Molecular Biology

Atypical DNA and RNA secondary structures play a crucial role in simple sequence repeat (SSR) diseases, which are associated with a class of neurological and neuromuscular disorders known as “anticipation diseases,” where the age of disease onset decreases and the severity of the disease is increased as the intergenerational expansion of the SSR increases. While the mechanisms underlying these diseases are complex and remain elusive, there is a consensus that stable, non-B-DNA atypical secondary structures play an important – if not causative – role. These structures include single-stranded DNA loops and hairpins, G-quartets, Z-DNA, triplex nucleic acid structures, and others. While all of these structures are of interest, structures based on nucleic acid triplexes have recently garnered increased attention as they have been implicated in gene regulation, gene repair, and gene engineering. Our work here focuses on the construction of DNA triplexes and RNA/DNA hybrids formed from GAA/TTC trinucleotide repeats, which underlie Friedreich’s ataxia. While there is some software, such as the Discovery Studio Visualizer, that can aid in the initial construction of DNA triple helices, the only option for the triple helix is constrained to be that of an antiparallel pyrimidine for the third strand. In this protocol, we illustrate how to build up more generalized DNA triplexes and DNA/RNA mixed hybrids. We make use of both the Discovery Studio Visualizer and the AMBER simulation package to construct the initial triplexes. Using the steps outlined here, one can – in principle – build up any triple nucleic acid helix with a desired sequence for large-scale molecular dynamics simulation studies.

Higher-order chromatin organization shaped by epigenetic modifications influence the chromatin environment and subsequently regulate gene expression. Direct visualization of the higher-order chromatin structure at their epigenomic states is of great importance for understanding chromatin compaction and its subsequent effect on gene expression and various cellular processes. With the recent advances in super-resolution microscopy, the higher-order chromatin structure can now be directly visualized in situ down to the scale of ~30 nm. This protocol provides detailed description of super-resolution imaging of higher-order chromatin structure using stochastic optical reconstruction microscopy (STORM). We discussed fluorescence staining methods of DNA and histone proteins and crucial technical factors to obtain high-quality super-resolution images.

Chromosome conformation capture sequencing (Hi-C) is a powerful method to comprehensively interrogate the three-dimensional positioning of chromatin in the nucleus. The development of Hi-C can be traced back to successive increases in the resolution and throughput of chromosome conformation capture (3C) (Dekker et al., 2002). The basic workflow of 3C consists of (i) fixation of intact chromatin, usually by formaldehyde, (ii) cutting the fixed chromatin with a restriction enzyme, (iii) religation of sticky ends under diluted conditions to favor ligations between cross-linked fragments or those between random fragments and (iv) quantifying the number of ligations events between pairs of genomic loci (de Wit and de Laat, 2012). In the original 3C protocol, ligation frequency was measured by amplification of selected ligation junctions corresponding to a small number of genomic loci (‘one versus one’) through semi-quantitative PCR (Dekker et al., 2002). The chromosome conformation capture-on-chip (4C) and chromosome conformation capture carbon copy (5C) technologies then extended 3C to count ligation events in a ‘one versus many’ or ‘many versus many’ manner, respectively. Hi-C (Lieberman-Aiden et al., 2009) finally combined 3C with next-generation sequencing (Metzker, 2010). Here, before religation sticky ends are filled in with biotin-labeled nucleotide analogs to enrich for fragments with a ligation junction in a later step. The Hi-C libraries are then subjected to high-throughput sequencing and the resultant reads mapped to a reference genome, allowing the determination of contact probabilities in a ‘many versus many’ way with a resolution that is limited only by the distribution of restriction sites and the read depth. The first application of Hi-C was the elucidation of global chromatin folding principles in the human genome (Lieberman-Aiden et al., 2009). Similar efforts have since been carried out in other eukaryotic model species such as yeast (Duan et al., 2010), Drosophila (Sexton et al., 2012) and Arabidopsis (Grob et al., 2014; Wang et al., 2015; Liu et al., 2016). Other uses of Hi-C include the study of chromatin looping at high-resolution (Rao et al., 2014; Liu et al., 2016), of chromatin reorganization along the cell cycle (Naumova et al., 2013) and of differences in chromatin organization in mutant individuals (Feng et al., 2014). The tethered conformation capture protocol (TCC) (Kalhor et al., 2011) described here is a variant of the original Hi-C method (Lieberman-Aiden et al., 2009) and was adapted to Triticeae.

Chromosome conformation capture (3C) techniques are crucial to understanding tissue-specific regulation of gene expression, but current methods generally require large numbers of cells. This protocol describes two new low-input Capture-C approaches that can generate high-quality 3C interaction profiles from 10,000-20,000 cells, depending on the resolution used for analysis.

