High-Level dCas9 Expression Induces Abnormal Cell Morphology in Escherichia coli

Integrating bacterial molecular genetics with chemical biology for renewed antibacterial drug discovery

The application of dyes to understanding the aetiology of infection inspired antimicrobial chemotherapy and the first wave of antibacterial drugs. The second wave of antibacterial drug discovery was driven by rapid discovery of natural products, now making up 69% of current antibacterial drugs. But now with the most prevalent natural products already discovered, ∼107 new soil-dwelling bacterial species must be screened to discover one new class of natural product. Therefore, instead of a third wave of antibacterial drug discovery, there is now a discovery bottleneck. Unlike natural products which are curated by billions of years of microbial antagonism, the vast synthetic chemical space still requires artificial curation through the therapeutics science of antibacterial drugs - a systematic understanding of how small molecules interact with bacterial physiology, effect desired phenotypes, and benefit the host. Bacterial molecular genetics can elucidate pathogen biology relevant to therapeutics development, but it can also be applied directly to understanding mechanisms and liabilities of new chemical agents with new mechanisms of action. Therefore, the next phase of antibacterial drug discovery could be enabled by integrating chemical expertise with systematic dissection of bacterial infection biology. Facing the ambitious endeavour to find new molecules from nature or new-to-nature which cure bacterial infections, the capabilities furnished by modern chemical biology and molecular genetics can be applied to prospecting for chemical modulators of new targets which circumvent prevalent resistance mechanisms.

In vivo cloning of PCR product via site-specific recombination in Escherichia coli

Over the past years, several methods have been developed for gene cloning. Choosing a cloning strategy depends on various factors, among which simplicity and affordability have always been considered. The aim of this study, on the one hand, is to simplify gene cloning by skipping in vitro assembly reactions and, on the other hand, to reduce costs by eliminating relatively expensive materials. We investigated a cloning system using Escherichia coli harboring two plasmids, pLP-AmpR and pScissors-CmR. The pLP-AmpR contains a landing pad (LP) consisting of two genes (λ int and λ gam) that allow the replacement of the transformed linear DNA using site-specific recombination. After the replacement process, the inducible expressing SpCas9 and specific sgRNA from the pScissors-CmR (CRISPR/Cas9) vector leads to the removal of non-recombinant pLP-AmpR plasmids. The function of LP was explored by directly transforming PCR products. The pScissors-CmR plasmid was evaluated for curing three vectors, including the origins of pBR322, p15A, and pSC101. Replacing LP with a PCR product and fast-eradicating pSC101 origin-containing vectors was successful. Recombinant colonies were confirmed following gene replacement and plasmid curing processes. The results made us optimistic that this strategy may potentially be a simple and inexpensive cloning method. KEY POINTS: •The in vivo cloning was performed by replacing the target gene with the landing pad. •Fast eradication of non-recombinant plasmids was possible by adapting key vectors. •This strategy is not dependent on in vitro assembly reactions and expensive materials.

CRISPRi functional genomics in bacteria and its application to medical and industrial research

SUMMARYFunctional genomics is the use of systematic gene perturbation approaches to determine the contributions of genes under conditions of interest. Although functional genomic strategies have been used in bacteria for decades, recent studies have taken advantage of CRISPR (clustered regularly interspaced short palindromic repeats) technologies, such as CRISPRi (CRISPR interference), that are capable of precisely modulating expression of all genes in the genome. Here, we discuss and review the use of CRISPRi and related technologies for bacterial functional genomics. We discuss the strengths and weaknesses of CRISPRi as well as design considerations for CRISPRi genetic screens. We also review examples of how CRISPRi screens have defined relevant genetic targets for medical and industrial applications. Finally, we outline a few of the many possible directions that could be pursued using CRISPR-based functional genomics in bacteria. Our view is that the most exciting screens and discoveries are yet to come.

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.

CRISPR-Cas tools for simultaneous transcription & translation control in bacteria

Robust control over gene translation at arbitrary mRNA targets is an outstanding challenge in microbial synthetic biology. The development of tools that can regulate translation will greatly expand our ability to precisely control genes across the genome. In Escherichia coli, most genes are contained in multi-gene operons, which are subject to polar effects where targeting one gene for repression leads to silencing of other genes in the same operon. These effects pose a challenge for independently regulating individual genes in multi-gene operons. Here, we use CRISPR-dCas13 to address this challenge. We find dCas13-mediated repression exhibits up to 6-fold lower polar effects compared to dCas9. We then show that we can selectively activate single genes in a synthetic multi-gene operon by coupling dCas9 transcriptional activation of an operon with dCas13 translational repression of individual genes within the operon. We also show that dCas13 and dCas9 can be multiplexed for improved biosynthesis of a medically-relevant human milk oligosaccharide. Taken together, our findings suggest that combining transcriptional and translational control can access effects that are difficult to achieve with either mode independently. These combined tools for gene regulation will expand our abilities to precisely engineer bacteria for biotechnology and perform systematic genetic screens.

Expanding the flexibility of base editing for high-throughput genetic screens in bacteria

Genome-wide screens have become powerful tools for elucidating genotype-to-phenotype relationships in bacteria. Of the varying techniques to achieve knockout and knockdown, CRISPR base editors are emerging as promising options. However, the limited number of available, efficient target sites hampers their use for high-throughput screening. Here, we make multiple advances to enable flexible base editing as part of high-throughput genetic screening in bacteria. We first co-opt the Streptococcus canis Cas9 that exhibits more flexible protospacer-adjacent motif recognition than the traditional Streptococcus pyogenes Cas9. We then expand beyond introducing premature stop codons by mutating start codons. Next, we derive guide design rules by applying machine learning to an essentiality screen conducted in Escherichia coli. Finally, we rescue poorly edited sites by combining base editing with Cas9-induced cleavage of unedited cells, thereby enriching for intended edits. The efficiency of this dual system was validated through a conditional essentiality screen based on growth in minimal media. Overall, expanding the scope of genome-wide knockout screens with base editors could further facilitate the investigation of new gene functions and interactions in bacteria.

A CRISPR interference system for engineering biological nitrogen fixation

A grand challenge for the next century is in facing a changing climate through bioengineering solutions. Biological nitrogen fixation, the globally consequential, nitrogenase-catalyzed reduction of atmospheric nitrogen to bioavailable ammonia, is a vital area of focus. Nitrogen fixation engineering relies upon extensive understanding of underlying genetics in microbial models, including the broadly utilized gammaproteobacterium, Azotobacter vinelandii (A. vinelandii). Here, we report the first CRISPR interference (CRISPRi) system for targeted gene silencing in A. vinelandii that integrates genomically via site-specific transposon insertion. We demonstrate that CRISPRi can repress transcription of an essential nitrogen fixation gene by ~60%. Further, we show that nitrogenase genes are suitably expressed from the transposon insertion site, indicating that CRISPRi and engineered nitrogen fixation genes can be co-integrated for combinatorial studies of gene expression and engineering. Our established CRISPRi system fills an important gap for engineering microbial nitrogen fixation for desired purposes.IMPORTANCEAll life on Earth requires nitrogen to survive. About 78% of the atmosphere alone is nitrogen, yet humans cannot use it directly. Instead, we obtain the nitrogen we need for our survival through the food we eat. For more than 100 years, a substantial portion of agricultural productivity has relied on industrial methods for nitrogen fertilizer synthesis, which consumes significant amounts of nonrenewable energy resources and exacerbates environmental degradation and human-induced climate change. Promising alternatives to these industrial methods rely on engineering the only biological pathway for generating bioaccessible nitrogen: microbial nitrogen fixation. Bioengineering strategies require an extensive understanding of underlying genetics in nitrogen-fixing microbes, but genetic tools for this critical goal remain lacking. The CRISPRi gene silencing system that we report, developed in the broadly utilized nitrogen-fixing bacterial model, Azotobacter vinelandii, is an important step toward elucidating the complexity of nitrogen fixation genetics and enabling their manipulation.

