Molecular Biology

An efficient and precise genome-editing approach is in high demand in any molecular biology or cell biology laboratory worldwide. However, despite a recent rapid progress in the toolbox tailored for precise genome-editing, including the base editors and prime editors, there is still a need for a cost-effective knock-in (KI) approach amenable for long donor DNA cargos with high efficiency. By harnessing the high-efficient double-strand break (DSB) repair pathway of microhomology-mediated end joining, we previously showed that a specially designed 3′-overhang double-strand DNA (odsDNA) donor harboring 50-nt homology arm (HA) allows high-efficient exogenous DNA KI when combined with CRISPR-Cas9 technology. The lengths of the 3′-overhangs of odsDNA donors could be manipulated by the five consecutive phosphorothioate (PT) modifications. In this protocol, we detail the stepwise procedures to conduct the LOCK (Long dsDNA with 3′-Overhangs mediated CRISPR Knock-in) method for gene-sized (~1–3 kb) KI in mammalian cells.

Graphical overview

Improvement of large DNA fragment knock-in rates by attaching odsDNA donors to Cas9-PCV2 fusion protein

Epigenetic modifications of DNA, and especially cytosine, play a crucial role in regulating basic cellular processes and thereby the overall cellular metabolism. Their levels change during organismic and cellular development, but especially also in pathogenic aberrations such as cancer. Levels of respective modifications are often addressed in bulk by specialized mass spectrometry techniques or by employing dedicated ChIP-seq protocols, with the latter giving information about the sequence context of the modification. However, to address modification levels on a single cell basis, high- or low-content microscopy techniques remain the preferred methodology. The protocol presented here describes a straightforward method to detect and quantify different DNA modifications in human cell lines, which can also be adapted to other cultured mammalian cell types. To this end, cells are immunostained against two different cytosine modifications in combination with DNA counterstaining. Image acquisition takes place on a confocal microscopy system. A semi-automated analysis pipeline helps to gather data in a fast and reliable fashion. The protocol is comparatively simple, fast, and cost effective. By employing methodologies that are often well established in most molecular biology laboratories, many researchers are able to apply the described protocol straight away in-house.

Nucleic acids in living organisms are more complex than the simple combinations of the four canonical nucleotides. Recent advances in biomedical research have led to the discovery of numerous naturally occurring nucleotide modifications and enzymes responsible for the synthesis of such modifications. In turn, these enzymes can be leveraged towards toolkits for DNA and RNA manipulation for epigenetic sequencing or other biotechnological applications. Here, we present the protocol to obtain purified 5-hydroxymethylcytosine carbamoyltransferase enzymes and the associated assays to convert 5-hydroxymethylcytosine to 5-carbamoyloxymethylcytosine in vitro. We include detailed assays using DNA, RNA, and single nucleotide/deoxynucleotide as substrates. These assays can be combined with downstream applications for genetic/epigenetic regulatory mechanism studies and next-generation sequencing purposes.

In recent years, DNA methylation research has been accelerated by the advent of nanopore sequencers. However, read length has been limited by the constraints of base conversion using the bisulfite method, making analysis of chromatin content difficult. The read length of the previous method combining bisulfite conversion and long-read sequencing was ~1.5 kb, even using targeted PCR. In this study, we have improved read length (~5 kb), by converting unmethylated cytosines to uracils with APOBEC enzymes, to reduce DNA fragmentation. The converted DNA was then sequenced using a PromethION nanopore sequencer. We have also developed a new analysis pipeline that accounts for base conversions, which are not present in conventional nanopore sequencing, as well as errors produced by nanopore sequencing.

