analyzed with mass spectrometry (Section I in Procedure). ⑥ Mass spectrometry output (.RAW) files are processed using MaxQuant software to calculate intensities (Section A in Data analysis). ⑦ The resulting data are pre-processed, filtered, and statistically analyzed using Perseus and R software (Sections B and C in Data analysis) ⑧ Further analysis is done using software or web tools such as Cytoscape or STRING to gain biological insights (Sections D and E in Data analysis).

The transmembrane receptor–ligand interactions play a vital role in the physiological and pathological processes of living cells, such as immune cell activation, neural synapse formation, or viral invasion into host cells. Mounting evidence suggests that these processes involve mechanosensing and mechanotransduction, which are directly mediated by the force-dependent transmembrane receptor–ligand interactions. Some single-molecule force spectroscopy techniques have been applied to investigate force-dependent kinetics of receptor–ligand interactions. Among these, the biomembrane force probe (BFP), a unique and powerful technique, can quantitatively and accurately determine the force-dependent parameters of transmembrane receptor–ligand interactions at the single-molecule level on living cells. The stiffness, spatial resolution, force, and bond lifetime range of BFP are 0.1–3 pN/nm, 2–3 nm, 1–10³ pN, and 5 × 10^-4–200 s, respectively. Therefore, this technique is very suitable for studying transient and weak interactions between transmembrane receptors and their ligands. Here, we share in detail the in situ characterization of the single-molecule force-dependent bond lifetime of transmembrane receptor–ligand interactions, based on a force-clamp assay with BFP.

Autophagy is an evolutionarily conserved intracellular degradation process. During autophagy, a set of autophagy-related (ATG) proteins orchestrate the formation of double-bound membrane vesicles called autophagosomes to engulf cytoplasmic material and deliver it to the vacuole for breakdown. Among ATG proteins, the ATG8 is the only one decorating mature autophagosomes and therefore is regarded as a bona fide autophagic marker; colocalization assays with ATG8 are wildly used as a reliable method to identify the components of autophagy machinery or autophagic substrates. Here, we describe a colocalization assay with fluorescent-tagged ATG8 using a tobacco (Nicotiana benthamiana)-based transient expression system.

Detecting protein-protein interactions (PPIs) is one of the most used approaches to reveal the molecular regulation of protein of interests (POIs). Immunoprecipitation of POIs followed by mass spectrometry or western blot analysis enables us to detect co-precipitated POI-binding proteins. However, some binding proteins are lost during cell lysis or immunoprecipitation if the protein binding affinity is weak. Crosslinking POI and its binding proteins stabilizes the PPI and increases the chance of detecting the interacting proteins. Here, we introduce the method of DSP (dithiobis(succinimidyl propionate))-mediated crosslinking, followed by tandem immunoprecipitation (FLAG and HA tags). The eluted proteins interacting with POI can be analyzed by mass spectrometry or western blotting. This method has the potential to be applied to various cytoplasmic proteins.

Graphical abstract: