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Development of a Recombineering System for the Acetogen Eubacterium limosum with Cas9 Counterselection for Markerless Genome Engineering
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  • Patrick A. Sanford
    Patrick A. Sanford
    Department of Chemical Engineering, Northeastern University, 360 Huntington Avenue, 223 Cullinane, Boston, Massachusetts 02115, United States
    More by Patrick A. Sanford
  • Benjamin M. Woolston*
    Benjamin M. Woolston
    Department of Chemical Engineering, Northeastern University, 360 Huntington Avenue, 223 Cullinane, Boston, Massachusetts 02115, United States
    *Email: [email protected]
    More by Benjamin M. Woolston
    Orcidhttps://orcid.org/0000-0002-6570-2236
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ACS Synthetic Biology

Cite this: ACS Synth. Biol. 2024, 13, 8, 2505–2514
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https://doi.org/10.1021/acssynbio.4c00253
Published July 21, 2024

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Abstract

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Eubacterium limosum is a Clostridial acetogen that efficiently utilizes a wide range of single-carbon substrates and contributes to metabolism of health-associated compounds in the human gut microbiota. These traits have led to interest in developing it as a platform for sustainable CO2-based biofuel production to combat carbon emissions, and for exploring the importance of the microbiota in human health. However, synthetic biology and metabolic engineering in E. limosum have been hindered by the inability to rapidly make precise genomic modifications. Here, we screened a diverse library of recombinase proteins to develop a highly efficient oligonucleotide-based recombineering system based on the viral recombinase RecT. Following optimization, the system is capable of catalyzing ssDNA recombination at an efficiency of up to 2%. Addition of a Cas9 counterselection system eliminated unrecombined cells, with up to 100% of viable cells encoding the desired mutation, enabling creation of genomic point mutations in a scarless and markerless manner. We deployed this system to create a clean knockout of the extracellular polymeric substance (EPS) gene cluster, generating a strain incapable of biofilm formation. This approach is rapid and simple, not requiring laborious homology arm cloning, and can readily be retargeted to almost any genomic locus. This work overcomes a major bottleneck in E. limosum genetic engineering by enabling precise genomic modifications, and provides both a roadmap and associated recombinase plasmid library for developing similar systems in other Clostridia of interest.

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Introduction

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Eubacterium limosum is an acetogenic bacterium of growing interest thanks to its metabolic flexibility and role as a member of the human gut microbiota. (1,2) Acetogens are capable of metabolizing single carbon substrates such as carbon dioxide at high efficiency, converting them into products such as acetate and ethanol with a carbon yield of up to 95%, providing an attractive avenue for greenhouse gas sequestration. (3−8) E. limosum is particularly suited to such applications due to its ability to metabolize an expanded range of substrates including methanol and formate. (9) Additionally, as a member of the gut microbiota, it has been found to demethylate a range of dietary compounds of relevance to human health, including flavonoids and carnitine, making it an organism of interest for studying microbiota-related pathologies. (10,11) Among acetogens, E. limosum possesses a growing engineering toolbox including an analyzed genome, an expanded range of selection markers, and libraries of genetic elements, all of which have been used primarily in plasmid-based engineering applications. (5,12−15) Recently, more additions have been made to the organism’s genome engineering repertoire including a CRISPR/dCas9 system for gene silencing and a toxin/antitoxin system for markerless knockout generation. (16,17) However, the organism’s engineering toolbox could benefit from rapid genome engineering techniques such as homologous recombination to further leverage its native strengths. We recently reported utilizing the microbe’s native homologous recombination machinery to perform genomic knock-ins and targeted gene deletions. (18) However, this method requires tedious cloning, is only capable of catalyzing recombination with very long overhangs at very low efficiency, and with the integration of a selectable marker, making it unsuitable for high-throughput engineering, creation of point mutations, or markerless knockouts. This is especially relevant since, as an acetogen, E. limosum lacks the nonhomologous end joining DNA repair pathway, further limiting its genome engineering capacity. (19) The development of more efficient recombineering systems has therefore been of growing interest in the acetogen synthetic biology community, as such tools would significantly streamline engineering efforts in these organisms. (20,21) Without these tools, engineering is limited to large edits, complex multistep workflows, and the integration of selection markers, rendering evaluation of the physiological impact of specific genomic point mutations or construction of complex genetic circuits challenging.
Homologous recombination (HR) in bacteria via phage-derived recombinases is one of the fundamental tools in the synthetic biology toolbox. (22) The most well-known examples of this are the lambda Red (in which the recombinase enzyme is β) and RecT (in which the recombinase enzyme is RecT) recombineering systems in Escherichia coli, which enable gene modification via point or large mutations, gene knockout, and random oligonucleotide (oligo) mediated mutagenesis. These tools have significantly contributed to E. coli ’s development as the de facto bacterial engineering host. (23−25) More recently, Cas9 has been added to these recombineering systems as a “proof-reading” counterselection agent, cleaving unmutated DNA and thereby boosting the proportion of viable recombined cells to 100%, often in a scarless and markerless manner. (26,27) While β from lambda Red is the most commonly used recombinase in E. coli, this protein family is not widespread and has limited host range. (28,29) Across all bacteria, recombinases from the phage protein superfamily RecT are the most broadly distributed. (30) RecT binds to single strand DNA and anneals it to homologous ssDNA in the host during DNA replication in a process independent of native RecA-mediated DNA repair. (31,32) Given their functional orthogonality to native DNA repair, it has been hypothesized that RecT recombinase may possess much broader host range than β, unconstrained by taxonomy. Indeed, a recent screen of a large library of RecT proteins in E. coli revealed a surprising lack of correlation between recombinase activity and phylogenetic similarity. (29) Consequently, RecT recombinases are a promising family to investigate for implementing recombineering in underdeveloped microbes. Here, to enhance the synthetic biology capacity of E. limosum, we developed a highly efficient RecT-based recombineering system with Cas9 counterselection for markerless genome engineering.

