Generalized bacterial genome editing using mobile group II introns and Cre-lox

Establishment of a RecET-Assisted CRISPR-Cas12a System for Large Deoxyribonucleic Acid-Fragment Manipulation in Zymomonas mobilis

The lack of effective and efficient genome-editing toolkits for large DNA-fragment manipulation impedes the development of robust cell factories to meet the needs of fast-growing biomanufacturing. Zymomonas mobilis is an important nonmodel polyploid industrial strain with excellent industrial characteristics. Although diverse CRISPR-Cas systems have been established in Z. mobilis for gene deletion, replacement, and ssDNA recombination, it is still challenging to achieve large DNA-fragment manipulation due to its low recombination and repair efficiencies for double-strand DNA breaks. In this study, a RecET-assisted CRISPR-Cas12a genome editing system was developed using a chromosome-borne cas12a and recET-encoded recombinase, as well as an all-in-one editing plasmid. Different promoters were used for recET and cas12a to determine optimal expression. The combination of PB-cas12a_Pt-recET had the highest efficiency of 97.92 ± 2.95% for 9-kb DNA-fragment deletion, which also had efficiencies about 100%, 80%, and 5%, respectively, for the deletion of 9-16, 20-25, and 30 kb DNA fragments. The RecET-assisted CRISPR-Cas12a was further applied for deletions of different large gene clusters and had the potential for efficient pathway knock-in. This study highlights the importance of the Cas12a nuclease expression levels and the combination of the RecET system in improving the double-strand DNA repair capability for large DNA-fragment manipulation in Z. mobilis. The RecET-assisted CRISPR-Cas12a system established in this study provides a versatile and powerful tool for large DNA-fragment manipulation in Z. mobilis, which is beneficial for functional genomic research, strain improvement, as well as the development of synthetic microbial chassis.

In silico analysis of non-conventional gene targets for genetic interventions to enhance fatty acid production: a review

Since the 1990s, fatty acids (FA) have drawn significant industrial attention due to their diverse applications creating a demand for biological systems capable of producing high FA titers. While various strategies have been explored to achieve this, many of the conventional approaches rely on extensive genetic manipulations, which often result in strain instability, thus limiting its potential to yield better FA titers. Moreover, stresses such as pH, osmotic, and oxidative imbalances generated during FA production aggravate these challenges, further limiting FA titers. Under stress conditions, the cellular system responds by regulating stress-response proteins to bring about homeostasis. Recent findings suggest that transmembrane proteins, regulators of two-component systems, and cytoplasmic regulators can be strategically leveraged to address the problems related to stress-induced strain instability. Thus, non-conventional genetic targets, like chaperones (e.g., heat shock proteins) and DNA-binding transcriptional regulators (e.g., RcdA), which are not directly involved in FA metabolism, represent promising candidates to enhance strain stability and FA yields. Tools like Opt-Box and Weighted Gene Co-expression Network Analysis (WGCNA) serve as excellent platforms for understanding the cross-talk between these regulators and downstream enzymes. This review emphasizes the need for a shift towards identifying novel genetic targets by employing advanced in silico analysis and explains several molecular techniques that can aid in strain construction. Lastly, it discusses certain non-conventional gene targets that can help to overcome strain instability arising due to various stresses generated during/due to FA production.

Shewanella oneidensis: Biotechnological Application of Metal-Reducing Bacteria

What is an unconventional organism in biotechnology? The γ-proteobacterium Shewanella oneidensis might fall into this category as it was initially established as a laboratory model organism for a process that was not seen as potentially interesting for biotechnology. The reduction of solid-state extracellular electron acceptors such as iron and manganese oxides is highly relevant for many biogeochemical cycles, although it turned out in recent years to be quite relevant for many potential biotechnological applications as well. Applications started with the production of nanoparticles and dramatically increased after understanding that electrodes in bioelectrochemical systems can also be used by these organisms. From the potential production of current and hydrogen in these systems and the development of biosensors, the field expanded to anode-assisted fermentations enabling fermentation reactions that were - so far - dependent on oxygen as an electron acceptor. Now the field expands further to cathode-dependent production routines. As a side product to all these application endeavors, S. oneidensis was understood more and more, and our understanding and genetic repertoire is at eye level to E. coli. Corresponding to this line of thought, this chapter will first summarize the available arsenal of tools in molecular biology that was established for working with the organism and thereafter describe so far established directions of application. Last but not least, we will highlight potential future directions of work with the unconventional model organism S. oneidensis.

Contributions of Gamma-Aminobutyric Acid (GABA) Produced by Lactic Acid Bacteria on Food Quality and Human Health: Current Applications and Future Prospects

The need to increase food safety and improve human health has led to a worldwide increase in interest in gamma-aminobutyric acid (GABA), produced by lactic acid bacteria (LABs). GABA, produced from glutamic acid in a reaction catalyzed by glutamate decarboxylase (GAD), is a four-carbon, non-protein amino acid that is increasingly used in the food industry to improve the safety/quality of foods. In addition to the possible positive effects of GABA, called a postbiotic, on neuroprotection, improving sleep quality, alleviating depression and relieving pain, the various health benefits of GABA-enriched foods such as antidiabetic, antihypertension, and anti-inflammatory effects are also being investigated. For all these reasons, it is not surprising that efforts to identify LAB strains with a high GABA productivity and to increase GABA production from LABs through genetic engineering to increase GABA yield are accelerating. However, GABA's contributions to food safety/quality and human health have not yet been fully discussed in the literature. Therefore, this current review highlights the synthesis and food applications of GABA produced from LABs, discusses its health benefits such as, for example, alleviating drug withdrawal syndromes and regulating obesity and overeating. Still, other potential food and drug interactions (among others) remain unanswered questions to be elucidated in the future. Hence, this review paves the way toward further studies.

Increasing the efficiency of gene editing with CRISPR-Cas9 via concurrent expression of the Beta protein

Escherichia coli has emerged as an important host for the production of biopharmaceuticals or other industrially relevant molecules. An efficient gene editing tool is indispensable for ensuring high production levels and optimal release of target products. However, in Escherichia coli, the CRISPR-Cas9 system has been shown to achieve gene modifications with relatively low frequency. Large-scale PCR screening is required, hindering the identification of positive clones. The beta protein, which weakly binds to single-stranded DNA but tightly associates with complementary strand annealing products, offers a promising solution to this issue. In the present study, we describe a targeted and continuous gene editing strategy for the Escherichia coli genome. This strategy involves the coexpression of the beta protein alongside the CRISPR-Cas9 system, enabling a variety of genome modifications such as gene deletion and insertion with an efficiency exceeding 80 %. The integrity of beta proteins is essential for the CRISPR-Cas9/Beta-based gene editing system. In this work, the deletion of either the N- or C-terminal domain significantly impaired system efficiency. Overall, our findings established the CRISPR-Cas9/Beta system as a suitable gene editing tool for various applications in Escherichia coli.

