Targeted protein degradation is an emerging concept of drug discovery to selectively eliminate the pathogenic proteins by activating their degradation in cells. The proteolysis-targeting chimeras (PROTACs) are bifunctional small molecules that induce the degradation of a protein of interest (POI) by proteasome. Here we describe a method that utilizes an ssrA-based BacPROTAC (bacterial PROTACs) to target the drug-resistant proteins CTX-M-14, a class A β-lactamase commonly found in extended-spectrum beta-lactamase (ESBL) plasmids of antimicrobial resistance (AMR) bacteria. This strategy could resensitize the drug-resistant bacteria and revive previously disregarded antibiotics, opens up a new avenue for therapeutic development of AMR bacteria.

Many organisms possess N-degron pathways that link the proteolytic stability of individual proteins to the identity of their N-terminal amino acid. The set of N-degrons recognized by proteolytic machinery, and the regulatory paradigms leading to N-degron exposure, have been well studied in Escherichia coli and several other bacterial species. However, N-degron landscapes remain unexplored in the majority of bacterial phyla, and the extent to which these pathways vary among clades remains unknown. To characterize N-degron proteolysis in Mycolicibacterium smegmatis, a member of Actinomycetota and a model system for pathogenic mycobacteria, we devised a cell-based assay incorporating a dual-fluorescence reporter. We evaluated the proteolysis of mCherry substrates with defined N-terminal sequences by comparing fluorescence levels to a stable GFP reference in plate reader and flow cytometry contexts. By examining proteolysis patterns in wild-type M. smegmatis and cells lacking the N-recognin ClpS, we established that canonical primary destabilizing residues (Leu, Phe, Tyr, Trp) are subject to ClpS-dependent N-degron proteolysis. This dual-fluorescence approach is broadly applicable to interrogation of N-degron pathways across bacterial species.

Fluorescent reporters provide a useful tool for studying degron motifs. Fusing a degron of interest to a fluorescent protein allows to accurately track protein levels overtime to characterize the degradation kinetics of studied degrons. Here, we describe a rapid and simple method to study degron peptides in Escherichia coli using plasmid-encoded eGFP-degron fusion constructs. The described methods provide an accessible workflow to evaluate degrons. We provide protocols for generation of pBAD plasmids encoding the studied constructs and two different methods for evaluating degrons: an end-point fluorescence measurement on agar plates and a kinetic measurement in liquid cultures in a 96-well format for high-throughput degron studies.

The ssrA degron is widely employed in fusion proteins to regulate protein stability in bacteria or to serve as an interaction module. These uses take advantage of the ssrA tag's dual functionality-its ability to bind the SspB adaptor and to recruit the ClpXP protease. A comprehensive comparison of these functions across a standardized set of degron variants can be helpful in the development of variants specifically optimized for particular applications and their use in targeted protein degradation systems. Here, we describe a simple and efficient purification protocol for a set of ssrA-based degrons and a degradation assay that can be used as the primary assay to compare the effectiveness of degradation of degron-tagged proteins.

Degrade green fluorescent protein (deGradFP) is a degron method that was originally developed in Drosophila melanogaster. It targets GFP-fusion proteins to an endogenous ubiquitin-proteasome pathway using an anti-GFP nanobody. We have adapted it to a kinetoplastid parasite Trypanosoma brucei to rapidly degrade kinetochore proteins that have low turnover rates. Here, we describe the workflow of deGradFP in T. brucei.

The ability to specifically degrade one protein in a cell provides an immediate insight into any resulting phenotype and thus function that may be occluded by secondary effects using other techniques. The auxin-inducible degron is a method developed to achieve this end and is effective in mammalian and yeast cells. The approach is dependent on the recognition of a tagged target protein by an endogenous E3 ubiquitin ligase modified by the expression of a plant F-box protein. This approach was initially unsuccessful in trypanosomes, probably due to a lack of interaction between the endogenous E3 ubiquitin ligase and the plant F-box protein. This was overcome by expressing the entire rice E3 ubiquitin ligase in trypanosomes, and here we describe the production of the auxin-inducible degron competent cell lines and tagging of target genes.

The auxin-inducible degradation (AID) system is a powerful tool in modern molecular genetics that allows for conditional protein depletion. In Caenorhabditis elegans, tools exist to allow for rapid, tissue-specific depletion of target proteins. The system requires tagging a gene of interest with an AID degron and a transgene expressing the Arabidopsis thaliana TIR1 F-box protein, which can form a functional SCF ubiquitin ligase with endogenous Skp and Cullin proteins. Here, I describe how to generate degron knock-ins by CRISPR, cross them to TIR1-expressing strains, and perform depletion experiments. I provide a description of our current methods and highlight alternative approaches.

The endoplasmic reticulum (ER) is the site where proteins that are synthesized and destined for the secretory pathway fold into their native conformations. Genetic mutations, oxidative stress, reduced glycosylation, and disruption to ER folding and quality control systems are all factors that have been demonstrated to impair this process, leading to the production of misfolded proteins. It is imperative for the survival of cells that these misfolded proteins are recognized by specific signals and directed towards degradation via ER-associated degradation (ERAD), ER-phagy, or related ER-to-lysosome/vacuole transport processes. A pivotal step in this process is the recognition of specific degrons and the distinction between terminally misfolded proteins that necessitate clearance and folding intermediates that have the potential to reach their final conformation. A significant proportion of proteins that traverse the secretory pathway in eukaryotes undergo N-glycosylation. The attached N-glycans not only facilitate the folding and quality control of glycoproteins, but can also be processed to display a glycan degron signal necessary to initiate ERAD of glycoproteins. Here, we describe methods for investigating glycan-related degrons that are key determinants for the ERAD of misfolded glycoproteins in plants.

