Molecular Biology

We established a step-by-step approach for generating a single-nucleotide mutation in the promoter region of an immune regulatory gene in human monocyte THP-1 cells by employing a plasmid-based CRISPR-Cas9 system delivered via transfection with a homology-directed repair template DNA (HDR). Key steps include designing a single-guide RNA (sgRNA), cloning it into a CRISPR plasmid encoding the Cas9 protein, transfection of the plasmid constructs along with single-stranded oligonucleotide repair template (ssODNs) into THP-1 cells, followed by selection and validation. This approach provides a precise and relevant model to investigate the role of single polymorphisms in the regulation of inflammatory gene expression in human monocytes. In addition to the rs1024611 single-nucleotide polymorphism (SNP), this CRISPR/Cas9-based strategy is broadly applicable to functional studies of noncoding and coding variants in innate immune genes.

Engineering of microbial cells, including E. coli, is essential in prototyping genetic designs used in numerous applications throughout synthetic biology. While many advanced genome editing tools, such as CRISPR-based tools, offer new capabilities with genetically recalcitrant organisms, these tools often do not offer an immediate advantage in readily manipulated microbes, such as E. coli, especially when scarless modifications are not critical. We describe a comprehensive recombineering tutorial that we commonly use for multiplex engineering of E. coli using antibiotic markers. We leverage a group of 15 antibiotic resistance cassettes, most of which can be readily included when designing double-stranded DNA donors intended for recombineering and purchased from several vendors. Using these methods, 10–15 defined modifications to a single host strain can be achieved in less than three weeks, using two-day editing cycles. We discuss sequences and protocols as well as the optimal design of genetic modifications and the associated DNA.

Labeling cells with reporter genes allows researchers to visually identify specific cells and observe how they interact with each other in dynamic biological systems. Even though various labeling methods are now available, a specific description of gene knock-in labeling methods for human trophoblast stem cells (hTSCs) has not been reported. Here, we present a streamlined protocol for labeling hTSCs with the green fluorescent protein (GFP) reporter gene via CRISPR/Cas9-mediated knock-in of the gene into the adeno-associated virus site 1 (AAVS1) safe harbor locus. A commonly used hTSC cell line, CT29, was transfected with a dual plasmid system encoding the Cas9 endonuclease and an AAVS1-targeted guide RNA in one plasmid and a donor plasmid encoding a puromycin resistance gene and GFP reporter gene flanked by AAVS1 homology arms. Puromycin-resistant clonal cells were isolated, and AAVS1 integration was confirmed via PCR and sequencing of the PCR products. The labeled cells are proliferative and can give rise to extravillous cytotrophoblast cells (EVT) and the syncytiotrophoblast (ST). To our knowledge, this is the first report using the CRISPR/Cas9 system for AAVS1 integration of a reporter gene in human trophoblast stem cells. It provides an efficient tool to facilitate the study of human trophoblast development and function in co-culture systems and will be highly useful in developing clinical gene therapy-related plasmid constructs.

The standard protocols for allelic exchange using homologous recombination deploy suicide vectors with negative selection markers. However, the use of multiple restriction enzymes to generate sticky ends in the vector and the insert for cloning is time-consuming, resource-intensive, and challenging. The advent of next-generation proofreading enzymes is enabling researchers to routinely carry out long-range PCR. Hence, amplifying 5–6 kb of complete low-complex DNA cloning vectors and 2–3 kb of complex genomic regions is much easier. Here, we report a simple, accurate, rapid, and unidirectional approach for chromosomal in-frame gene deletion and complementation by reconstitution of the full-length gene without using any restriction enzymes. The method requires long-range PCR using Phusion polymerase to linearize the vector and amplify the target gene to create a recombinant vector (pRM1) and further inverse PCR amplification of pRM1 to create a recombinant vector (pRM4) with a deleted version of the gene. The cloning steps involve the use of kinase and ligase for phosphorylation and ligation steps, respectively. The recombinant plasmid, pRM4, is finally transformed into electrocompetent cells of Xanthomonas sontii, a gram-negative phytobacterium, for final genomic integration/excision to obtain an in-frame gene deletion mutant (PPL1RM15). Gene reconstitution for complementation is carried out by electroporating the deletion mutant with the recombinant plasmid (pRM1) carrying the wild-type allele. Clean gene mutation, allele restoration, and plasmid excision are confirmed using whole-genome sequencing.

