分子生物学

Chromatin immunoprecipitation (ChIP) maps, on a genome-wide scale, transcription factor binding sites, and the distribution of other chromatin-associated proteins and their modifications. As such, it provides valuable insights into mechanisms of gene regulation. However, successful ChIP experiments are dependent on the availability of a high-quality antibody against the target of interest. Using antibodies with poor sensitivity and specificity can yield misleading results. This can be partly circumvented by using epitope-tagged systems (e.g., HA, Myc, His), but these approaches are still antibody-dependent. HaloTag^® is a modified dehalogenase enzyme, which covalently binds synthetic ligands. This system can be used for imaging and purification of HaloTag^® fusion proteins, and has been used for ChIP in vitro. Here, we present a protocol for using the HaloTag^® system for ChIP in vivo, to map, with sensitivity and specificity, the cistrome of a dynamic mouse transcription factor expressed at its endogenous locus.

Graphical abstract:

Characterizing the molecular mechanisms regulating gene expression is crucial for understanding the regulatory processes underlying physiological responses to environmental and developmental signals in eukaryotes. The covalent modification of histones contributes to the compaction levels of chromatin, as well as the recruitment of the transcriptional machinery to specific loci, facilitating metastable changes in gene activity. ChIP-seq (Chromatin Immunoprecipitation followed by sequencing) has become the gold standard method for determining histone modification profiles among different organisms, tissues, and genotypes. In the current protocol, we describe a highly robust method for performing ChIP-seq of histone modifications in Arabidopsis thaliana plantlets. Besides its robustness, this method uses in-house-prepared buffers for chromatin extraction, immunoprecipitation, washing, and elusion, making it cost-effective in contrast to commercial kits.

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.

CRISPR-Cas9 has transformed biomedical research and medicine through convenient and targeted manipulation of DNA. Time- and spatially-resolved control over Cas9 activity through the recently developed very fast CRISPR (vfCRISPR) system have facilitated comprehensive studies of DNA damage and repair. Understanding the fundamental principles of Cas9 binding and cleavage behavior is essential before the widespread use of these systems and can be readily accomplished in vitro through both cleavage and electrophoretic mobility shift assays (EMSA). The protocol for in vitro cleavage consists of Cas9 with guide RNA (gRNA) ribonucleoprotein (RNP) formation, followed by incubation with target DNA. For EMSA, this reaction is directly loaded onto an agarose gel for visualization of the target DNA band that is shifted due to binding by the Cas9 RNP. To assay for cleavage, Proteinase K is added to degrade the RNP, allowing target DNA (cleaved and/or uncleaved) to migrate consistently with its molecular weight. Heating at 95°C rapidly inactivates the RNP on demand, allowing time-resolved measurements of Cas9 cleavage kinetics. This protocol facilitates the characterization of the light-activation mechanism of photocaged vfCRISPR gRNA.

Protein translocation on DNA represents the key biochemical activity of ssDNA translocases (aka helicases) and dsDNA translocases such as chromatin remodelers. Translocation depends on DNA binding but is a distinct process as it typically involves multiple DNA binding states, which are usually dependent on nucleotide binding/hydrolysis and are characterized by different affinities for the DNA. Several translocation assays have been described to distinguish between these two modes of action, simple binding as opposed to directional movement on dsDNA. Perhaps the most widely used is the triplex-forming oligonucleotide displacement assay. Traditionally, this assay relies on the formation of a DNA triplex from a dsDNA segment and a short radioactively labeled oligonucleotide. Upon translocation of the protein of interest along the DNA substrate, the third DNA strand is destabilized and eventually released off the DNA duplex. This process can be visualized and quantitated by polyacrylamide electrophoresis. Here, we present an effective, sensitive, and convenient variation of this assay that utilizes a fluorescently labeled oligonucleotide, eliminating the need to radioactively label DNA. In short, our protocol provides a safe and user-friendly alternative.