Transcription factors are an exciting class of drug targets. Recent developments in small molecule-induced proximity allow for small molecule control of their activity by recruitment of epigenetic proteins. In this chapter, we describe the development of a transcriptional activity assay for the transcription factor BCL6 with a small molecule activator, TCIP1, in HEK293T cells [1]. The assay is based on the Bio-Glo Luciferase assay system and uses transient transfection of a BCL6-expressing plasmid, as well as a reporter plasmid where a BCL6 DNA-binding sequence controls expression of a luciferase protein. The method described here is specific to BCL6 but can easily be optimized to monitor the activity of other transcription factors by altering the DNA-binding sequence of the reporter.

Precise modulation of protein levels is essential for investigating the dynamics of signaling pathways. Degradation-based approaches allow rapid reduction of protein abundance, and their combination with optogenetic regulation enables controlled and reversible manipulation. In this chapter, we describe an optogenetic strategy that stabilizes a protein of interest by inhibiting its degradation. As a proof of concept, we demonstrate light-dependent stabilization of MKP3 and constitutively active MEK and examine the resulting changes in ERK signaling. We also provide a step-by-step protocol for measuring protein stabilization kinetics. This method integrates a potent degradation tag with optogenetic control to achieve tunable and reversible regulation of protein levels in living cells.

mRNA molecules are critical for relaying genetic information from the genome into functional proteins for maintaining cellular function. Real-time imaging of mRNA in living cells reveals the spatiotemporal dynamics of RNA and offers insights into RNA-mediated regulation of gene expression and cellular function. Here, we describe a method of using RNA-regulated destabilization domains to simultaneously image multiple mRNAs in living cells. In this method, three mRNAs of interest are engineered to contain three distinct RNA imaging tags. Each tag specifically binds to and stabilizes its cognate RNA-regulated destabilization domain fused to fluorescent proteins, resulting in selective activation of fluorescence on the corresponding mRNA for imaging.

Scientific research in drug discovery relies on robust, versatile, and well-established methodologies. One effective strategy for targeting disease-relevant proteins, such as E3 ligases, involves rational drug design using degron peptides as starting points for compound development. Here, we present a comprehensive set of complementary assays for the evaluation of degron-based binders, enabling their integration into an efficient drug discovery pipeline. We introduce screening strategies such as Differential Scanning Fluorimetry (DSF) and Fluorescence Polarization (FP), alongside both solution-based and immobilization-based biophysical techniques, including Isothermal Titration Calorimetry (ITC) and Surface Plasmon Resonance (SPR) for reliable affinity determination. To assess intracellular interactions with full-length target proteins, we also employ Nano Bioluminescence Resonance Energy Transfer (NanoBRET). Together, these methods establish a robust framework for the discovery and characterization of degron-based peptidomimetic compounds.

Activity-based ubiquitin probes serve as crucial tools for capturing and elucidating the molecular mechanisms of E3 ligase-catalyzed substrate ubiquitination. We recently reported a chemoenzymatic strategy for generating ubiquitin chain extension probes, which offers advantages such as mild reaction conditions, simple operation, and high specificity, thereby providing new opportunities for E3 enzyme research. This article details the experimental protocol for obtaining a K48-linked di-ubiquitin extension probe based on a substrate-Type II degron using the chemoenzymatic approach.

The C-terminal cyclic imide modification is recognized as a degron by the E3 ligase adapter cereblon (CRBN). Reliable methods to generate and measure this modification on CRBN substrates are essential for studying its biological function. In this chapter, we describe two complementary approaches for producing C-terminal cyclic imide-modified peptides and proteins: sortase-mediated transpeptidation and enzymatic conversion by protein carboxymethyltransferase (PCMT1). Additionally, we present analytical strategies, including a time-resolved Förster resonance energy transfer (TR-FRET) assay for measuring binding of substrates to CRBN in vitro and mass spectrometry methods for confirming cyclic imide formation. Together, these protocols provide practical methods for investigating the chemistry and biology of C-terminal cyclic imides.

Ubiquitination is a complex post-translational modification that regulates a wide range of cellular processes through the covalent attachment of ubiquitin to substrate proteins. While canonical ubiquitination occurs on lysine residues, non-canonical modifications on serine, threonine, and the protein N-terminus are increasingly recognized. This chapter describes the UbiSite approach, a refined strategy for site-specific mapping of global ubiquitination using mass spectrometry-based proteomics. The method relies on digestion of the proteome of interest with LysC endopeptidase. LysC digestion leaves a 13 amino acid remnant on the ubiquitinated site of modified peptides, and this remnant is recognized by the monoclonal UbiSite antibody allowing selective enrichment of ubiquitinated peptides. Tens of thousands modification sites are readily identified from cells or tissue extracts in a single UbiSite experiment with the workflow described here. The 13 amino acid remnant is unique to ubiquitin and the UbiSite antibody thereby discriminates ubiquitin from other ubiquitin-like modifiers and enables detection of both canonical and non-canonical ubiquitination. The UbiSite procedure can be automated and combined with label-free quantification as well. UbiSite offers a robust and versatile platform for comprehensive ubiquitinome profiling with enhanced specificity and depth, enabling new insights into the regulatory roles of ubiquitin across diverse biological contexts.