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.