Model-guided metabolic rewiring to bypass pyruvate oxidation for pyruvate derivative synthesis by minimizing carbon loss

Engineering microbial hosts to synthesize pyruvate derivatives depends on blocking pyruvate oxidation, thereby causing severe growth defects in aerobic glucose-based bioprocesses. To decouple pyruvate metabolism from cell growth to improve pyruvate availability, a genome-scale metabolic model combined with constraint-based flux balance analysis, geometric flux balance analysis, and flux variable analysis was used to identify genetic targets for strain design. Using translation elements from a ~3,000 cistronic library to modulate fxpK expression in a bicistronic cassette, a bifido shunt pathway was introduced to generate three molecules of non-pyruvate-derived acetyl-CoA from one molecule of glucose, bypassing pyruvate oxidation and carbon dioxide generation. The dynamic control of flux distribution by T7 RNAP-mediated synthetic small RNA decoupled pyruvate catabolism from cell growth. Adaptive laboratory evolution and multi-omics analysis revealed that a mutated isocitrate dehydrogenase functioned as a metabolic switch to activate the glyoxylate shunt as the only C4 anaplerotic pathway to generate malate from two molecules of acetyl-CoA input and bypass two decarboxylation reactions in the tricarboxylic acid cycle. A chassis strain for pyruvate derivative synthesis was constructed to reduce carbon loss by using the glyoxylate shunt as the only C4 anaplerotic pathway and the bifido shunt as a non-pyruvate-derived acetyl-CoA synthetic pathway and produced 22.46, 27.62, and 6.28 g/L of l-leucine, l-alanine, and l-valine by a controlled small RNA switch, respectively. Our study establishes a novel metabolic pattern of glucose-grown bacteria to minimize carbon loss under aerobic conditions and provides valuable insights into cell design for manufacturing pyruvate-derived products.IMPORTANCEBio-manufacturing from biomass-derived carbon sources using microbes as a cell factory provides an eco-friendly alternative to petrochemical-based processes. Pyruvate serves as a crucial building block for the biosynthesis of industrial chemicals; however, it is different to improve pyruvate availability in vivo due to the coupling of pyruvate-derived acetyl-CoA with microbial growth and energy metabolism via the oxidative tricarboxylic acid cycle. A genome-scale metabolic model combined with three algorithm analyses was used for strain design. Carbon metabolism was reprogrammed using two genetic control tools to fine-tune gene expression. Adaptive laboratory evolution and multi-omics analysis screened the growth-related regulatory targets beyond rational design. A novel metabolic pattern of glucose-grown bacteria is established to maintain growth fitness and minimize carbon loss under aerobic conditions for the synthesis of pyruvate-derived products. This study provides valuable insights into the design of a microbial cell factory for synthetic biology to produce industrial bio-products of interest.

Genomic editing in Burkholderia multivorans by CRISPR/Cas9

Burkholderia cepacia complex bacteria have emerged as opportunistic pathogens in patients with cystic fibrosis and immunocompromised individuals, causing life-threatening infections. Because of the relevance of these microorganisms, genetic manipulation is crucial for explaining the genetic mechanisms leading to pathogenesis. Despite the availability of allelic exchange tools to obtain unmarked gene deletions in Burkholderia, these require a step of merodiploid formation and another of merodiploid resolution through two independent homologous recombination events, making the procedure long-lasting. The CRISPR/Cas9-based system could ease this constraint, as only one step is needed for allelic exchange. Here, we report the modification of a two-plasmid system (pCasPA and pACRISPR) for genome editing in Burkholderia multivorans. Several modifications were implemented, including selection marker replacement, the optimization of araB promoter induction for the expression of Cas9 and λ-Red system encoding genes, and the establishment of plasmid curing procedures based on the sacB gene or growth at a sub-optimal temperature of 18°C-20°C with serial passages. We have shown the efficiency of this CRISPR/Cas9 method in the precise and unmarked deletion of different genes (rpfR, bceF, cepR, and bcsB) from two strains of B. multivorans, as well as its usefulness in the targeted insertion of the gfp gene encoding the green fluorescence protein into a precise genome location. As pCasPA was successfully introduced in other Burkholderia cepacia complex species, this study opens up the possibility of using CRISPR/Cas9-based systems as efficient tools for genome editing in these species, allowing faster and more cost-effective genetic manipulation.IMPORTANCEBurkholderia encompasses different species of bacteria, some of them pathogenic to animals and plants, but others are beneficial by promoting plant growth through symbiosis or as biocontrol agents. Among these species, Burkholderia multivorans, a member of the Burkholderia cepacia complex, is one of the predominant species infecting the lungs of cystic fibrosis patients, often causing respiratory chronic infections that are very difficult to eradicate. Since the B. multivorans species is understudied, we have developed a genetic tool based on the CRISPR/Cas9 system to delete genes efficiently from the genomes of these strains. We could also insert foreign genes that can be precisely placed in a chosen genomic region. This method, faster than other conventional strategies based on allelic exchange, will have a major contribution to understanding the virulence mechanisms in B. multivorans, but it can likely be extended to other Burkholderia species.

A mobile CRISPRi collection enables genetic interaction studies for the essential genes of Escherichia coli

Advances in gene editing, in particular CRISPR interference (CRISPRi), have enabled depletion of essential cellular machinery to study the downstream effects on bacterial physiology. Here, we describe the construction of an ordered E. coli CRISPRi collection, designed to knock down the expression of 356 essential genes with the induction of a catalytically inactive Cas9, harbored on the conjugative plasmid pFD152. This mobile CRISPRi library can be conjugated into other ordered genetic libraries to assess combined effects of essential gene knockdowns with non-essential gene deletions. As proof of concept, we probed cell envelope synthesis with two complementary crosses: (1) an Lpp deletion into every CRISPRi knockdown strain and (2) the lolA knockdown plasmid into the Keio collection. These experiments revealed a number of notable genetic interactions for the essential phenotype probed and, in particular, showed suppressing interactions for the loci in question.

Improved prediction of bacterial CRISPRi guide efficiency from depletion screens through mixed-effect machine learning and data integration

CRISPR interference (CRISPRi) is the leading technique to silence gene expression in bacteria; however, design rules remain poorly defined. We develop a best-in-class prediction algorithm for guide silencing efficiency by systematically investigating factors influencing guide depletion in genome-wide essentiality screens, with the surprising discovery that gene-specific features substantially impact prediction. We develop a mixed-effect random forest regression model that provides better estimates of guide efficiency. We further apply methods from explainable AI to extract interpretable design rules from the model. This study provides a blueprint for predictive models for CRISPR technologies where only indirect measurements of guide activity are available.

De Novo Synthesis of a Conjugative System from Human Gut Metagenomic Data for Targeted Delivery of Cas9 Antimicrobials

Metagenomic sequences represent an untapped source of genetic novelty, particularly for conjugative systems that could be used for plasmid-based delivery of Cas9-derived antimicrobial agents. However, unlocking the functional potential of conjugative systems purely from metagenomic sequences requires the identification of suitable candidate systems as starting scaffolds for de novo DNA synthesis. Here, we developed a bioinformatics approach that searches through the metagenomic "trash bin" for genes associated with conjugative systems present on contigs that are typically excluded from common metagenomic analysis pipelines. Using a human metagenomic gut data set representing 2805 taxonomically distinct units, we identified 1598 contigs containing conjugation genes with a differential distribution in human cohorts. We synthesized de novo an entire Citrobacter spp. conjugative system of 54 kb containing at least 47 genes and assembled it into a plasmid, pCitro. We found that pCitro conjugates from Escherichia coli to Citrobacter rodentium with a 30-fold higher frequency than to E. coli, and is compatible with Citrobacter resident plasmids. Mutations in the traV and traY conjugation components of pCitro inhibited conjugation. We showed that pCitro can be repurposed as an antimicrobial delivery agent by programming it with the TevCas9 nuclease and Citrobacter-specific sgRNAs to kill C. rodentium. Our study reveals a trove of uncharacterized conjugative systems in metagenomic data and describes an experimental framework to animate these large genetic systems as novel target-adapted delivery vectors for Cas9-based editing of bacterial genomes.

