The chromatin immunoprecipitation followed by sequencing (ChIP-seq) assay is an instrumental and accurate method for understanding chromatin dynamics in eukaryotic cells. It provides critical insights into the regulation of gene expression and enables identification of regulatory elements, patterns of histone modifications, and chromatin states in health and disease conditions. Although cell cultures are great models to study molecular mechanisms associated with pathologies, studying tissues provides a physiologically native environment that reflects the cellular heterogeneity and spatial organization that are missing in an in vitro model. Several ChIP-seq protocols have been published; however, performing ChIP-seq in tissues remains a challenge in many settings due to the heterogeneity of tissues, complexity of cell matrices, low input material and intricacy of chromatin fragmentation and handling. Here, we present an optimized ChIP-seq protocol for solid tissues, with a focus on colorectal cancer. In this article, we incorporate simplified and efficient procedures for tissue preparation, chromatin extraction, immunoprecipitation, and library construction. The refined protocols overcome common limitations related to tissue processing and allows for highly reproducible, sensitive, and scalable analysis of disease-relevant chromatin states in vivo. © 2025 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol 1: Frozen tissues preparation Basic Protocol 2: Chromatin immunoprecipitation from tissues Basic Protocol 3: Library construction for DNA sequencing Basic Protocol 4: DNA nanoballs preparation for the DNBSEQ-G99RS sequencing platform and data quality control.
Histone deacetylases (HDACs) and sirtuins (SIRTs) play essential roles in regulating chromatin structure and gene expression by catalyzing the removal of acyl groups from histone lysine residues. Accurate characterization of their deacetylation kinetics is critical for understanding their enzymatic mechanisms and for guiding inhibitor or activator development. Given the complexity of enzyme-nucleosome core particle (NCP) interactions, including the influence of histone composition, post-translational modifications, and DNA context, NCP-based assays provide a more physiologically relevant platform than those performed on peptide or free histone substrates. Here, we present optimized protocols for assessing HDAC and SIRT deacetylation kinetics using NCP substrates, including determination of Michaelis-Menten parameters, evaluation of inhibitor and activator potency (IC50 or EC50), and instructions for ensuring assay reproducibility. These methods enable robust comparison of small-molecule modulators under conditions that better mimic the native chromatin environment, supporting both mechanistic studies and drug discovery efforts. © 2025 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol 1: Assay of HDAC complex or SIRT deacylation on nucleosome substrates Basic Protocol 2: Assay of HDAC complex or SIRT deacylation on free histone proteins Basic Protocol 3: KM measurement for deacylation assay on nucleosome or cofactor Basic Protocol 4: Assessment of deacylation inhibitor or activator effects on NCP substrates.
Targeted protein degradation (TPD) has revolutionized the way we think of drug discovery and has the potential for substantial therapeutic benefits. Traditional mechanisms rely on taking advantage of the cells endogenous protein degradation pathway known as the ubiquitin proteasome system (UPS). Traditional proteolysis targeting chimeras (PROTACs) rely on this mechanism by developing heterobifunctional molecules, which are compounds that contain two different ligands that bind two different proteins linked together with varying linker lengths. These compounds typically contain a ligand to a protein of interest that is to be degraded and a linker to the other ligand that binds to an E3 ligase. Once these compounds bind both proteins of interest with the proper confirmation, the E-ligase complex can facilitate the ubiquitination of the protein, leading to its recognition by the proteasome for degradation. This approach has been effective at developing degraders for a wide variety of proteins, yet there remain several challenges, such as limited ligands to E3 ligases, selectivity, and degrading proteins that cannot be ubiquitinated. To overcome these limitations, we developed a new targeted protein degradation approach that can bypass the need for E3 ligases and ubiquitination that we have named ByeTACs. This was accomplished by developing a bifunctional molecule that recruits proteins directly to the 26S proteasome, no longer requiring the E ligase cascade. The protocols presented here describe the synthesis and application of a ByeTAC targeting bromodomain-containing protein 4 (BRD4), that can be generalized to other POIs to assess their "ByeTACability." © 2025 Wiley Periodicals LLC. Basic Protocol 1: Synthesis and characterization of a ByeTAC library targeting BRD4 Basic Protocol 2: Assessing degradation of BRD4 ByeTACs in cells Basic Protocol 3: Validating BRD4 ByeTACs mechanism of action.