Chloroplast genomes present an alternative strategy for large-scale engineering of photosynthetic eukaryotes. Prior to our work, the chloroplast genomes of Chlamydomonas reinhardtii (204 kb) and Zea mays (140 kb) had been cloned using bacterial and yeast artificial chromosome (BAC/YAC) libraries, respectively. These methods lack design flexibility as they are reliant upon the random capture of genomic fragments during BAC/YAC library creation; additionally, both demonstrated a low efficiency (≤ 10%) for correct assembly of the genome in yeast. With this in mind, we sought to create a highly flexible and efficient approach for assembling the 117 kb chloroplast genome of Phaeodactylum tricornutum, a photosynthetic marine diatom. Our original article demonstrated a PCR-based approach for cloning the P. tricornutum chloroplast genome that had 90%–100% efficiency when screening as few as 10 yeast colonies following assembly. In this article, we will discuss this approach in greater depth as we believe this technique could be extrapolated to other species, particularly those with a similar chloroplast genome size and architecture.

This protocol describes the generation of chimeric mice in which the Y chromosome is deleted from a proportion of blood cells. This model recapitulates the phenomenon of hematopoietic mosaic loss of Y chromosome (mLOY), which is frequently observed in the blood of aged men. To construct mice with hematopoietic Y chromosome loss, lineage-negative cells are isolated from the bone marrow of ROSA26-Cas9 knock-in mice. These cells are transduced with a lentivirus vector encoding a guide RNA (gRNA) that targets multiple repeats of the Y chromosome centromere, effectively removing the Y chromosome. These cells are then transplanted into lethally irradiated wildtype C57BL6 mice. Control gRNAs are designed to target either no specific region or the fourth intron of Actin gene. Transduced cells are tracked by measuring the fraction of blood cells expressing the virally encoded reporter gene tRFP. This model represents a clinically relevant model of hematopoietic mosaic loss of Y chromosome, which can be used to study the impact of mLOY on various age-related diseases.

Graphical overview

The CRISPR/Cas9 system is a powerful tool for gene repair that holds great potential for gene therapy to cure monogenic diseases. Despite intensive improvement, the safety of this system remains a major clinical concern. In contrast to Cas9 nuclease, Cas9 nickases with a pair of short-distance (38–68 bp) PAM-out single-guide RNAs (sgRNAs) preserve gene repair efficiency while strongly reducing off-target effects. However, this approach still leads to efficient unwanted on-target mutations that may cause tumorigenesis or abnormal hematopoiesis. We establish a precise and safe spacer-nick gene repair approach that combines Cas9^D10A nickase with a pair of PAM-out sgRNAs at a distance of 200–350 bp. In combination with adeno-associated virus (AAV) serotype 6 donor templates, this approach leads to efficient gene repair with minimal unintended on- and off-target mutations in human hematopoietic stem and progenitor cells (HSPCs). Here, we provide detailed protocols to use the spacer-nick approach for gene repair and to assess the safety of this system in human HSPCs. The spacer-nick approach enables efficient gene correction for repair of disease-causing mutations with increased safety and suitability for gene therapy.

Graphical overview

The ongoing COVID-19 pandemic is caused by severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2). As this virus is classified as a biosafety level-3 (BSL-3) agent, the development of countermeasures and basic research methods is logistically difficult. Recently, using reverse genetics, we developed a BSL-2 cell culture system for production of transcription- and replication-component virus-like-particles (trVLPs) by genetic transcomplementation. The system consists of two parts: SARS-CoV-2 GFP/ΔN genomic RNA, in which the nucleocapsid (N) gene, a critical gene for virion packaging, is replaced by a GFP reporter gene; and a packaging cell line for ectopic expression of N (Caco-2-N). The complete viral life cycle can be recapitulated and confined to Caco-2-N cells, with GFP positivity serving as a surrogate readout for viral infection. In addition, we utilized an intein-mediated protein splicing technique to split the N gene into two independent vectors and generated the Caco-2-N^intein cells as a packaging cell line to further enhance the security of this cell culture model. Altogether, this system provides for a safe and convenient method to produce trVLPs in BSL-2 laboratories. These trVLPs can be modified to incorporate desired mutations, permitting high-throughput screening of antiviral compounds and evaluation of neutralizing antibodies. This protocol describes the details of the trVLP cell culture model to make SARS-CoV-2 research more readily accessible.