Next generation probiotics: Engineering live biotherapeutics

The population dynamics of the human microbiome have been associated with inflammatory bowel disease, cancer, obesity, autoimmune diseases, and many other human disease states. An emerging paradigm in treatment is the administration of live engineered organisms, also called next-generation probiotics. However, the efficacy of these microbial therapies can be limited by the organism's overall performance in the harsh and nutrient-limited environment of the gut. In this review, we summarize the current state of the art use of bacterial and yeast strains as probiotics, highlight the recent development of genetic tools for engineering new therapeutic functions in these organisms, and report on the latest therapeutic applications of engineered probiotics, including recent clinical trials. We also discuss the supplementation of prebiotics as a method of manipulating the microbiome and improving the overall performance of engineered live biotherapeutics.

Genetic Engineering of Resident Bacteria in the Gut Microbiome

Techniques by which to genetically manipulate members of the microbiota enable both the evaluation of host-microbe interactions and an avenue by which to monitor and modulate human physiology. Genetic engineering applications have traditionally focused on model gut residents, such as Escherichia coli and lactic acid bacteria. However, emerging efforts by which to develop synthetic biology toolsets for "nonmodel" resident gut microbes could provide an improved foundation for microbiome engineering. As genome engineering tools come online, so too have novel applications for engineered gut microbes. Engineered resident gut bacteria facilitate investigations of the roles of microbes and their metabolites on host health and allow for potential live microbial biotherapeutics. Due to the rapid pace of discovery in this burgeoning field, this minireview highlights advancements in the genetic engineering of all resident gut microbes.

SURE editing: combining oligo-recombineering and programmable insertion/deletion of selection markers to efficiently edit the Mycoplasma pneumoniae genome

The development of advanced genetic tools is boosting microbial engineering which can potentially tackle wide-ranging challenges currently faced by our society. Here we present SURE editing, a multi-recombinase engineering rationale combining oligonucleotide recombineering with the selective capacity of antibiotic resistance via transient insertion of selector plasmids. We test this method in Mycoplasma pneumoniae, a bacterium with a very inefficient native recombination machinery. Using SURE editing, we can seamlessly generate, in a single step, a wide variety of genome modifications at high efficiencies, including the largest possible deletion of this genome (30 Kb) and the targeted complementation of essential genes in the deletion of a region of interest. Additional steps can be taken to remove the selector plasmid from the edited area, to obtain markerless or even scarless edits. Of note, SURE editing is compatible with different site-specific recombinases for mediating transient plasmid integration. This battery of selector plasmids can be used to select different edits, regardless of the target sequence, which significantly reduces the cloning load associated to genome engineering projects. Given the proven functionality in several microorganisms of the machinery behind the SURE editing logic, this method is likely to represent a valuable advance for the synthetic biology field.

Delineation of the Ancestral Tus-Dependent Replication Fork Trap

In Escherichia coli, DNA replication termination is orchestrated by two clusters of Ter sites forming a DNA replication fork trap when bound by Tus proteins. The formation of a ‘locked’ Tus–Ter complex is essential for halting incoming DNA replication forks. However, the absence of replication fork arrest at some Ter sites raised questions about their significance. In this study, we examined the genome-wide distribution of Tus and found that only the six innermost Ter sites (TerA–E and G) were significantly bound by Tus. We also found that a single ectopic insertion of TerB in its non-permissive orientation could not be achieved, advocating against a need for ‘back-up’ Ter sites. Finally, examination of the genomes of a variety of Enterobacterales revealed a new replication fork trap architecture mostly found outside the Enterobacteriaceae family. Taken together, our data enabled the delineation of a narrow ancestral Tus-dependent DNA replication fork trap consisting of only two Ter sites.

Prokaryotic reverse transcriptases: from retroelements to specialized defense systems

Reverse transcriptases (RTs) catalyze the polymerization of DNA from an RNA template. These enzymes were first discovered in RNA tumor viruses in 1970, but it was not until 1989 that they were found in prokaryotes as a key component of retrons. Apart from RTs encoded by the 'selfish' mobile retroelements known as group II introns, prokaryotic RTs are extraordinarily diverse, but their function has remained elusive. However, recent studies have revealed that different lineages of prokaryotic RTs, including retrons, those associated with CRISPR-Cas systems, Abi-like RTs and other yet uncharacterized RTs, are key components of different lines of defense against phages and other mobile genetic elements. Prokaryotic RTs participate in various antiviral strategies, including abortive infection (Abi), in which the infected cell is induced to commit suicide to protect the host population, adaptive immunity, in which a memory of previous infection is used to build an efficient defense, and other as yet unidentified mechanisms. These prokaryotic enzymes are attracting considerable attention, both for use in cutting-edge technologies, such as genome editing, and as an emerging research topic. In this review, we discuss what is known about prokaryotic RTs, and the exciting evidence for their domestication from retroelements to create specialized defense systems.

Multiplexed characterization of rationally designed promoter architectures deconstructs combinatorial logic for IPTG-inducible systems

A crucial step towards engineering biological systems is the ability to precisely tune the genetic response to environmental stimuli. In the case of Escherichia coli inducible promoters, our incomplete understanding of the relationship between sequence composition and gene expression hinders our ability to predictably control transcriptional responses. Here, we profile the expression dynamics of 8269 rationally designed, IPTG-inducible promoters that collectively explore the individual and combinatorial effects of RNA polymerase and LacI repressor binding site strengths. We then fit a statistical mechanics model to measured expression that accurately models gene expression and reveals properties of theoretically optimal inducible promoters. Furthermore, we characterize three alternative promoter architectures and show that repositioning binding sites within promoters influences the types of combinatorial effects observed between promoter elements. In total, this approach enables us to deconstruct relationships between inducible promoter elements and discover practical insights for engineering inducible promoters with desirable characteristics.

