Multiplex genome editing of microorganisms using CRISPR-Cas

The Influence of Homologous Arm Length on Homologous Recombination Gene Editing Efficiency Mediated by SSB/CRISPR-Cas9 in Escherichia coli

The precise editing of genes mediated by CRISPR-Cas9 necessitates the application of donor DNA with appropriate lengths of homologous arms and fragment sizes. Our previous development, SSB/CRISPR-Cas9, has demonstrated high efficiency in homologous recombination and non-homologous end joining gene editing within bacteria. In this study, we optimized the lengths and sizes of homologous arms of the donor DNA within this system. Two sets of donor DNA constructs were generated: one set comprised donors with only 10-100 bp homologous arms, while the other set included donors with homologous arms ranging from 10-100 bp, between which was a tetracycline resistance expression cassette (1439 bp). These donor constructs were transformed into Escherichia coli MG1655 cells alongside pCas-SSB/pTargetF-lacZ. Notably, when the homologous arms ranged from 10 to 70 bp, the transformation efficiency of non-selectable donors was significantly higher than that of selectable donors. However, within the range of 10-100 bp homologous arm lengths, the homologous recombination rate of selectable donors was significantly higher than that of non-selectable donors, with the gap narrowing as the homologous arm length increased. For selectable donor DNA with homologous arm lengths of 10-60 bp, the homologous recombination rate increased linearly, reaching a plateau when the homologous arm length was between 60-100 bp. Conversely, for non-selectable donor DNA, the homologous recombination rate increased linearly with homologous arm lengths of 10-90 bp, plateauing at 90-100 bp. Editing two loci simultaneously with 100 bp homologous arms, whether selectable or non-selectable, showed no difference in transformation or homologous recombination rates. Editing three loci simultaneously with 100 bp non-selectable homologous arms resulted in a 45% homologous recombination rate. These results suggest that efficient homologous recombination gene editing mediated by SSB/CRISPR-Cas9 can be achieved using donor DNA with 90-100 bp non-selectable homologous arms or 60-100 bp selectable homologous arms.

Multiplex CRISPR-Cas Genome Editing: Next-Generation Microbial Strain Engineering

Genome editing is a crucial technology for obtaining desired phenotypes in a variety of species, ranging from microbes to plants, animals, and humans. With the advent of CRISPR-Cas technology, it has become possible to edit the intended sequence by modifying the target recognition sequence in guide RNA (gRNA). By expressing multiple gRNAs simultaneously, it is possible to edit multiple targets at the same time, allowing for the simultaneous introduction of various functions into the cell. This can significantly reduce the time and cost of obtaining engineered microbial strains for specific traits. In this review, we investigate the resolution of multiplex genome editing and its application in engineering microorganisms, including bacteria and yeast. Furthermore, we examine how recent advancements in artificial intelligence technology could assist in microbial genome editing and engineering. Based on these insights, we present our perspectives on the future evolution and potential impact of multiplex genome editing technologies in the agriculture and food industry.

A comprehensive report on valorization of waste to single cell protein: strategies, challenges, and future prospects

The food insecurity due to a vertical increase in the global population urgently demands substantial advancements in the agricultural sector and to identify sustainable affordable sources of nutrition, particularly proteins. Single-cell protein (SCP) has been revealed as the dried biomass of microorganisms such as algae, yeast, and bacteria cultivated in a controlled environment. Production of SCP is a promising alternative to conventional protein sources like soy and meat, due to quicker production, minimal land requirement, and flexibility to various climatic conditions. In addition to protein production, it also contributes to waste management by converting it into food and feed for both human and animal consumption. This article provides an overview of SCP production, including its benefits, safety, acceptability, and cost, as well as limitations that constrains its maximum use. Furthermore, this review criticizes the downstream processing of SCP, encompassing cell wall disruption, removal of nucleic acid, harvesting of biomass, drying, packaging, storage, and transportation. The potential applications of SCP, such as in food and feed as well as in the production of bioplastics, emulsifiers, and as flavoring agents for baked food, soup, and salad, are also discussed.

Enhanced fatty acid storage combined with the multi-factor optimization of fermentation for high-level production of docosahexaenoic acid in Schizochytrium sp

Schizochytrium sp. hasreceived much attention for itsability to synthesize and accumulate high-level docosahexaenoic acid (DHA), which can reach nearly 40 % of total fatty acids. In this study, the titer of DHA in Schizochytrium sp. was successfully improved by enhancing DHA storage through overexpressing the diacylglycerol acyltransferase (ScDGAT2C) gene, as well as optimizing the supply of precursors and cofactors required for DHA synthesis by response surface methodology. Notably, malic acid, citric acid, and biotin showed synergistic and time-dependent effects on DHA accumulation. The maximum lipid and DHA titers of the engineered Schizochytrium sp. strain reached 84.28 ± 1.02 g/L and 42.23 ± 0.69 g/L, respectively, with the optimal concentration combination (1.62 g/L malic acid + 0.37 g/L citric acid + 8.28 mg/L biotin) were added 48 h after inoculation. This study provides an effective strategy for improving lipid and DHA production in Schizochytrium sp.

Reconstructing curcumin biosynthesis in yeast reveals the implication of caffeoyl-shikimate esterase in phenylpropanoid metabolic flux

Curcumin is a polyphenolic natural product from the roots of turmeric (Curcuma longa). It has been a popular coloring and flavoring agent in food industries with known health benefits. The conventional phenylpropanoid pathway is known to proceed from phenylalanine via p-coumaroyl-CoA intermediate. Although hydroxycinnamoyl-CoA: shikimate hydroxycinnamoyl transferase (HCT) plays a key catalysis in the biosynthesis of phenylpropanoid products at the downstream of p-coumaric acid, a recent discovery of caffeoyl-shikimate esterase (CSE) showed that an alternative pathway exists. Here, the biosynthetic efficiency of the conventional and the alternative pathway in producing feruloyl-CoA was examined using curcumin production in yeast. A novel modular multiplex genome-edit (MMG)-CRISPR platform was developed to facilitate rapid integrations of up to eight genes into the yeast genome in two steps. Using this MMG-CRISPR platform and metabolic engineering strategies, the alternative CSE phenylpropanoid pathway consistently showed higher titers (2-19 folds) of curcumin production than the conventional pathway in engineered yeast strains. In shake flask cultures using a synthetic minimal medium without phenylalanine, the curcumin production titer reached up to 1.5 mg/L, which is three orders of magnitude (∼4800-fold) improvement over non-engineered base strain. This is the first demonstration of de novo curcumin biosynthesis in yeast. Our work shows the critical role of CSE in improving the metabolic flux in yeast towards the phenylpropanoid biosynthetic pathway. In addition, we showcased the convenience and reliability of modular multiplex CRISPR/Cas9 genome editing in constructing complex synthetic pathways in yeast.