Results and Discussion

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Recombinase Library Screening

Given the unclear importance of the phylogenetic similarity between recombinase source and target host, we assembled a library of 57 unique recombinases for assessment in E. limosum selected from a broad range of sources including both characterized and putative enzymes (29,30) (Table S1). We began constructing the recombinase library by selecting a group of published and/or characterized recombinases including GP35 from Bacillus subtilis, β from E. coli, and RecTs from Clostridium perfringens, Pseudomonas syringae, and E. coli. (29,30,33,34) Next, since strong bacterial recombinases typically originate from phages, we utilized the PHASTER tool to search E. limosum strains ATCC 8486, SA11, and 81C1 for prophage regions. (35) This analysis revealed several putative prophages in the genomes of these strains, with the 81C1 and SA11 strains each containing one gene annotated as recT, which we selected for testing. Following on from this analysis, PHASTER also identified the original phages likely responsible for these infections, from which we identified the original recT genes and added them to the library. These were checked for genes annotated as recT as inferred from homology, of which, across the three E. limosum strains, phages PLE3, phiCT19406A, phiCT19406B, phiCTC2A, phiCTC2B, and phiCT19406C encoded suitable genes. Proteins with their original sequence as found in their native phages were added because prophages within bacterial genomes are commonly inactivated via a plethora of mechanisms including point mutations, regulatory deactivation, and gene shuffling including operon inversion, among others, making it possible for a prophage-derived RecT to be nonfunctional. (36)
Next, we searched for recombinase genes in related organisms in the NCBI genome database, screening the class Clostridia and phylum Firmicutes in alignment with E. limosum’s taxonomy. From this screen, we selected seven additional proteins annotated as RecT. Then, based on recent data indicating that phylogenetic similarity may not be a factor in recombinase performance, we expanded our criteria and selected 29 additional recT genes from a broad range of bacterial backgrounds. (29) Finally, we applied the same approach to phages and selected 10 recT genes from a diverse range of bacteriophages. The 57 recombinase genes were refactored for expression in E. limosum and synthesized into plasmid-borne expression cassettes under the control of the inducible p2TetO1 promoter. (15,37)
We assessed recombinase performance in a fashion similar to procedures reported previously by creating a base “recombinase test strain” (RTS) encoding a mutated genomic copy of ermB (ermB′) for inactivated clarithromycin resistance with three stop codons inserted early in the sequence to prevent expression (29,33) (Figure S1). We transformed this strain individually with each of the recombinase expression constructs, then pooled the resulting strains into a single library culture (Table S2). We transformed the library culture with 120 bp recombinogenic oligos targeting the stop codons in ermB′ to return them to the original coding sequence and restore antibiotic resistance function (Figure 1a, Table S3). This initial library test was performed with oligos in both the leading and lagging strand orientations as well as on the RTS strain without a RecT recombinase to assess the efficacy of the native RecA recombinase in catalyzing oligo-length ssDNA recombination. While it is generally accepted that recombinases function at much higher efficiency annealing heterologous ssDNA to the lagging strand of genomic DNA during DNA replication, it is possible for them to anneal foreign ssDNA to the leading strand as well. (22) So, while lagging strand ssDNA oligos are typically used for recombineering, we included leading strand orientation oligos as well as this orientation has occasionally been known to recombine. (38)

Figure 1

Figure 1. Screening a recombinase library reveals a highly active RecT protein from the E. limosum SA11 prophage. (a) Scheme for testing recombinase library by activating a mutated ermB gene containing early stop codons. (b) Number of clarithromycin-resistant colonies obtained by transforming recombinase library or no-library control with 120 bp ssDNA recombinogenic oligos targeting ermB′. Error bars indicate standard deviation of biological triplicate transformations. (c) Proportion of clarithromycin resistant colonies, from a sample of 100 colonies each, encoding any given recombinase from conditions treated with either leading or lagging strand oligos for ermB′ correction.

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These transformations revealed that the library possessed significant recombinase function with a strong preference for lagging strand ssDNA, resulting in about 28,000 recombined colonies for lagging strand and 308 for leading strand conformations per transformation (Figure 1b). Additionally, the native RecA recombinase is capable of catalyzing some very low level of recombination with ssDNA oligos, but at an efficiency many orders of magnitude below the RecT recombinases. Sequencing of 100 resistant colonies showed that the RecT from the E. limosum SA11 prophage was the most abundant library member, being responsible for 41 ± 9.6% (α = 0.05) of recombined colonies, followed by the RecT from the E. limosum 81C1 prophage at 23 ± 8.2% (α = 0.05) with lagging strand ssDNA (Figure 1c). Based on these findings, we selected SA11 RecT and lagging strand ssDNA as our recombinase and DNA orientation for further optimization.
The difference in efficiency between the DNA conformations is perhaps unsurprising given the high availability of the lagging strand in the genomic DNA during replication and that this phenomenon is observed across other recombinases. (22,39) While orientation specificity was as expected, the distribution of recombinases is interesting in that it seems to indicate that phylogenetic similarity between source and host is an important factor for recombinase function in E. limosum, which is in contrast to recent results in E. coli. (29) In fact, the two most efficient recombinases in E. limosum ATCC 8486 were sourced from prophages in the genomes of two other strains of E. limosum: SA11 and 81C1, accounting for nearly two-thirds of all recombined colonies. RecT from another closely related source, Clostridium tertium, also contributed significantly, and ultimately about 73% of all recombined colonies possessed recT genes sourced from the class Clostridia. However, this is not to say that source similarity is the only determinant of recombinase function for E. limosum, as recombinases isolated from distantly related Sneathia vaginalis and E. coli both catalyzed efficient recombination. Surprisingly, the prophage from SA11 which encoded the best RecT enzyme originated from Lactobacillus phage PLE3, a phage whose typical host is not closely related to E. limosum. This finding adds more complexity to the question of whether phylogenetic similarity is important in recombinase selection given this phage’s normal host range.

Recombineering System Optimization

Recombination efficiency has previously been optimized in other organisms by a variety of means including varying DNA concentration and overhang length, modifying DNA structure, mutating the recombinase enzyme, supplementing the reaction with small-molecule enhancers, and changing recovery and growth conditions. (40−42) Total homology length is an essential consideration in recombinase function, as some recombinases work best with specific lengths of homology between heterologous DNA and genomic template. (40) Additionally, the concentration at which the heterologous DNA is provided is an important consideration as this is known to have a significant impact on transformation efficiency in general. (43) Therefore, we began optimizing our system by assessing the impact of varying oligo lengths and concentrations utilizing 120, 60, and 30 bp oligos spanning a final concentration of 1–1000 ng/μL in the transformation reaction. Maximum recombination efficiency was achieved using 60 bp oligos at a concentration of 1000 ng/μL (Figure 2a). We were unable to examine any higher DNA concentrations due to arcing during electroporation. Interestingly, we obtained very few colonies for any concentration of 30 bp oligo DNA. These oligos encode a total of 26 bp homology with the host gDNA, which implies that this length is insufficient for recombination catalyzed by RecT. However, maximum efficiency was observed with the 60 bp oligos encoding a total homology of 56 bp, which indicates that the minimum homology length for recombination is between 26 and 56 bp. Additionally, increasing the oligo length beyond 60 bp did not significantly improve efficiency for any concentration, however, efficiency did significantly decrease with the 120 bp oligos at the highest concentration of 1000 ng/μL.