Synthetic Biology Toolbox for Nitrogen-Fixing Soil Microbes

The soil environment adjacent to plant roots, termed the rhizosphere, is home to a wide variety of microorganisms that can significantly affect the physiology of nearby plants. Microbes in the rhizosphere can provide nutrients, secrete signaling compounds, and inhibit pathogens. These processes could be manipulated with synthetic biology to enhance the agricultural performance of crops grown for food, energy, or environmental remediation, if methods can be implemented in these nonmodel microbes. A common first step for domesticating nonmodel organisms is the development of a set of genetic engineering tools, termed a synthetic biology toolbox. A toolbox comprises transformation protocols, replicating vectors, genome engineering (e.g., CRISPR/Cas9), constitutive and inducible promoter systems, and other gene expression control elements. This work validated synthetic biology toolboxes in three nitrogen-fixing soil bacteria: Azotobacter vinelandii, Stutzerimonas stutzeri (Pseudomonas stutzeri), and a new isolate of Klebsiella variicola. All three organisms were amenable to transformation and reporter protein expression, with several functional inducible systems available for each organism. S. stutzeri and K. variicola showed more reliable plasmid-based expression, resulting in successful Cas9 recombineering to create scarless deletions and insertions. Using these tools, we generated mutants with inducible nitrogenase activity and introduced heterologous genes to produce resorcinol products with relevant biological activity in the rhizosphere.

Construction of a CRISPR Interference System for Gene Knockdown in Stenotrophomonas maltophilia AGS-1 from Aerobic Granular Sludge

To identify the function of attachment genes involved in biofilm formation in Stenotrophomonas maltophilia AGS-1 isolated from aerobic granular sludge, an effective gene molecular tool is needed. We developed a two-plasmid CRISPRi system in Stenotrophomonas maltophilia AGS-1. One plasmid expressed dCas9 protein with the l-arabinose inducible promoter, and the other plasmid contained the sgRNA cassette complementary to the target gene. Under control of the araC-inducible promoter, this system exhibited little leaky basal expression and highly induced expression that silenced endogenous and exogenous genes with reversible knockdown. This system achieved up to 211-fold suppression for mCherry expression on the nontemplate strand compared to the template strand (91-fold). The utility of the developed CRISPRi platform was also characterized by suppressing the xanA and rpfF genes. The expression of these two genes was rapidly depleted and the adhesion ability decreased, which demonstrated that the modulation of either gene was an important factor for biofilm formation of the AGS-1 strain. The system also tested the ability to simultaneously silence transcriptional suppression of multiple targeted genes, an entire operon, or part of it. Lastly, the use of CRISPRi allowed us to dissect the gene intricacies involved in flagellar biosynthesis. Collectively, these results demonstrated that the CRISPRi system was a simple, feasible, and controllable manipulation system of gene expression in the AGS-1 strain.

Synthetic Gene Circuits Combining CRISPR Interference and CRISPR Activation in E. coli: Importance of Equal Guide RNA Binding Affinities to Avoid Context-Dependent Effects

Gene expression control based on clustered regularly interspaced short palindromic repeats (CRISPR) has emerged as a powerful approach for constructing synthetic gene circuits. While the use of CRISPR interference (CRISPRi) is already well-established in prokaryotic circuits, CRISPR activation (CRISPRa) is less mature, and a combination of the two in the same circuits is only just emerging. Here, we report that combining CRISPRi with SoxS-based CRISPRa in Escherichia coli can lead to context-dependent effects due to different affinities in the formation of CRISPRa and CRISPRi complexes, resulting in loss of predictable behavior. We show that this effect can be avoided by using the same scaffold guide RNA structure for both complexes.

Interspecific competition can drive plasmid loss from a focal species in a microbial community

Plasmids are key disseminators of antimicrobial resistance genes and virulence factors, and it is therefore critical to predict and reduce plasmid spread within microbial communities. The cost of plasmid carriage is a key metric that can be used to predict plasmids' ecological fate, and it is unclear whether plasmid costs are affected by growth partners in a microbial community. We carried out competition experiments and tracked plasmid maintenance using a model system consisting of a synthetic and stable five-species community and a broad host-range plasmid, engineered to carry different payloads. We report that both the cost of plasmid carriage and its long-term maintenance in a focal strain depended on the presence of competitors, and that these interactions were species specific. Addition of growth partners increased the cost of a high-payload plasmid to a focal strain, and accordingly, plasmid loss from the focal species occurred over a shorter time frame. We propose that the destabilising effect of interspecific competition on plasmid maintenance may be leveraged in clinical and natural environments to cure plasmids from focal strains.

Effective gene silencing using type I-E CRISPR system in the multiploid, radiation-resistant bacterium Deinococcus radiodurans

The extremely radiation-resistant bacterium, Deinococcus radiodurans, is a microbe of importance, both, for studying stress tolerance mechanisms and as a chassis for industrial biotechnology. However, the molecular tools available for use in this organism continue to be limiting, with its multiploid genome presenting an additional challenge. In view of this, the clustered regularly interspaced short palindromic repeat (CRISPR)-Cas tools provide a large repertoire of applications for gene manipulation. We show the utility of the type I-E Cascade system for knocking down gene expression in this organism. A single-vector system was designed for the expression of the Cascade components as well as the crRNA. The type I-E Cascade system was better tolerated than the type II-A dCas9 system in D. radiodurans. An assayable acid phosphatase gene, phoN integrated into the genome of this organism could be knocked down to 10% of its activity using the Cascade system. Cascade-based knockdown of ssb, a gene important for radiation resistance resulted in poor recovery post-irradiation. Targeting the Radiation and Desiccation Response Motif (RDRM), upstream of the ssb, prevented de-repression of its expression upon radiation exposure. In addition to this, multi-locus targeting was demonstrated on the deinococcal genome, by knocking down both phoN and ssb expression simultaneously. The programmable CRISPR interference tool developed in this study will facilitate the study of essential genes, hypothetical genes, and cis-elements involved in radiation response as well as enable metabolic engineering in this organism. Further, the tool can be extended for implementing high-throughput approaches in such studies. IMPORTANCE Deinococcus radiodurans is a microbe that exhibits a very high degree of radiation resistance. In addition, it is also identified as an organism of industrial importance. We report the development of a gene-knockdown system in this organism by engineering a type I-E clustered regularly interspaced short palindromic repeat (CRISPR)-Cascade system. We used this system to silence an assayable acid phosphatase gene, phoN to 10% of its activity. The study further shows the application of the Cascade system to target an essential gene ssb, that caused poor recovery from radiation. We demonstrate the utility of CRISPR-Cascade to study the role of a regulatory cis-element in radiation response as well as for multi-gene silencing. This easy-to-implement CRISPR interference system would provide an effective tool for better understanding of complex phenomena such as radiation response in D. radiodurans and may also enhance the potential of this microbe for industrial application.