High-throughput protein expression and purification can now take advantage of the copious expansion of genes from extensive genomics and metagenomic surveillance programs. This article outlines a pipeline that involves strategic selection of a large repertoire of protein targets for cloning by commercial synthetic services, followed by high-throughput transformation, expression, and solubility screening. This pipeline will be applicable to structural and functional genomics approaches, as well as any project that requires large-scale screening for production of soluble proteins in a time-efficient manner. The protocols involve transformation, protein expression and solubility determination in a high-throughput fashion in a 96-well plate format. Proteins screened in this pipeline are then able to be purified for any assay needed. © 2025 The Author(s). Current Protocols published by Wiley Periodicals LLC. Basic Protocol 1: Target optimization Basic Protocol 2: High-throughput transformation Basic Protocol 3: High-throughput expression and solubility screening.
Lysine methylation is an important post-translational modification (PTM) that regulates diverse cellular processes. Proteomic analysis is a robust method to study PTMs, but a lack of good enrichment tool limits current understanding of lysine methylation. In a previous study, we demonstrated that aryl diazonium containing 2,6-dimethoxy substitutions can conjugate monomethyllysine-modified (Kme1) peptides with high selectivity and that the reaction is reversible under acidic conditions, allowing the release of Kme1 peptides. Therefore, such a warhead has great potential for the enrichment of low-abundance Kme1 peptides from biological samples. Here, we report the preparation of aryl diazonium-functionalized resins as enrichment tools and their application for mass spectrometry-based proteomic studies of Kme1 peptides. In this procedure, aniline with a PEG linker as a precursor is synthesized and then coupled to a hydrophilic solid phase. After preparation of tryptic peptides from cell samples, the aniline groups on the resin are converted to aryl diazonium for Kme1 peptide capture. After sufficient treatment and washing, the covalent linkage is broken under acidic conditions to release the original Kme1 peptides from the resin. Finally, the enriched samples are processed by mass spectrometry scanning and data analysis to identify ∼10,000 Kme1 sites in cell or mouse tissue samples. Herein, we demonstrate an efficient Kme1 peptides enrichment strategy for deep coverage of the Kme1 proteome in biological samples. © 2025 Wiley Periodicals LLC. Basic Protocol 1: Preparation of aryl aniline-functionalized Sepharose resin Basic Protocol 2: Preparation of tryptic peptides from biological samples Basic Protocol 3: Enrichment of Kme1 peptides from whole-cell lysate tryptic peptides Basic Protocol 4: Data acquisition and analysis for Kme1 proteomics.
Multiplex experimentation that can assay multiple cellular signaling pathways in the same cells requires orthogonal genetically encoded reporters that report over large dynamic ranges. Luciferases are cost-effective, versatile candidates whose output signals can be sensitively detected in a multiplex fashion. Commonly used dual luciferase reporter assays detect one luciferase that is coupled to a single cellular pathway and a second that is coupled to a control pathway for normalization purposes. We have expanded this approach to multiplex hextuple luciferase assays that can report on five cellular signaling pathways and one control, each of which is encoded by a unique luciferase. Light emission by the six luciferases can be distinguished by the use of two distinct substrates, each specific for three luciferases, followed by spectral decomposition of the light emitted by each of the three luciferase enzymes with bandpass filters. Here, we present detailed protocols on how to perform multiplex hextuple luciferase assaying to monitor pathway fluxes through transcriptional response elements for five specific signaling pathways (i.e., c-Myc, NF-κβ, TGF-β, p53, and MAPK/JNK) using the constitutive CMV promoter as normalization control. Protocols are provided for preparing reporter vector plasmids for multiplex reporter assaying, performing cell culture and multiplex luciferase reporter vector plasmid transfection, executing multiplex luciferase assays, and analyzing and interpreting data obtained by a plate reader appropriately equipped to detect the different luminescences. © 2020 Wiley Periodicals LLC. Basic Protocol 1 : Preparation of vectors for multiplex hextuple luciferase assaying Basic Protocol 2 : Cell culture work for multiplex hextuple luciferase assays Basic Protocol 3 : Transfection of luciferase reporter plasmids followed by drug and recombinant protein treatments Basic Protocol 4 : Performing the multiplex hextuple luciferase assay