CRISPR-Cas9-Mediated Genome Editing in Paenibacillus polymyxa

In recent years, the clustered regularly interspaced palindromic repeats-Cas (CRISPR-Cas) technology has become the method of choice for precision genome editing in many organisms due to its simplicity and efficacy. Multiplex genome editing, point mutations, and large genomic modifications are attractive features of the CRISPR-Cas9 system. These applications facilitate both the ease and velocity of genetic manipulations and the discovery of novel functions. In this protocol chapter, we describe the use of a CRISPR-Cas9 system for multiplex integration and deletion modifications, and deletions of large genomic regions by the use of a single guide RNA (sgRNA), and, finally, targeted point mutation modifications in Paenibacillus polymyxa.

A randomized multiplex CRISPRi-Seq approach for the identification of critical combinations of genes

Identifying virulence-critical genes from pathogens is often limited by functional redundancy. To rapidly interrogate the contributions of combinations of genes to a biological outcome, we have developed a multiplex, randomized CRISPR interference sequencing (MuRCiS) approach. At its center is a new method for the randomized self-assembly of CRISPR arrays from synthetic oligonucleotide pairs. When paired with PacBio long-read sequencing, MuRCiS allowed for near-comprehensive interrogation of all pairwise combinations of a group of 44 Legionella pneumophila virulence genes encoding highly conserved transmembrane proteins for their role in pathogenesis. Both amoeba and human macrophages were challenged with L. pneumophila bearing the pooled CRISPR array libraries, leading to the identification of several new virulence-critical combinations of genes. lpg2888 and lpg3000 were particularly fascinating for their apparent redundant functions during L. pneumophila human macrophage infection, while lpg3000 alone was essential for L. pneumophila virulence in the amoeban host Acanthamoeba castellanii. Thus, MuRCiS provides a method for rapid genetic examination of even large groups of redundant genes, setting the stage for application of this technology to a variety of biological contexts and organisms.

A randomized multiplex CRISPRi-Seq approach for the identification of critical combinations of genes

Identifying virulence-critical genes from pathogens is often limited by functional redundancy. To rapidly interrogate the contributions of combinations of genes to a biological outcome, we have developed a multiplex, randomized CRISPR interference sequencing (MuRCiS) approach. At its center is a new method for the randomized self-assembly of CRISPR arrays from synthetic oligonucleotide pairs. When paired with PacBio long-read sequencing, MuRCiS allowed for near-comprehensive interrogation of all pairwise combinations of a group of 44 Legionella pneumophila virulence genes encoding highly conserved transmembrane proteins for their role in pathogenesis. Both amoeba and human macrophages were challenged with L. pneumophila bearing the pooled CRISPR array libraries, leading to the identification of several new virulence-critical combinations of genes. lpg2888 and lpg3000 were particularly fascinating for their apparent redundant functions during L. pneumophila human macrophage infection, while lpg3000 alone was essential for L. pneumophila virulence in the amoeban host Acanthamoeba castellanii. Thus, MuRCiS provides a method for rapid genetic examination of even large groups of redundant genes, setting the stage for application of this technology to a variety of biological contexts and organisms.

An outlook to sophisticated technologies and novel developments for metabolic regulation in the Saccharomyces cerevisiae expression system

Saccharomyces cerevisiae is one of the most extensively used biosynthetic systems for the production of diverse bioproducts, especially biotherapeutics and recombinant proteins. Because the expression and insertion of foreign genes are always impaired by the endogenous factors of Saccharomyces cerevisiae and nonproductive procedures, various technologies have been developed to enhance the strength and efficiency of transcription and facilitate gene editing procedures. Thus, the limitations that block heterologous protein secretion have been overcome. Highly efficient promoters responsible for the initiation of transcription and the accurate regulation of expression have been developed that can be precisely regulated with synthetic promoters and double promoter expression systems. Appropriate codon optimization and harmonization for adaption to the genomic codon abundance of S. cerevisiae are expected to further improve the transcription and translation efficiency. Efficient and accurate translocation can be achieved by fusing a specifically designed signal peptide to an upstream foreign gene to facilitate the secretion of newly synthesized proteins. In addition to the widely applied promoter engineering technology and the clear mechanism of the endoplasmic reticulum secretory pathway, the innovative genome editing technique CRISPR/Cas (clustered regularly interspaced short palindromic repeats/CRISPR-associated system) and its derivative tools allow for more precise and efficient gene disruption, site-directed mutation, and foreign gene insertion. This review focuses on sophisticated engineering techniques and emerging genetic technologies developed for the accurate metabolic regulation of the S. cerevisiae expression system.

Non-homologous end-joining-deficient filamentous fungal strains mitigate the impact of off-target mutations during the application of CRISPR/Cas9

CRISPR/Cas9 genome editing technology has been implemented in almost all living organisms. Its editing precision appears to be very high and therefore could represent a big change from conventional genetic engineering approaches. However, guide RNA binding to nucleotides similar to the target site could result in undesired off-target mutations. Despite this, evaluating whether mutations occur is rarely performed in genome editing studies. In this study, we generated CRISPR/Cas9-derived filamentous fungal strains and analyzed them for the occurrence of mutations, and to which extent genome stability affects their occurrence. As a test case, we deleted the (hemi-)cellulolytic regulator-encoding gene xlnR in two Aspergillus niger strains: a wild type (WT) and a non-homologous end-joining (NHEJ)-deficient strain ΔkusA. Initial phenotypic analysis suggested a much higher prevalence of mutations in the WT compared to NHEJ-deficient strains, which was confirmed and quantified by whole-genome sequencing analysis. Our results clearly demonstrate that CRISPR/Cas9 applied to an NHEJ-deficient strain is an efficient strategy to avoid unwanted mutations. IMPORTANCE Filamentous fungi are commonly used biofactories for the production of industrially relevant proteins and metabolites. Often, fungal biofactories undergo genetic development (genetic engineering, genome editing, etc.) aimed at improving production yields. In this context, CRISPR/Cas9 has gained much attention as a genome editing strategy due to its simplicity, versatility, and precision. However, despite the high level of accuracy reported for CRISPR/Cas9, in some cases unintentional cleavages in non-targeted loci-known as off-target mutations-could arise. While biosafety should be a central feature of emerging biotechnologies to minimize unintended consequences, few studies quantitatively evaluate the risk of off-target mutations. This study demonstrates that the use of non-homologous end-joining-deficient fungal strains drastically reduces the number of unintended genomic mutations, ensuring that CRISPR/Cas9 can be safely applied for strain development.