LptD depletion disrupts morphological homeostasis and upregulates carbohydrate metabolism in Escherichia coli

In a previous in silico study, we identified an essential outer membrane protein (LptD) as an attractive target for development of novel antibiotics. Here, we characterized the effects of LptD depletion on Escherichia coli physiology and morphology. An E. coli CRISPR interference (CRISPRi) strain was constructed to allow control of lptD expression. Induction of the CRISPRi system led to ∼440-fold reduction of gene expression. Dose-dependent growth inhibition was observed, where strong knockdown effectively inhibited initial growth but partial knockdown exhibited maximum overall killing after 24 h. LptD depletion led to morphological changes where cells exhibited long, filamentous cell shapes and cytoplasmic accumulation of lipopolysaccharide (LPS). Transcriptional profiling by RNA-Seq showed that LptD knockdown led to upregulation of carbohydrate metabolism, especially in the colanic acid biosynthesis pathway. This pathway was further overexpressed in the presence of sublethal concentrations of colistin, an antibiotic targeting LPS, indicating a specific transcriptional response to this synergistic envelope damage. Additionally, exposure to colistin during LptD depletion resulted in downregulation of pathways related to motility and chemotaxis, two important virulence traits. Altogether, these results show that LptD depletion (i) affects E. coli survival, (ii) upregulates carbohydrate metabolism, and (iii) synergizes with the antimicrobial activity of colistin.

OptoCRISPRi-HD: Engineering a Bacterial Green-Light-Activated CRISPRi System with a High Dynamic Range

The ability to modulate gene expression is crucial for studying gene function and programming cell behaviors. Combining the reliability of CRISPRi and the precision of optogenetics, the optoCRISPRi technique is emerging as an advanced tool for live-cell gene regulation. Since previous versions of optoCRISPRi often exhibit no more than a 10-fold dynamic range due to the leakage activity, they are not suitable for targets that are sensitive to such leakage or critical for cell growth. Here, we describe a green-light-activated CRISPRi system with a high dynamic range (40 fold) and the flexibility of changing targets in Escherichia coli. Our optoCRISPRi-HD system can efficiently repress essential genes, nonessential genes, or inhibit the initiation of DNA replication. Providing a regulative system with high resolution over space-time and extensive targets, our study would facilitate further research involving complex gene networks, metabolic flux redirection, or bioprinting.

Screening and identification of genes involved in β-alanine biosynthesis in Bacillus subtilis

β-alanine is the only naturally occurring β-amino acid, which is widely used in medicine, food, and feed fields, and generally produced through synthetic biological methods based on engineered strains of Escherichia coli or Corynebacterium glutamicum. However, the β-alanine biosynthesis in Bacillus subtilis, a traditional industrial model microorganism of food safety grade, has not been thoroughly explored. In this study, the native l-aspartate-α-decarboxylase was overexpressed in B. subtilis 168 to obtain an increase of 842% in β-alanine production. A total of 16 single-gene knockout strains were constructed to block the competitive consumption pathways to identify a total of 6 genes (i.e., ptsG, fbp, ydaP, yhfS, mmgA, and pckA) involved in β-alanine synthesis, while the multigene knockout of these 6 genes obtained an increased β-alanine production by 40.1%. Ten single-gene suppression strains with the competitive metabolic pathways inhibited revealed that the inhibited expressions of genes glmS, accB, and accA enhanced the β-alanine production. The introduction of heterologous phosphoenolpyruvate carboxylase increased the β-alanine production by 81.7%, which was 17-fold higher than that of the original strain. This was the first study using multiple molecular strategies to investigate the biosynthetic pathway of β-alanine in B. subtilis and to identify the genetic factors limiting the excessive synthesis of β-alanine by microorganisms.

Removal of AMR plasmids using a mobile, broad host-range CRISPR-Cas9 delivery tool

Antimicrobial resistance (AMR) genes are widely disseminated on plasmids. Therefore, interventions aimed at blocking plasmid uptake and transfer may curb the spread of AMR. Previous studies have used CRISPR-Cas-based technology to remove plasmids encoding AMR genes from target bacteria, using either phage- or plasmid-based delivery vehicles that typically have narrow host ranges. To make this technology feasible for removal of AMR plasmids from multiple members of complex microbial communities, an efficient, broad host-range delivery vehicle is needed. We engineered the broad host-range IncP1-plasmid pKJK5 to encode cas9 programmed to target an AMR gene. We demonstrate that the resulting plasmid pKJK5::csg has the ability to block the uptake of AMR plasmids and to remove resident plasmids from Escherichia coli. Furthermore, due to its broad host range, pKJK5::csg successfully blocked AMR plasmid uptake in a range of environmental, pig- and human-associated coliform isolates, as well as in isolates of two species of Pseudomonas. This study firmly establishes pKJK5::csg as a promising broad host-range CRISPR-Cas9 delivery tool for AMR plasmid removal, which has the potential to be applied in complex microbial communities to remove AMR genes from a broad range of bacterial species.

Boosting genome editing efficiency in human cells and plants with novel LbCas12a variants

BACKGROUND

Cas12a (formerly known as Cpf1), the class II type V CRISPR nuclease, has been widely used for genome editing in mammalian cells and plants due to its distinct characteristics from Cas9. Despite being one of the most robust Cas12a nucleases, LbCas12a in general is less efficient than SpCas9 for genome editing in human cells, animals, and plants.

RESULTS

To improve the editing efficiency of LbCas12a, we conduct saturation mutagenesis in E. coli and identify 1977 positive point mutations of LbCas12a. We selectively assess the editing efficiency of 56 LbCas12a variants in human cells, identifying an optimal LbCas12a variant (RVQ: G146R/R182V/E795Q) with the most robust editing activity. We further test LbCas12a-RV, LbCas12a-RRV, and LbCas12a-RVQ in plants and find LbCas12a-RV has robust editing activity in rice and tomato protoplasts. Interestingly, LbCas12a-RRV, resulting from the stacking of RV and D156R, displays improved editing efficiency in stably transformed rice and poplar plants, leading to up to 100% editing efficiency in T₀ plants of both plant species. Moreover, this high-efficiency editing occurs even at the non-canonical TTV PAM sites.

CONCLUSIONS

Our results demonstrate that LbCas12a-RVQ is a powerful tool for genome editing in human cells while LbCas12a-RRV confers robust genome editing in plants. Our study reveals the tremendous potential of these LbCas12a variants for advancing precision genome editing applications across a wide range of organisms.

Targeted and high-throughput gene knockdown in diverse bacteria using synthetic sRNAs

Synthetic sRNAs allow knockdown of target genes at translational level, but have been restricted to a limited number of bacteria. Here, we report the development of a broad-host-range synthetic sRNA (BHR-sRNA) platform employing the RoxS scaffold and the Hfq chaperone from Bacillus subtilis. BHR-sRNA is tested in 16 bacterial species including commensal, probiotic, pathogenic, and industrial bacteria, with >50% of target gene knockdown achieved in 12 bacterial species. For medical applications, virulence factors in Staphylococcus epidermidis and Klebsiella pneumoniae are knocked down to mitigate their virulence-associated phenotypes. For metabolic engineering applications, high performance Corynebacterium glutamicum strains capable of producing valerolactam (bulk chemical) and methyl anthranilate (fine chemical) are developed by combinatorial knockdown of target genes. A genome-scale sRNA library covering 2959 C. glutamicum genes is constructed for high-throughput colorimetric screening of indigoidine (natural colorant) overproducers. The BHR-sRNA platform will expedite engineering of diverse bacteria of both industrial and medical interest.

An overview of genome engineering in plants, including its scope, technologies, progress and grand challenges

Genome editing is a useful, adaptable, and favored technique for both functional genomics and crop enhancement. Over the years, rapidly evolving genome editing technologies, including clustered regularly interspaced short palindromic repeats/CRISPR-associated protein (CRISPR/Cas), transcription activator-like effector nucleases (TALENs), and zinc finger nucleases (ZFNs), have shown broad application prospects in gene function research and improvement of critical agronomic traits in many crops. These technologies have also opened up opportunities for plant breeding. These techniques provide excellent chances for the quick modification of crops and the advancement of plant science in the future. The current review describes various genome editing techniques and how they function, particularly CRISPR/Cas9 systems, which can contribute significantly to the most accurate characterization of genomic rearrangement and plant gene functions as well as the enhancement of critical traits in field crops. To accelerate the use of gene-editing technologies for crop enhancement, the speed editing strategy of gene-family members was designed. As it permits genome editing in numerous biological systems, the CRISPR technology provides a valuable edge in this regard that particularly captures the attention of scientists.