Inducible degron systems are widely used to specifically and rapidly deplete proteins of interest in cell lines and organisms. An advantage of inducible degradation is that the biological system under study remains intact and functional until perturbation, a feature that necessitates that the endogenous levels of the protein are maintained. However, endogenous tagging of genes with auxin-inducible degrons (AID) can result in chronic, auxin-independent proteasome-mediated degradation. The ARF-AID (auxin-response factor–auxin-inducible degron) system is a re-engineered auxin-inducible protein degradation system. The additional expression of the ARF-PB1 domain prevents chronic, auxin-independent degradation of AID-tagged proteins while preserving rapid auxin-induced degradation of tagged proteins. Here, we describe the protocol for engineering human cell lines to implement the ARF-AID system for specific and inducible protein degradation. These methods are adaptable and can be extended from cell lines to organisms. © 2020 The Authors. Basic Protocol 1 : Generation of ARF-P2A-TIR1 progenitor cells Basic Protocol 2 : Designing, cloning, and testing of a gene-specific sgRNA Basic Protocol 3 : Design and amplification of a homology-directed repair construct (C-terminal tagging) Alternate Protocol 1 : Design and amplification of a homology-directed repair construct (N-terminal tagging) Basic Protocol 4 : Tagging of a gene of interest with AID Alternate Protocol 2 : Establishment of an ARF-AID clamp system Basic Protocol 5 : Testing of auxin-mediated degradation of the AID-tagged protein
The lentivirus system enables efficient genetic modification of both dividing and non-dividing cells and therefore is a useful tool for elucidating developmental processes and disease pathogenesis. The development of third-generation lentiviruses has resulted in improved biosafety, low immunogenicity, and substantial packaging capabilities. However, because third-generation lentiviruses require successful co-transfection with four plasmids, this typically means that lower titers are attained. This is problematic, as it is often desirable to produce purified lentiviruses with high titers (>1 × 108 TU/ml), especially for in vivo applications. The manufacturing process for lentiviruses involves several critical experimental factors that can influence titer, purity, and transduction efficiency. Here, we describe a straightforward, stepwise protocol for the reproducible manufacture of high-titer third-generation lentiviruses (1 × 108 to 1 × 109 TU/ml). This optimized protocol enhances transgene expression by use of Lipofectamine transfection and optimized serum replacement medium, a single ultracentrifugation step, use of a sucrose cushion, and addition of a histone deacetylation inhibitor. Furthermore, we provide alternate methods for titration analyses, including functional and genomic integration analyses, using common laboratory techniques such as FACS as well as genomic DNA extraction and qPCR. These optimized methods will be beneficial for investigating developmental processes and disease pathogenesis in vitro and in vivo. © 2020 The Authors. Basic Protocol 1: Lentivirus production Support Protocol: Lentivirus concentration Basic Protocol 2: Lentivirus titration Alternate Protocol 1: Determination of viral titration by FACS analysis Alternate Protocol 2: Determination of viral titration by genome integration analysis.
Long read sequencing technologies now allow high-quality sequencing of RNAs (or their cDNAs) that are hundreds to thousands of nucleotides long. Long read sequences of nascent RNA provide single-nucleotide-resolution information about co-transcriptional RNA processing events-e.g., splicing, folding, and base modifications. Here, we describe how to isolate nascent RNA from mammalian cells through subcellular fractionation of chromatin-associated RNA, as well as how to deplete poly(A)+ RNA and rRNA, and, finally, how to generate a full-length cDNA library for use on long read sequencing platforms. This approach allows for an understanding of coordinated splicing status across multi-intron transcripts by revealing patterns of splicing or other RNA processing events that cannot be gained from traditional short read RNA sequencing. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Subcellular fractionation Basic Protocol 2: Nascent RNA isolation and adapter ligation Basic Protocol 3: cDNA amplicon preparation.
During the course of their life cycle, most RNAs move between several cellular environments where they associate with different RNA binding proteins (RBPs). Reciprocally, a significant portion of RBPs reside in more than a single cellular compartment, where they can interact with discrete RNAs and even exert distinct biological roles. Proximity-CLIP combines proximity biotinylation of proteins with photoactivatable ribonucleoside-enhanced protein-RNA crosslinking to simultaneously profile the proteome, including RBPs and the RBP-bound transcriptome, in any given subcellular compartment. Here we provide a detailed experimental protocol for Proximity-CLIP along with a simplified non-radioactive, small-RNA cDNA library preparation protocol. Published 2020 U.S. Government. Basic Protocol 1: Cell culture, 4SU labeling, proximity biotinylation, and crosslinking Basic Protocol 2: Cell extraction, streptavidin affinity purification, and on-beads trypsinization Basic Protocol 3: RNA footprints cDNA library preparation Support Protocol: Preparation of RNA-seq libraries from intact RNA.