Engineering S. oneidensis for Performance Improvement of Microbial Fuel Cell-a Mini Review

Microbial fuel cell (MFC) is a promising technology that utilizes exoelectrogens cultivated in the form of biofilm to generate power from various types of sources supplied. A metal-reducing pathway is utilized by these organisms to transfer electrons obtained from the metabolism of substrate from anaerobic respiration extracellularly. A widely established model organism that is capable of extracellular electron transfer (EET) is Shewanella oneidensis. This review highlights the strategies used in the transformation of S. oneidensis and the recent development of MFC in terms of intervention through genetic modifications. S. oneidensis was genetically engineered for several aims including the study on the underlying mechanisms of EET, and the enhancement of power generation and wastewater treating potential when used in an MFC. Through engineering S. oneidensis, genes responsible for EET are identified and strategies on enhancing the EET efficiency are studied. Overexpressing genes related to EET to enhance biofilm formation, mediator biosynthesis, and respiration appears as one of the common approaches.

Precision design of stable genetic circuits carried in highly-insulated E. coli genomic landing pads

Genetic circuits have many applications, from guiding living therapeutics to ordering process in a bioreactor, but to be useful they have to be genetically stable and not hinder the host. Encoding circuits in the genome reduces burden, but this decreases performance and can interfere with native transcription. We have designed genomic landing pads in Escherichia coli at high-expression sites, flanked by ultrastrong double terminators. DNA payloads >8 kb are targeted to the landing pads using phage integrases. One landing pad is dedicated to carrying a sensor array, and two are used to carry genetic circuits. NOT/NOR gates based on repressors are optimized for the genome and characterized in the landing pads. These data are used, in conjunction with design automation software (Cello 2.0), to design circuits that perform quantitatively as predicted. These circuits require fourfold less RNA polymerase than when carried on a plasmid and are stable for weeks in a recA+ strain without selection. This approach enables the design of synthetic regulatory networks to guide cells in environments or for applications where plasmid use is infeasible.

A double-locus scarless genome editing system in Escherichia coli

OBJECTIVE

To develop a convenient double-locus scarless genome editing system in Escherichia coli, based on the type II Streptococcus pyogenes CRISPR/Cas9 and λ Red recombination cassette.

RESULTS

A two-plasmid genome editing system was constructed. The large-sized plasmid harbors the cas9 and λ Red recombination genes (gam, bet, and exo), while the small-molecular plasmid can simultaneously express two different gRNAs (targeting genome RNAs). The recombination efficiency was tested by targeting the galK, lacZ, and dbpA genes in E. coli with ssDNA or dsDNA. Resulting concurrent double-locus recombination efficiencies were 88 ± 5.5% (point mutation), 39.7 ± 4.3% (deletion/insertion), and 57.8 ± 3.4%-58.5 ± 4.1% (mixed point and deletion/insertion mutation), depending on 30 (ssDNA) or 40 bp (dsDNA) homologous side arms employed. In addition, the curing efficiency of the guide plasmid expressing gRNAs for negative selection was higher (96 ± 3% in 4 h) than the help plasmid carrying cas9 and λ Red (92 ± 2% in 9 h).

CONCLUSIONS

The new editing system is convenient and efficient for simultaneous double-locus recombination in the genome and should be favorable for high-throughput multiplex genome editing in synthetic biology and metabolic engineering.

Reconstitution of TCA cycle involving l-isoleucine dioxygenase for hydroxylation of l-isoleucine in Escherichia coli using CRISPR-Cas9

l-isoleucine dioxygenase (IDO) is an Fe (II)/α-ketoglutarate (α-KG)-dependent dioxygenase that specifically converts l-isoleucine (l-Ile) to (2S, 3R, 4S)-4-hydroxyisoleucine (4-HIL). 4-HIL is an important drug for the treatment and prevention of type 1 and type 2 diabetes but the yields using current methods are low. In this study, the CRISPR-Cas9 gene editing system was used to knockout sucAB and aceAK gene in the TCA cycle pathway of Escherichia coli (E. coli). For single-gene knockout, the whole process took approximately 7 days. However, the manipulation time was reduced by 2 days for each round of gene modification for multigene editing. Using the genome-edited recombinant strain E. coli BL21(DE3) ΔsucABΔaceAK/pET-28a(+)-ido (2Δ-ido), the bioconversion ratio of L-Ile to 4-HIL was enhanced by about 15% compared to E. coli BL21(DE3)/pET-28a(+)-ido [BL21(DE3)-ido] strain. The CRISPR-Cas9 editing strategy has the potential in modifying multiple genes more rapidly and in optimizing strains for industrial production.

A Targetron-Recombinase System for Large-Scale Genome Engineering of Clostridia

Clostridia are a group of Gram-positive anaerobic bacteria of medical and industrial importance for which limited genetic methods are available. Here, we demonstrate an approach to make large genomic deletions and insertions in the model Clostridium phytofermentans by combining designed group II introns (targetrons) and Cre recombinase. We apply these methods to delete a 50-gene prophage island by programming targetrons to position markerless lox66 and lox71 sites, which mediate deletion of the intervening 39-kb DNA region using Cre recombinase. Gene expression and growth of the deletion strain showed that the prophage genes contribute to fitness on nonpreferred carbon sources. We also inserted an inducible fluorescent reporter gene into a neutral genomic site by recombination-mediated cassette exchange (RMCE) between genomic and plasmid-based tandem lox sites bearing heterospecific spacers to prevent intracassette recombination. These approaches generally enable facile markerless genome engineering in clostridia to study their genome structure and regulation.IMPORTANCE Clostridia are anaerobic bacteria with important roles in intestinal and soil microbiomes. The inability to experimentally modify the genomes of clostridia has limited their study and application in biotechnology. Here, we developed a targetron-recombinase system to efficiently make large targeted genomic deletions and insertions using the model Clostridium phytofermentans We applied this approach to reveal the importance of a prophage to host fitness and introduce an inducible reporter by recombination-mediated cassette exchange.