Graphical abstract:

Figure 1. Schematic of the triplex-forming oligonucleotide displacement assay.

In bacteria, the restart of stalled DNA replication forks requires the DNA helicase PriA. PriA can recognize and remodel abandoned DNA replication forks, unwind DNA in the 3'-to-5' direction, and facilitate the loading of the helicase DnaB onto the DNA to restart replication. ssDNA-binding protein (SSB) is typically present at the abandoned forks, protecting the ssDNA from nucleases. Research that is based on the assays for junction dissociation, surface plasmon resonance, single-molecule FRET, and x-ray crystal structure has revealed the helicase activity of PriA, the SSB-PriA interaction, and structural information of PriA helicase. Here, we used Atomic Force Microscopy (AFM) to visualize the interaction between PriA and DNA substrates with or without SSB in the absence of ATP to delineate the substrate recognition pattern of PriA before its ATP-catalyzed DNA-unwinding reaction. The protocol describes the steps to obtain high-resolution AFM images and the details of data analysis and presentation.

Chromatin immunoprecipitation coupled with quantitative PCR (ChIP-qPCR) or high-throughput sequencing (ChIP-seq) has become the gold standard for the identification of binding sites of DNA binding proteins and the localization of histone modification on a locus-specific or genome-wide scale, respectively. ChIP experiments can be divided into seven critical steps: (A) sample collection, (B) crosslinking of proteins to DNA, (C) nuclear extraction, (D) chromatin isolation and fragmentation by sonication, (E) immunoprecipitation of histone marks by appropriate antibodies, (F) DNA recovery, and (G) identification of precipitated protein-associated DNA by qPCR or high-throughput sequencing. Here, we describe a time-efficient protocol that can be used for ChIP-qPCR experiments to study the localization of histone modifications in young inflorescences of the model plants Arabidopsis thaliana.

Identification of specific DNA binding sites of transcription factors is important in understanding their functions. Recent techniques allow us to investigate genome-wide in vivo binding positions by chromatin immunoprecipitation combined with high-throughput sequencing. However, to further explore the binding motifs of transcription factors, in-depth biochemical analysis is required. Here, we describe an efficient protocol of protein-DNA interactions based on a combination of our in vitro transcription/translation system and AlphaScreen^® technology. The in vitro transcription/translation system supports an efficient and quick way of protein synthesis by alleviating cumbersome cloning steps. In addition, AlphaScreen^® system provides a highly sensitive, quick, and easy handling platform to investigate the protein-DNA interactions in vitro. Thus, our method largely contributes to comprehensive analysis of the biochemical properties of transcription factors.

Aptamers have emerged as a novel category in the field of bioreceptors due to their wide applications ranging from biosensing to therapeutics. Several variations of their screening process, called SELEX have been reported which can yield sequences with desired properties needed for their final use. We report a facile microtiter plate-based Cell-SELEX method for a gram-negative bacteria E. coli. The optimized protocol allows the reduction of number of rounds for SELEX by offering higher surface area and longer retention times. In addition, this protocol can be modified for other prokaryotic and eukaryotic cells, and glycan moieties as target for generation of high affinity bio-receptors in a short course of time in-vitro.

Nitrogen is an essential nutrient for all living organisms. In cyanobacteria, a group of oxygenic photosynthetic bacteria, nitrogen homeostasis is maintained by an intricate regulatory network around the transcription factor NtcA. Although mechanisms controlling NtcA activity appear to be well understood, the sets of genes under its control (i.e., its regulon) remain poorly defined. In this protocol, we describe the procedure for chromatin immunoprecipitation using NtcA antibodies, followed by DNA sequencing analysis (ChIP-seq) during early acclimation to nitrogen starvation in the cyanobacterium Synechocystis sp. PCC 6803 (hereafter Synechocystis). This protocol can be extended to analyze any DNA-binding protein in cyanobacteria for which suitable antibodies exist.