Biochemical investigations into DNA-binding and DNA-cutting proteins often benefit from the specific attachment of a radioactive label to one of the two DNA termini. In many cases, it is essential to perform two versions of the same experiment: one with the 5′ DNA end labeled and one with the 3′ DNA end labeled. While homogeneous 5′-radiolabeling can be accomplished using a single kinase-catalyzed phosphorylation step, existing procedures for 3′-radiolabeling often result in probe heterogeneity, prohibiting precise DNA fragment identification in downstream experiments. We present here a new protocol to efficiently attach a ³²P-phosphate to the 3′ end of a DNA oligonucleotide of arbitrary sequence, relying on inexpensive DNA oligonucleotide modifications (2′-O-methylribonucleotide and ribonucleotide sugar substitutions), two enzymes (T4 polynucleotide kinase and T4 RNA ligase 2), and the differential susceptibility of DNA and RNA to hydroxide treatment. Radioactive probe molecules produced by this protocol are homogeneous and oxidant-compatible, and they can be used for precise cleavage-site mapping in the context of both DNase enzyme characterization and DNA footprinting assays.

Graphic abstract

Mosaic analysis in Drosophila, an important tool to assess cellular phenotypes of mutants in an otherwise heterozygous background, relies on mitosis. Hence, it cannot be used to inactivate gene function in mitotically inactive, terminally differentiated cells such as neurons. To address this issue, we developed “Flip-flop”, a novel, Flippase-dependent in vivo cassette-inversion method that functions independent of mitosis, and therefore can be used for gene inactivation in both mitotic as well as post-mitotic cells. This method allows tagging protein-coding genes with EGFP and generates mutant cells that are marked with mCherry upon cassette inversion. Here, we describe protocols for generation and validation of fly lines that can be used for conditional gene inactivation in mitotic as well as post-mitotic cells. We provide typical examples of Flip-flop mediated mosaic analysis in SNF4Aγ and Trim9. Use of Flip-flop mediated functional analysis will permit a detailed investigation of the role of genes previously recalcitrant to mosaic analysis.

Powerful and general methods that can enhance gene expression are useful to systematically study gene function. To date, compared with the methods in generating loss-of-function mutants, methods to achieve gain-of-function are limited. The entire field in Drosophila has relied heavily on the Gal4/UAS:cDNA overexpression system developed over two decades ago. It is laborious and expensive to clone the coding DNA sequence (CDS) of a gene, especially those of large size. In addition, side effects of this method are often observed because of the ectopic expression. Also, simultaneous activation of two genes with the traditional method is often time-consuming, and few are achievable for three or more genes. In this protocol, we describe how to build an effective and convenient targeting activator system, flySAM, to activate endogenous genes in Drosophila melanogaster based on the structure-guided engineering of CRISPR-Cas9 complex.

In recent years, great progress has been made in the research of genome editing systems, one of which is the CRISPR-Cas9 system, a powerful technology that is applied to edit animal genome. Here, we describe a CRISPR-Cas9 mediated mutation protocol for efficiently and specifically editing genes in Drosophila. In this optimized system, the mutant progeny can be generated by only injecting a DNA plasmid encoding synthetic guide RNA (sgRNA) under the control of the U6b promoter into transgenic fly embryos in which Cas9 is specifically expressed in the progenitor cells, thus the gene of interest can be edited by the CRISPR in germ cells, with high rate of heritable mutations and few side effects.

The smut fungus Ustilago maydis is an established model organism for elucidating how biotrophic pathogens colonize plants and how gene families contribute to virulence. Here we describe a step by step protocol for the generation of CRISPR plasmids for single and multiplexed gene editing in U. maydis. Furthermore, we describe the necessary steps required for generating edited clonal populations, losing the Cas9 containing plasmid, and for selecting the desired clones.

The discovery of the CRISPR-Cas9 system from Streptococcus pyogenes has allowed the development of genome engineering tools in a variety of organisms. A frequent limitation in CRISPR-Cas9 function is adequate expression levels of sgRNA. This protocol provides a strategy to construct hybrid RNA polymerase III (Pol III) promoters that facilitate high expression of sgRNA and improved CRISPR-Cas9 function. We provide selection criteria of Pol III promoters, efficient promoter construction methods, and a sample screening technique to test the efficiency of the hybrid promoters. A hybrid promoter system developed for Yarrowia lipolytica will serve as a model.