Base-editing technologies enable the introduction of point mutations at targeted genomic sites in mammalian cells, with higher efficiency and precision than traditional genome-editing methods that use DNA double-strand breaks, such as zinc finger nucleases (ZFNs), transcription-activator-like effector nucleases (TALENs), and the clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR-associated protein 9 (CRISPR-Cas9) system. This allows the generation of single-nucleotide-variant isogenic cell lines (i.e., cell lines whose genomic sequences differ from each other only at a single, edited nucleotide) in a more time- and resource-effective manner. These single-nucleotide-variant clonal cell lines represent a powerful tool with which to assess the functional role of genetic variants in a native cellular context. Base editing can therefore facilitate genotype-to-phenotype studies in a controlled laboratory setting, with applications in both basic research and clinical applications. Here, we provide optimized protocols (including experimental design, methods, and analyses) to design base-editing constructs, transfect adherent cells, quantify base-editing efficiencies in bulk, and generate single-nucleotide-variant clonal cell lines. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Design and production of plasmids for base-editing experiments Basic Protocol 2: Transfection of adherent cells and harvesting of genomic DNA Basic Protocol 3: Genotyping of harvested cells using Sanger sequencing Alternate Protocol 1: Next-generation sequencing to quantify base editing Basic Protocol 4: Single-cell isolation of base-edited cells using FACS Alternate Protocol 2: Single-cell isolation of base-edited cells using dilution plating Basic Protocol 5: Clonal expansion to generate isogenic cell lines and genotyping of clones.
The biochemical and biophysical investigation of proteins, nucleic acids, and the assemblies that they form yields essential information to understand complex systems. Analytical ultracentrifugation (AUC) represents a broadly applicable and information-rich method for investigating macromolecular characteristics such as size, shape, stoichiometry, and binding properties, all in the true solution-state environment that is lacking in most orthogonal methods. Despite this, AUC remains underutilized relative to its capabilities and potential in the fields of biochemistry and molecular biology. Although there has been a rapid development of computing power and AUC analysis tools in this millennium, fewer advancements have occurred in development of new applications of the technique, leaving these powerful instruments underappreciated and underused in many research institutes. With AUC previously limited to absorbance and Rayleigh interference optics, the addition of fluorescence detection systems has greatly enhanced the applicability of AUC to macromolecular systems that are traditionally difficult to characterize. This overview provides a resource for novices, highlighting the potential of AUC and encouraging its use in their research, as well as for current users, who may benefit from our experience. We discuss the strengths of fluorescence-detected AUC and demonstrate the power of even simple AUC experiments to answer practical and fundamental questions about biophysical properties of macromolecular assemblies. We address the development and utility of AUC, explore experimental design considerations, present case studies investigating properties of biological macromolecules that are of common interest to researchers, and review popular analysis approaches. © 2020 The Authors.
Nucleosomes are substrates for a broad range of factors, including those involved in transcription or chromosome maintenance/reorganization and enzymes that covalently modify histones. Given the heterogeneous nature of nucleosomes in vivo (i.e., varying histone composition, post-translational modifications, DNA sequence register), understanding the specificity and activities of chromatin-interacting factors has required in vitro studies using well-defined nucleosome substrates. Here, we provide detailed methods for large-scale PCR preparation of DNA, assembly of nucleosomes from purified DNA and histones, and purification of DNA and mononucleosomes. Such production of well-defined nucleosomes for biochemical and biophysical studies is key for studying numerous proteins and protein complexes that bind and/or alter nucleosomes and for revealing inherent characteristics of nucleosomes. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Large-scale PCR amplification of DNA Basic Protocol 2: DNA and nucleosome purification using a Bio-Rad Mini Prep Cell/Prep Cell Basic Protocol 3: Nucleosome reconstitution via linear gradient salt dialysis.
