Genome Engineering Methods Research Articles

Patterns of Aneuploidy and Signaling Consequences in Cancer.

Aneuploidy, or a change in the number of whole chromosomes or chromosome arms, is a near-universal feature of cancer. Chromosomes affected by aneuploidy are not random, with observed cancer-specific and tissue-specific patterns. Recent advances in genome engineering methods have allowed the creation of models with targeted aneuploidy events. These models can be used to uncover the downstream effects of individual aneuploidies on cancer phenotypes including proliferation, apoptosis, metabolism, and immune signaling. Here, we review the current state of research into the patterns of aneuploidy in cancer and their impact on signaling pathways and biological processes.

Graded-CRISPRi, a Tool for Tuning the Strengths of CRISPRi-Mediated Knockdowns in Vibrio natriegens Using gRNA Libraries.

In recent years, the fast-growing bacterium Vibrio natriegens has gained increasing attention as it has the potential to become a next-generation chassis for synthetic biology. A wide range of genetic parts and genome engineering methods have already been developed. However, there is still a need for a well-characterized tool to effectively and gradually reduce the expression levels of native genes. To bridge this gap, we created graded-CRISPRi, a system utilizing gRNA variants that lead to varying levels of repression strength. By incorporating multiple gRNA sequences into our design, we successfully extended this concept to simultaneously repress four distinct reporter genes. Furthermore, we demonstrated the capability of using graded-CRISPRi to target native genes, thereby examining the effect of various knockdown levels on growth.

Comparing Methods to Genetically Engineer Bacteriophage and Increase Host Range.

Antibacterial resistance is an emerging problem in military medicine. Disruptions to the health care systems in war-torn countries that result from ongoing conflict can potentially exacerbate this problem and increase the risk to U.S. forces in the deployed environment. Therefore, novel therapies are needed to mitigate the impact of these potentially devastating infections on military operations. Bacteriophages are viruses that infect and kill bacteria. They can be delivered as therapeutic agents and offer a promising alternative to traditional antibiotic chemotherapy. There are several potential benefits to their use, including high specificity and comparative ease of use in the field setting. However, the process of engineering phages for military medical applications can be a laborious and time-consuming endeavor. This review examines available techniques and compares their efficacy. This review evaluates the scientific literature on the development and application of four methods of bacteriophage genome engineering and their consideration in the context of military applications. Preffered Reporting Items for Systematic Reviews and Meta-Analyses (PRISMA) guidelines were followed for a systematic review of available literature that met criteria for analysis and inclusion. The research completed for this review article originated from the United States Military Academy's library "Scout" search engine, which compiles results from 254 available databases (including PubMed, Google Scholar, and SciFinder). Particular attention was focused on identifying useful mechanistic insight into the nature of the engineering technique, the ease of use, and the applicability of the technique to countering the problem of antimicrobial resistance in the military setting. A total of 52 studies were identified that met inclusion criteria following PRISMA guidelines. The bioengineering techniques analyzed included homologous recombination (12 articles), in vivo recombineering (9 articles), bacteriophage recombineering of electroporated DNA (7 articles), and the CRISPR-Cas system (10 articles). Rates of success and fidelity varied across each platform, and comparative benefits and drawbacks are considered. Each of the phage engineering techniques addressed herein varies in amount of effort and overall success rate. CRISPR-Cas-facilitated modification of phage genomes presents a highly efficient method that does not require a lengthy purification and screening process. It therefore appears to be the method best suited for military medical applications.

Structural Basis for the Interaction of Redβ Single-Strand Annealing Protein with Escherichia coli Single-Stranded DNA-Binding Protein