Single-Stranded DNA-Binding Proteins Mediate DSB Repair and Effectively Improve CRISPR/Cas9 Genome Editing in Escherichia coli and Pseudomonas

Single-stranded DNA-binding proteins (SSBs) are essential for all living organisms. Whether SSBs can repair DNA double-strand breaks (DSBs) and improve the efficiency of CRISPR/Cas9-mediated genome editing has not been determined. Here, based on a pCas/pTargetF system, we constructed pCas-SSB and pCas-T4L by replacing the λ-Red recombinases with Escherichia coli SSB and phage T4 DNA ligase in pCas, respectively. Inactivation of the E. coli lacZ gene with homologous donor dsDNA increased the gene editing efficiency of pCas-SSB/pTargetF by 21.4% compared to pCas/pTargetF. Inactivation of the E. coli lacZ gene via NHEJ increased the gene editing efficiency of pCas-SSB/pTargetF by 33.2% compared to pCas-T4L/pTargetF. Furthermore, the gene-editing efficiency of pCas-SSB/pTargetF in E. coli (ΔrecA, ΔrecBCD, ΔSSB) with or without donor dsDNA did not differ. Additionally, pCas-SSB/pTargetF with donor dsDNA successfully deleted the wp116 gene in Pseudomonas sp. UW4. These results demonstrate that E. coli SSB repairs DSBs caused by CRISPR/Cas9 and effectively improves CRISPR/Cas9 genome editing in E. coli and Pseudomonas.

Whole-Cell P450 Biocatalysis Using Engineered Escherichia coli with Fine-Tuned Heme Biosynthesis

By exploiting versatile P450 enzymes, whole-cell biocatalysis can be performed to synthesize valuable compounds in Escherichia coli. However, the insufficient supply of heme limits the whole-cell P450 biocatalytic activity. Here a strategy for improving intracellular heme biosynthesis to enhance the catalytic efficiencies of P450s is reported. After comparing the effects of improving heme transport and biosynthesis on P450 activities, intracellular heme biosynthesis is optimized through the integrated expression of necessary synthetic genes at proper ratios and the assembly of rate-limiting enzymes using DNA-guided scaffolds. The intracellular heme level is fine-tuned by the combined use of mutated heme-sensitive biosensors and small regulatory RNA systems. The catalytic efficiencies of three different P450s, BM3, sca-2, and CYP105D7, are enhanced through fine-tuning heme biosynthesis for the synthesis of hydroquinone, pravastatin, and 7,3',4'-trihydroxyisoflavone as example products of chemical intermediate, drug, and natural product, respectively. This strategy of fine-tuned heme biosynthesis will be generally useful for developing whole-cell biocatalysts involving hemoproteins.

Metabolically-targeted dCas9 expression in bacteria

The ability to restrict gene expression to a relevant bacterial species in a complex microbiome is an unsolved problem. In the context of the human microbiome, one desirable target metabolic activity are glucuronide-utilization enzymes (GUS) that are implicated in the toxic re-activation of glucuronidated compounds in the human gastrointestinal (GI) tract, including the chemotherapeutic drug irinotecan. Here, we take advantage of the variable distribution of GUS enzymes in bacteria as a means to distinguish between bacteria with GUS activity, and re-purpose the glucuronide-responsive GusR transcription factor as a biosensor to regulate dCas9 expression in response to glucuronide inducers. We fused the Escherichia coli gusA regulatory region to the dCas9 gene to create pGreg-dCas9, and showed that dCas9 expression is induced by glucuronides, but not other carbon sources. When conjugated from E. coli to Gammaproteobacteria derived from human stool, dCas9 expression from pGreg-dCas9 was restricted to GUS-positive bacteria. dCas9-sgRNAs targeted to gusA specifically down-regulated gus operon transcription in Gammaproteobacteria, with a resulting ∼100-fold decrease in GusA activity. Our data outline a general strategy to re-purpose bacterial transcription factors responsive to exogenous metabolites for precise ligand-dependent expression of genetic tools such as dCas9 in diverse bacterial species.

Expanding the Scope of Bacterial CRISPR Activation with PAM-Flexible dCas9 Variants

CRISPR-Cas transcriptional tools have been widely applied for programmable regulation of complex biological networks. In comparison to eukaryotic systems, bacterial CRISPR activation (CRISPRa) has stringent target site requirements for effective gene activation. While genes may not always have an NGG protospacer adjacent motif (PAM) at the appropriate position, PAM-flexible dCas9 variants can expand the range of targetable sites. Here we systematically evaluate a panel of PAM-flexible dCas9 variants for their ability to activate bacterial genes. We observe that dxCas9-NG provides a high dynamic range of gene activation for sites with NGN PAMs while dSpRY permits modest activity across almost any PAM. Similar trends were observed for heterologous and endogenous promoters. For all variants tested, improved PAM-flexibility comes with the trade-off that CRISPRi-mediated gene repression becomes less effective. Weaker CRISPR interference (CRISPRi) gene repression can be partially rescued by expressing multiple sgRNAs to target many sites in the gene of interest. Our work provides a framework to choose the most effective dCas9 variant for a given set of gene targets, which will further expand the utility of CRISPRa/i gene regulation in bacterial systems.

Repurposing the Endogenous CRISPR-Cas9 System for High-Efficiency Genome Editing in Lacticaseibacillus paracasei

Lactobacilli such as Lacticaseibacillus (Lcb) paracasei are generally regarded as safe and health-promoting microbes, and have been widely applied in food and pharmaceutical industries. However, the genetic bases of their beneficial properties were mostly uncertain because of the lack of effective genetic manipulation tools. The type II CRISPR-Cas9 system is the largest family present in lactobacilli, but none of them yet have been developed for genetic modifications. Here, we establish the first endogenous CRISPR-Cas9 genome-editing system in lactobacilli. With a validated protospacer adjacent motif (PAM) and customized single guide RNA (sgRNA) expression cassette, the native CRISPR-Cas9 system was reprogrammed to achieve gene deletion and chromosomal insertion at over 90% efficiency, as well as nucleotide substitution at ≥50% efficiency. We also effectively accomplished deletions of large genomic fragments (5-10 kb) and simultaneous deletion of multiple genes at distal loci, both of which are the first cases in lactobacilli when either endogenous or exogenous CRISPR-Cas systems were employed. In addition, we designed a controllable plasmid-targeting sgRNA expression module and integrated it into the editing plasmid. The all-in-one vector realized gene deletion and plasmid curing at high efficiency (>90%). Collectively, the present study develops a convenient and precise genetic tool in Lcb. paracasei and contributes to the genetics and engineering of lactobacilli.

From DNA-protein interactions to the genetic circuit design using CRISPR-dCas systems

In the last decade, the CRISPR-Cas technology has gained widespread popularity in different fields from genome editing and detecting specific DNA/RNA sequences to gene expression control. At the heart of this technology is the ability of CRISPR-Cas complexes to be programmed for targeting particular DNA loci, even when using catalytically inactive dCas-proteins. The repertoire of naturally derived and engineered dCas-proteins including fusion proteins presents a promising toolbox that can be used to construct functional synthetic genetic circuits. Rational genetic circuit design, apart from having practical relevance, is an important step towards a deeper quantitative understanding of the basic principles governing gene expression regulation and functioning of living organisms. In this minireview, we provide a succinct overview of the application of CRISPR-dCas-based systems in the emerging field of synthetic genetic circuit design. We discuss the diversity of dCas-based tools, their properties, and their application in different types of genetic circuits and outline challenges and further research directions in the field.