Figure 2

Figure 2. Oligo and process modifications affect recombineering efficiency. (a) Effect of oligo size and concentration on recombineering efficiency (% viable cells recombined). (b) Effect of oligo modifications on recombineering efficiency. “S-oligo” refers to oligos with phosphonothioate bases in the first and final three positions of the oligo, “wobble” refers to oligos encoding consecutive wobble-base codon mutations in addition to the original targeted mutation, and “generic” refers to oligos without either of these alterations. (c) Effect of recovery time on recombineering efficiency. In all cases error bars represent standard deviation of biological triplicate transformations. ND: not detected.

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Next, we used the optimized conditions of 60 bp oligos at a concentration of 1000 ng/μL to assess the impact of oligo modifications on recombination efficiency by two strategies: mutating consecutive wobble-bases, and reducing the deleterious effect of exonucleases. (40,44) It has previously been reported for recombineering systems in other organisms that efficiency can be improved by mutating consecutive wobble-bases, a modification which is hypothesized to enable evasion of native DNA proofreading machinery. (40) It has further been shown that oligo degradation by exonucleases can reduce recombination efficiency by diminishing the effective concentration of recombinogenic DNA. (44) We obtained oligos encoding mutations to consecutive wobble bases in addition to the repair for ermB′, and also obtained oligos with phosphorothioate bases in the first and final three positions to minimize exonuclease degradation (45) (Table S3). Contrary to expectations, these modifications had no beneficial effect on recombination efficiency, and in fact the additional wobble base mutations reduced efficiency (Figure 2b). This may indicate that, in E. limosum, the potential enhancing effect of evading mismatch repair is outcompeted by a reduced efficiency of recombination. It is also likely that the concentration of 1000 ng/μL is so high that it overwhelms the cell’s exonucleases, rendering the phosphorothioate modification unnecessary.
Finally, we assessed the effect of recovery time on recombination efficiency, since it has been hypothesized that differences in cell survival and growth between recombined and unrecombined strains following transformation affect overall transformation efficiency observed after plating for single colonies. (40) We tested a panel of recovery times spanning 1–6 h and observed only minor differences across that range, with efficiency relatively stable from 3 h onward (Figure 2c). Optimal efficiency was observed in a 3–6 h recovery time window, which is consistent with the optimal recovery time for general transformations in E. limosum. (15) Using optimized conditions, the recombineering system is capable of up to 2% recombineering efficiency (1/50 cells possess the desired mutation from recombinase activity), which in most cases equates to more than 2 × 106 recombined colonies per transformation. This should be sufficient for library-based engineering approaches given careful screening criteria, however, in most cases, it is not sufficient for markerless genome engineering.

Implementation of Cas9 Counterselection

To enable high efficiency markerless genome engineering, we combined our recombineering system with Cas9 counterselection, a strategy which has enabled near perfect recombineering efficiency in E. coli. (46) We tied the recombinase-mediated mutation to the deletion of a PAM site and in tandem expressed Cas9 with an sgRNA targeting that region in unmutated strains (Table S4). Since the PAM site is essential for Cas9 activity, the Cas9 enzyme will only bind and cleave unrecombined genomes and therefore only correctly recombined strains will survive Cas9 expression. Additionally, a second sgRNA was included to target the recombinase plasmid to reduce the probability of aberrant recombination after obtaining the desired mutation (Figure 3). (47) To demonstrate the applicability of this method in E. limosum, we used a strain with the colorimetric reporter gene bgaB integrated into the genome (Figure S2). In the presence of S-Gal and ammonium ferric citrate, cells expressing the β-galactosidase enzyme BgaB produce an insoluble black product which makes them easily differentiated by visual screening. (15) Analysis of a homology model of BgaB identified Glu-303 as the catalytic nucleophile (Figure S3), consistent with Glu-537 in E. coli β-galactosidase which, when mutated, abolishes activity of that enzyme. (48) To eliminate function of BgaB, we designed a recombinogenic oligo, oPAS518, to mutate this codon to instead code for alanine (Table S3). (49) The oligo additionally possessed a silent mutation to base 912 of the bgaB gene from guanine to adenine to remove the PAM site adjacent to the coding mutation.

Figure 3

Figure 3. Scheme for improved recombineering efficiency using Cas9 counterselection. Plasmid encoding RecT and two sgRNAs-one targeting a PAM site near the desired mutation and one targeting the RecT plasmid-is transformed into E. limosum expressing BgaB for a black phenotype. That strain is again transformed with Cas9 plasmid and recombinogenic oligos targeting inactivation of BgaB and selected on tetracycline. Expression of Cas9 kills unrecombined colonies and destroys the recombinase plasmid. Finally, surviving colonies are subcultured to cure the Cas9 plasmid resulting in a strain with a markerless and scarless genomic point mutation to inactivate BgaB, leaving cells with a white phenotype.