Construction of a bacteriophage-derived recombinase system in Bacillus licheniformis for gene deletion

Bacillus licheniformis and its related strains have found extensive applications in diverse industries, agriculture, and medicine. However, the current breeding methods for this strain primarily rely on natural screening and traditional mutagenesis. The limited availability of efficient genetic engineering tools, particularly recombination techniques, has hindered further advancements in its applications. In this study, we conducted a comprehensive investigation to identify and characterize a recombinase, RecT, derived from a Bacillus phage. Remarkably, the recombinase exhibited a 105-fold enhancement in the recombination efficiency of the strain. To facilitate genome editing, we developed a system based on the conditional expression of RecT using a rhamnose-inducible promoter (Prha). The efficacy of this system was evaluated by deleting the amyL gene, which encodes an α-amylase. Our findings revealed that the induction time and concentration of rhamnose, along with the generation time of the strain, significantly influenced the editing efficiency. Optimal conditions for genome editing were determined as follows: the wild-type strain was initially transformed with the genome editing plasmid, followed by cultivation and induction with 1.5% rhamnose for 8 h. Subsequently, the strain was further cultured for an additional 24 h, equivalent to approximately three generations. Consequently, the recombination efficiency reached an impressive 16.67%. This study represents a significant advancement in enhancing the recombination efficiency of B. licheniformis through the utilization of a RecT-based recombination system. Moreover, it provides a highly effective genome editing tool for genetic engineering applications in this strain.

Strategies for the Development of Industrial Fungal Producing Strains

The use of microorganisms in industry has enabled the (over)production of various compounds (e.g., primary and secondary metabolites, proteins and enzymes) that are relevant for the production of antibiotics, food, beverages, cosmetics, chemicals and biofuels, among others. Industrial strains are commonly obtained by conventional (non-GMO) strain improvement strategies and random screening and selection. However, recombinant DNA technology has made it possible to improve microbial strains by adding, deleting or modifying specific genes. Techniques such as genetic engineering and genome editing are contributing to the development of industrial production strains. Nevertheless, there is still significant room for further strain improvement. In this review, we will focus on classical and recent methods, tools and technologies used for the development of fungal production strains with the potential to be applied at an industrial scale. Additionally, the use of functional genomics, transcriptomics, proteomics and metabolomics together with the implementation of genetic manipulation techniques and expression tools will be discussed.

Multiplex Single-Nucleotide Microbial Genome Editing Achieved by CRISPR-Cas9 Using 5'-End-Truncated sgRNAs

Multiplex genome editing with CRISPR-Cas9 offers a cost-effective solution for time and labor savings. However, achieving high accuracy remains a challenge. In an Escherichia coli model system, we achieved highly efficient single-nucleotide level simultaneous editing of the galK and xylB genes using the 5'-end-truncated single-molecular guide RNA (sgRNA) method. Furthermore, we successfully demonstrated the simultaneous editing of three genes (galK, xylB, and srlD) at single-nucleotide resolution. To showcase practical application, we targeted the cI857 and ilvG genes in the genome of E. coli. While untruncated sgRNAs failed to produce any edited cells, the use of truncated sgRNAs allowed us to achieve simultaneous and accurate editing of these two genes with an efficiency of 30%. This enabled the edited cells to retain their lysogenic state at 42 °C and effectively alleviated l-valine toxicity. These results suggest that our truncated sgRNA method holds significant potential for widespread and practical use in synthetic biology.

Base editing enables duplex point mutagenesis in Clostridium autoethanogenum at the price of numerous off-target mutations

Base editors are recent multiplex gene editing tools derived from the Cas9 nuclease of Streptomyces pyogenes. They can target and modify a single nucleotide in the genome without inducing double-strand breaks (DSB) of the DNA helix. As such, they hold great potential for the engineering of microbes that lack effective DSB repair pathways such as homologous recombination (HR) or non-homologous end-joining (NHEJ). However, few applications of base editors have been reported in prokaryotes to date, and their advantages and drawbacks have not been systematically reported. Here, we used the base editors Target-AID and Target-AID-NG to introduce nonsense mutations into four different coding sequences of the industrially relevant Gram-positive bacterium Clostridium autoethanogenum. While up to two loci could be edited simultaneously using a variety of multiplexing strategies, most colonies exhibited mixed genotypes and most available protospacers led to undesired mutations within the targeted editing window. Additionally, fifteen off-target mutations were detected by sequencing the genome of the resulting strain, among them seven single-nucleotide polymorphisms (SNP) in or near loci bearing some similarity with the targeted protospacers, one 15 nt duplication, and one 12 kb deletion which removed uracil DNA glycosylase (UDG), a key DNA repair enzyme thought to be an obstacle to base editing mutagenesis. A strategy to process prokaryotic single-guide RNA arrays by exploiting tRNA maturation mechanisms is also illustrated.

Recombineering: Genetic Engineering in Escherichia coli Using Homologous Recombination

The bacterial chromosome and bacterial plasmids can be engineered in vivo by homologous recombination using either PCR products or synthetic double-stranded DNA (dsDNA) or single-stranded DNA as substrates. Multiple linear dsDNA molecules can be assembled into an intact plasmid. The technology of recombineering is possible because bacteriophage-encoded recombination proteins efficiently recombine sequences with homologies as short as 35 to 50 bases. Recombineering allows DNA sequences to be inserted or deleted without regard to the location of restriction sites and can also be used in combination with CRISPR/Cas targeting systems. © 2023 Wiley Periodicals LLC. This article has been contributed to by U.S. Government employees and their work is in the public domain in the USA. Basic Protocol: Making electrocompetent cells and transforming with linear DNA Support Protocol 1: Selection/counter-selections for genome engineering Support Protocol 2: Creating and screening for oligo recombinants by PCR Support Protocol 3: Other methods of screening for unselected recombinants Support Protocol 4: Curing recombineering plasmids containing a temperature-sensitive replication function Support Protocol 5: Removal of the prophage by recombineering Alternate Protocol 1: Using CRISPR/Cas9 as a counter-selection following recombineering Alternate Protocol 2: Assembly of linear dsDNA fragments into functional plasmids Alternate Protocol 3: Retrieval of alleles onto a plasmid by gap repair Alternate Protocol 4: Modifying multicopy plasmids with recombineering Support Protocol 6: Screening for unselected plasmid recombinants Alternate Protocol 5: Recombineering with an intact λ prophage Alternate Protocol 6: Targeting an infecting λ phage with the defective prophage strains.

Locus-Specific Detection of DNA Methylation: The Advance, Challenge, and Perspective of CRISPR-Cas Assisted Biosensors

Deoxyribonucleic acid (DNA) methylation is one of the epigenetic characteristics that result in heritable and revisable phenotype changes but without sequence changes in DNA. Aberrant methylation occurring at a specific locus was reported to be associated with cancers, insulin resistance, obesity, Alzheimer's disease, Parkinson's disease, etc. Therefore, locus-specific DNA methylation can serve as a valuable biomarker for disease diagnosis and therapy. Recently, Clustered regularly interspaced short palindromic repeats (CRISPR)-Cas systems are applied to develop biosensors for DNA, ribonucleic acid, proteins, and small molecules detection. Because of their highly specific binding ability and signal amplification capacity, CRISPR-Cas assisted biosensor also serve as a potential tool for locus-specific detection of DNA methylation. In this perspective, based on the detection principle, a detailed classification and comprehensive discussion of recent works about the latest advances in locus-specific detection of DNA methylation using CRISPR-Cas systems are provided. Furthermore, current challenges and future perspectives of CRISPR-based locus-specific detection of DNA methylation are outlined.