CRISPRi-based programmable logic inverter cascade for antibiotic-free selection and maintenance of multiple plasmids

Antibiotics have been widely used for plasmid-mediated cell engineering. However, continued use of antibiotics increases the metabolic burden, horizontal gene transfer risks, and biomanufacturing costs. There are limited approaches to maintaining multiple plasmids without antibiotics. Herein, we developed an inverter cascade using CRISPRi by building a plasmid containing a single guide RNA (sgRNA) landing pad (pSLiP); this inhibited host cell growth by repressing an essential cellular gene. Anti-sgRNAs on separate plasmids restored cell growth by blocking the expression of growth-inhibitory sgRNAs in pSLiP. We maintained three plasmids in Escherichia coli with a single antibiotic selective marker. To completely avoid antibiotic use and maintain the CRISPRi-based logic inverter cascade, we created a novel d-glutamate auxotrophic E. coli. This enabled the stable maintenance of the plasmid without antibiotics, enhanced the production of the terpenoid, (-)-α-bisabolol, and generation of an antibiotic-resistance gene-free plasmid. CRISPRi is therefore widely applicable in genetic circuits and may allow for antibiotic-free biomanufacturing.

CRISPRactivation-SMS, a message for PAM sequence independent gene up-regulation in Escherichia coli

Governance of the endogenous gene regulatory network enables the navigation of cells towards beneficial traits for recombinant protein production. CRISPRactivation and interference provides the basis for gene expression modulation but is primarily applied in eukaryotes. Particularly the lack of wide-ranging prokaryotic CRISPRa studies might be attributed to intrinsic limitations of bacterial activators and Cas9 proteins. While bacterial activators need accurate spatial orientation and distancing towards the target promoter to be functional, Cas9-based CRISPR tools only bind sites adjacent to NGG PAM sequences. These circumstances hampered Cas9-guided activators from mediating the up-regulation of endogenous genes at precise positions in bacteria. We could overcome this limitation by combining the PAM independent Cas9 variant SpRY and a CRISPRa construct using phage protein MCP fused to transcriptional activator SoxS. This CRISPRa construct, referred to as SMS, was compared with previously reported CRISPRa constructs and showed up-regulation of a reporter gene library independent of its PAM sequence in Escherichia coli. We also demonstrated down-regulation and multi-gene expression control with SMS at non-NGG PAM sites. Furthermore, we successfully applied SMS to up-regulate endogenous genes, and transgenes at non-NGG PAM sites, which was impossible with the previous CRISPRa construct.

Overcoming Leak Sensitivity in CRISPRi Circuits Using Antisense RNA Sequestration and Regulatory Feedback

The controlled binding of the catalytically dead CRISPR nuclease (dCas) to DNA can be used to create complex, programmable transcriptional genetic circuits, a fundamental goal of synthetic biology. This approach, called CRISPR interference (CRISPRi), is advantageous over existing methods because the programmable nature of CRISPR proteins in principle enables the simultaneous regulation of many different targets without crosstalk. However, the performance of dCas-based genetic circuits is limited by both the sensitivity to leaky repression within CRISPRi logic gates and retroactive effects due to a shared pool of dCas proteins. By utilizing antisense RNAs (asRNAs) to sequester gRNA transcripts as well as CRISPRi feedback to self-regulate asRNA production, we demonstrate a mechanism that suppresses unwanted repression by CRISPRi and improves logical gene circuit function in Escherichia coli. This improvement is particularly pronounced during stationary expression when CRISPRi circuits do not achieve the expected regulatory dynamics. Furthermore, the use of dual CRISPRi/asRNA inverters restores the logical performance of layered circuits such as a double inverter. By studying circuit induction at the single-cell level in microfluidic channels, we provide insight into the dynamics of antisense sequestration of gRNA and regulatory feedback on dCas-based repression and derepression. These results demonstrate how CRISPRi inverters can be improved for use in more complex genetic circuitry without sacrificing the programmability and orthogonality of dCas proteins.

Bacterial genome reductions: Tools, applications, and challenges

Bacterial cells are widely used to produce value-added products due to their versatility, ease of manipulation, and the abundance of genome engineering tools. However, the efficiency of producing these desired biomolecules is often hindered by the cells’ own metabolism, genetic instability, and the toxicity of the product. To overcome these challenges, genome reductions have been performed, making strains with the potential of serving as chassis for downstream applications. Here we review the current technologies that enable the design and construction of such reduced-genome bacteria as well as the challenges that limit their assembly and applicability. While genomic reductions have shown improvement of many cellular characteristics, a major challenge still exists in constructing these cells efficiently and rapidly. Computational tools have been created in attempts at minimizing the time needed to design these organisms, but gaps still exist in modelling these reductions in silico. Genomic reductions are a promising avenue for improving the production of value-added products, constructing chassis cells, and for uncovering cellular function but are currently limited by their time-consuming construction methods. With improvements to and the creation of novel genome editing tools and in silico models, these approaches could be combined to expedite this process and create more streamlined and efficient cell factories.

CRISPR/dCas9-based metabolic pathway engineering for the systematic optimization of exopolysaccharide biosynthesis in Streptococcus thermophilus

Streptococcus thermophilus is used extensively in the dairy industry and has shown great promise as a chassis cell for the biosynthesis of high-value metabolites. However, metabolic engineering in S. thermophilus lacks effective genetic modification tools to modulate gene expression to relieve metabolic burden and maximize the production of desired compounds. Here, we developed a clustered regularly interspaced short palindromic repeats interference (CRISPRi) system for efficient gene transcriptional modulation in S. thermophilus. Our CRISPRi system typically achieved 66 to 98% knockdown of single or multiple gene expression. We used CRISPRi for the biosynthesis of a new exopolysaccharide (EPS) as a paradigm model. Repression of galK at module of uridine diphosphate glucose sugar metabolism and overexpression of epsA and epsE at EPS synthesis module resulted in an approximately 2-fold increase in EPS titer (277 mg/L) when compared with a control strain. This study demonstrated the effectiveness of CRISPRi as a powerful metabolic engineering tool and synthetic biology strategy for S. thermophilus.

Mitigating Host Burden of Genetic Circuits by Engineering Autonegatively Regulated Parts and Improving Functional Prediction

Mitigating unintended interferences between circuits and host cells is key to realize applications of synthetic regulatory systems both for bacteria and mammalian cells. Here, we demonstrated that growth burden and circuit dysregulation occurred in a concentration-dependent manner for specific transcription factors (CymR*/CymR) in E.coli, and direct negative feedback modules were able to control the concentration of CymR*/CymR, mitigate growth burden, and restore circuit functions. A quantitative design scheme was developed for circuits embedded with autorepression modules. Four key parameters were theoretically identified to determine the performance of autoregulated switches and were experimentally modified by fine-tuning promoter architectures and cooperativity. Using this strategy, we synthesized a number of switches and demonstrated its improvement of product titers and host growth controlling the complex deoxyviolacein biosynthesis pathway. Furthermore, we restored functions of a dysregulated multilayer NOR gate by integrating autorepression modules. Our work provides a blueprint for engineering host-adaptable synthetic systems.

