Molecular Biology

8-oxo-7,8-dihydro-2′-deoxyguanosine (8-oxodG) is considered to be a premutagenic DNA lesion generated by 2'-deoxyguanosine (dG) oxidation due to reactive oxygen species (ROS). In recent years, the 8-oxodG distribution in human, mouse, and yeast genomes has been underlined using various next-generation sequencing (NGS)–based strategies. The present study reports the OxiDIP-Seq protocol, which combines specific 8-oxodG immuno-precipitation of single-stranded DNA with NGS, and the pipeline analysis that allows the genome-wide 8-oxodG distribution in mammalian cells. The development of this OxiDIP-Seq method increases knowledge on the oxidative DNA damage/repair field, providing a high-resolution map of 8-oxodG in human cells.

Chromatin accessibility is a key determinant of gene expression that can be altered under different physiological and disease conditions. Skeletal muscle is made up of myofibers that are highly plastic and adaptive. Therefore, assessing the genome-wide chromatin state of myofibers under various conditions is very important to gain insight into the epigenetic state of myonuclei. The rigid nature of myofibers, as well as the low number of myonuclei that they contain, have rendered genome-wide studies with myofibers challenging. In recent years, ATAC-Seq from whole muscle and single nucleus ATAC-Seq have been performed. However, these techniques cannot distinguish between different fiber and cell types present in the muscle. In addition, due to the limited depth capacity obtained from single nucleus ATAC-Seq, an extensive comparative analysis cannot be performed. Here, we introduce a protocol where we combine the isolation of a single myofiber with OMNI ATAC-Seq. This protocol allows for genome-wide analysis of accessible chromatin regions of a selected single myofiber at a sufficient depth for comparative analysis under various physiological and disease conditions. This protocol can also allow for a specific myofiber to be selected, such as a regenerating myofiber. In the future, this protocol can help identify global changes in chromatin state under various conditions, as well as between different types of myofibers.

Graphical abstract:

The majority of biopsies in both basic research and translational cancer studies are preserved in the format of archived formalin-fixed paraffin-embedded (FFPE) samples. Profiling histone modifications in archived FFPE tissues is critically important to understand gene regulation in human disease. The required input for current genome-wide histone modification profiling studies from FFPE samples is either 10–20 tissue sections or whole tissue blocks, which prevents better resolved analyses. Nevertheless, it is desirable to consume a minimal amount of FFPE tissue sections in the analysis as clinical tissue of interest are limited. Here, we present FFPE tissue with antibody-guided chromatin tagmentation with sequencing (FACT-seq), highly sensitive method to efficiently profile histone modifications in FFPE tissue by combining a novel fusion protein of hyperactive Tn5 transposase and protein A (T7-pA-Tn5) transposition and T7 in vitro transcription. FACT-seq generates high-quality chromatin profiles from different histone modifications with low number of FFPE nuclei. We showed a very small piece of FFPE tissue section containing ~4000 nuclei is sufficient to decode H3K27ac modifications with FACT-seq. In archived FFPE human colorectal and human glioblastoma cancer tissue, H3K27ac FACT-seq revealed disease specific super enhancers. In summary, FACT-seq allows researchers to decode histone modifications like H3K27ac and H3K27me3 in archival FFPE tissues with high sensitivity, thus allowing us to understand epigenetic regulation.

Graphical abstract:

(i) FFPE tissue section; (ii) Isolated nuclei; (iii) Primary antibody, secondary antibody and T7-pA-Tn5 bind to targets; (iv) DNA purification; (v) In vitro transcription and sequencing library preparation; (vi) Sequencing

ATAC-seq (assay for transposase-accessible chromatin with high-throughput sequencing) is a powerful method to evaluate chromatin accessibility and nucleosome positioning at a genome-wide scale. This assay uses a hyperactive Tn5 transposase, to simultaneously cut open chromatin and insert adapter sequences. After sequencing, the reads generated through this technique are generally indicative of transcriptional regulatory elements that are located in accessible chromatin. This method was originally developed by Buenrostro et al. (2013), and since then it has been improved by the same authors several times, until their last update called OMNI ATAC-seq (Corces et al., 2017). Here, we describe an ATAC-seq protocol based on the OMNI-ATAC method, with a special focus on the initial steps of thawing cryopreserved cells, and the final steps of library purification using magnetic beads. This protocol can be of interest for laboratories working in a fast-paced environment.

Graphic abstract:

Flowchart of the protocol

Gene expression depends on the binding of transcription factors with DNA regulatory sequences. The level of accessibility for these sequences varies between cells and cell types. Until recently, using the Tn5 assay for transposase-accessible chromatin for sequencing (ATAC-seq) technology allowed assessing the profiles of chromatin from an entire organ or, when coupled with the isolation of nuclei tagged in specific cell types (INTACT) method, from a cell-type. Recently, ATAC-seq experiments were conducted at the level of individual plant nuclei. Applying single nuclei ATAC-seq (sNucATAC-seq) technology to thousands of individual cells revealed more finely tuned profiles of chromatin accessibility. In this manuscript, we describe a method to isolate nuclei fom plant roots and green tissues, permeabilize the nuclear membrane using detergent to allow the penetration of the Tn5 transposase, and re-suspend them in a nuclei resuspension buffer compatible with the construction of sNucATAC-seq libraries using the 10× Genomic’s Chromium technology. This protocol was successfully applied on Arabidopsis thaliana and Glycine max root nuclei.