Construction and evaluation of gRNA arrays for multiplex CRISPR-Cas9

Endonuclease system CRISPR-Cas9 represents a powerful toolbox for the budding yeast's Saccharomyces cerevisiae genome perturbation. The resulting double-strand breaks are preferentially repaired via highly efficient homologous recombination, which subsequently leads to marker-free genome editing. The goal of this study was to evaluate precise targeting of multiple loci simultaneously. To construct an array of independently expressing guide RNAs (gRNAs), the genes encoding them were assembled through a BioBrick construction procedure. We designed a multiplex CRISPR-Cas9 system for targeting 6 marker genes, whereby the gRNA array was expressed from a single plasmid. To evaluate the performance of the gRNA array, the activity of the designed system was assessed by the success rate of the introduction of perturbations within the target loci: successful gRNA expression, followed by target DNA double-strand breaks formation and their repair by homologous recombination led to premature termination of the coding sequence of the marker genes, resulting in the prevention of growth of the transformants on the corresponding selection media. In conclusion, we successfully introduced up to five simultaneous perturbations within single cells of yeast S. cerevisiae using the multiplex CRISPR-Cas9 system. While this has been done before, we here present an alternative sequential BioBrick assembly with the capability to accommodate many highly similar gRNA-expression cassettes, and an exhaustive evaluation of their performance.

Zebrafish Models for Skeletal Muscle Senescence: Lessons from Cell Cultures and Rodent Models

Human life expectancy has markedly increased over the past hundred years. Consequently, the percentage of elderly people is increasing. Aging and sarcopenic changes in skeletal muscles not only reduce locomotor activities in elderly people but also increase the chance of trauma, such as bone fractures, and the incidence of other diseases, such as metabolic syndrome, due to reduced physical activity. Exercise therapy is currently the only treatment and prevention approach for skeletal muscle aging. In this review, we aimed to summarize the strategies for modeling skeletal muscle senescence in cell cultures and rodents and provide future perspectives based on zebrafish models. In cell cultures, in addition to myoblast proliferation and myotube differentiation, senescence induction into differentiated myotubes is also promising. In rodents, several models have been reported that reflect the skeletal muscle aging phenotype or parts of it, including the accelerated aging models. Although there are fewer models of skeletal muscle aging in zebrafish than in mice, various models have been reported in recent years with the development of CRISPR/Cas9 technology, and further advancements in the field using zebrafish models are expected in the future.

Inducible CRISPR/Cas9 Allows for Multiplexed and Rapidly Segregated Single-Target Genome Editing in Synechocystis Sp. PCC 6803

Establishing various synthetic biology tools is crucial for the development of cyanobacteria for biotechnology use, especially tools that allow for precise and markerless genome editing in a time-efficient manner. Here, we describe a riboswitch-inducible CRISPR/Cas9 system, contained on a single replicative vector, for the model cyanobacterium Synechocystis sp. PCC 6803. A theophylline-responsive riboswitch allowed tight control of Cas9 expression, which enabled reliable transformation of the CRISPR/Cas9 vector intoSynechocystis. Induction of the CRISPR/Cas9 mediated various types of genomic edits, specifically deletions and insertions of varying size. The editing efficiency varied depending on the target and intended edit; smaller edits performed better, reaching, e.g., 100% for insertion of a FLAG-tag onto rbcL. Importantly, the single-vector CRISPR/Cas9 system mediated multiplexed editing of up to three targets in parallel inSynechocystis. All single-target and several double-target mutants were also fully segregated after the first round of induction. Lastly, a vector curing system based on the nickel-inducible expression of the toxic mazF (from Escherichia coli) was added to the CRISPR/Cas9 vector. This inducible system allowed for curing of the vector in 25-75% of screened colonies, enabling edited mutants to become markerless.

Potential use of microbial engineering in single-cell protein production

Single-cell proteins (SCPs) have been widely used in human food and animal feed applications, still, there are challenges in their production and commercialization. Recently, advances in microbial synthetic biology, genomic engineering, and biofoundry technologies have offered capabilities to effectively and rapidly engineer microorganisms for improving the productivity, nutritional, and functional quality of SCPs. In this review, we discuss various synthetic biology, genomic engineering, and biofoundry tools that can be harnessed for SCP production and genetic modification. We also describe the current and potential applications of genetic modification in producing intermediate feedstocks, as well as biomass-based and multifunctional SCPs. Finally, we discuss the technological and policy-control related challenges encountered when deploying genetic modification in SCP production for animal feed and human food applications.

CRISPR-Cas9-mediated Large Cluster Deletion and Multiplex Genome Editing in Paenibacillus polymyxa

The use of molecular tools based on the clustered regularly interspaced short palindromic repeats-Cas (CRISPR-Cas) systems has rapidly advanced genetic engineering. These molecular biological tools have been applied for different genetic engineering purposes in multiple organisms, including the quite rarely explored Paenibacillus polymyxa. However, only limited studies on large cluster deletion and multiplex genome editing have been described for this highly interesting and versatile bacterium. Here, we demonstrate the utilization of a Cas9-based system to realize targeted deletions of four biosynthetic gene clusters in the range of 12-41 kb by the use of a single targeting sgRNA. Furthermore, we also harnessed the system for multiplex editing of genes and large genomic regions. Multiplex deletion was achieved with more than 80% efficiency, while simultaneous integration at two distantly located sites was obtained with 58% efficiency. The findings reported in this study are anticipated to accelerate future research in P. polymyxa and related species.

Production of polyunsaturated fatty acids by Schizochytrium (Aurantiochytrium) spp

Diverse health benefits are associated with dietary consumption of omega-3 long-chain polyunsaturated fatty acids (ω-3 LC-PUFA), particularly docosahexaenoic acid (DHA) and eicosapentaenoic acid (EPA). Traditionally, these fatty acids have been obtained from fish oil, but limited supply, variably quality, and an inability to sustainably increase production for a rapidly growing market, are driving the quest for alternative sources. DHA derived from certain marine protists (heterotrophic thraustochytrids) already has an established history of commercial production for high-value dietary use, but is too expensive for use in aquaculture feeds, a much larger potential market for ω-3 LC-PUFA. Sustainable expansion of aquaculture is prevented by its current dependence on wild-caught fish oil as the source of ω-3 LC-PUFA nutrients required in the diet of aquacultured animals. Although several thraustochytrids have been shown to produce DHA and EPA, there is a particular interest in Schizochytrium spp. (now Aurantiochytrium spp.), as some of the better producers. The need for larger scale production has resulted in development of many strategies for improving productivity and production economics of ω-3 PUFA in Schizochytrium spp. Developments in fermentation technology and metabolic engineering for enhancing LC-PUFA production in Schizochytrium spp. are reviewed.

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.