CRISPR-Based Approaches for Gene Regulation in Non-Model Bacteria

CRISPR interference (CRISPRi) and CRISPR activation (CRISPRa) have become ubiquitous approaches to control gene expression in bacteria due to their simple design and effectiveness. By regulating transcription of a target gene(s), CRISPRi/a can dynamically engineer cellular metabolism, implement transcriptional regulation circuitry, or elucidate genotype-phenotype relationships from smaller targeted libraries up to whole genome-wide libraries. While CRISPRi/a has been primarily established in the model bacteria Escherichia coli and Bacillus subtilis, a growing numbering of studies have demonstrated the extension of these tools to other species of bacteria (here broadly referred to as non-model bacteria). In this mini-review, we discuss the challenges that contribute to the slower creation of CRISPRi/a tools in diverse, non-model bacteria and summarize the current state of these approaches across bacterial phyla. We find that despite the potential difficulties in establishing novel CRISPRi/a in non-model microbes, over 190 recent examples across eight bacterial phyla have been reported in the literature. Most studies have focused on tool development or used these CRISPRi/a approaches to interrogate gene function, with fewer examples applying CRISPRi/a gene regulation for metabolic engineering or high-throughput screens and selections. To date, most CRISPRi/a reports have been developed for common strains of non-model bacterial species, suggesting barriers remain to establish these genetic tools in undomesticated bacteria. More efficient and generalizable methods will help realize the immense potential of programmable CRISPR-based transcriptional control in diverse bacteria.

Gene Silencing through CRISPR Interference in Mycoplasmas

Mycoplasmas are pathogenic, genome-reduced bacteria. The development of such fields of science as system and synthetic biology is closely associated with them. Despite intensive research of different representatives of this genus, genetic manipulations remain challenging in mycoplasmas. Here we demonstrate a single-plasmid transposon-based CRISPRi system for the repression of gene expression in mycoplasmas. We show that selected expression determinants provide a level of dCas9 that does not lead to a significant slow-down of mycoplasma growth. For the first time we describe the proteomic response of genome-reduced bacteria to the expression of exogenous dcas9. The functionality of the resulting vector is confirmed by targeting the three genes coding transcription factors-fur, essential spxA, whiA, and histone-like protein hup1 in Mycoplasma gallisepticum. As a result, the expression level of each gene was decreased tenfold and influenced the mRNA level of predicted targets of transcription factors. To illustrate the versatility of this vector, we performed a knockdown of metabolic genes in a representative member of another cluster of the Mycoplasma genus-Mycoplasma hominis. The developed CRISPRi system is a powerful tool to discover the functioning of genes that are essential, decipher regulatory networks and that can help to identify novel drug targets to control Mycoplasma infections.

Developing a CRISPR-assisted base-editing system for genome engineering of Pseudomonas chlororaphis

Pseudomonas chlororaphis is a non‐pathogenic, plant growth‐promoting rhizobacterium that secretes phenazine compounds with broad‐spectrum antibiotic activity. Currently available genome‐editing methods for P. chlororaphis are based on homologous recombination (HR)‐dependent allelic exchange, which requires both exogenous DNA repair proteins (e.g. λ‐Red–like systems) and endogenous functions (e.g. RecA) for HR and/or providing donor DNA templates. In general, these procedures are time‐consuming, laborious and inefficient. Here, we established a CRISPR‐assisted base‐editing (CBE) system based on the fusion of a rat cytidine deaminase (rAPOBEC1), enhanced‐specificity Cas9 nickase (eSpCas9pp^D10A) and uracil DNA glycosylase inhibitor (UGI). This CBE system converts C:G into T:A without DNA strands breaks or any donor DNA template. By engineering a premature STOP codon in target spacers, the hmgA and phzO genes of P. chlororaphis were successfully interrupted at high efficiency. The phzO‐inactivated strain obtained by base editing exhibited identical phenotypic features as compared with a mutant obtained by HR‐based allelic exchange. The use of this CBE system was extended to other P. chlororaphis strains (subspecies LX24 and HT66) and also to P. fluorescens 10586, with an equally high editing efficiency. The wide applicability of this CBE method will accelerate bacterial physiology research and metabolic engineering of non‐traditional bacterial hosts.

Gradients in gene essentiality reshape antibacterial research

Essential genes encode the processes that are necessary for life. Until recently, commonly applied binary classifications left no space between essential and non-essential genes. In this review, we frame bacterial gene essentiality in the context of genetic networks. We explore how the quantitative properties of gene essentiality are influenced by the nature of the encoded process, environmental conditions and genetic background, including a strain's distinct evolutionary history. The covered topics have important consequences for antibacterials, which inhibit essential processes. We argue that the quantitative properties of essentiality can thus be used to prioritize antibacterial cellular targets and desired spectrum of activity in specific infection settings. We summarize our points with a case study on the core essential genome of the cystic fibrosis pathobiome and highlight avenues for targeted antibacterial development.

Characterization of CRISPR-Cas systems in Bifidobacterium breve

The clustered regularly interspaced short palindromic repeat (CRISPR)-CRISPR-associated protein (Cas) system is an important adaptive immune system for bacteria to resist foreign DNA infection, which has been widely used in genotyping and gene editing. To provide a theoretical basis for the application of the CRISPR-Cas system in Bifidobacterium breve, the occurrence and diversity of CRISPR-Cas systems were analysed in 150 B. breve strains. Specifically, 47 % (71/150) of B. breve genomes possessed the CRISPR-Cas system, and type I-C CRISPR-Cas system was the most widely distributed among those strains. The spacer sequences present in B. breve can be used as a genotyping marker. Additionally, the phage assembly-related proteins were important targets of the type I-C CRISPR-Cas system in B. breve, and the protospacer adjacent motif sequences were further characterized in B. breve type I-C system as 5'-TTC-3'. All these results might provide a molecular basis for the development of endogenous genome editing tools in B. breve.

Synthetic 3'-UTR valves for optimal metabolic flux control in Escherichia coli

As the design of genetic circuitry for synthetic biology becomes more sophisticated, diverse regulatory bioparts are required. Despite their importance, well-characterized 3′-untranslated region (3′-UTR) bioparts are limited. Thus, transcript 3′-ends require further investigation to understand the underlying regulatory role and applications of the 3′-UTR. Here, we revisited the use of Term-Seq in the Escherichia coli strain K-12 MG1655 to enhance our understanding of 3′-UTR regulatory functions and to provide a diverse collection of tunable 3′-UTR bioparts with a wide termination strength range. Comprehensive analysis of 1,629 transcript 3′-end positions revealed multiple 3′-termini classes generated through transcription termination and RNA processing. The examination of individual Rho-independent terminators revealed a reduction in downstream gene expression over a wide range, which led to the design of novel synthetic metabolic valves that control metabolic fluxes in branched pathways. These synthetic metabolic valves determine the optimal balance of heterologous pathways for maximum target biochemical productivity. The regulatory strategy using 3′-UTR bioparts is advantageous over promoter- or 5′-UTR-based transcriptional control as it modulates gene expression at transcription levels without trans-acting element requirements (e.g. transcription factors). Our results provide a foundational platform for 3′-UTR engineering in synthetic biology applications.

Spacer2PAM: A computational framework to guide experimental determination of functional CRISPR-Cas system PAM sequences

RNA-guided nucleases from CRISPR-Cas systems expand opportunities for precise, targeted genome modification. Endogenous CRISPR-Cas systems in many prokaryotes are attractive to circumvent expression, functionality, and unintended activity hurdles posed by heterologous CRISPR-Cas effectors. However, each CRISPR-Cas system recognizes a unique set of protospacer adjacent motifs (PAMs), which requires identification by extensive screening of randomized DNA libraries. This challenge hinders development of endogenous CRISPR-Cas systems, especially those based on multi-protein effectors and in organisms that are slow-growing or have transformation idiosyncrasies. To address this challenge, we present Spacer2PAM, an easy-to-use, easy-to-interpret R package built to predict and guide experimental determination of functional PAM sequences for any CRISPR-Cas system given its corresponding CRISPR array as input. Spacer2PAM can be used in a 'Quick' method to generate a single PAM prediction or in a 'Comprehensive' method to inform targeted PAM libraries small enough to screen in difficult to transform organisms. We demonstrate Spacer2PAM by predicting PAM sequences for industrially relevant organisms and experimentally identifying seven PAM sequences that mediate interference from the Spacer2PAM-informed PAM library for the type I-B CRISPR-Cas system from Clostridium autoethanogenum. We anticipate that Spacer2PAM will facilitate the use of endogenous CRISPR-Cas systems for industrial biotechnology and synthetic biology.