Atomic force microscopy (AFM) is a powerful tool to image macromolecular complexes with nanometer resolution and exquisite single-molecule sensitivity. While AFM imaging is well-established to investigate DNA and nucleoprotein complexes, AFM studies are often limited by small datasets and manual image analysis that is slow and prone to user bias. Recently, we have shown that a combination of large scale AFM imaging and automated image analysis of nucleosomes can overcome these previous limitations of AFM nucleoprotein studies. Using our high-throughput imaging and analysis pipeline, we have resolved nucleosome wrapping intermediates with five base pair resolution and revealed how distinct nucleosome variants and environmental conditions affect the unwrapping pathways of nucleosomal DNA. Here, we provide a detailed protocol of our workflow to analyze DNA and nucleosome conformations focusing on practical aspects and experimental parameters. We expect our protocol to drastically enhance AFM analyses of DNA and nucleosomes and to be readily adaptable to a wide variety of other protein and protein-nucleic acid complexes.

We previously introduced Cleavage Under Targets & Tagmentation (CUT&Tag), an epigenomic profiling method in which antibody tethering of the Tn5 transposase to a chromatin epitope of interest maps specific chromatin features in small samples and single cells. With CUT&Tag, intact cells or nuclei are permeabilized, followed by successive addition of a primary antibody, a secondary antibody, and a chimeric Protein A-Transposase fusion protein that binds to the antibody. Addition of Mg⁺⁺ activates the transposase and inserts sequencing adapters into adjacent DNA in situ. We have since adapted CUT&Tag to also map chromatin accessibility by simply modifying the transposase activation conditions when using histone H3K4me2, H3K4me3, or Serine-5-phosphorylated RNA Polymerase II antibodies. Using these antibodies, we redirect the tagmentation of accessible DNA sites to produce chromatin accessibility maps with exceptionally high signal-to-noise and resolution. All steps from nuclei to amplified sequencing-ready libraries are performed in single PCR tubes using non-toxic reagents and inexpensive equipment, making our simplified strategy for simultaneous chromatin profiling and accessibility mapping suitable for the lab, home workbench, or classroom.

Chromatin consists of compacted DNA in complex with proteins and contributes to the organization of DNA and its stability. Furthermore, chromatin plays key roles in regulating cellular processes such as DNA replication, transcription, DNA repair, and mitosis. Chromatin assumes more compact (inaccessible) or decondensed (accessible) conformations depending on the function that is being supported in the genome, either locally or globally. The activity of nucleases has been used previously to assess the accessibility of specific genomic regions in vitro, such as origins of replication at varying points in the cell cycle. Here, we provide an assay to determine the accessibility of specific human genomic regions (example used herein: Lamin B2 origin of DNA replication) by measuring the effect of DNase I nuclease on qPCR signal from the studied site. This assay provides a powerful method to interrogate the molecular mechanisms that regulate chromatin accessibility, and how these processes affect various cellular functions involving the human genome that require manipulation of chromatin conformation.

Regulation of gene expression involves dynamic changes in chromatin organization, where in many cases open chromatin structure correlates with gene activation. Several methods enable monitoring changes in chromatin accessibility, including ATAC-seq, FAIRE-seq, MNase-seq and DNAse-seq methods, which involve Next-generation-sequencing (NGS). Focusing on the adult Drosophila differentiated gut enterocytes (ECs) we used a sequencing-free method that enables visualizing and semi-quantifying large-scale changes in chromatin structure using in vitro methylation assay with the bacterial CpG Methyltransferase, M. Sssl, that determine chromatin accessibility. In brief, as CpG methylation is minimal in differentiated somatic Drosophila cells, we used the bacterial M. SssI enzyme to methylate CpG dinucleotides in situ depending on their chromatin accessibility. The methylated dinucleotides are detected using 5mCytosine monoclonal antibody and nuclei are visualized microscopically. Thus, the 5mC method enables to monitor large-scale chromatin changes in heterogenic cellular tissue focusing on the cell type of interest and without the need for cell purification or NGS.

Transcription regulation is a key aspect of cellular identity established during development and maintained into adulthood. Molecular and biochemical assays that probe the genome are critical tools in exploring mechanisms of transcription regulation and cell type identity. The mammalian brain is composed of a huge diversity of cell types with distinct properties and functions. To understand these specific roles, it is necessary to selectively target cell populations for study. However, the need to selectively study restricted cell populations poses a challenge in neurobiology. It is often difficult to collect sufficient cellular input for many standard biochemical and molecular assays. Recently, important advances have been made to scale assays down, opening up new frontiers to explore molecular mechanisms in neurons. Concurrently, methodologies for preparing neurons for such assays has advanced taking into consideration specific methods to preserve the cell biology meant to be assayed. Here we describe a method for preparing live neurons from adult brain tissue for the Assay for Transposase Accessible Chromatin (ATAC).