Comprehensive Genome Engineering Toolbox for Microalgae Nannochloropsis oceanica Based on CRISPR-Cas Systems

Microalgae can produce industrially relevant metabolites using atmospheric CO₂ and sunlight as carbon and energy sources, respectively. Developing molecular tools for high-throughput genome engineering could accelerate the generation of tailored strains with improved traits. To this end, we developed a genome editing strategy based on Cas12a ribonucleoproteins (RNPs) and homology-directed repair (HDR) to generate scarless and markerless mutants of the microalga Nannochloropsis oceanica. We also developed an episomal plasmid-based Cas12a system for efficiently introducing indels at the target site. Additionally, we exploited the ability of Cas12a to process an associated CRISPR array to perform multiplexed genome engineering. We efficiently targeted three sites in the host genome in a single transformation, thereby making a major step toward high-throughput genome engineering in microalgae. Furthermore, a CRISPR interference (CRISPRi) tool based on Cas9 and Cas12a was developed for effective downregulation of target genes. We observed up to 85% reduction in the transcript levels upon performing CRISPRi with dCas9 in N. oceanica. Overall, these developments substantially accelerate genome engineering efforts in N. oceanica and potentially provide a general toolbox for improving other microalgal strains.

A highly effective and self-transmissible CRISPR antimicrobial for elimination of target plasmids without antibiotic selection

The use of CRISPR/Cas (Clustered Regularly Interspaced Short Palindromic Repeats/CRISPR associated protein) for sequence-specific elimination of bacteria or resistance genes is a powerful tool for combating antibiotic resistance. However, this approach requires efficient delivery of CRISPR/Cas DNA cassette(s) into the targeted bacterial population. Compared to phage transduction, plasmid conjugation can deliver DNA to a broader host range but often suffers from low delivery efficiency. Here, we developed multi-plasmid conjugation systems for efficient CRISPR/Cas delivery, target DNA elimination and plasmid replacement. The CRISPR/Cas system, delivered via a broad-host-range R1162 mobilizable plasmid, specifically eliminated the targeted plasmid in recipient cells. A self-transmissible RK2 helper plasmid facilitated the spread of mobilizable CRISPR/Cas. The replacement of the target plasmid with another plasmid from the same compatibility group helped speed up target plasmid elimination especially when the target plasmid was also mobilizable. Together, we showed that up to 100% of target plasmid from the entire recipient population could be replaced even at a low (1:180) donor-to-recipient ratio and in the absence of transconjugant selection. Such an ability to modify genetic content of microbiota efficiently in the absence of selection will be critical for future development of CRISPR antimicrobials as well as genetic tools for in situ microbiome engineering.

Multigene editing: current approaches and beyond

CRISPR/Cas9 multigene editing is an active and widely studied topic in the fields of biomedicine and biology. It involves a simultaneous participation of multiple single-guide RNAs (sgRNAs) to edit multiple target genes in a way that each gene is edited by one of these sgRNAs. There are possibly numerous sgRNA candidates capable of on-target editing on each of these genes with various efficiencies. Meanwhile, each of these sgRNA candidates may cause unwanted off-target editing at many other genes. Therefore, selection optimization of these multiple sgRNAs is demanded so as to minimize the number of sgRNAs and thus reduce the collective negative effects caused by the off-target editing. This survey reviews wet-laboratory approaches to the implementation of multigene editing and their needs of computational tools for better design. We found that though off-target editing is unavoidable during the gene editing, those disfavored cuttings by some target genes' sgRNAs can potentially become on-target editing sites for some other genes of interests. This off-to-on role conversion is beneficial to optimize the sgRNA selection in multigene editing. We present a preference cutting score to assess those beneficial off-target cutting sites, which have a few mismatches with their host genes' on-target editing sites. These potential sgRNAs can be prioritized for recommendation via ranking their on-target average cutting efficiency, the total off-target site number and their average preference cutting score. We also present case studies on cancer-associated genes to demonstrate tremendous usefulness of the new method.

Tailoring in fungi for next generation cellulase production with special reference to CRISPR/CAS system

Cellulose is the utmost plenteous source of biopolymer in our earth, and fungi are the most efficient and ubiquitous organism in degrading the cellulosic biomass by synthesizing cellulases. Tailoring through genetic manipulation has played a substantial role in constructing novel fungal strains towards improved cellulase production of desired traits. However, the traditional methods of genetic manipulation of fungi are time-consuming and tedious. With the availability of the full-genome sequences of several industrially relevant filamentous fungi, CRISPR-CAS (clustered regularly interspaced short palindromic repeats/CRISPR-associated protein) technology has come into the focus for the proficient development of manipulated strains of filamentous fungi. This review summarizes the mode of action of cellulases, transcription level regulation for cellulase expression, various traditional strategies of genetic manipulation with CRISPR-CAS technology to develop modified fungal strains for a preferred level of cellulase production, and the futuristic trend in this arena of research.

The Role of Metabolic Engineering Technologies for the Production of Fatty Acids in Yeast

Simple Summary

Metabolic engineering involves the sustainable production of high-value products. E. coli and yeast, in particular, are used for such processes. Using metabolic engineering, the biosynthetic pathways of these cells are altered to obtain a high production of desired products. Fatty acids (FAs) and their derivatives are products produced using metabolic engineering. However, classical methods used for engineering yeast metabolic pathways for the production of fatty acids and their derivatives face problems such as the low supply of key precursors and product tolerance. This review introduces the different ways FAs are being produced in E. coli and yeast and the genetic manipulations for enhanced production of FAs. The review also summarizes the latest techniques (i.e., CRISPR–Cas and synthetic biology) for developing FA-producing yeast cell factories.

Abstract

Metabolic engineering is a cutting-edge field that aims to produce simple, readily available, and inexpensive biomolecules by applying different genetic engineering and molecular biology techniques. Fatty acids (FAs) play an important role in determining the physicochemical properties of membrane lipids and are precursors of biofuels. Microbial production of FAs and FA-derived biofuels has several advantages in terms of sustainability and cost. Conventional yeast Saccharomyces cerevisiae is one of the models used for FA synthesis. Several genetic manipulations have been performed to enhance the citrate accumulation and its conversation into acetyl-CoA, a precursor for FA synthesis. Success has been achieved in producing different chemicals, including FAs and their derivatives, through metabolic engineering. However, several hurdles such as slow growth rate, low oleaginicity, and cytotoxicity are still need to be resolved. More robust research needs to be conducted on developing microbes capable of resisting diverse environments, chemicals, and cost-effective feed requirements. Redesigning microbes to produce FAs with cutting-edge synthetic biology and CRISPR techniques can solve these problems. Here, we reviewed the technological progression of metabolic engineering techniques and genetic studies conducted on S. cerevisiae, making it suitable as a model organism and a great candidate for the production of biomolecules, especially FAs.