Uncovering the Distinct Properties of a Bacterial Type I-E CRISPR Activation System

Synthetic gene regulators based upon CRISPR-Cas systems offer programmable technologies to control gene expression in bacteria. Bacterial CRISPR activators (CRISPRa) have been developed that use engineered type II CRISPR-dCas9 to localize transcription activation domains near promoter elements. However, several reports have demonstrated distance-dependent requirements and periodical activation patterns that overall limit their flexibility. Here, we demonstrate the potential of using an alternative type I-E CRISPR-Cas system to create a CRISPRa with distinct and expanded regulatory properties. Furthermore, we create the first bacterial CRISPRa system based upon a type I-E CRISPR-Cas and characterize the distance-dependent activation patterns to reveal a distinct and more frequent periodicity of activation.

CRISPR Interference Modules as Low-Burden Logic Inverters in Synthetic Circuits

CRISPR and CRISPRi systems have revolutionized our biological engineering capabilities by enabling the editing and regulation of virtually any gene, via customization of single guide RNA (sgRNA) sequences. CRISPRi modules can work as programmable logic inverters, in which the dCas9-sgRNA complex represses a target transcriptional unit. They have been successfully used in bacterial synthetic biology to engineer information processing tasks, as an alternative to the traditionally adopted transcriptional regulators. In this work, we investigated and modulated the transfer function of several model systems with specific focus on the cell load caused by the CRISPRi logic inverters. First, an optimal expression cassette for dCas9 was rationally designed to meet the low-burden high-repression trade-off. Then, a circuit collection was studied at varying levels of dCas9 and sgRNAs targeting three different promoters from the popular tet, lac and lux systems, placed at different DNA copy numbers. The CRISPRi NOT gates showed low-burden properties that were exploited to fix a high resource-consuming circuit previously exhibiting a non-functional input-output characteristic, and were also adopted to upgrade a transcriptional regulator-based NOT gate into a 2-input NOR gate. The obtained data demonstrate that CRISPRi-based modules can effectively act as low-burden components in different synthetic circuits for information processing.

Repurposing CRISPR-Cas Systems as Genetic Tools for the Enterobacteriales

Over the last decade, the study of CRISPR-Cas systems has progressed from a newly discovered bacterial defense mechanism to a diverse suite of genetic tools that have been applied across all domains of life. While the initial applications of CRISPR-Cas technology fulfilled a need to more precisely edit eukaryotic genomes, creative "repurposing" of this adaptive immune system has led to new approaches for genetic analysis of microorganisms, including improved gene editing, conditional gene regulation, plasmid curing and manipulation, and other novel uses. The main objective of this review is to describe the development and current state-of-the-art use of CRISPR-Cas techniques specifically as it is applied to members of the Enterobacteriales. While many of the applications covered have been initially developed in Escherichia coli, we also highlight the potential, along with the limitations, of this technology for expanding the availability of genetic tools in less-well-characterized non-model species, including bacterial pathogens.

CRISPR/Cas Technologies and Their Applications in Escherichia coli

The clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein (Cas) systems have revolutionized genome editing and greatly promoted the development of biotechnology. However, these systems unfortunately have not been developed and applied in bacteria as extensively as in eukaryotic organism. Here, the research progress on the most widely used CRISPR/Cas tools and their applications in Escherichia coli is summarized. Genome editing based on homologous recombination, non-homologous DNA end-joining, transposons, and base editors are discussed. Finally, the state of the art of transcriptional regulation using CRISPRi is briefly reviewed. This review provides a useful reference for the application of CRISPR/Cas systems in other bacterial species.

Detection of pks Island mRNAs Using Toehold Sensors in Escherichia coli

Synthetic biologists have applied biomolecular engineering approaches toward the goal of novel biological devices and have shown progress in diverse areas of medicine and biotechnology. Especially promising is the application of synthetic biological devices towards a novel class of molecular diagnostics. As an example, a de-novo-designed riboregulator called toehold switch, with its programmability and compatibility with field-deployable devices showed promising in vitro applications for viral RNA detection such as Zika and Corona viruses. However, the in vivo application of high-performance RNA sensors remains challenging due to the secondary structure of long mRNA species. Here, we introduced ‘Helper RNAs’ that can enhance the functionality of toehold switch sensors by mitigating the effect of secondary structures around a target site. By employing the helper RNAs, previously reported mCherry mRNA sensor showed improved fold-changes in vivo. To further generalize the Helper RNA approaches, we employed automatic design pipeline for toehold sensors that target the essential genes within the pks island, an important target of biomedical research in connection with colorectal cancer. The toehold switch sensors showed fold-changes upon the expression of full-length mRNAs that apparently depended sensitively on the identity of the gene as well as the predicted local structure within the target region of the mRNA. Still, the helper RNAs could improve the performance of toehold switch sensors in many instances, with up to 10-fold improvement over no helper cases. These results suggest that the helper RNA approaches can further assist the design of functional RNA devices in vivo with the aid of the streamlined automatic design software developed here. Further, our solutions for screening and stabilizing single-stranded region of mRNA may find use in other in vivo mRNA-sensing applications such as cas13 crRNA design, transcriptome engineering, and trans-cleaving ribozymes.

Bacterial resistance to CRISPR-Cas antimicrobials

In the age of antibiotic resistance and precise microbiome engineering, CRISPR-Cas antimicrobials promise to have a substantial impact on the way we treat diseases in the future. However, the efficacy of these antimicrobials and their mechanisms of resistance remain to be elucidated. We systematically investigated how a target E. coli strain can escape killing by episomally-encoded CRISPR-Cas9 antimicrobials. Using Cas9 from Streptococcus pyogenes (SpCas9) we studied the killing efficiency and resistance mutation rate towards CRISPR-Cas9 antimicrobials and elucidated the underlying genetic alterations. We find that killing efficiency is not correlated with the number of cutting sites or the type of target. While the number of targets did not significantly affect efficiency of killing, it did reduce the emergence of chromosomal mutations conferring resistance. The most frequent target of resistance mutations was the plasmid-encoded SpCas9 that was inactivated by bacterial genome rearrangements involving translocation of mobile genetic elements such as insertion elements. This resistance mechanism can be overcome by re-introduction of an intact copy of SpCas9. The work presented here provides a guide to design strategies that reduce resistance and improve the activity of CRISPR-Cas antimicrobials.

A transferrable and integrative type I-F Cascade for heterologous genome editing and transcription modulation

The Class 1 type I CRISPR–Cas systems represent the most abundant and diverse CRISPR systems in nature. However, their applications for generic genome editing have been hindered due to difficulties of introducing the class-specific, multi-component effectors (Cascade) in heterologous hosts for functioning. Here we established a transferrable Cascade system that enables stable integration and expression of a highly active type I-F Cascade in heterologous bacterial hosts for various genetic exploitations. Using the genetically recalcitrant Pseudomonas species as a paradigm, we show that the transferred Cascade displayed substantially higher DNA interference activity and greater editing capacity than both the integrative and plasmid-borne Cas9 systems, and enabled deletion of large fragments such as the 21-kb integrated cassette with efficiency and simplicity. An advanced I-F-λ_red system was further developed to enable editing in genotypes with poor homologous recombination capacity, clinical isolates lacking sequence information, and cells containing anti-CRISPR elements Acrs. Lastly, an ‘all-in-one’ I-F Cascade-mediated CRISPRi platform was developed for transcription modulation by simultaneous introduction of the Cascade and the programmed mini-CRISPR array in one-step. This study provides a framework for expanding the diverse type I Cascades for widespread, heterologous genome editing and establishment of editing techniques in ‘non-model’ bacterial species.