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We constructed plasmid pEL11-BgaB encoding RecT, inducibly expressed from the p2TetO1 promoter, along with two sgRNAs: one targeting the plasmid itself, and the other targeting bgaB in the genome (Tables S3 and S4). (15,37) We transformed the E. limosum strain encoding genomic bgaB with this plasmid, and prepared competent cells with RecT induced during growth. We transformed this strain with the Cas9-bearing plasmid along with the recombinogenic oligo encoding the E303A BgaB mutation and selected on tetracycline plates. To reduce total exposure to Cas9, we added anhydrotetracycline for induction only during recovery and did not include it in agar plates. We performed expression analysis to confirm that the concentration of tetracycline used for selection was not sufficient to induce expression from the p2TetO1 promoter, and therefore selection could not inadvertently induce expression of Cas9 or RecT (Figure S4). Since the Cas9 plasmid, pRec-Cas2, encodes tetracycline resistance, only cells which received the Cas9 plasmid survived plating. Consequently, these cells also received the Cas9 counterselection, killing those which were not recombined by RecT, leaving only properly recombined cells. Following transformation, we picked colonies from each condition and treated them with S-Gal and ammonium ferric citrate to assess BgaB activity. The experimental condition with RecT, the recombinogenic oligo, and Cas9 counterselection resulted in all colonies containing inactive BgaB as evidenced by their uniform white color. In contrast, control transformations lacking either RecT, oligos, or Cas9, retained their black phenotype (Figure 4a,b). These results were also confirmed by DNA sequencing, which showed a white colony possessed the two targeted point mutations, while a black colony retained the unmutated sequence (Figure S5). Additionally, the absence of colonies on tetracycline plates for conditions lacking the recombinogenic oligo but transformed with Cas9 plasmid encoding Tet(M) confirms that Cas9 counterselection killed unrecombined colonies, with no escapees (Figure 4c). While it is sometimes possible for escapees to evade Cas9 counterselection, we did not observe the phenomenon here. It is interesting to note that the number of colonies observed on tetracycline plates for the experimental condition treated with RecT, Cas9, and recombinogenic oligos was far lower than would normally be expected for plasmid transformations. In this strategy, a single cell must receive both the plasmid and recombinogenic oligo during the transformation in order to obtain the mutation and survive on the selection plate. Receiving either one of these is already unlikely, and the necessity of receiving both compounds this unlikeliness, resulting in far fewer correct colonies.

Figure 4

Figure 4. Cas9 counterselection boosts proportion of viable recombined cells to 100%. Error bars represent standard deviation of biological triplicates. (a) Recombination efficiency as determined by colony color, (b) representative 96-well plates for colonies picked from condition treated with RecT and recombinogenic oligos but lacking Cas9 and sgRNA (left) and condition with Cas9 and sgRNA (right). Arrows on the left plate indicate recombined strains. Every colony on the right plate lacks BgaB activity. (c) Number of colonies obtained on tetracycline and nonselective agar plates showing that on the order of hundreds of recombined colonies may be obtained from a transformation. (d) Colony PCR results showing removal of pEL11-BgaB (plasmid encoding RecT and sgRNAs) and pRec-Cas2 (Plasmid encoding Cas9) following counterselection and curing. ND: not detected.

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We desired to eliminate the recombinase plasmid after achieving the targeted mutation, to remove the selection marker and prepare the strain for future genetic engineering work. The additional sgRNA targeting the pEL11 backbone results in Cas9 cleavage of the RecT expression plasmid. Additionally, the Cas9 expression plasmid is maintained by RepH, which has a low copy number in E. limosum and is easily lost by curing. (15) Following selection on tetracycline plates, we grew recombined strains for two generations in liquid culture without selection to cure out the Cas9 plasmid, resulting in a final strain encoding the desired genomic point mutation with no plasmids or selection markers (Figures 4d and S6). The design of the system makes it easy to swap out guide RNAs on the recombinase plasmid by simply PCR amplifying the full plasmid and encoding a new sgRNA on oligo overhangs. This enables quick turnaround for genome engineering workflows in a strategy which can be applied to any arbitrary target in the genome with a suitable nearby PAM site. RecT recombination with Cas9 counterselection enables rapid and simple generation of genome point mutations in E. limosum with up to 100% of viable cells successfully recombined, resulting in a final strain containing no plasmids or selection markers.

Recombinase-Mediated Genome Engineering

As a practical demonstration of our system, we applied the optimized recombineering/Cas9 counterselection scheme to create an E. limosum knockout strain incapable of biofilm formation. Extracellular polymeric substance (EPS) in E. limosum forms a thick, slimy mass which makes it challenging to manipulate by pipetting and centrifugation and hinders its applications. (50) We previously identified the genes responsible for EPS production and created a knockout strain using RecA-mediated HR, however this process left the catP selection marker in the genome, significantly reducing the strain’s usefulness in an engineering context. (18) Here, we generated an analogous strain without any selection markers left in the genome utilizing RecT recombineering and a 120 bp lagging strand oligo designed for a complete knockout of the gene cluster, with the oligo encoding 43 bp homology arms from either side of the gene cluster and flanking a 34 bp noncoding region for integration and replacement of the cluster. We constructed plasmid pEL11-eps encoding RecT and sgRNA targeting the eps gene cluster in the genome along with sgRNA for plasmid removal and transformed this into E. limosum along with pRec-Cas2 and the recombinogenic oligo in a strategy analogous to that utilized for BgaB inactivation (Figure 5a).

Figure 5

Figure 5. Recombination for biofilm abolition with Cas9 counterselection. (a) Scheme for eps gene cluster inactivation. (b) Colony PCR results for amplification across eps gene cluster region +150 bp on either side. Bands show full length gene cluster of ∼5 kb for WT strain and removal of operon for engineered strain leaving just the ∼150 bp on either side. (c) Centrifugation analysis demonstrating Δeps strain pellets at low speed (3500g). Both strains are fully suspended before centrifugation (left), but only the Δeps strain pellets after centrifugation (right).

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Following induction, transformation, and selection, we obtained EPS-deficient strains and performed colony PCR and sequencing to confirm the correct genomic mutation (Figure 5b). Since EPS production prevents pelleting by centrifugation at low speed, we utilized this as a readout to assess our knockout strain. As expected, the Δeps strain pelleted fully whereas no pelleting was observed with a wild type culture centrifuged under the same conditions (Figure 5c), confirming that we were able to regenerate the Δeps strain using a clean approach leaving no selection markers in the genome.
Rapid genomic modification has been a major bottleneck in the synthetic biology of acetogens, and Clostridia more broadly. The development here of high-efficiency recombineering and markerless genome engineering will significantly streamline the development of engineered E. limosum strains for production of an expanded array of products from single-carbon substrates, and facilitate fundamental investigations into acetogen physiology and metabolism. This tool is an advancement beyond commonly used CRISPR/Cas systems in Clostridia as it opens new genome engineering approaches like library-based or random oligo mutagenesis-based approaches. Additionally, it does not require cloning long homology arms, which can be laborious in Clostridial engineering systems. Beyond E. limosum, we believe the workflow presented here-specifically the mining of prophages and their original phages for highly active RecT’s-may serve as a roadmap for the broader Clostridial synthetic biology community. Additionally, the recombinase library (sequences provided in Supporting Information) we generated employs a widely used Clostridial plasmid architecture (pIP404 ori, catP for thiamphenicol resistance, p2tetO1 promoter) and encodes a broad range of RecT’s (including several from Clostridia), facilitating its testing in other Clostridia of industrial significance and importance in human health. Given the wide host range of RecT enzymes, such screening will likely accelerate development of genetic tools for this notoriously recalcitrant class of organisms.

