New Frontiers: Precise Editing of Allergen Genes Using CRISPR

Genome engineering with clustered regularly interspaced short palindromic repeats (CRISPR) technology offers the unique potential for unequivocally deleting allergen genes at the source. Compared to prior gene editing approaches, CRISPR boasts substantial improvements in editing efficiency, throughput, and precision. CRISPR has demonstrated success in several clinical applications such as sickle cell disease and β-thalassemia, and preliminary knockout studies of allergenic proteins using CRISPR editing show promise. Given the advantages of CRISPR, as well as specific DNA targets in the allergen genes, CRISPR gene editing is a viable approach for tackling allergy, which may lead to significant disease improvement. This review will highlight recent applications of CRISPR editing of allergens, particularly cat allergen Fel d 1, and will discuss the advantages and limitations of this approach compared to existing treatment options.

Minimal genetically encoded tags for fluorescent protein labeling in living neurons

Modern light microscopy, including super-resolution techniques, has brought about a demand for small labeling tags that bring the fluorophore closer to the target. This challenge can be addressed by labeling unnatural amino acids (UAAs) with bioorthogonal click chemistry. The minimal size of the UAA and the possibility to couple the fluorophores directly to the protein of interest with single-residue precision in living cells make click labeling unique. Here, we establish click labeling in living primary neurons and use it for fixed-cell, live-cell, dual-color pulse–chase, and super-resolution microscopy of neurofilament light chain (NFL). We also show that click labeling can be combined with CRISPR/Cas9 genome engineering for tagging endogenous NFL. Due to its versatile nature and compatibility with advanced multicolor microscopy techniques, we anticipate that click labeling will contribute to novel discoveries in the neurobiology field.

Knockout of ABC Transporter ABCG4 Gene Confers Resistance to Cry1 Proteins in Ostrinia furnacalis

Ostrinia furnacalis is an important borer on maize. Long-term and large-scale planting of transgenic corn has led O. furnacalis evolving resistance and reducing the control effect. Recently, high levels of resistance to Bt Cry1 toxins have been reported to be genetically linked to the mutation or down-regulation of ABC transporter subfamily G gene ABCG4 in O. furnacalis. In order to further determine the relationship between ABCG4 gene and the resistance to Cry1 toxins in O. furnacalis, the novel CRISPR/Cas9 genome engineering system was utilized to successfully construct ABCG4-KO knockout homozygous strain. Bioassay results indicated that an ABCG4-KO strain had a higher resistance to Cry1 proteins compared with a susceptible strain (ACB-BtS). The result indicates that the ABCG4 gene may act as a receptor of the Bt Cry1 toxin in O. furnacalis. Furthermore, the development time was significantly changed in the early stage ABCG4-KO larvae, and the population parameters were also significantly changed. In summary, our CRISPR/Cas9-mediated genome editing study presents evidence that ABCG4 gene is a functional receptor for Bt Cry1 toxins, laying the foundation for further clarification of the Bt resistance mechanism.

Allele-specific endogenous tagging and quantitative analysis of β-catenin in colorectal cancer cells

Wnt signaling plays important roles in development, homeostasis, and tumorigenesis. Mutations in β-catenin that activate Wnt signaling have been found in colorectal and hepatocellular carcinomas. However, the dynamics of wild-type and mutant forms of β-catenin are not fully understood. Here, we genome-engineered fluorescently tagged alleles of endogenous β-catenin in a colorectal cancer cell line. Wild-type and oncogenic mutant alleles were tagged with different fluorescent proteins, enabling the analysis of both variants in the same cell. We analyzed the properties of both β-catenin alleles using immunoprecipitation, immunofluorescence, and fluorescence correlation spectroscopy approaches, revealing distinctly different biophysical properties. In addition, activation of Wnt signaling by treatment with a GSK3β inhibitor or a truncating APC mutation modulated the wild-type allele to mimic the properties of the mutant β-catenin allele. The one-step tagging strategy demonstrates how genome engineering can be employed for the parallel functional analysis of different genetic variants.

Targeted editing and evolution of engineered ribosomes in vivo by filtered editing

Genome editing technologies introduce targeted chromosomal modifications in organisms yet are constrained by the inability to selectively modify repetitive genetic elements. Here we describe filtered editing, a genome editing method that embeds group 1 self-splicing introns into repetitive genetic elements to construct unique genetic addresses that can be selectively modified. We introduce intron-containing ribosomes into the E. coli genome and perform targeted modifications of these ribosomes using CRISPR/Cas9 and multiplex automated genome engineering. Self-splicing of introns post-transcription yields scarless RNA molecules, generating a complex library of targeted combinatorial variants. We use filtered editing to co-evolve the 16S rRNA to tune the ribosome’s translational efficiency and the 23S rRNA to isolate antibiotic-resistant ribosome variants without interfering with native translation. This work sets the stage to engineer mutant ribosomes that polymerize abiological monomers with diverse chemistries and expands the scope of genome engineering for precise editing and evolution of repetitive DNA sequences.

