分子生物学

CRISPR-Cas9 technology has become an essential tool for plant genome editing. Recent advancements have significantly improved the ability to target multiple genes simultaneously within the same genetic background through various strategies. Additionally, there has been significant progress in developing methods for inducible or tissue-specific editing. These advancements offer numerous possibilities for tailored genome modifications. Building upon existing research, we have developed an optimized and modular strategy allowing the targeting of several genes simultaneously in combination with the synchronized expression of the Cas9 endonuclease in the egg cell. This system allows significant editing efficiency while avoiding mosaicism. In addition, the versatile system we propose allows adaptation to inducible and/or tissue-specific edition according to the promoter chosen to drive the expression of the Cas9 gene. Here, we describe a step-by-step protocol for generating the binary vector necessary for establishing Arabidopsis edited lines using a versatile cloning strategy that combines Gateway^® and Golden Gate technologies. We describe a versatile system that allows the cloning of as many guides as needed to target DNA, which can be multiplexed into a polycistronic gene and combined in the same construct with sequences for the expression of the Cas9 endonuclease. The expression of Cas9 is controlled by selecting from among a collection of promoters, including constitutive, inducible, ubiquitous, or tissue-specific promoters. Only one vector containing the polycistronic gene (tRNA-sgRNA) needs to be constructed. For that, sgRNA (composed of protospacers chosen to target the gene of interest and sgRNA scaffold) is cloned in tandem with the pre-tRNA sequence. Then, a single recombination reaction is required to assemble the promoter, the zCas9 coding sequence, and the tRNA-gRNA polycistronic gene. Each element is cloned in an entry vector and finally assembled according to the Multisite Gateway^® Technology. Here, we detail the process to express zCas9 under the control of egg cell promoter fused to enhancer sequence (EC1.2en-EC1.1p) and to simultaneously target two multiple C2 domains and transmembrane region protein genes (MCTP3 and MCTP4, respectively at3g57880 and at1g51570), using one or two sgRNA per gene.

Human babesiosis is a tick-borne disease caused by Babesia pathogens. The disease, which presents with malaria-like symptoms, can be life-threatening, especially in individuals with weakened immune systems and the elderly. The worldwide prevalence of human babesiosis has been gradually rising, prompting alarm among public health experts. In other pathogens, genetic techniques have proven to be valuable tools for conducting functional studies to understand the importance of specific genes in development and pathogenesis as well as to validate novel cellular targets for drug discovery. Genetic manipulation methods have been established for several non-human Babesia and Theileria species and, more recently, have begun to be developed for human Babesia parasites. We have previously reported the development of a method for genetic manipulation of the human pathogen Babesia duncani. This method is based on positive selection using the hDHFR gene as a selectable marker, whose expression is regulated by the ef-1aB promoter, along with homology regions that facilitate integration into the gene of interest through homologous recombination. Herein, we provide a detailed description of the steps needed to implement this strategy in B. duncani to study gene function. It is anticipated that the implementation of this method will significantly improve our understanding of babesiosis and facilitate the development of novel and more effective therapeutic strategies for the treatment of human babesiosis.

Foot-and-mouth disease (FMD) is a severe and extremely contagious viral disease of cloven-hoofed domestic and wild animals, which leads to serious economic losses to the livestock industry globally. FMD is caused by the FMD virus (FMDV), a positive-strand RNA virus that belongs to the genus Aphthovirus, within the family Picornaviridae. Early detection and characterization of FMDV strains are key factors to control new outbreaks and prevent the spread of the disease. Here, we describe a direct RNA sequencing method using Oxford Nanopore Technology (ONT) Flongle flow cells on MinION Mk1C (or GridION) to characterize FMDV. This is a rapid, low cost, and easily deployed point of care (POC) method for a near real-time characterization of FMDV in endemic areas or outbreak investigation sites.

Single-cell RNA sequencing (scRNA-seq) stands as a cutting-edge technology widely used in biological and biomedical research. Existing scRNA-seq methods rely on reverse transcription (RT) and second-strand synthesis (SSS) to convert RNA to cDNA before amplification. However, these methods often suffer from limited RT/SSS efficiency, which compromises the sensitivity of RNA detection. Here, we develop a new method, linearly amplified single-stranded RNA-derived transcriptome sequencing (LAST-seq), which directly amplifies the original single-stranded RNA without prior RT and SSS and offers high-sensitivity RNA detection and a low level of technical noise in single-cell transcriptome analysis. LAST-seq has been applied to quantify transcriptional bursting kinetics in human cells, advancing our understanding of chromatin organization’s role in regulating gene expression.