Redβ is a protein from bacteriophage λ that binds to single-stranded DNA (ssDNA) to promote the annealing of complementary strands. Together with λ-exonuclease (λ-exo), Redβ is part of a two-component DNA recombination system involved in multiple aspects of genome maintenance. The proteins have been exploited in powerful methods for bacterial genome engineering in which Redβ can anneal an electroporated oligonucleotide to a complementary target site at the lagging strand of a replication fork. Successful annealing in vivo requires the interaction of Redβ with E. coli single-stranded DNA-binding protein (SSB), which coats the ssDNA at the lagging strand to coordinate access of numerous replication proteins. Previous mutational analysis revealed that the interaction between Redβ and SSB involves the C-terminal domain (CTD) of Redβ and the C-terminal tail of SSB (SSB-Ct), the site for binding of numerous host proteins. Here, we have determined the x-ray crystal structure of Redβ CTD in complex with a peptide corresponding to the last nine residues of SSB (MDFDDDIPF). Formation of the complex is predominantly mediated by hydrophobic interactions between two phenylalanine side chains of SSB (Phe-171 and Phe-177) and an apolar groove on the CTD, combined with electrostatic interactions between the C-terminal carboxylate of SSB and Lys-214 of the CTD. Mutation of any of these residues to alanine significantly disrupts the interaction of full-length Redβ and SSB proteins. Structural knowledge of this interaction will help to expand the utility of Redβ-mediated recombination to a wider range of bacterial hosts for applications in synthetic biology.

Omics-driven exploration and mining of key functional genes for the improvement of food and fiber crops.

The deployment of omics technologies has obtained an incredible boost over the past few decades with the advances in next-generation sequencing (NGS) technologies, innovative bioinformatics tools, and the deluge of available biological information. The major omics technologies in the limelight are genomics, transcriptomics, proteomics, metabolomics, and phenomics. These biotechnological advances have modernized crop breeding and opened new horizons for developing crop varieties with improved traits. The genomes of several crop species are sequenced, and a huge number of genes associated with crucial economic traits have been identified. These identified genes not only provide insights into the understanding of regulatory mechanisms of crop traits but also decipher practical grounds to assist in the molecular breeding of crops. This review discusses the potential of omics technologies for the acquisition of biological information and mining of the genes associated with important agronomic traits in important food and fiber crops, such as wheat, rice, maize, potato, tomato, cassava, and cotton. Different functional genomics approaches for the validation of these important genes are also highlighted. Furthermore, a list of genes discovered by employing omics approaches is being represented as potential targets for genetic modifications by the latest genome engineering methods for the development of climate-resilient crops that would in turn provide great impetus to secure global food security.

Targeted Gene Editing of Nuclear-Encoded Plastid Proteins in Phaeodactylum tricornutum via CRISPR/Cas9.

Genome modifications in microalgae have emerged as a crucial and indispensable tool for research in fundamental and applied biology. In particular, CRISPR/Cas9 has gained significant recognition as a highly effective method for genome engineering in these photosynthetic organisms, enabling the targeted induction of mutations in specific regions of the genome. Here, we present a comprehensive protocol for generating knock-out mutants in the model diatom Phaeodactylum tricornutum using CRISPR/Cas9 by both biolistic transformation and bacterial conjugation. Our protocol outlines the step-by-step procedures and experimental conditions required to achieve successful genome editing, including the design and construction of guide RNAs, the delivery of CRISPR/Cas9 components into the algae cells, and the selection of the generated knockout mutants. Through the implementation of this protocol, researchers can harness the potential of CRISPR/Cas9 in P. tricornutum to advance the understanding of diatom biology and explore their potential applications in various fields.

Use of Cas9 Targeting and Red Recombination for Designer Phage Engineering.

Bacteriophages (phages) are natural antibiotics and biological nanoparticles, whose application is significantly boosted by recent advances of synthetic biology tools. Designer phages are synthetic phages created by genome engineering in a way to increase the benefits or decrease the drawbacks of natural phages. Here we report the development of a straightforward genome engineering method to efficiently obtain engineered phages in a model bacterial pathogen, Pseudomonas aeruginosa. This was achieved by eliminating the wild type phages based on the Streptococcus pyogenes Cas9 (SpCas9) and facilitating the recombinant generation based on the Red recombination system of the coliphage λ (λRed). The producer (PD) cells of P. aeruginosa strain PAO1 was created by miniTn7-based chromosomal integration of the genes for SpCas9 and λRed under an inducible promoter. To validate the efficiency of the recombinant generation, we created the fluorescent phages from a temperate phage MP29. A plasmid bearing the single guide RNA (sgRNA) gene for selectively targeting the wild type gp35 gene and the editing template for tagging the Gp35 with superfolder green fluorescent protein (sfGFP) was introduced into the PD cells by electroporation. We found that the targeting efficiency was affected by the position and number of sgRNA. The fluorescent phage particles were efficiently recovered from the culture of the PD cells expressing dual sgRNA molecules. This protocol can be used to create designer phages in P. aeruginosa for both application and research purposes.