Changes in protein synthesis occur under diverse physiological and pathological conditions. For example, translation can increase in response to growth signals or decrease in response to pathological states. Such changes have traditionally been measured by tracking the incorporation of radiolabeled amino acids. However, use of radioactivity is increasingly disfavored, and a simple and efficient puromycin-based, non-radioactive method called the SUnSET assay has gained popularity for measuring protein synthesis in diverse cell types and tissues. Here, we describe the principles, procedures, and troubleshooting steps for measuring protein synthesis using the SUnSET assay in cultured cells and mouse tissues. © 2020 Wiley Periodicals LLC Basic Protocol 1: Measuring protein synthesis in cultured cells by western blotting Support Protocol 1: Ponceau staining Support Protocol 2: Testing the specificity of the anti-puromycin antibody Basic Protocol 2: Measuring protein synthesis in cultured cells by immunofluorescence Basic Protocol 3: Measuring protein synthesis in mouse tissues by western blotting.
Transmembrane proteins are responsible for many critical cellular functions and represent one of the largest families of drug targets. However, these proteins, especially multipass transmembrane proteins, are difficult to study because they must be embedded in a lipid bilayer to maintain their native conformations. The development of the virion display (VirD) technology enables transmembrane proteins to be integrated into the viral envelope of herpes simplex virus 1 (HSV-1). Combining high-throughput cloning, expression, and purification techniques, VirD technology has been applied to the largest set of human transmembrane proteins, namely G-protein-coupled receptors, and has allowed the identification of interactions that are both specific and functional. This article describes the procedures to integrate an open reading frame for any transmembrane protein into the HSV-1 genome and produce recombinant HSV-1 virus to ultimately generate pure VirD virions for biological and pharmaceutical studies. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Gateway cloning of transmembrane proteins Support Protocol 1: Ethanol precipitation of bacterial artificial chromosomal DNA Support Protocol 2: Preparation of competent cells Basic Protocol 2: Production of recombinant HSV-1 virions.
High-throughput screening is one of the pillars of drug development. Unbiased transcriptome profiling is now widely used for a deeper understanding of a drug's mechanisms of action, off target effects, and cytotoxicity. Although currently available high-throughput RNA-Seq (HT RNA-Seq) methods such as PLATE-Seq, DRUG-Seq, and BRB-Seq serve these purposes, the inherent nature of these methods does not allow sample-wise sequencing library quality control. Here, we describe an HTR method called High-throughput CellulAr RNA Sequencing (HiCAR-Seq). HiCAR-Seq was optimized to work directly on cultured cells (as little as 1,000 cells) or 10 ng of total RNA. HiCAR-Seq involves reverse transcription from cultured cells or total RNA using oligo-dT primers followed by the PCR amplification of full-length cDNAs using sample-specific barcode primers in individual plate wells. Amplification of cDNA from every sample can be verified using Bioanalyzer. This step not only reveals cDNA amplification but also provides greater precision for pooling equal concentrations of cDNA from different samples. A single pooled cDNA library is made suitable for sequencing on Illumina sequencers using a tagmentation kit. Because HiCAR-Seq targets a small region at the 3' of the mRNAs, as little as 3 to 4 million reads/sample are enough to infer changes in gene expression in human or mouse cells. We believe that HiCAR-Seq represents a robust and competitive addition to the existing set of transcriptome-based high-throughput screening methods. © 2020 Wiley Periodicals LLC. Basic Protocol 1: cDNA synthesis and barcoding/enrichment PCR Basic Protocol 2: Nextera tagmentation/amplification, quantification, and sequencing.
Sensitive simultaneous assessment of multiple signaling pathways within the same cells requires orthogonal reporters that can assay over large dynamic ranges. Luciferases are such genetically encoded candidates due to their sensitivity, versatility, and cost-effectiveness. We expand luciferase multiplexing in post-lysis endpoint luciferase assays from two to six. Light emissions are distinguished by a combination of distinct substrates and emission spectra deconvolution. All six luciferase reporter units are stitched together into one plasmid facilitating delivery of all reporter units through a process we termed solotransfection, minimizing experimental errors. We engineer a multiplex hextuple luciferase assay to probe pathway fluxes through five transcriptional response elements against a control constitutive promoter. We can monitor effects of siRNA, ligand, and chemical compound treatments on their target pathways along with the four other probed cellular pathways. We demonstrate the effectiveness and adaptiveness of multiplex luciferase assaying, and its broad application across different research fields.