CRISPR/Cas9/sgRNA-mediated targeted gene modification confirms the cause-effect relationship between gyrA mutation and quinolone resistance in Escherichia coli

Quinolones are broad-spectrum antibiotics that have been used for decades in treating bacterial infections in humans and animals, and subsequently bacterial resistance to these agents has increased. While studies indicated the relationship between gyrA mutations and bacterial resistance to quinolones, CRISPR/Cas9 was used in this study to investigate causal role of gyrA mutation in the quinolone resistance. In this study, 818 clinical Escherichia coli isolates were analyzed for gyrA mutations and their resistance to quinolones. The CRISPR/Cas9 system was used to generate gyrA mutations in quinolone-susceptible E. coli ATCC 25922, and quinolone-resistant clinical E. coli. The antimicrobial resistance prevalence rate in E. coli against nalidixic acid, ciprofloxacin and enrofloxacin was 77.1% (631/818), 51.1% (418/818) and 49.8% (407/818), respectively. The gyrA mutations were identified in nucleotide positions 248, 255, 259, 260, 261, 273 and 300, and mutations at positions 248 and 259 resulting in amino acid changes at positions 83 and 87 were associated with quinolone resistance. Double-site amino acid mutations increase resistance to quinolones. The gyrA mutations causing changes at amino acids 83 and 87 reversed the features of quinolone resistance in ATCC and clinical strains, verifying the causal role of gyrA mutation in the quinolone resistance of E. coli.

TargeTron Technology Applicable in Solventogenic Clostridia: Revisiting 12 Years' Advances

Clostridium has great potential in industrial application and medical research. But low DNA repair capacity and plasmids transformation efficiency severely delay development and application of genetic tools based on homologous recombination (HR). TargeTron is a gene editing technique dependent on the mobility of group II introns, rather than homologous recombination, which makes it very suitable for gene disruption of Clostridium. The application of TargeTron technology in solventogenic Clostridium is academically reported in 2007 and this tool has been introduced in various clostridia as it is easy to operate, time saving, and reliable. TargeTron has made great progress in solventogenic Clostridium in the aspects of acetone-butanol-ethanol (ABE) fermentation pathway modification, important functional genes identification, and xylose metabolic pathway analysis and reconstruction. In the review, 12 years' advances of TargeTron technology applicable in solventogenic Clostridium, including its principle, technical characteristics, application, and efforts to expand its capabilities, or to avoid potential drawbacks, are revisisted. Some other technologies as putative competitors or collaborators are also discussed. It is believed that TargeTron combined with CRISPR/Cas-assisted gene/base editing and gene-expression regulation system will make a better future for clostridial genetic modification.

Construction of a novel Escherichia coli expression system: relocation of lpxA from chromosome to a constitutive expression vector

The selective marker in the plasmid-based expression system is usually a gene that encodes an antibiotic-resistant protein; therefore, the antibiotic has to add to maintain the plasmid when growing the bacteria. This antibiotic addition would lead to increase of production cost and the environment contamination. In this study, a novel Escherichia coli expression system, the lpxA deletion mutant harboring an lpxA-carrying vector, was developed. To develop this system, three plasmids pCas9Cre, pTF-A-UD, and pRSFCmlpxA were constructed. The plasmid pCas9Cre produces enzymes Cas9, λ-Red, and Cre and can be cured by growing at 42 °C; pTF-A-UD contains several DNA fragments required for deleting the chromosomal lpxA and can be cured by adding isopropyl-D-thiogalactopyranoside; pRSFCmlpxA contains the lpxA mutant lpxA123 and CamR. When E. coli were transformed with these three plasmids, the chromosomal lpxA and the CamR in pRSFCmlpxA can be efficiently removed, resulting in an E. coli lpxA mutant harboring pRSFlpxA. The lpxA is essential for the growth of E. coli; its relocation from chromosome to a constitutive expression vector is an ideal strategy to maintain the vector without antibiotic addition. The lpxA123 in pRSFlpxA can complement the deletion of the chromosomal lpxA and provide a strong selective pressure to maintain the plasmid pRSFlpxA. This study provides an experimental evidence that this novel expression system is convenient and efficient to use and can be used to improve L-threonine biosynthesis in the wild type E. coli MG1655 and an L-threonine producing E. coli TWF006.

Synthetic evolution

The combination of modern biotechnologies such as DNA synthesis, λ red recombineering, CRISPR-based editing and next-generation high-throughput sequencing increasingly enables precise manipulation of genes and genomes. Beyond rational design, these technologies also enable the targeted, and potentially continuous, introduction of multiple mutations. While this might seem to be merely a return to natural selection, the ability to target evolution greatly reduces fitness burdens and focuses mutation and selection on those genes and traits that best contribute to a desired phenotype, ultimately throwing evolution into fast forward.

Group II Intron RNPs and Reverse Transcriptases: From Retroelements to Research Tools

Group II introns, self-splicing retrotransposons, serve as both targets of investigation into their structure, splicing, and retromobility and a source of tools for genome editing and RNA analysis. Here, we describe the first cryo-electron microscopy (cryo-EM) structure determination, at 3.8-4.5 Å, of a group II intron ribozyme complexed with its encoded protein, containing a reverse transcriptase (RT), required for RNA splicing and retromobility. We also describe a method called RIG-seq using a retrotransposon indicator gene for high-throughput integration profiling of group II introns and other retrotransposons. Targetrons, RNA-guided gene targeting agents widely used for bacterial genome engineering, are described next. Finally, we detail thermostable group II intron RTs, which synthesize cDNAs with high accuracy and processivity, for use in various RNA-seq applications and relate their properties to a 3.0-Å crystal structure of the protein poised for reverse transcription. Biological insights from these group II intron revelations are discussed.

Insertional Mutagenesis Protocol for Constructing Single or Sequential Mutations

Genetic mutation enables the study of the function of specific genes, particularly when a mutant is compared against its isogenic parent. In Proteus mirabilis bacteria, traditional allelic exchange mutation is labor-intensive and has a high failure rate in some strains. Likewise, there is no working protocol for lambda red recombinase-based mutation in P. mirabilis. Here we describe an alternative method of insertional mutagenesis based on retargeting of group II introns. The protocol includes steps to generate single or multiple mutations, with the possibility to delete intervening sequences of DNA.