Application of Nanotechnology in Plant Genetic Engineering.

The ever-increasing food requirement with globally growing population demands advanced agricultural practices to improve grain yield, to gain crop resilience under unpredictable extreme weather, and to reduce production loss caused by insects and pathogens. To fulfill such requests, genome engineering technology has been applied to various plant species. To date, several generations of genome engineering methods have been developed. Among these methods, the new mainstream technology is clustered regularly interspaced short palindromic repeats (CRISPR) with nucleases. One of the most important processes in genome engineering is to deliver gene cassettes into plant cells. Conventionally used systems have several shortcomings, such as being labor- and time-consuming procedures, potential tissue damage, and low transformation efficiency. Taking advantage of nanotechnology, the nanoparticle-mediated gene delivery method presents technical superiority over conventional approaches due to its high efficiency and adaptability in different plant species. In this review, we summarize the evolution of plant biomolecular delivery methods and discussed their characteristics as well as limitations. We focused on the cutting-edge nanotechnology-based delivery system, and reviewed different types of nanoparticles, preparation of nanomaterials, mechanism of nanoparticle transport, and advanced application in plant genome engineering. On the basis of established methods, we concluded that the combination of genome editing, nanoparticle-mediated gene transformation and de novo regeneration technologies can accelerate crop improvement efficiently in the future.

Precision Genome Engineering in Streptococcus suis Based on a Broad-Host-Range Vector and CRISPR-Cas9 Technology.

Streptococcussuis is an important zoonotic pathogen that causes severe invasive disease in pigs and humans. Current methods for genome engineering of S. suis rely on the insertion of antibiotic resistance markers, which is time-consuming and labor-intensive and does not allow the precise introduction of small genomic mutations. Here we developed a system for CRISPR-based genome editing in S. suis, utilizing linear DNA fragments for homologous recombination (HR) and a plasmid-based negative selection system for bacteria not edited by HR. To enable the use of this system in other bacteria, we engineered a broad-host-range replicon in the CRISPR plasmid. We demonstrated the utility of this system to rapidly introduce multiple gene deletions in successive rounds of genome editing and to make precise nucleotide changes in essential genes. Furthermore, we characterized a mechanism by which S. suis can escape killing by a targeted Cas9-sgRNA complex in the absence of HR. A characteristic of this new mechanism is the presence of very slow-growing colonies in a persister-like state that may allow for DNA repair or the introduction of mutations, alleviating Cas9 pressure. This does not impact the utility of CRISPR-based genome editing because the escape colonies are easily distinguished from genetically edited clones due to their small colony size. Our CRISPR-based editing system is a valuable addition to the genetic toolbox for engineering of S. suis, as it accelerates the process of mutant construction and simplifies the removal of antibiotic markers between successive rounds of genome editing.

Bacteriophage genome engineering for phage therapy to combat bacterial antimicrobial resistance as an alternative to antibiotics.

Bacteriophages (phages) are viruses that mainly infect bacteria and are ubiquitously distributed in nature, especially to their host. Phage engineering involves nucleic acids manipulation of phage genome for antimicrobial activity directed against pathogens through the applications of molecular biology techniques such as synthetic biology methods, homologous recombination, CRISPY-BRED and CRISPY-BRIP recombineering, rebooting phage-based engineering, and targeted nucleases including CRISPR/Cas9, zinc-finger nucleases (ZFNs) and transcription activator-like effector nucleases (TALENs). Management of bacteria is widely achieved using antibiotics whose mechanism of action has been shown to target both the genetic dogma and the metabolism of pathogens. However, the overuse of antibiotics has caused the emergence of multidrug-resistant (MDR) bacteria which account for nearly 5million deaths as of 2019 thereby posing threats to the public health sector, particularly by 2050. Lytic phages have drawn attention as a strong alternative to antibiotics owing to the promising efficacy and safety of phage therapy in various models in vivo and human studies. Therefore, harnessing phage genome engineering methods, particularly CRISPR/Cas9 to overcome the limitations such as phage narrow host range, phage resistance or any potential eukaryotic immune response for phage-based enzymes/proteins therapy may designate phage therapy as a strong alternative to antibiotics for combatting bacterial antimicrobial resistance (AMR). Here, the current trends and progress in phage genome engineering techniques and phage therapy are reviewed.