High-throughput cell-based screening assays are valuable tools in the discovery of chemical probes and therapeutic agents. Such assays are designed to examine the effects of small compounds on targets, pathways, or phenotypes participating in normal and disease processes. While most cell-based assays measure single quantities, multiplexed assays seek to address these limitations by obtaining multiple simultaneous measurements. The signals from such measurements should be independently detectable and cover large dynamic ranges. Luciferases are good candidates for generation of such signals. They are genetically encoded, versatile, and cost-effective, and their output signals can be sensitively detected. We recently developed a multiplex luciferase assay that allows monitoring the activity of five experimental pathways against one control simultaneously. We used synthetic assembly cloning to assemble all six luciferase reporter units into a single vector over eight stitching rounds. Because all six reporters are on a single piece of DNA, a single vector ensures stoichiometric ratios of each transcriptional unit in each transfected cell, resulting in lower experimental variation. Our proof-of-concept multiplex hextuple luciferase assay was designed to simultaneously monitor the p53, TGF-β, NF-κβ, c-Myc, and MAPK/JNK signaling pathways. The same synthetic assembly cloning pipeline allows the stitching of numerous other cellular pathway luciferase reporters. Here we present an improved three-step synthetic assembly protocol to quickly and efficiently generate multiplex hextuple luciferase reporter plasmids for other signaling pathways of interest. This improved assembly protocol provides the opportunity to analyze any five desired pathways at once much more quickly. Protocols are provided on how to prepare DNA components and destination vector plasmids, design synthetic DNA, perform assembly cloning of new transcriptional reporter elements, implement multipartite synthetic assembly cloning of single-pathway luciferase reporters, and carry out one-step assembly of final multiplex hextuple luciferase vectors. We present protocols on how to perform multiplex hextuple luciferase in an accompanying Current Protocols in Molecular Biology article. © 2020 Wiley Periodicals LLC. Basic Protocol 1: Preparation of DNA parts and destination vectors for synthetic assembly cloning Basic Protocol 2: DNA synthesis and assembly cloning of a typical transcriptional reporter element Alternate Protocol: DNA synthesis and assembly cloning of a challenging transcriptional reporter element Basic Protocol 3: Multipartite synthetic assembly cloning of individual pathway luciferase reporters Basic Protocol 4: One step assembly into final multiplex hextuple luciferase vectors Support Protocol: Generation of home-made chemocompetent E. coli DH10B-T1R cells.
Antibodies are widely used in therapeutic, diagnostic, and research applications, and antibody derivatives such as F(ab')2 fragments are used when only a particular antibody region is required. F(ab')2 can be produced through antibody engineering, but some applications require F(ab')2 produced from an original formulated antibody or directly from a polyclonal antibody pool. The cysteine protease immunoglobulin-degrading enzyme (IdeS) from Streptococcus pyogenes digests immunoglobulin G (IgG) specifically and efficiently to produce F(ab')2 . Here we detail the production and purification of recombinant IdeS; its utilization to digest monoclonal or polyclonal antibodies to F(ab')2 fragments; and F(ab')2 purification through consecutive affinity chromatography steps. The resultant F(ab')2 exhibit high purity, retain antigen-binding functionality, and are readily utilizable in various downstream applications. © 2020 by John Wiley & Sons, Inc. Basic Protocol: Production and purification of F(ab')2 fragments from monoclonal and polyclonal antibodies using IdeS Alternate Protocol: Purification of polyclonal antigen-specific F(ab')2 fragments from human serum or secretions Support Protocol: Production and purification of IdeS.
The ARF-AID (Auxin Response Factor-Auxin Inducible Degron) system is a re-engineered auxin-inducible protein degradation system. Inducible degron systems are widely used to specifically and rapidly deplete proteins of interest in cell lines and organisms. An advantage of inducible degradation is that the biological system under study remains intact and functional until perturbation. This feature necessitates that the endogenous levels of the protein are maintained. However, endogenous tagging of genes with AID can result in chronic, auxin-independent proteasome-mediated degradation. The additional expression of the ARF-PB1 domain in the re-engineered ARF-AID system prevents chronic degradation of AID-tagged proteins while preserving rapid degradation of tagged proteins. Here we describe the protocol for engineering human cell lines to implement the ARF-AID system for specific and inducible protein degradation. These methods are adaptable and can be extended from cell lines to organisms.