Materials and Methods

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Reagents, Bacterial Strains, and Growth Conditions

All cloning enzymes were obtained from New England Biolabs (NEB), all oligos 60 bp or shorter were obtained from Genewiz/Azenta and all oligos 61 bp+ or containing phosphorothioate bases were obtained from Sigma-Aldrich. Precloned plasmid constructs utilizing synthesized DNA were obtained from the Joint Genome Institute (JGI). PCR cleanup and miniprep kits were obtained from Zymo Research, gDNA prep kit from NEB, and all other reagents from Fisher Scientific. Cloning and plasmid work were carried out in either E. coli NEB5α or stable competent E. coli, both from NEB and all work in E. limosum was conducted in strain ATCC 8486. E. coli was grown at 37 °C in liquid LB, or LB agar containing 50 μg/mL ampicillin or chloramphenicol. E. limosum was grown anaerobically at 37 °C without agitation in DSMZ 135 medium without sulfide for liquid culture and reinforced clostridial medium (RCM) for agar plates. Thiamphenicol was used at a concentration of 5 μg/mL, clarithromycin and tetracycline at 0.5 μg/mL, and aTc at 30 ng/mL. All anaerobic work was done in a Coy Lab Products flexible anaerobic chamber with a 10% CO2, 5% H2, 85% N2 gas mix. Electroporation was conducted using a Bio-Rad GenePulser XCell electroporator following a protocol and electrocompetency preparation reported previously. (15) Optical density and DNA concentration measurements were taken with a Molecular Devices QuickDrop spectrophotometer.

Cloning and Construct Preparation

Plasmids pEL6-pEL67 were obtained precloned and codon harmonized from JGI, each with an identical architecture using the IP404 origin of replication, catP for selection on thiamphenicol, and p2TetO1 driving the recombinase. Plasmid pRTS3 was cloned via a 5-piece NEB HiFi assembly using backbone amplified from pUC19 with oligos oPAS222 and 223, overhangs for genomic integration amplified from E. limosum ATCC 8486 gDNA using oligos oPAS328 and 271 for the lefthand side overhang (LHS) and oPAS269 and 329 for the righthand side overhang (RHS), Tet(M) amplified from pVEL4 using oPAS270 and 227, and ermB′ amplified from synthesized DNA using oPAS267 and 268. Linear DNA for integration into the E. limosum genome was amplified from the assembled plasmid using oPAS330 and 331. Control plasmid pRTS3.(+) was constructed in the same way with the exception of using functional ermB amplified from pCL2.1 with oPAS267 and 268 rather than mutated ermB′. (15)
Plasmid pRec-Cas1 was cloned via a 3-piece T4 ligation reaction using backbone amplified from pUC19 with oligos oPAS620 and 621, RepH amplified from pMTL83151 using oPAS622 and 623, and Tet(M) amplified from pVEL4 using oPAS624 and 625. (51) Plasmid pRec-Cas2 was then cloned by a 2-piece T4 ligation reaction using Cas9 amplified from synthesized DNA with oPAS698 and 676 and digested with NheI and BsrGI and backbone amplified from pRec-Cas1 with oPA696 and 697 before digest with NheI and BsrGI. pEL11-BgaB was in turn cloned by digesting pEL11 with NheI to linearize and performing an NEB HiFi assembly reaction with a combined sgRNA expression cassette targeting BgaB and pEL11. Individual sgRNA fragments driven by the J23119 promoter targeting BgaB and pEL11 were synthesized and then combined using overlap extension PCR by amplifying pEL11 sgRNA with oPAS682 and 683 and BgaB sgRNA with 684 and 685 and then combining them and amplifying the double cassette with oPAS717 and 718. Plasmid pEL11-eps was cloned by amplifying pEL11-BgaB with oEPS53 and 54 to replace the BgaB-targeting sgRNA with one targeting the eps operon and then religated in a one-piece NEB-HiFi assembly reaction. Plasmid pBgaBg was cloned via a 2-piece NEB HiFi assembly using a backbone from pRTS3.(+) amplified with oPAS496 and 497, and bgaB driven by nifJp amplified from pELIM2.2 using oPAS498 and 499. (15) Linear DNA for genomic knock-in transformation in E. limosum was amplified from the plasmid using oligos oPAS330 and 331.

Recombinase System Development

E. limosum genomes ATCC 8486, SA11, and 81C1 were screened for prophage regions using PHASTER (accession numbers NZ_CP019962.1, NZ_CP011914.1, NZ_LR215983.1 respectively). We selected phages based on gene hit count as determined by PHASTER, assessing ones with four or more clustered gene hits in a single genome region. Phages PLE3, phiCT19406A, phiCT19406B, phiCTC2A, phiCTC2B, and phiCT19406C have accession numbers NC_031125, NC_030950.1, NC_030947.1, NC_030949.1, NC_030951.1, NC_029006.1 respectively. Strain RTS was generated by PCR amplifying linear DNA encoding ermB′ from pRTS3 and transforming it into WT E. limosum utilizing RecA HR for genomic integration as described previously and plating on clarithromycin. (18) Competent cells were prepared from strain RTS and transformed with recombinase plasmids. Individual strains were selected on thiamphenicol, picked into liquid cultures, grown to an OD600 of 0.5, and pooled into a single culture. Any deviations from OD600 0.5 between cultures were normalized to 0.5 with DSMZ 135 media before 750 μL of each were combined into a single culture from which competent cells were prepared. Transformations were performed with 50 μL competent cells and 100 ng/μL DNA with 120 bp oligos in both leading and lagging strand orientations as well as a no DNA control and plated in a dilution series 1:10, 1:100, 1:1000, 1:10000 onto clarithromycin plates. Plates were allowed to grow for 6 days before 100 single colonies were picked from each condition. Recombinases encoded by individual resistant colonies were determined by colony PCR to amplify the recombinase gene, and sanger sequencing to determine recombinase identity. Any sequencing samples with a quality score of 24 or below were not included in the data set. The frequency of a given recombinase was calculated at the 95% confidence level (Z = 1.96) based on sample size and population size for each condition. Recombineering optimization was performed solely on strain RTS + pEL11 but otherwise in an analogous manner. All transformations were performed in biological triplicate.