Genome Engineering of Human Urine-Derived Stem Cells to Express Lactoferrin and Deoxyribonuclease.

Urine-derived stem cells (USCs) are adult kidney cells that have been isolated from a urine sample and propagated in tissue culture on gelatin-coated plates. Urine is a practical and completely painless source of cells for gene and cell therapy applications. We have isolated, expanded, and optimized transfection of USCs to develop regenerative therapies based on piggyBac transposon modification. USCs from a healthy donor sample were isolated according to established protocols. Within 2 months, 10 clones had been expanded, analyzed, and frozen. Fluorescence-activated cell sorting analysis of individual clones revealed that all 10 clones expressed characteristic USC markers (97-99% positive for CD44, CD73, CD90, and CD146; negative for CD31, CD34, and CD45). The isolated USCs were successfully differentiated along the osteogenic, adipogenic, and chondrogenic lineages, suggesting multipotent differentiation capacity. Additionally, the USCs were differentiated into podocytes positive for NEPHRIN (NPHS1), podocalyxin, and Wilms tumor 1 (WT1). Transfection of USCs with a strongly expressing Green fluorescent protein plasmid was optimized to achieve 61% efficiency in live cells using several commercially available lipophilic reagents. Transgene promoters were compared in five luciferase-expressing piggyBac transposons by live animal imaging. The CMV promoter produced the highest luciferase signal, followed by EF1-α. Finally, HEK-293 and USCs were transfected with piggyBac transposons expressing lactoferrin and DNase1 for treatment of acute kidney injury associated with rhabdomyolysis. We found that both proteins were expressed in USCs and that lactoferrin was successfully secreted into the cell culture media. In conclusion, USCs represent a clinically relevant cell type that can express nonviral transgenes. Impact statement Acute kidney injury (AKI) affects over 13 million people worldwide each year, with hospitalization rates on the rise. There are no therapies that directly regenerate the kidney after AKI. Each human kidney contains approximately one million nephrons that process ∼100 L of urinary filtrate each day. Thousands of kidney cells become detached and are excreted in the urine. A small percentage of these cells can be clonally derived into urine-derived stem cells. We have optimized methods for genome engineering of adult human urine-derived stem cells for future applications in regenerative approaches to treat kidney injury.

Editing of Phage Genomes - Recombineering-Assisted SpCas9 Modification of Model Coliphages T7, T5, and T3

Bacteriophages-viruses that infect bacterial cells - are the most abundant biological entities on Earth. The use of phages in fundamental research and industry requires tools for precise manipulation of their genomes. Yet, compared to bacterial genome engineering, modification of phage genomes is challenging because of the lack of selective markers and thus requires laborious screenings of recombinant/mutated phage variants. The development of the CRISPR-Cas technologies allowed to solve this issue by the implementation of negative selection that eliminates the parental phage genomes. In this manuscript, we summarize current methods of phage genome engineering and their coupling with CRISPR-Cas technologies. We also provide examples of our successful application of these methods for introduction of specific insertions, deletions, and point mutations in the genomes of model Escherichia coli lytic phages T7, T5, and T3.

Editing of Phage Genomes—Recombineering-assisted SpCas9 Modification of Model Coliphages T7, T5, and T3

Abstract Bacteriophages—viruses that infect bacterial cells—are the most abundant biological entities on Earth. The use of phages in fundamental research and industry requires tools for precise manipulation of their genomes. Yet, compared to bacterial genome engineering, modification of phage genomes is challenging because of the lack of selective markers and thus requires laborious screenings of recombinant/mutated phage variants. The development of the CRISPR-Cas technologies allowed to solve this issue by the implementation of negative selection that eliminates the parental phage genomes. In this manuscript, we summarize current methods of phage genome engineering and their coupling with CRISPR-Cas technologies. We also provide examples of our successful application of these methods for introduction of specific insertions, deletions, and point mutations in the genomes of model Escherichia coli lytic phages T7, T5, and T3.