We have developed locus-specific chromatin immunoprecipitation (locus-specific ChIP) technologies consisting of insertional ChIP (iChIP) and engineered DNA-binding molecule-mediated ChIP (enChIP). Locus-specific ChIP is a method to isolate a genomic region of interest from cells while it also identifies what binds to this region using mass spectrometry (for protein) or next generation sequencing (for RNA or DNA) as described in Fujita et al. (2016a). Recently, we identified genomic regions that physically interact with a locus using an updated form of enChIP, in vitro enChIP, in combination with NGS (in vitro enChIP-Seq) (Fujita et al., 2017a). Here, we describe a protocol on in vitro enChIP to isolate a target locus for identification of genomic regions that physically interact with the locus.

The basic unit of chromatin is the nucleosome, a histone octamer with 147 base pairs of DNA wrapped around it. Positions of nucleosomes relative to each other and to DNA elements have a strong impact on chromatin structure and gene activity and are tightly regulated at multiple levels, i.e., DNA sequence, transcription factor binding, histone modifications and variants, and chromatin remodeling enzymes (Bell et al., 2011; Hughes and Rando, 2014). Nucleosome positions in cells or isolated nuclei can be detected by partial nuclease digestion of native or cross-linked chromatin followed by ligation-mediated polymerase chain reaction (LM-PCR) (McPherson et al., 1993; Soutoglou and Talianidis, 2002). This protocol describes a nucleosome positioning assay using Micrococcal Nuclease (MNase) digestion of formaldehyde-fixed chromatin followed by LM-PCR. We exemplify the nucleosome positioning assay for the promoter of genes encoding ribosomal RNA (rRNA genes or rDNA) in mice, which has two mutually exclusive configurations. The rDNA promoter harbors either an upstream nucleosome (NucU) covering nucleotides -157 to -2 relative to the transcription start site, or a downstream nucleosome (NucD) at position -132 to +22 (Li et al., 2006; Xie et al., 2012). Radioactive labeling of LM-PCR products followed by denaturing urea-polyacrylamide gel electrophoresis allows resolution and relative quantification of both configurations. As depicted in the diagram in Figure 1, the nucleosome positioning assay is a versatile low to medium throughput method to map discrete nucleosome positions with high precision in a semi-quantitative manner.


Figure 1. Flow chart depicting the nucleosome positioning assay. The diagram shows how the assay is used to detect the ratio between upstream (NucU) and downstream (NucD) nucleosome positions at the mouse rDNA promoter. After all steps have been performed, the LM-PCR yields two radiolabeled products that differ in size and correspond to NucU and NucD. Signal intensities of the bands reflect the relative abundance of each nucleosome position in the original sample.

Somatic chromosomes are usually studied from the root tip cells of the plants for cytological investigations. In dioecious plant species like Coccinia grandis, it is very difficult to get meristematic root tip cells from the mature plants of the respective sex forms. In this report, young leaves of the respective sexual phenotypes were used as tissue samples for mitotic chromosome analysis. For meiotic preparation, flower buds of appropriate size were selected for chromosomal studies. Following protocols could be effectively used for routine chromosome preparations in other plant species as well.

Ultraviolet (UV) irradiation induces helix distorting photolesions such as cyclobutane pyrimidine dimers (CPD) and pyrimidine-pyrimidone (6-4) photoproducts (6-4PP) which threaten genomic integrity if unrepaired. In mammals, nucleotide excision repair (NER) is the only pathway that removes UV-induced DNA damages. Here we describe DNA slot blot repair assay for quantitative detection of NER activity using DNA damage specific antibodies such as anti-CPD and anti-6-4PP. Briefly, genomic DNA irradiated with UV was isolated from cells, and the genomic DNA was vacuum-transferred to a nitrocellulose membrane using a Bio-Dot SF microfiltration apparatus (Bio-Rad). A monoclonal antibody that recognizes CPD or 6-4PP was applied to detect the remaining amount of photolesions in the genomic DNA. For loading control of even loading, DNA onto the membrane can be further analyzed by SYBR-gold staining.

In this protocol, we describe a simple and efficient method for meiotic chromosome spread preparation in rice pollen mother cells. Meiotic chromosome preparation by spreading itself is an important technique for plant cytogenetics (Higgins et al., 2004; Chelysheva et al., 2012; Wang et al., 2009); furthermore, it is a crucial step for applying other cytogenetic methods including Fluorescence in situ hybridization (FISH) and immunostaining.

This protocol is a more detailed version of previous protocols (Yang et al., 2011; Bolaños-Villegas et al., 2013) developed for the examination of meiotic chromosome spreads. Meiotic chromosome spreads are useful to determine the presence of defects in chromosome pairing and segregation. The protocol also describes how to perform fluorescent in situ hybridization experiments with a centromere probe used to label chromosomes.