Implementation of Cas9 Counterselection

A homology model of the Geobacillus stearothermophilus β-galactosidase (BgaB) was made with the SWISS-MODEL server, using the crystal structure of the galactose-bound Bifidobacterium animalis enzyme (4UNI) as template. (52,53) Conserved active site residues were identified by overlaying the structures in PYMOL. Strain E. limosum pEL11-BgaB/BgaBg was created by performing a knock-in transformation to place BgaB with ermB for selection into the genome in a fashion and location identical to RTS. (18) Next, competent cells were prepared from this strain and transformed with pEL11-BgaB and selected on thiamphenicol. Finally, competent cells were prepared from this strain with aTc induction and used for recombineering transformations. Recombineering transformations were performed in biological triplicates with 50 μL of competent cells and 10 μL of DNA for a total volume of 60 μL according to Table 1. Recombinogenic oligos were used at a final concentration of 1000 ng/μL and plasmid DNA at 4 ng/μL. aTc was added to the recovery media. Colonies were counted after 6 days of growth and viability determined by comparison to transform B6. 96 colonies were selected from each transformation plate for conditions B1–B6 and grown in 200 μL 96-well plate cultures overnight. Cells were spotted onto tetracycline, thiamphenicol, and nonselective plates and then 250 μg/mL S-Gal and 400 μg/mL ammonium ferric citrate added to each well in the 96-well culture plates and allowed to incubate at 37 °C for 1 h before visually assessing BgaB activity. After correctly recombined strains were identified, they were subcultured twice in nonselective liquid media before plating and colony PCR analysis to confirm loss of pEL11-BgaB and pRec-Cas2.
Table 1. Transformations for Cas9 Counterselection Demonstration Using the BgaB Reporter
transformstrainDNAplating
B1pEL11/BgaBgoPAS518RCM + tetracycline & RCM only
B2pEL11-BgaB/BgaBgoPAS518RCM + tetracycline & RCM only
pRec-Cas2
B3pEL11/BgaBgoPAS518RCM + tetracycline & RCM only
pRec-Cas2
B4pEL11-BgaB/BgaBgpRec-Cas2RCM + tetracycline & RCM only
B5pEL11-BgaB/BgaBgoPAS518RCM + tetracycline & RCM only
B6pEL11/BgaBgNoneRCM + tetracycline & RCM only

Recombinase-Mediated Genome Engineering

Wild type competent cells were transformed with pEL11-eps, selected, cultured, induced, and made competent. 50 μL pEL11-eps competent cells were transformed with 5 μL oligo oEPS55 at a final concentration of 500 ng/μL and plasmid pRec-Cas2 at 4 ng/μL with aTc added to the recovery media for cas9 expression before plating on tetracycline plates. This volume was used rather than the 10 μL used previously in order to keep the total additional volume in the transformation to 10 μL since 5 μL of plasmid was also added. After 6 days of growth, single colonies were picked from agar plates into a well plate with 150 μL DSMZ 135 media per well and allowed to grow overnight. Following growth, each well was pipetted to qualitatively assess EPS production, and cells from a well without any biofilm were passaged into a 5 mL culture and allowed to grow overnight. Cultures were then centrifuged at 3500 xg for 5 min to assess pelleting. Colony PCR was performed on the knockout region using oligos oPAS763 and 764.

Data Availability

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The data underlying this study are available in the published article and its Supporting Information. All plasmids and strains used in this work are available by e-mail request to the corresponding author, or through www.woolstonlab.org/resources.

Supporting Information

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The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acssynbio.4c00253.

  • Tables of the recombinases, plasmids, oligos, and guide RNAs used in the study, along with enzyme assay data, sequencing, plating, and colony PCR data to further validate results presented in the main text (PDF)

  • Nucleotide sequences for the recombinase library members (XLSX)

Terms & Conditions

Most electronic Supporting Information files are available without a subscription to ACS Web Editions. Such files may be downloaded by article for research use (if there is a public use license linked to the relevant article, that license may permit other uses). Permission may be obtained from ACS for other uses through requests via the RightsLink permission system: http://pubs.acs.org/page/copyright/permissions.html.

Author Information

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  • Corresponding Author
  • Author
    • Patrick A. Sanford - Department of Chemical Engineering, Northeastern University, 360 Huntington Avenue, 223 Cullinane, Boston, Massachusetts 02115, United States
  • Author Contributions

    P.A.S. devised and performed experiments and engineering strategies, wrote the manuscript, and created the figures and graphics. Both authors analyzed data. B.M.W. conceived of the concept and edited the manuscript and figures.

  • Notes
    The authors declare no competing financial interest.

Acknowledgments

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This work was financially supported by Northeastern University startup funds and Department of Energy, Advanced Research Projects Agency-Energy ECOSynBIO under award number #DE-AR0001511. The work (proposal: 10.46936/10.25585/60008422) conducted by the U.S. Department of Energy Joint Genome Institute (https://ror.org/04xm1d337), a DOE Office of Science User Facility, is supported by the Office of Science of the U.S. Department of Energy operated under Contract No. DE-AC02-05CH11231.