Comprehending the evolution of gene editing platforms for crop trait improvement.

CRISPR (Clustered Regularly Interspaced Short Palindromic Repeats)/Cas (CRISPR-associated) system was initially discovered as an underlying mechanism for conferring adaptive immunity to bacteria and archaea against viruses. Over the past decade, this has been repurposed as a genome-editing tool. Numerous gene editing-based crop improvement technologies involving CRISPR/Cas platforms individually or in combination with next-generation sequencing methods have been developed that have revolutionized plant genome-editing methodologies. Initially, CRISPR/Cas nucleases replaced the earlier used sequence-specific nucleases (SSNs), such as zinc-finger nucleases (ZFNs) and transcription activator-like effector nucleases (TALENs), to address the problem of associated off-targets. The adaptation of this platform led to the development of concepts such as epigenome editing, base editing, and prime editing. Epigenome editing employed epi-effectors to manipulate chromatin structure, while base editing uses base editors to engineer precise changes for trait improvement. Newer technologies such as prime editing have now been developed as a “search-and-replace” tool to engineer all possible single-base changes. Owing to the availability of these, the field of genome editing has evolved rapidly to develop crop plants with improved traits. In this review, we present the evolution of the CRISPR/Cas system into new-age methods of genome engineering across various plant species and the impact they have had on tweaking plant genomes and associated outcomes on crop improvement initiatives.

Mobile Translation Systems Generate Genomically Engineered Escherichia coli Cells with Improved Growth Phenotypes.

Cellular translation is responsible for the synthesis of proteins, a highly diverse class of macromolecules that form the basis of biological function. In Escherichia coli, harnessing and engineering of the biomolecular components of translation, such as ribosomes, transfer RNAs (tRNAs), and aminoacyl-tRNA synthetases, has led to both biotechnology products and an expanded genetic code. However, the engineering potential of molecular translation is hampered by the limited capabilities for rapidly sampling the large genomic space necessary to evolve well-coordinated synthetic translation networks inside cells. To address this limitation, we developed a genome engineering method inspired by the action of mobile genetic elements termed mobilization. Mobilization utilizes the stochastic action of the recombinase flippase (FLP) to generate up to ∼400 million genomic insertions, deletions, or rearrangements at flippase recognition target sites per milliliter of culture per OD in living E. coli cells. As a model, we applied our approach to evolve faster-growing E. coli strains living exclusively off genomically expressed tethered ribosomes. In an iterative "pulse-passaging scheme," we generated genomic libraries of cells via induction of FLP recombinase (pulse) followed by passaging the population without induction of FLP to enrich the resulting population for cells with higher fitness. We observed large structural genomic diversity across these cells, with the fastest growing strains exhibiting a 71% increase in growth rate compared to the ancestral strain. We anticipate that both these strains and the mobilization method will be useful tools for synthetic biology efforts to engineer translation systems.

Genome Engineering of the Fast-Growing Mycoplasma feriruminatoris toward a Live Vaccine Chassis.

Development of a new generation of vaccines is a key challenge for the control of infectious diseases affecting both humans and animals. Synthetic biology methods offer new ways to engineer bacterial chassis that can be used as vectors to present heterologous antigens and train the immune system against pathogens. Here, we describe the construction of a bacterial chassis based on the fast-growing Mycoplasma feriruminatoris, and the first steps toward its application as a live vaccine against contagious caprine pleuropneumonia (CCPP). To do so, the M. feriruminatoris genome was cloned in yeast, modified by iterative cycles of Cas9-mediated deletion of loci encoding virulence factors, and transplanted back in Mycoplasma capricolum subsp. capricolum recipient cells to produce the designed M. feriruminatoris chassis. Deleted genes encoded the glycerol transport and metabolism systems GtsABCD and GlpOKF and the Mycoplasma Ig binding protein-Mycoplasma Ig protease (MIB-MIP) immunoglobulin cleavage system. Phenotypic assays of the M. feriruminatoris chassis confirmed the corresponding loss of H2O2 production and IgG cleavage activities, while growth remained unaltered. The resulting mycoplasma chassis was further evaluated as a platform for the expression of heterologous surface proteins. A genome locus encoding an inactivated MIB-MIP system from the CCPP-causative agent Mycoplasma capricolum subsp. capripneumoniae was grafted in replacement of its homolog at the original locus in the chassis genome. Both heterologous proteins were detected in the resulting strain using proteomics, confirming their expression. This study demonstrates that advanced genome engineering methods are henceforth available for the fast-growing M. feriruminatoris, facilitating the development of novel vaccines, in particular against major mycoplasma diseases.