Cytosine Base Editor-Mediated Multiplex Genome Editing to Accelerate Discovery of Novel Antibiotics in Bacillus subtilis and Paenibacillus polymyxa

Genome-based identification of new antibiotics is emerging as an alternative to traditional methods. However, uncovering hidden antibiotics under the background of known antibiotics remains a challenge. To over this problem using a quick and effective genetic approach, we developed a multiplex genome editing system using a cytosine base editor (CBE). The CBE system achieved simultaneous double, triple, quadruple, and quintuple gene editing with efficiencies of 100, 100, 83, and 75%, respectively, as well as the 100% editing efficiency of single targets in Bacillus subtilis. Whole-genome sequencing of the edited strains showed that they had an average of 8.5 off-target single-nucleotide variants at gRNA-independent positions. The CBE system was used to simultaneously knockout five known antibiotic biosynthetic gene clusters to leave only an uncharacterized polyketide biosynthetic gene cluster in Paenibacillus polymyxa E681. The polyketide showed antimicrobial activities against gram-positive bacteria, but not gram-negative bacteria and fungi. Therefore, our findings suggested that the CBE system might serve as a powerful tool for multiplex genome editing and greatly accelerating the unraveling of hidden antibiotics in Bacillus and Paenibacillus species.

gEL DNA: A Cloning- and Polymerase Chain Reaction-Free Method for CRISPR-Based Multiplexed Genome Editing

Even for the genetically accessible yeast Saccharomyces cerevisiae, the CRISPR-Cas DNA editing technology has strongly accelerated and facilitated strain construction. Several methods have been validated for fast and highly efficient single editing events, and diverse approaches for multiplex genome editing have been described in the literature by means of SpCas9 or FnCas12a endonucleases and their associated guide RNAs (gRNAs). The gRNAs used to guide the Cas endonuclease to the editing site are typically expressed from plasmids using native Pol II or Pol III RNA polymerases. These gRNA expression plasmids require laborious, time-consuming cloning steps, which hampers their implementation for academic and applied purposes. In this study, we explore the potential of expressing gRNA from linear DNA fragments using the T7 RNA polymerase (T7RNAP) for single and multiplex genome editing in Saccharomyces cerevisiae. Using FnCas12a, this work demonstrates that transforming short, linear DNA fragments encoding gRNAs in yeast strains expressing T7RNAP promotes highly efficient single and duplex DNA editing. These DNA fragments can be custom ordered, which makes this approach highly suitable for high-throughput strain construction. This work expands the CRISPR toolbox for large-scale strain construction programs in S. cerevisiae and promises to be relevant for other less genetically accessible yeast species.

Recent advances of Cas12a applications in bacteria

Clustered regularly interspaced short palindromic repeats (CRISPR)-mediated genome engineering and related technologies have revolutionized biotechnology over the last decade by enhancing the efficiency of sophisticated biological systems. Cas12a (Cpf1) is an RNA-guided endonuclease associated to the CRISPR adaptive immune system found in many prokaryotes. Contrary to its more prominent counterpart Cas9, Cas12a recognizes A/T rich DNA sequences and is able to process its corresponding guide RNA directly, rendering it a versatile tool for multiplex genome editing efforts and other applications in biotechnology. While Cas12a has been extensively used in eukaryotic cell systems, microbial applications are still limited. In this review, we highlight the mechanistic and functional differences between Cas12a and Cas9 and focus on recent advances of applications using Cas12a in bacterial hosts. Furthermore, we discuss advantages as well as current challenges and give a future outlook for this promising alternative CRISPR-Cas system for bacterial genome editing and beyond. KEY POINTS: • Cas12a is a powerful tool for genome engineering and transcriptional perturbation • Cas12a causes less toxic side effects in bacteria than Cas9 • Self-processing of crRNA arrays facilitates multiplexing approaches.

Recent Advances in Genome Editing Tools in Medical Mycology Research

Manipulating fungal genomes is an important tool to understand the function of target genes, pathobiology of fungal infections, virulence potential, and pathogenicity of medically important fungi, and to develop novel diagnostics and therapeutic targets. Here, we provide an overview of recent advances in genetic manipulation techniques used in the field of medical mycology. Fungi use several strategies to cope with stress and adapt themselves against environmental effectors. For instance, mutations in the 14 alpha-demethylase gene may result in azole resistance in Aspergillusfumigatus strains and shield them against fungicide's effects. Over the past few decades, several genome editing methods have been introduced for genetic manipulations in pathogenic fungi. Application of restriction enzymes to target and cut a double-stranded DNA in a pre-defined sequence was the first technique used for cloning in Aspergillus and Candida. Genome editing technologies, including zinc-finger nucleases (ZFNs) and transcriptional activator-like effector nucleases (TALENs), have been also used to engineer a double-stranded DNA molecule. As a result, TALENs were considered more practical to identify single nucleotide polymorphisms. Recently, Class 2 type II Clustered Regularly Interspaced Short Palindromic Repeat (CRISPR)/Cas9 technology has emerged as a more useful tool for genome manipulation in fungal research.

Novel Strategy to Combat Antibiotic Resistance: A Sight into the Combination of CRISPR/Cas9 and Nanoparticles

Antibiotic resistance is a significant crisis that threatens human health and safety worldwide. There is an urgent need for new strategies to control multidrug-resistant (MDR) bacterial infections. The latest breakthrough in gene-editing tools based on CRISPR/Cas9 has potential application in combating MDR bacterial infections because of their high targeting ability to specifically disrupt the drug resistance genes that microbes use for infection or to kill the pathogen directly. Despite the potential that CRISPR/Cas9 showed, its further utilization has been hampered by undesirable delivery efficiency in vivo. Nanotechnology offers an alternative way to overcome the shortcomings of traditional delivery methods of therapeutic agents. Advances in nanotechnology can improve the efficacy and safety of CRISPR/Cas9 components by using customized nanoparticle delivery systems. The combination of CRISPR/Cas9 and nanotechnology has the potential to open new avenues in the therapy of MDR bacterial infections. This review describes the recent advances related to CRISPR/Cas9 and nanoparticles for antimicrobial therapy and gene delivery, including the improvement in the packaging and localizing efficiency of the CRISPR/Cas9 components in the NP (nanoparticle)/CRISPR system. We pay particular attention to the strengths and limitations of the nanotechnology-based CRISPR/Cas9 delivery system to fight nosocomial pathogens.We highlight the need for more scientific research to explore the combinatorial efficacy of various nanoparticles and CRISPR technology to control and prevent antimicrobial resistance.

Genetic Modification of Sodalis Species by DNA Transduction

Bacteriophages (phages) are ubiquitous in nature. These viruses play a number of central roles in microbial ecology and evolution by, for instance, promoting horizontal gene transfer (HGT) among bacterial species. The ability of phages to mediate HGT through transduction has been widely exploited as an experimental tool for the genetic study of bacteria. As such, bacteriophage P1 represents a prototypical generalized transducing phage with a broad host range that has been extensively employed in the genetic manipulation of Escherichia coli and a number of other model bacterial species. Here we demonstrate that P1 is capable of infecting, lysogenizing, and promoting transduction in members of the bacterial genus Sodalis, including the maternally inherited insect endosymbiont Sodalis glossinidius. While establishing new tools for the genetic study of these bacterial species, our results suggest that P1 may be used to deliver DNA to many Gram-negative endosymbionts in their insect host, thereby circumventing a culturing requirement to genetically manipulate these organisms.