References

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    Bacterial recombineering: Genome engineering via phage-based homologous recombination
    Pines, Gur; Freed, Emily F.; Winkler, James D.; Gill, Ryan T.
    ACS Synthetic Biology (2015), 4 (11), 1176-1185CODEN: ASBCD6; ISSN:2161-5063. (American Chemical Society)
    A review. The ability to specifically modify bacterial genomes in a precise and efficient manner is highly desired in various fields, ranging from mol. genetics to metabolic engineering and synthetic biol. Much has changed from the initial realization that phage-derived genes may be employed for such tasks to today, where recombineering enables complex genetic edits within a genome or a population. Here, we review the major developments leading to recombineering becoming the method of choice for in situ bacterial genome editing while highlighting the various applications of recombineering in pushing the boundaries of synthetic biol. We also present the current understanding of the mechanism of recombineering. Finally, we discuss in detail issues surrounding recombineering efficiency and future directions for recombineering-based genome editing.
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    Murphy, K. C.; Campellone, K. G. Lambda Red-mediated recombinogenic engineering of enterohemorrhagic and enteropathogenic E. coli. BMC Mol. Biol. 2003, 4, 1– 12,  DOI: 10.1186/1471-2199-4-11
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    Wang, H. H.; Isaacs, F. J.; Carr, P. A. Programming cells by multiplex genome engineering and accelerated evolution. Nature 2009, 460 (7257), 894– 898,  DOI: 10.1038/nature08187
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    Programming cells by multiplex genome engineering and accelerated evolution
    Wang, Harris H.; Isaacs, Farren J.; Carr, Peter A.; Sun, Zachary Z.; Xu, George; Forest, Craig R.; Church, George M.
    Nature (London, United Kingdom) (2009), 460 (7257), 894-898CODEN: NATUAS; ISSN:0028-0836. (Nature Publishing Group)
    The breadth of genomic diversity found among organisms in nature allows populations to adapt to diverse environments. However, genomic diversity is difficult to generate in the lab. and new phenotypes do not easily arise on practical timescales. Although in vitro and directed evolution methods have created genetic variants with usefully altered phenotypes, these methods are limited to laborious and serial manipulation of single genes and are not used for parallel and continuous directed evolution of gene networks or genomes. Here, we describe multiplex automated genome engineering (MAGE) for large-scale programming and evolution of cells. MAGE simultaneously targets many locations on the chromosome for modification in a single cell or across a population of cells, thus producing combinatorial genomic diversity. Because the process is cyclical and scalable, we constructed prototype devices that automate the MAGE technol. to facilitate rapid and continuous generation of a diverse set of genetic changes (mismatches, insertions, deletions). We applied MAGE to optimize the 1-deoxy-d-xylulose-5-phosphate (DXP) biosynthesis pathway in Escherichia coli to overproduce the industrially important isoprenoid lycopene. Twenty-four genetic components in the DXP pathway were modified simultaneously using a complex pool of synthetic DNA, creating over 4.3 billion combinatorial genomic variants per day. We isolated variants with more than fivefold increase in lycopene prodn. within 3 days, a significant improvement over existing metabolic engineering techniques. Our multiplex approach embraces engineering in the context of evolution by expediting the design and evolution of organisms with new and improved properties.
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  25. 25
    Kolodner, R.; Hall, S. D.; Luisi-DeLuca, C. Homologous pairing proteins encoded by the Escherichia coli recE and recT genes. Mol. Microbiol. 1994, 11 (1), 23– 30,  DOI: 10.1111/j.1365-2958.1994.tb00286.x
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    Pyne, M. E.; Moo-Young, M.; Chung, D. A.; Chou, C. P. Coupling the CRISPR/Cas9 system with lambda red recombineering enables simplified chromosomal gene replacement in Escherichia coli. Appl. Environ. Microbiol. 2015, 81 (15), 5103– 5114,  DOI: 10.1128/AEM.01248-15
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    26
    Coupling the CRISPR/Cas9 system with lambda Red recombineering enables simplified chromosomal gene replacement in Escherichia coli
    Pyne, Michael E.; Moo-Young, Murray; Chung, Duane A.; Chou, C. Perry
    Applied and Environmental Microbiology (2015), 81 (15), 5103-5114CODEN: AEMIDF; ISSN:1098-5336. (American Society for Microbiology)
    To date, most genetic engineering approaches coupling the type II Streptococcus pyogenes clustered regularly interspaced short palindromic repeat (CRISPR)/Cas9 system to lambda Red recombineering have involved minor single nucleotide mutations. Here we show that procedures for carrying out more complex chromosomal gene replacements in Escherichia coli can be substantially enhanced through implementation of CRISPR/Cas9 genome editing. We developed a three-plasmid approach that allows not only highly efficient recombination of short single-stranded oligonucleotides but also replacement of multigene chromosomal stretches of DNA with large PCR products. By systematically challenging the proposed system with respect to the magnitude of chromosomal deletion and size of DNA insertion, we demonstrated DNA deletions of up to 19.4 kb, encompassing 19 nonessential chromosomal genes, and insertion of up to 3 kb of heterologous DNA with recombination efficiencies permitting mutant detection by colony PCR screening. Since CRISPR/Cas9-coupled recombineering does not rely on the use of chromosome- encoded antibiotic resistance, or flippase recombination for antibiotic marker recycling, our approach is simpler, less labor-intensive, and allows efficient prodn. of gene replacement mutants that are both markerless and "scar"-less.
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    Reisch, C. R.; Prather, K. L. J. The no-SCAR (Scarless Cas9 Assisted Recombineering) system for genome editing in Escherichia coli. Sci. Rep. 2015, 5, 15096  DOI: 10.1038/srep15096
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    27
    The no-SCAR (Scarless Cas9 Assisted Recombineering) system for genome editing in Escherichia coli
    Reisch, Chris R.; Prather, Kristala L. J.
    Scientific Reports (2015), 5 (), 15096CODEN: SRCEC3; ISSN:2045-2322. (Nature Publishing Group)
    Genome engineering methods in E. coli allow for easy to perform manipulations of the chromosome in vivo with the assistance of the λ-Red recombinase system. These methods generally rely on the insertion of an antibiotic resistance cassette followed by removal of the same cassette, resulting in a two-step procedure for genomic manipulations. Here we describe a method and plasmid system that can edit the genome of E. coli without chromosomal markers. This system, known as Scarless Cas9 Assisted Recombineering (no-SCAR), uses λ-Red to facilitate genomic integration of donor DNA and double stranded DNA cleavage by Cas9 to counter select against wild-type cells. We show that point mutations, gene deletions, and short sequence insertions were efficiently performed in several genomic loci in a single-step with regards to the chromosome and did not leave behind scar sites. The single-guide RNA encoding plasmid can be easily cured due to its temp. sensitive origin of replication, allowing for iterative chromosomal manipulations of the same strain, as is often required in metabolic engineering. In addn., we demonstrate the ability to efficiently cure the second plasmid in the system by targeting with Cas9, leaving the cells plasmid-free.