Systematic Dissection of Sequence Elements Controlling σ70 Promoters Using a Genomically Encoded Multiplexed Reporter Assay in Escherichia coli

Promoters are the key drivers of gene expression and are largely responsible for the regulation of cellular responses to time and environment. In Escherichia coli, decades of studies have revealed most, if not all, of the sequence elements necessary to encode promoter function. Despite our knowledge of these motifs, it is still not possible to predict the strength and regulation of a promoter from primary sequence alone. Here we develop a novel multiplexed assay to study promoter function in E. coli by building a site-specific genomic recombination-mediated cassette exchange system that allows for the facile construction and testing of large libraries of genetic designs integrated into precise genomic locations. We build and test a library of 10898 σ70 promoter variants consisting of all combinations of a set of eight -35 elements, eight -10 elements, three UP elements, eight spacers, and eight backgrounds. We find that the -35 and -10 sequence elements can explain approximately 74% of the variance in promoter strength within our data set using a simple log-linear statistical model. Simple neural network models explain >95% of the variance in our data set by capturing nonlinear interactions with the spacer, background, and UP elements.

Floxed-Cassette Allelic Exchange Mutagenesis Enables Markerless Gene Deletion in Chlamydia trachomatis and Can Reverse Cassette-Induced Polar Effects

As obligate intracellular bacteria, Chlamydia spp. have evolved numerous, likely intricate, mechanisms to create and maintain a privileged intracellular niche. Recent progress in elucidating and characterizing these processes has been bolstered by the development of techniques enabling basic genetic tractability. Florescence-reported allelic exchange mutagenesis (FRAEM) couples chromosomal gene deletion with the insertion of a selection cassette encoding antibiotic resistance and green fluorescent protein (GFP). Similar to other bacteria, many chlamydial genes exist within polycistronic operons, raising the possibility of polar effects mediated by insertion cassettes. Indeed, FRAEM-mediated deletion of Chlamydia trachomatis tmeA negatively impacts the expression of tmeB We have adapted FRAEM technology by employing a gfp-bla cassette flanked by loxP sites. Conditional expression of Cre recombinase in Chlamydia tmeA containing a floxed cassette resulted in deletion of the marker and restoration of tmeB expression.IMPORTANCE C. trachomatis infections represent a significant burden to human health. The ability to genetically manipulate Chlamydia spp. is overcoming historic confounding barriers that have impeded rapid progress in understanding overall chlamydial pathogenesis. The current state of genetic manipulation in Chlamydia spp. requires further development, including mechanisms to generate markerless gene disruption. We leveraged a stepwise Cre-lox approach to excise selection marker genes from a deleted gene locus. We found this process to be efficient, and the removal of extraneous elements resulted in the reversal of a negative polar effect on a downstream gene. This technique facilitates a more direct assessment of gene function and adds to the Chlamydia molecular toolbox by facilitating the deletion of genes within operons.

A Multiplex Genome Editing Method for Escherichia coli Based on CRISPR-Cas12a

Various methods for editing specific sites in the Escherichia coli chromosome are available, and gene-size (∼1 kb) integration into a single site or to introduce deletions, short insertions or point mutations into multiple sites can be conducted in a short period of time. However, a method for rapidly integrating multiple gene-size sequences into different sites has not been developed yet. Here, we describe a method and plasmid system that makes it possible to simultaneously insert genes into multiple specific loci of the E. coli genome without the need for chromosomal markers. The method uses a CRISPR-Cas12a system to eliminate unmodified cells by double-stranded DNA cleavage in conjunction with the phage-derived λ-Red recombinases to facilitate recombination between the chromosome and the donor DNA. We achieved the insertion of up to 3 heterologous genes in one round of recombination and selection. To demonstrate the practical application of this gene-insertion method, we constructed a recombinant E. coli producing an industrially useful chemical, 5-aminolevulinic acid (ALA), with high-yield. Moreover, a similar two-plasmid system was built to edit the genome of the extremophile Halomonas bluephagenesis.

Genetic Engineering of Bee Gut Microbiome Bacteria with a Toolkit for Modular Assembly of Broad-Host-Range Plasmids

Engineering the bacteria present in animal microbiomes promises to lead to breakthroughs in medicine and agriculture, but progress is hampered by a dearth of tools for genetically modifying the diverse species that comprise these communities. Here we present a toolkit of genetic parts for the modular construction of broad-host-range plasmids built around the RSF1010 replicon. Golden Gate assembly of parts in this toolkit can be used to rapidly test various antibiotic resistance markers, promoters, fluorescent reporters, and other coding sequences in newly isolated bacteria. We demonstrate the utility of this toolkit in multiple species of Proteobacteria that are native to the gut microbiomes of honey bees ( Apis mellifera) and bumble bees (B ombus sp.). Expressing fluorescent proteins in Snodgrassella alvi, Gilliamella apicola, Bartonella apis, and Serratia strains enables us to visualize how these bacteria colonize the bee gut. We also demonstrate CRISPRi repression in B. apis and use Cas9-facilitated knockout of an S. alvi adhesion gene to show that it is important for colonization of the gut. Beyond characterizing how the gut microbiome influences the health of these prominent pollinators, this bee microbiome toolkit (BTK) will be useful for engineering bacteria found in other natural microbial communities.

Synthetic Biology to Improve the Production of Lipases and Esterases (Review)

Synthetic biology is an emergent field of research whose aim is to make biology an engineering discipline, thus permitting to design, control, and standardize biological processes. Synthetic biology is therefore expected to boost the development of biotechnological processes such as protein production and enzyme engineering, which can be significantly relevant for lipases and esterases.

Managing the SOS Response for Enhanced CRISPR-Cas-Based Recombineering in E. coli through Transient Inhibition of Host RecA Activity

Phage-derived "recombineering" methods are utilized for bacterial genome editing. Recombineering results in a heterogeneous population of modified and unmodified chromosomes, and therefore selection methods, such as CRISPR-Cas9, are required to select for edited clones. Cells can evade CRISPR-Cas-induced cell death through recA-mediated induction of the SOS response. The SOS response increases RecA dependent repair as well as mutation rates through induction of the umuDC error prone polymerase. As a result, CRISPR-Cas selection is more efficient in recA mutants. We report an approach to inhibiting the SOS response and RecA activity through the expression of a mutant dominant negative form of RecA, which incorporates into wild type RecA filaments and inhibits activity. Using a plasmid-based system in which Cas9 and recA mutants are coexpressed, we can achieve increased efficiency and consistency of CRISPR-Cas9-mediated selection and recombineering in E. coli, while reducing the induction of the SOS response. To date, this approach has been shown to be independent of recA genotype and host strain lineage. Using this system, we demonstrate increased CRISPR-Cas selection efficacy with over 10 000 guides covering the E. coli chromosome. The use of dominant negative RecA or homologues may be of broad use in bacterial CRISPR-Cas-based genome editing where the SOS pathways are present.