Lipid nanoparticle (LNP)-based drug delivery systems (DDSs) are widely recognized for their ability to enhance efficient and precise delivery of therapeutic agents, including nucleic acids like DNA and mRNA. Despite this acknowledgment, there is a notable knowledge gap regarding the systemic biodistribution and organ accumulation of these nanoparticles. The ability to track LNPs in vivo is crucial for understanding their fate within biological systems. Fluorescent labeling of LNPs facilitates real-time tracking, quantification, and visualization of their behavior within biological systems, providing valuable insights into biodistribution, cellular uptake, and the optimization of drug delivery strategies. Our prior research established reversely engineered LNPs as an exceptional mRNA delivery platform for treating ulcerative colitis. This study presents a detailed protocol for labeling interleukin-22 (IL-22) mRNA-loaded LNPs, their oral administration to mice, and visualization of DiR-labeled LNPs biodistribution in the gastrointestinal tract using IVIS spectrum. This fluorescence-based approach will assist researchers in gaining a dynamic understanding of nanoparticle fate in other models of interest.

The cell–cell adhesion molecule E-cadherin has been intensively studied due to its prevalence in tissue function and its spatiotemporal regulation during epithelial-to-mesenchymal cell transition. Nonetheless, regulating and studying the dynamics of it has proven challenging. We developed a photoswitchable version of E-cadherin, named opto-E-cadherin, which can be toggled OFF with blue light illumination and back ON in the dark. Herein, we describe easy-to-use methods to test and characterise opto-E-cadherin cell clones for downstream experiments.

Two-dimensional (2D) agarose gel electrophoresis is the method of choice to analyze DNA topology. The possibility to use E. coli strains with different genetic backgrounds in combination with nicking enzymes and different concentrations of norfloxacin improves the resolution of 2D gels to study the electrophoretic behavior of three different families of DNA topoisomers: supercoiled DNA molecules, post-replicative catenanes, and knotted DNA molecules. Here, we describe the materials and procedures required to optimize their separation by 2D gels. Understanding the differences in their electrophoretic behavior can help explain some important physical characteristics of these different types of DNA topoisomers.

Pseudouridine (Ψ), the most prevalent modified base in cellular RNAs, has been mapped to numerous sites not only in rRNAs, tRNAs, and snRNAs but also mRNAs. Although there have been multiple techniques to identify Ψs, due to the recent development of sequencing technologies some reagents are not compatible with the current sequencer. Here, we show the updated Pseudo-seq, a technique enabling the genome-wide identification of pseudouridylation sites with single-nucleotide precision. We provide a comprehensive description of Pseudo-seq, covering protocols for RNA isolation from human cells, library preparation, and detailed data analysis procedures. The methodology presented is easily adaptable to any cell or tissue type with high-quality mRNA isolation. It can be used for discovering novel pseudouridylation sites, thus constituting a crucial initial step toward understanding the regulation and function of this modification.

The field of oligonucleotide therapeutics is rapidly advancing, particularly for combating orphan diseases and cancer. However, the intrinsic instability of oligonucleotides, especially RNA, poses a substantial challenge in the face of the harsh conditions encountered intracellularly and in circulation. Therefore, evaluating the stability of oligos in serum is of great significance when developing oligonucleotide therapeutics. This protocol outlines a dependable and reproducible method for preparing oligonucleotide duplexes, coupled with confirmation by gel electrophoresis. Subsequently, the protocol defines a mechanism to assess the stability of the oligo duplexes in serum. This protocol seeks to establish a standardized reference for researchers, enabling them to compare the impact of various modifications on oligo stability and assess the degradation kinetics effectively.

DNA methylation is a key epigenetic mechanism underlying many biological processes, and its aberrant regulation has been tightly associated with various human diseases. Precise manipulation of DNA methylation holds the promise to advance our understanding of this critical mechanism and to develop novel therapeutic methods. Previously, we were only able to alter genome-wide DNA methylation by treating with small molecules (e.g., 5-Aza-2-deoxycytidine) or perturbing relevant genes (e.g., DNA methyltransferase) targetlessly, which makes it challenging to investigate the functional significance of this epigenetic mark at specific genomic loci. By fusing the catalytic domain of a key enzyme in the DNA demethylation process (Ten-eleven translocation dioxygenases 1, Tet1) with a reprogrammable sequence-specific DNA-targeting molecular protein, dCas9, we developed a DNA methylation editing tool (dCas9-Tet1) to demethylate specific genomic loci in a targeted manner. This dCas9-Tet1 system allows us to study the role of DNA methylation at almost any given loci with only the replacement of a single-guide RNA. Here, we describe a protocol that enables modular and scalable manipulation of DNA methylation at specific genomic loci in various cell cultures with high efficiency and specificity using the dCas9-Tet1 system.