Efficient Large-Scale and Scarless Genome Engineering Enables the Construction and Screening of Bacillus subtilis Biofuel Overproducers.

Bacillus subtilis is a versatile microbial cell factory that can produce valuable proteins and value-added chemicals. Long fragment editing techniques are of great importance for accelerating bacterial genome engineering to obtain desirable and genetically stable host strains. Herein, we develop an efficient CRISPR-Cas9 method for large-scale and scarless genome engineering in the Bacillus subtilis genome, which can delete up to 134.3 kb DNA fragments, 3.5 times as long as the previous report, with a positivity rate of 100%. The effects of using a heterologous NHEJ system, linear donor DNA, and various donor DNA length on the engineering efficiencies were also investigated. The CRISPR-Cas9 method was then utilized for Bacillus subtilis genome simplification and construction of a series of individual and cumulative deletion mutants, which are further screened for overproducer of isobutanol, a new generation biofuel. These results suggest that the method is a powerful genome engineering tool for constructing and screening engineered host strains with enhanced capabilities, highlighting the potential for synthetic biology and metabolic engineering.

CRISPR somatic genome engineering and cancer modeling in the mouse pancreas and liver.

Genetically engineered mouse models (GEMMs) transformed the study of organismal disease phenotypes but are limited by their lengthy generation in embryonic stem cells. Here, we describe methods for rapid and scalable genome engineering in somatic cells of the liver and pancreas through delivery of CRISPR components into living mice. We introduce the spectrum of genetic tools, delineate viral and nonviral CRISPR delivery strategies and describe a series of applications, ranging from gene editing and cancer modeling to chromosome engineering or CRISPR multiplexing and its spatio-temporal control. Beyond experimental design and execution, the protocol describes quantification of genetic and functional editing outcomes, including sequencing approaches, data analysis and interpretation. Compared to traditional knockout mice, somatic GEMMs face an increased risk for mouse-to-mouse variability because of the higher experimental demands of the procedures. The robust protocols described here will help unleash the full potential of somatic genome manipulation. Depending on the delivery method and envisaged application, the protocol takes 3-5 weeks.

Viral Generation, Packaging, and Transduction on a Digital Microfluidic Platform.

Viral-based systems are a popular delivery method for introducing exogenous genetic material into mammalian cells. Unfortunately, the preparation of lentiviruses containing the machinery to edit the cells is labor-intensive, with steps requiring optimization and sensitive handling. To mitigate these challenges, we introduce the first microfluidic method that integrates lentiviral generation, packaging, and transduction. The new method allows the production of viral titers between 106 and 107 (similar to macroscale production) and high transduction efficiency for hard-to-transfect cell lines. We extend the technique for gene editing applications and show how this technique can be used to knock out and knock down estrogen receptor gene─a gene prominently responsible for 70% of breast cancer cases. This new technique is automated with multiplexing capabilities, which have the potential to standardize the methods for viral-based genome engineering.

English

The genome engineering methods have gained much appreciation for hitting plant disease resistance panorama since the last decade. The in vitro regeneration system is the foundation part of any research related to genetic engineering in plants. Hence, we established a tissue culture regime for efficient in vitro regeneration response of potato cultivar Lady Rosetta tagged with low to no resistance to late blight on foliage and medium to low resistance to late blight on tubers in potato. In this study, five callus induction media and seven regeneration media were investigated to get the best in vitro regeneration response in Lady Rosetta. Based on a statistical analysis of recorded data, the selected combination was CIM2-28 days old calli-RM7. The genetic fidelity of in vitro regenerated plants on a selected combination was assessed using the SSR markers. The wild-type and in vitro regenerated plants showed the same DNA profile. Moreover, a minimal inhibitory dose of basta was optimized for selecting genetically transformed plants in potato, and a 9mg /L basta dose was selected.