A novel and versatile dual fluorescent reporter tool for the study of gene expression and regulation in multi- and single copy number

To unravel intricate mechanisms of gene regulation it is imperative to work in physiologically relevant conditions and therefore preferentially in single copy constructs, which are not always easy to manipulate. Such in vivo studies are generally based on enzymatic assays, microarrays, RNA-seq, qRT-PCR, or multicopy reporter gene systems, frequently with β-galactosidase, luciferase or a fluorescent protein as reporter. Each method has its advantages and shortcomings and may require validation. Enzyme assays are generally reliable but may be quite complex, time consuming, and require a (expensive) substrate. Microarrays and RNA-seq provide a genome wide view of gene expression but may rapidly become expensive and time consuming especially for detailed studies with large numbers of mutants, different growth conditions and multiple time points. Multicopy reporter gene systems are handy to generate numerous constructs but may not provide accurate information due to titration effects of trans-acting regulatory elements. Therefore and in spite of the existence of various reporter systems, there is still need for an efficient and user-friendly tool for detailed studies and high throughput screenings. Here we develop and validate a novel and versatile fluorescent reporter tool to study gene regulation in single copy mode that enables real-time measurement. This tool bears two independent fluorescent reporters that allow high throughput screening and standardization, and combines modern efficient cloning methods (multicopy, in vitro manipulation) with classical genetics (in vivo homologous recombination with a stable, self-transmissible episome) to generate multi- and single copy reporter systems. We validate the system with constitutive and differentially regulated promoters and show that the tool can equally be used with heterologous transcription factors. The flexibility and versatility of this dual reporter tool in combination with an easy conversion from a multicopy plasmid to a stable, single copy reporter system makes this system unique and attractive for a variety of applications. Examples are in vivo studies of DNA-binding transcription factors (single copy) or screening of promoter and RBS libraries (multicopy) for synthetic biology purposes.

Construction of an easy-to-use CRISPR-Cas9 system by patching a newly designed EXIT circuit

Background

Plasmid-borne genetic editing tools, including the widely used CRISPR-Cas9 system, have greatly facilitated bacterial programming to obtain novel functionalities. However, the lack of effective post-editing plasmid elimination methods impedes follow-up genetic manipulation or application. Conventional strategies including exposure to physical and chemical treatments, or exploiting temperature-sensitive replication origins have several drawbacks (e.g., they are limited for efficiency and are time-consuming). Therefore, the demand is apparent for easy and rapid elimination of the tool plasmids from their bacterial hosts after genetic manipulation.

Results

To bridge this gap, we designed a novel EXIT circuit with the homing endonuclease, which can be exploited for rapid and efficient elimination of various plasmids with diverse replication origins. As a proof of concept, we validated the EXIT circuit in Escherichia coli by harnessing homing endonuclease I-SceI and its cleavage site. When integrated into multiple plasmids with different origins, the EXIT circuit allowed them to be eliminated from the host cells, simultaneously. By combining the widely used plasmid-borne CRISPR-Cas9 system and the EXIT circuit, we constructed an easy-to-use CRISPR-Cas9 system that eliminated the Cas9- and the single-guide RNA (sgRNA)-encoding plasmids in one-step. Within 3 days, we successfully constructed an atrazine-degrading E. coli strain, thus further demonstrating the advantage of this new CRISPR-Cas9 system for bacterial genome editing.

Conclusions

Our novel EXIT circuit, which exploits the homing endonuclease I-SceI, enables plasmid(s) with different replication origins to be eliminated from their host cells rapidly and efficiently. We also developed an easy-to-use CRISPR-Cas9 system with the EXIT circuit, and this new system can be widely applied to bacterial genome editing.

Electronic supplementary material

The online version of this article (doi:10.1186/s13036-017-0072-5) contains supplementary material, which is available to authorized users.

An efficient system for deletion of large DNA fragments in Escherichia coli via introduction of both Cas9 and the non-homologous end joining system from Mycobacterium smegmatis

Accompanied with the internal non-homologous end joining (NHEJ) system, Cas9 can be used to easily inactivate a gene or delete a fragment through introduction of DNA double-stranded breaks (DSBs) in eukaryotic cells. While in most prokaryotes (e.g. Escherichia coli), due to the lack of NHEJ, homologous recombination (HR) is required for repair of DSBs, which is less convenient. Here, a markerless system was developed for rapid gene inactivation or fragment deletion in E. coli via introduction of both Cas9 and a bacterial NHEJ system. Three bacterial NHEJ systems, i.e. Mycobacterium smegmatis (Msm), Mycobacterium tuberculosis (Mtb) and Bacillus subtilis (Bs), were tested in E. coli, and the MsmNHEJ system showed the best efficiency. With the employment of Cas9 and MsmNHEJ, we efficiently mutated lacZ gene, deleted glnALG operon and two large DNA fragments (67 kb and 123 kb) in E. coli, respectively. Moreover, the system was further designed to allow for continuous inactivation of genes or deletion of DNA fragments in E. coli. We envision this system can be extended to other bacteria, especially those with low HR efficiency.

Use of Group II Intron Technology for Targeted Mutagenesis in Chlamydia trachomatis

Dissecting the contribution of genes to virulence in fulfillment of Molecular Koch's postulates is essential for developing prevention and treatment strategies for bacterial pathogens. This chapter will discuss the application of a targeted, intron-based insertional mutagenesis method for creating mutants in the obligate, intracellular bacterial pathogen Chlamydia trachomatis. The methods employed for intron targeting, mutant selection, and mutant verification will be outlined including available selection markers, gene targeting strategies, and potential pitfalls.

Bacteriophage-based tools: recent advances and novel applications

Bacteriophages (phages) are viruses that infect bacterial hosts, and since their discovery over a century ago they have been primarily exploited to control bacterial populations and to serve as tools in molecular biology. In this commentary, we highlight recent diverse advances in the field of phage research, going beyond bacterial control using whole phage, to areas including biocontrol using phage-derived enzybiotics, diagnostics, drug discovery, novel drug delivery systems and bionanotechnology.