Double nicking by RNA-directed Cascade-nCas3 for high-efficiency large-scale genome engineering

New CRISPR-based genome editing technologies are developed to continually drive advances in life sciences, which, however, are predominantly derived from systems of Type II CRISPR-Cas9 and Type V CRISPR-Cas12a for eukaryotes. Here we report a novel CRISPR-n(nickase)Cas3 genome editing tool established upon a Type I-F system. We demonstrate that nCas3 variants can be created by alanine-substituting any catalytic residue of the Cas3 helicase domain. While nCas3 overproduction via plasmid shows severe cytotoxicity, an in situ nCas3 introduces targeted double-strand breaks, facilitating genome editing without visible cell killing. By harnessing this CRISPR-nCas3 in situ gene insertion, nucleotide substitution and deletion of genes or genomic DNA stretches can be consistently accomplished with near-100% efficiencies, including simultaneous removal of two large genomic fragments. Our work describes the first establishment of a CRISPR-nCas3-based genome editing technology, thereby offering a simple, yet useful approach to convert the naturally most abundantly occurring Type I systems into advanced genome editing tools to facilitate high-throughput prokaryotic engineering.

Beyond Mendelian Inheritance: Genetic Buffering and Phenotype Variability

Understanding the way genes work amongst individuals and across generations to shape form and function is a common theme for many genetic studies. The recent advances in genetics, genome engineering and DNA sequencing reinforced the notion that genes are not the only players that determine a phenotype. Due to physiological or pathological fluctuations in gene expression, even genetically identical cells can behave and manifest different phenotypes under the same conditions. Here, we discuss mechanisms that can influence or even disrupt the axis between genotype and phenotype; the role of modifier genes, the general concept of genetic redundancy, genetic compensation, the recently described transcriptional adaptation, environmental stressors, and phenotypic plasticity. We furthermore highlight the usage of induced pluripotent stem cells (iPSCs), the generation of isogenic lines through genome engineering, and sequencing technologies can help extract new genetic and epigenetic mechanisms from what is hitherto considered ‘noise’.

Genetic Engineering of Streptomyces ghanaensis ATCC14672 for Improved Production of Moenomycins

Streptomycetes are soil-dwelling multicellular microorganisms famous for their unprecedented ability to synthesize numerous bioactive natural products (NPs). In addition to their rich arsenal of secondary metabolites, Streptomyces are characterized by complex morphological differentiation. Mostly, industrial production of NPs is done by submerged fermentation, where streptomycetes grow as a vegetative mycelium forming pellets. Often, suboptimal growth peculiarities are the major bottleneck for industrial exploitation. In this work, we employed genetic engineering approaches to improve the production of moenomycins (Mm) in Streptomyces ghanaensis, the only known natural direct inhibitors of bacterial peptidoglycan glycosyltransferses. We showed that in vivo elimination of binding sites for the pleiotropic regulator AdpA in the oriC region strongly influences growth and positively correlates with Mm accumulation. Additionally, a marker- and “scar”-less deletion of moeH5, encoding an amidotransferase from the Mm gene cluster, significantly narrows down the Mm production spectrum. Strikingly, antibiotic titers were strongly enhanced by the elimination of the pleiotropic regulatory gene wblA, involved in the late steps of morphogenesis. Altogether, we generated Mm overproducers with optimized growth parameters, which are useful for further genome engineering and chemoenzymatic generation of novel Mm derivatives. Analogously, such a scheme can be applied to other Streptomyces spp.

Multiplexed CRISPR-mediated engineering of protein secretory pathway genes in the thermotolerant methylotrophic yeast Ogataea thermomethanolica

CRISPR multiplex gRNA systems have been employed in genome engineering in various industrially relevant yeast species. The thermotolerant methylotrophic yeast Ogataea thermomethanolica TBRC 656 is an alternative host for heterologous protein production. However, the limited secretory capability of this yeast is a bottleneck for protein production. Here, we refined CRISPR-based genome engineering tools for simultaneous mutagenesis and activation of multiple protein secretory pathway genes to improve heterologous protein secretion. We demonstrated that multiplexed CRISPR-Cas9 mutation of up to four genes (SOD1, VPS1, YPT7 and YPT35) in one single cell is practicable. We also developed a multiplexed CRISPR-dCas9 system which allows simultaneous activation of multiple genes in this yeast. 27 multiplexed gRNA combinations were tested for activation of three genes (SOD1, VPS1 and YPT7), three of which were demonstrated to increase the secretion of fungal xylanase and phytase up to 29% and 41%, respectively. Altogether, our study provided a toolkit for mutagenesis and activation of multiple genes in O. thermomethanolica, which could be useful for future strain engineering to improve heterologous protein production in this yeast.