IMPORTANCE A large number of economically important insects maintain intimate associations with maternally inherited endosymbiotic bacteria. Due to the inherent nature of these associations, insect endosymbionts cannot be usually isolated in pure culture or genetically manipulated. Here we use a broad-host-range bacteriophage to deliver exogenous DNA to an insect endosymbiont and a closely related free-living species. Our results suggest that broad-host-range bacteriophages can be used to genetically alter insect endosymbionts in their insect host and, as a result, bypass a culturing requirement to genetically alter these bacteria.

A multiplex CRISPR interference tool for virulence gene interrogation in Legionella pneumophila

Catalytically inactive dCas9 imposes transcriptional gene repression by sterically precluding RNA polymerase activity at a given gene to which it was directed by CRISPR (cr)RNAs. This gene silencing technology, known as CRISPR interference (CRISPRi), has been employed in various bacterial species to interrogate genes, mostly individually or in pairs. Here, we developed a multiplex CRISPRi platform in the pathogen Legionella pneumophila capable of silencing up to ten genes simultaneously. Constraints on precursor-crRNA expression were overcome by combining a strong promoter with a boxA element upstream of a CRISPR array. Using crRNAs directed against virulence protein-encoding genes, we demonstrated that CRISPRi is fully functional not only during growth in axenic media, but also during macrophage infection, and that gene depletion by CRISPRi recapitulated the growth defect of deletion strains. By altering the position of crRNA-encoding spacers within the CRISPR array, our platform achieved the gradual depletion of targets that was mirrored by the severity in phenotypes. Multiplex CRISPRi thus holds great promise for probing large sets of genes in bulk in order to decipher virulence strategies of L. pneumophila and other bacterial pathogens.

Development of a Cas12a-Based Genome Editing Tool for Moderate Thermophiles

The ability of CRISPR-Cas12a nucleases to function reliably in a wide range of species has been key to their rapid adoption as genome engineering tools. However, so far, Cas12a nucleases have been limited for use in organisms with growth temperatures up to 37 °C. Here, we biochemically characterize three Cas12a orthologs for their temperature stability and activity. We demonstrate that Francisella novicida Cas12a (FnCas12a) has great biochemical potential for applications that require enhanced stability, including use at temperatures >37°C. Furthermore, by employing the moderate thermophilic bacterium Bacillus smithii as our experimental platform, we demonstrate that FnCas12a is active in vivo at temperatures up to 43°C. Subsequently, we develop a single-plasmid FnCas12a-based genome editing tool for B. smithii, combining the FnCas12a targeting system with plasmid-borne homologous recombination (HR) templates that carry the desired modifications. Culturing of B. smithii cells at 45°C allows for the uninhibited realization of the HR-based editing step, while a subsequent culturing step at reduced temperatures induces the efficient counterselection of the non-edited cells by FnCas12a. The developed gene-editing tool yields gene-knockout mutants within 3 days, and does not require tightly controllable expression of FnCas12a to achieve high editing efficiencies, indicating its potential for other (thermophilic) bacteria and archaea, including those with minimal genetic toolboxes. Altogether, our findings provide new biochemical insights into three widely used Cas12a nucleases, and establish the first Cas12a-based bacterial genome editing tools for moderate thermophilic microorganisms.

Multiplex Genome Editing in Yeast by CRISPR/Cas9 - A Potent and Agile Tool to Reconstruct Complex Metabolic Pathways

Numerous important pharmaceuticals and nutraceuticals originate from plant specialized metabolites, most of which are synthesized via complex biosynthetic pathways. The elucidation of these pathways is critical for the applicable uses of these compounds. Although the rapid progress of the omics technology has revolutionized the identification of candidate genes involved in these pathways, the functional characterization of these genes remains a major bottleneck. Baker's yeast (Saccharomyces cerevisiae) has been used as a microbial platform for characterizing newly discovered metabolic genes in plant specialized metabolism. Using yeast for the investigation of numerous plant enzymes is a streamlined process because of yeast's efficient transformation, limited endogenous specialized metabolism, partially sharing its primary metabolism with plants, and its capability of post-translational modification. Despite these advantages, reconstructing complex plant biosynthetic pathways in yeast can be time intensive. Since its discovery, CRISPR/Cas9 has greatly stimulated metabolic engineering in yeast. Yeast is a popular system for genome editing due to its efficient homology-directed repair mechanism, which allows precise integration of heterologous genes into its genome. One practical use of CRISPR/Cas9 in yeast is multiplex genome editing aimed at reconstructing complex metabolic pathways. This system has the capability of integrating multiple genes of interest in a single transformation, simplifying the reconstruction of complex pathways. As plant specialized metabolites usually have complex multigene biosynthetic pathways, the multiplex CRISPR/Cas9 system in yeast is suited well for functional genomics research in plant specialized metabolism. Here, we review the most advanced methods to achieve efficient multiplex CRISPR/Cas9 editing in yeast. We will also discuss how this powerful tool has been applied to benefit the study of plant specialized metabolism.

Multiplexed Genome Engineering with Cas12a

Genome engineering technologies based on CRISPR-Cas systems are fueling efforts to study genotype-phenotype relationships in a high-throughput and multiplexed fashion. While many genome engineering technologies exist and provide a means to efficiently manipulate one or a few genes in a singular context-knockout, inhibition, or activation in a constitutive, conditional, or inducible manner-progress towards engineering complex cellular programs has been hampered by the lack of technologies that can integrate these functions within a unified framework. To address this challenge, our lab created single transcript CRISPR-Cas12a (SiT-Cas12a), which enables conditional, inducible, orthogonal, and massively multiplexed genome engineering of dozens, to potentially hundreds, of genomic targets in eukaryotic cells simultaneously-providing a novel way to interrogate and engineer complex genetic programs. In this chapter, we outline the utility of SiT-Cas12a in human cells and describe experimental procedures for executing massively multiplexed genome engineering experiments-including strategies for designing and assembling customized multiplexed CRISPR guide RNA arrays as well as validating and analyzing CRISPR guide RNA array processing and genome engineering outcomes.