Novel Technologies for Optimal Strain Breeding

The implementation of a knowledge-based bioeconomy requires the rapid development of highly efficient microbial production strains that are able to convert renewable carbon sources to value-added products, such as bulk and fine chemicals, pharmaceuticals, or proteins at industrial scale. Starting from classical strain breeding by random mutagenesis and screening in the 1950s via rational design by metabolic engineering initiated in the 1970s, a range of powerful new technologies have been developed in the past two decades that can revolutionize future strain engineering. In particular, next-generation sequencing technologies combined with new methods of genome engineering and high-throughput screening based on genetically encoded biosensors have allowed for new concepts. In this chapter, selected new technologies relevant for breeding microbial production strains with a special emphasis on amino acid producers will be summarized.

Bacterial genome engineering and synthetic biology: combating pathogens

BACKGROUND

The emergence and prevalence of multidrug resistant (MDR) pathogenic bacteria poses a serious threat to human and animal health globally. Nosocomial infections and common ailments such as pneumonia, wound, urinary tract, and bloodstream infections are becoming more challenging to treat due to the rapid spread of MDR pathogenic bacteria. According to recent reports by the World Health Organization (WHO) and Centers for Disease Control and Prevention (CDC), there is an unprecedented increase in the occurrence of MDR infections worldwide. The rise in these infections has generated an economic strain worldwide, prompting the WHO to endorse a global action plan to improve awareness and understanding of antimicrobial resistance. This health crisis necessitates an immediate action to target the underlying mechanisms of drug resistance in bacteria.

RESEARCH

The advent of new bacterial genome engineering and synthetic biology (SB) tools is providing promising diagnostic and treatment plans to monitor and treat widespread recalcitrant bacterial infections. Key advances in genetic engineering approaches can successfully aid in targeting and editing pathogenic bacterial genomes for understanding and mitigating drug resistance mechanisms. In this review, we discuss the application of specific genome engineering and SB methods such as recombineering, clustered regularly interspaced short palindromic repeats (CRISPR), and bacterial cell-cell signaling mechanisms for pathogen targeting. The utility of these tools in developing antibacterial strategies such as novel antibiotic production, phage therapy, diagnostics and vaccine production to name a few, are also highlighted.

CONCLUSIONS

The prevalent use of antibiotics and the spread of MDR bacteria raise the prospect of a post-antibiotic era, which underscores the need for developing novel therapeutics to target MDR pathogens. The development of enabling SB technologies offers promising solutions to deliver safe and effective antibacterial therapies.

Phage Therapy in the Era of Synthetic Biology

For more than a century, bacteriophage (or phage) research has enabled some of the most important discoveries in biological sciences and has equipped scientists with many of the molecular biology tools that have advanced our understanding of replication, maintenance, and expression of genetic material. Phages have also been recognized and exploited as natural antimicrobial agents and nanovectors for gene therapy, but their potential as therapeutics has not been fully exploited in Western medicine because of challenges such as narrow host range, bacterial resistance, and unique pharmacokinetics. However, increasing concern related to the emergence of bacteria resistant to multiple antibiotics has heightened interest in phage therapy and the development of strategies to overcome hurdles associated with bacteriophage therapeutics. Recent progress in sequencing technologies, DNA manipulation, and synthetic biology allowed scientists to refactor the entire bacterial genome of Mycoplasma mycoides, thereby creating the first synthetic cell. These new strategies for engineering genomes may have the potential to accelerate the construction of designer phage genomes with superior therapeutic potential. Here, we discuss the use of phage as therapeutics, as well as how synthetic biology can create bacteriophage with desirable attributes.

Rapid and Efficient One-Step Metabolic Pathway Integration in E. coli

Methods for importing heterologous genes into genetically tractable hosts are among the most desired tools of synthetic biology. Easy plug-and-play construction methods to rapidly test genes and pathways stably in the host genome would expedite synthetic biology and metabolic engineering applications. Here, we describe a CRISPR-based strategy that allows highly efficient, single step integration of large pathways in Escherichia coli. This strategy allows high efficiency integration in a broad range of homology arm sizes and genomic positions, with efficiencies ranging from 70 to 100% in 7 distinct loci. To demonstrate the large size capability, we integrated a 10 kb construct to implement isobutanol production in a single day. The ability to efficiently integrate entire metabolic pathways in a rapid and markerless manner will facilitate testing and engineering of novel pathways using the E. coli genome as a stable testing platform.

An Overview of Tyrosine Site-specific Recombination: From an Flp Perspective

Tyrosine site-specific recombinases (YRs) are widely distributed among prokaryotes and their viruses, and were thought to be confined to the budding yeast lineage among eukaryotes. However, YR-harboring retrotransposons (the DIRS and PAT families) and DNA transposons (Cryptons) have been identified in a variety of eukaryotes. The YRs utilize a common chemical mechanism, analogous to that of type IB topoisomerases, to bring about a plethora of genetic rearrangements with important physiological consequences in their respective biological contexts. A subset of the tyrosine recombinases has provided model systems for analyzing the chemical mechanisms and conformational features of the recombination reaction using chemical, biochemical, topological, structural, and single molecule-biophysical approaches. YRs with simple reaction requirements have been utilized to bring about programmed DNA rearrangements for addressing fundamental questions in developmental biology. They have also been employed to trace the topological features of DNA within high-order DNA interactions established by protein machines. The directed evolution of altered specificity YRs, combined with their spatially and temporally regulated expression, heralds their emergence as vital tools in genome engineering projects with wide-ranging biotechnological and medical applications.

Emancipating Chlamydia: Advances in the Genetic Manipulation of a Recalcitrant Intracellular Pathogen

Chlamydia species infect millions of individuals worldwide and are important etiological agents of sexually transmitted disease, infertility, and blinding trachoma. Historically, the genetic intractability of this intracellular pathogen has hindered the molecular dissection of virulence factors contributing to its pathogenesis. The obligate intracellular life cycle of Chlamydia and restrictions on the use of antibiotics as selectable markers have impeded the development of molecular tools to genetically manipulate these pathogens. However, recent developments in the field have resulted in significant gains in our ability to alter the genome of Chlamydia, which will expedite the elucidation of virulence mechanisms. In this review, we discuss the challenges affecting the development of molecular genetic tools for Chlamydia and the work that laid the foundation for recent advancements in the genetic analysis of this recalcitrant pathogen.

Site-specific recombinases: molecular machines for the Genetic Revolution

The fields of molecular genetics, biotechnology and synthetic biology are demanding ever more sophisticated molecular tools for programmed precise modification of cell genomic DNA and other DNA sequences. This review presents the current state of knowledge and development of one important group of DNA-modifying enzymes, the site-specific recombinases (SSRs). SSRs are Nature's 'molecular machines' for cut-and-paste editing of DNA molecules by inserting, deleting or inverting precisely defined DNA segments. We survey the SSRs that have been put to use, and the types of applications for which they are suitable. We also discuss problems associated with uses of SSRs, how these problems can be minimized, and how recombinases are being re-engineered for improved performance and novel applications.

CRMAGE: CRISPR Optimized MAGE Recombineering

A bottleneck in metabolic engineering and systems biology approaches is the lack of efficient genome engineering technologies. Here, we combine CRISPR/Cas9 and λ Red recombineering based MAGE technology (CRMAGE) to create a highly efficient and fast method for genome engineering of Escherichia coli. Using CRMAGE, the recombineering efficiency was between 96.5% and 99.7% for gene recoding of three genomic targets, compared to between 0.68% and 5.4% using traditional recombineering. For modulation of protein synthesis (small insertion/RBS substitution) the efficiency was increased from 6% to 70%. CRMAGE can be multiplexed and enables introduction of at least two mutations in a single round of recombineering with similar efficiencies. PAM-independent loci were targeted using degenerate codons, thereby making it possible to modify any site in the genome. CRMAGE is based on two plasmids that are assembled by a USER-cloning approach enabling quick and cost efficient gRNA replacement. CRMAGE furthermore utilizes CRISPR/Cas9 for efficient plasmid curing, thereby enabling multiple engineering rounds per day. To facilitate the design process, a web-based tool was developed to predict both the λ Red oligos and the gRNAs. The CRMAGE platform enables highly efficient and fast genome editing and may open up promising prospective for automation of genome-scale engineering.

Targeted Large-Scale Deletion of Bacterial Genomes Using CRISPR-Nickases

Programmable CRISPR-Cas systems have augmented our ability to produce precise genome manipulations. Here we demonstrate and characterize the ability of CRISPR-Cas derived nickases to direct targeted recombination of both small and large genomic regions flanked by repetitive elements in Escherichia coli. While CRISPR directed double-stranded DNA breaks are highly lethal in many bacteria, we show that CRISPR-guided nickase systems can be programmed to make precise, nonlethal, single-stranded incisions in targeted genomic regions. This induces recombination events and leads to targeted deletion. We demonstrate that dual-targeted nicking enables deletion of 36 and 97 Kb of the genome. Furthermore, multiplex targeting enables deletion of 133 Kb, accounting for approximately 3% of the entire E. coli genome. This technology provides a framework for methods to manipulate bacterial genomes using CRISPR-nickase systems. We envision this system working synergistically with preexisting bacterial genome engineering methods.

Genomes by design

Next-generation DNA sequencing has revealed the complete genome sequences of numerous organisms, establishing a fundamental and growing understanding of genetic variation and phenotypic diversity. Engineering at the gene, network and whole-genome scale aims to introduce targeted genetic changes both to explore emergent phenotypes and to introduce new functionalities. Expansion of these approaches into massively parallel platforms establishes the ability to generate targeted genome modifications, elucidating causal links between genotype and phenotype, as well as the ability to design and reprogramme organisms. In this Review, we explore techniques and applications in genome engineering, outlining key advances and defining challenges.

Efficient Genome Editing in Clostridium cellulolyticum via CRISPR-Cas9 Nickase

The CRISPR-Cas9 system is a powerful and revolutionary genome-editing tool for eukaryotic genomes, but its use in bacterial genomes is very limited. Here, we investigated the use of the Streptococcus pyogenes CRISPR-Cas9 system in editing the genome of Clostridium cellulolyticum, a model microorganism for bioenergy research. Wild-type Cas9-induced double-strand breaks were lethal to C. cellulolyticum due to the minimal expression of nonhomologous end joining (NHEJ) components in this strain. To circumvent this lethality, Cas9 nickase was applied to develop a single-nick-triggered homologous recombination strategy, which allows precise one-step editing at intended genomic loci by transforming a single vector. This strategy has a high editing efficiency (>95%) even using short homologous arms (0.2 kb), is able to deliver foreign genes into the genome in a single step without a marker, enables precise editing even at two very similar target sites differing by two bases preceding the seed region, and has a very high target site density (median interval distance of 9 bp and 95.7% gene coverage in C. cellulolyticum). Together, these results establish a simple and robust methodology for genome editing in NHEJ-ineffective prokaryotes.

Mining Environmental Plasmids for Synthetic Biology Parts and Devices

The scientific and technical ambition of contemporary synthetic biology is the engineering of biological objects with a degree of predictability comparable to those made through electric and industrial manufacturing. To this end, biological parts with given specifications are sequence-edited, standardized, and combined into devices, which are assembled into complete systems. This goal, however, faces the customary context dependency of biological ingredients and their amenability to mutation. Biological orthogonality (i.e., the ability to run a function in a fashion minimally influenced by the host) is thus a desirable trait in any deeply engineered construct. Promiscuous conjugative plasmids found in environmental bacteria have evolved precisely to autonomously deploy their encoded activities in a variety of hosts, and thus they become excellent sources of basic building blocks for genetic and metabolic circuits. In this article we review a number of such reusable functions that originated in environmental plasmids and keep their properties and functional parameters in a variety of hosts. The properties encoded in the corresponding sequences include inter alia origins of replication, DNA transfer machineries, toxin-antitoxin systems, antibiotic selection markers, site-specific recombinases, effector-dependent transcriptional regulators (with their cognate promoters), and metabolic genes and operons. Several of these sequences have been standardized as BioBricks and/or as components of the SEVA (Standard European Vector Architecture) collection. Such formatting facilitates their physical composability, which is aimed at designing and deploying complex genetic constructs with new-to-nature properties.

Multigene editing in the Escherichia coli genome via the CRISPR-Cas9 system

An efficient genome-scale editing tool is required for construction of industrially useful microbes. We describe a targeted, continual multigene editing strategy that was applied to the Escherichia coli genome by using the Streptococcus pyogenes type II CRISPR-Cas9 system to realize a variety of precise genome modifications, including gene deletion and insertion, with a highest efficiency of 100%, which was able to achieve simultaneous multigene editing of up to three targets. The system also demonstrated successful targeted chromosomal deletions in Tatumella citrea, another species of the Enterobacteriaceae, with highest efficiency of 100%.