Multiplex Genome Engineering Methods for Yeast Cell Factory Development

As biotechnological applications of synthetic biology tools including multiplex genome engineering are expanding rapidly, the construction of strategically designed yeast cell factories becomes increasingly possible. This is largely due to recent advancements in genome editing methods like CRISPR/Cas tech and high-throughput omics tools. The model organism, baker's yeast (Saccharomyces cerevisiae) is an important synthetic biology chassis for high-value metabolite production. Multiplex genome engineering approaches can expedite the construction and fine tuning of effective heterologous pathways in yeast cell factories. Numerous multiplex genome editing techniques have emerged to capitalize on this recently. This review focuses on recent advancements in such tools, such as delta integration and rDNA cluster integration coupled with CRISPR-Cas tools to greatly enhance multi-integration efficiency. Examples of pre-placed gate systems which are an innovative alternative approach for multi-copy gene integration were also reviewed. In addition to multiple integration studies, multiplexing of alternative genome editing methods are also discussed. Finally, multiplex genome editing studies involving non-conventional yeasts and the importance of automation for efficient cell factory design and construction are considered. Coupling the CRISPR/Cas system with traditional yeast multiplex genome integration or donor DNA delivery methods expedites strain development through increased efficiency and accuracy. Novel approaches such as pre-placing synthetic sequences in the genome along with improved bioinformatics tools and automation technologies have the potential to further streamline the strain development process. In addition, the techniques discussed to engineer S. cerevisiae, can be adapted for use in other industrially important yeast species for cell factory development.

Molecular mechanisms of psychiatric diseases

For most psychiatric diseases, pathogenetic concepts as well as paradigms underlying neuropsychopharmacologic approaches currently revolve around neurotransmitters such as dopamine, serotonin, and norepinephrine. However, despite the fact that several generations of neurotransmitter-based psychotropics including atypical antipsychotics, selective serotonin reuptake inhibitors, and serotonin-norepinephrine reuptake inhibitors are available, the effectiveness of these medications is limited, and relapse rates in psychiatric diseases are relatively high, indicating potential involvement of other pathogenetic pathways. Indeed, recent high-throughput studies in genetics and molecular biology have shown that pathogenesis of major psychiatric illnesses involves hundreds of genes and numerous pathways via such fundamental processes as DNA methylation, transcription, and splicing. Current review summarizes these and other molecular mechanisms of such psychiatric illnesses as schizophrenia, major depressive disorder, and alcohol use disorder and suggests a conceptual framework for future studies.

Synthetic Biology on Acetogenic Bacteria for Highly Efficient Conversion of C1 Gases to Biochemicals

Synthesis gas, which is mainly produced from fossil fuels or biomass gasification, consists of C1 gases such as carbon monoxide, carbon dioxide, and methane as well as hydrogen. Acetogenic bacteria (acetogens) have emerged as an alternative solution to recycle C1 gases by converting them into value-added biochemicals using the Wood-Ljungdahl pathway. Despite the advantage of utilizing acetogens as biocatalysts, it is difficult to develop industrial-scale bioprocesses because of their slow growth rates and low productivities. To solve these problems, conventional approaches to metabolic engineering have been applied; however, there are several limitations owing to the lack of required genetic bioparts for regulating their metabolic pathways. Recently, synthetic biology based on genetic parts, modules, and circuit design has been actively exploited to overcome the limitations in acetogen engineering. This review covers synthetic biology applications to design and build industrial platform acetogens.

Multiplexed genomic encoding of non-canonical amino acids for labeling large complexes

Stunning advances in the structural biology of multicomponent biomolecular complexes (MBCs) have ushered in an era of intense, structure-guided mechanistic and functional studies of these complexes. Nonetheless, existing methods to site-specifically conjugate MBCs with biochemical and biophysical labels are notoriously impracticable and/or significantly perturb MBC assembly and function. To overcome these limitations, we have developed a general, multiplexed method in which we genomically encode non-canonical amino acids (ncAAs) into multiple, structure-informed, individual sites within a target MBC; select for ncAA-containing MBC variants that assemble and function like the wildtype MBC; and site-specifically conjugate biochemical or biophysical labels to these ncAAs. As a proof-of-principle, we have used this method to generate unique single-molecule fluorescence resonance energy transfer (smFRET) signals reporting on ribosome structural dynamics that have thus far remained inaccessible to smFRET studies of translation.

CRISPR with a Happy Ending: Non-Templated DNA Repair for Prokaryotic Genome Engineering

The exploration of microbial metabolism is expected to support the development of a sustainable economy and tackle several problems related to the burdens of human consumption. Microorganisms have the potential to catalyze processes that are currently unavailable, unsustainable and/or inefficient. Their metabolism can be optimized and further expanded using tools like the clustered regularly interspaced short palindromic repeats and their associated proteins (CRISPR-Cas) systems. These tools have revolutionized the field of biotechnology, as they greatly streamline the genetic engineering of organisms from all domains of life. CRISPR-Cas and other nucleases mediate double-strand DNA breaks, which must be repaired to prevent cell death. In prokaryotes, these breaks can be repaired through either homologous recombination, when a DNA repair template is available, or through template-independent end joining, of which two major pathways are known. These end joining pathways depend on different sets of proteins and mediate DNA repair with different outcomes. Understanding these DNA repair pathways can be advantageous to steer the results of genome engineering experiments. In this review, we discuss different strategies for the genetic engineering of prokaryotes through either non-homologous end joining (NHEJ) or alternative end joining (AEJ), both of which are independent of exogenous DNA repair templates.

CRISPR-Cas9/CRISPRi tools for cell factory construction in E. coli

The innovative CRISPR-Cas based genome editing technology provides some functionality and advantages such as the high efficiency and specificity as well as ease of handling. Both aspects of the CRISPR-Cas9 system including genetic engineering and gene regulation are advantageously applicable to the construction of microbial cell factories. As one of the most extensively used cell factories, E. coli has been engineered to produce various high value-added chemical compounds such as pharmaceuticals, biochemicals, and biofuels. Therefore, to improve the production of valuable metabolites, many investigations have been performed by focusing on CRISPR-Cas- based metabolic engineering of this host. In the current review, the biology underlying CRISPR-Cas9 system was briefly explained and then the applications of CRISPR-Cas9/CRISPRi tools were considered for cell factory construction in E. coli.

One-Day Construction of Multiplex Arrays to Harness Natural CRISPR-Cas Systems

CRISPR-Cas systems are prokaryotic immune systems that have proliferated widely not only in bacteria and archaea, but also much more recently, in human biological research and applications. Much work to date has utilized synthetic sgRNAs along with the CRISPR nuclease Cas9, but the discovery of array-processing nucleases now allows the use of more compact, natural CRISPR arrays in heterologous hosts, in addition to organisms with endogenous systems. Unfortunately, the construction of multiplex natural CRISPR arrays remains technically challenging, expensive, and/or time-consuming. This limitation hampers research involving natural CRISPR arrays in both native and heterologous hosts. To address this problem, we present a method to assemble CRISPR arrays that is simple, rapid, affordable, and highly scalable-we assembled 9-spacer arrays with 1 day's worth of work. We used this method to harness the endogenous CRISPR-Cas system of the highly competent bacterium Acinetobacter baylyi, showing that while single spacers are not always completely effective at blocking DNA acquisition through natural competence, multiplex natural CRISPR arrays enable both nearly complete DNA exclusion and genome editing, including with multiple targets for both. In addition to demonstrating a CRISPR array assembly method that will benefit a variety of applications, we also find a potential bet-hedging strategy for balancing CRISPR defense versus DNA acquisition in naturally competent A. baylyi.