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    Ellis, H. M.; Yu, D.; DiTizio, T.; Court, D. L. High efficiency mutagenesis, repair, and engineering of chromosomal DNA using single-stranded oligonucleotides. Proc. Natl. Acad. Sci. U.S.A. 2001, 98 (12), 6742– 6746,  DOI: 10.1073/pnas.121164898
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    28
    High efficiency mutagenesis, repair, and engineering of chromosomal DNA using single-stranded oligonucleotides
    Ellis, Hilary M.; Yu, Daiguan; DiTizio, Tina; Court, Donald L.
    Proceedings of the National Academy of Sciences of the United States of America (2001), 98 (12), 6742-6746CODEN: PNASA6; ISSN:0027-8424. (National Academy of Sciences)
    Homologous DNA recombination is a fundamental, regenerative process within living organisms. However, in most organisms, homologous recombination is a rare event, requiring a complex set of reactions and extensive homol. We demonstrate in this paper that Beta protein of phage λ generates recombinants in chromosomal DNA by using synthetic single-stranded DNAs (ssDNA) as short as 30 bases long. This ssDNA recombination can be used to mutagenize or repair the chromosome with efficiencies that generate up to 6% recombinants among treated cells. Mechanistically, it appears that Beta protein, a Rad52-like protein, binds and anneals the ssDNA donor to a complementary single-strand near the DNA replication fork to generate the recombinant. This type of homologous recombination with ssDNA provides new avenues for studying and modifying genomes ranging from bacterial pathogens to eukaryotes. Beta protein and ssDNA may prove generally applicable for repairing DNA in many organisms.
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    Wannier, T. M.; Nyerges, A.; Kuchwara, H. M. Improved bacterial recombineering by parallelized protein discovery. Proc. Natl. Acad. Sci. U.S.A. 2020, 117 (24), 13689– 13698,  DOI: 10.1073/pnas.2001588117
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    29
    Improved bacterial recombineering by parallelized protein discovery
    Wannier, Timothy M.; Nyerges, Akos; Kuchwara, Helene M.; Czikkely, Marton; Balogh, David; Filsinger, Gabriel T.; Borders, Nathaniel C.; Gregg, Christopher J.; Lajoie, Marc J.; Rios, Xavier; Pal, Csaba; Church, George M.
    Proceedings of the National Academy of Sciences of the United States of America (2020), 117 (24), 13689-13698CODEN: PNASA6; ISSN:0027-8424. (National Academy of Sciences)
    Exploiting bacteriophage-derived homologous recombination processes has enabled precise, multiplex editing of microbial genomes and the construction of billions of customized genetic variants in a single day. The techniques that enable this, multiplex automated genome engineering (MAGE) and directed evolution with random genomic mutations (DIvERGE), are however, currently limited to a handful of microorganisms for which single-stranded DNA-annealing proteins (SSAPs) that promote efficient recombineering have been identified. Thus, to enable genome-scale engineering in new hosts, efficient SSAPs must first be found. Here the authors introduce a high-throughput method for SSAP discovery that the authors call 'serial enrichment for efficient recombineering' (SEER). By performing SEER in Escherichia coli to screen hundreds of putative SSAPs, the authors identify highly active variants PapRecT and CspRecT. CspRecT increases the efficiency of single-locus editing to as high as 50% and improves multiplex editing by 5-10-fold in E. coli, while PapRecT enables efficient recombineering in Pseudomonas aeruginosa, a concerning human pathogen. CspRecT and PapRecT are also active in other, clin. and biotechnol. relevant enterobacteria. The authors envision that the deployment of SEER in new species will pave the way toward pooled interrogation of genotype-to-phenotype relations in previously intractable bacteria.
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  30. 30
    Datta, S.; Costantino, N.; Zhou, X.; Court, D. L. Identification and analysis of recombineering functions from Gram-negative and Gram-positive bacteria and their phages. Proc. Natl. Acad. Sci. U.S.A. 2008, 105 (5), 1626– 1631,  DOI: 10.1073/pnas.0709089105
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    30
    Identification and analysis of recombineering functions from Gram-negative and Gram-positive bacteria and their phages
    Datta, Simanti; Costantino, Nina; Zhou, Xiaomei; Court, Donald L.
    Proceedings of the National Academy of Sciences of the United States of America (2008), 105 (5), 1626-1631CODEN: PNASA6; ISSN:0027-8424. (National Academy of Sciences)
    We report the identification and functional anal. of nine genes from Gram-pos. and Gram-neg. bacteria and their phages that are similar to lambda (λ) bet or Escherichia coli recT. Beta and RecT are single-strand DNA annealing proteins, referred to here as recombinases. Each of the nine other genes when expressed in E. col carries out oligonucleotide-mediated recombination. To our knowledge, this is the first study showing single-strand recombinase activity from diverse bacteria. Similar to bet and recT, most of these other recombinases were found to be assocd. with putative exonuclease genes. Beta and RecT in conjunction with their cognate exonucleases carry out recombination of linear double-strand DNA. Among four of these foreign recombinase/exonuclease pairs tested for recombination with double-strand DNA, three had activity, albeit barely detectable. Thus, although these recombinases can function in E. col to catalyze oligonucleotide recombination, the double-strand DNA recombination activities with their exonuclease partners were inefficient. This study also demonstrated that Gam, by inhibiting host RecBCD nuclease activity, helps to improve the efficiency of λ Red-mediated recombination with linear double-strand DNA, but Gam is not absolutely essential. Thus, in other bacterial species where Gam analogs have not been identified, double-strand DNA recombination may still work in the absence of a Gam-like function. We anticipate that at least some of the recombineering systems studied here will potentiate oligonucleotide and double-strand DNA-mediated recombineering in their native or related bacteria.
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    Noirot, P.; Kolodner, R. D. DNA strand invasion promoted by Escherichia coli RecT protein. J. Biol. Chem. 1998, 273 (20), 12274– 12280,  DOI: 10.1074/jbc.273.20.12274
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    Roca, A.; Cox, M.; Brenner, S. The RecA Protein: Structure and Function. Crit. Rev. Biochem. Mol. Biol. 1990, 25 (6), 415– 456,  DOI: 10.3109/10409239009090617
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    Dong, H.; Tao, W.; Gong, F.; Li, Y.; Zhang, Y. A functional recT gene for recombineering of Clostridium. J. Biotechnol. 2014, 173 (1), 65– 67,  DOI: 10.1016/j.jbiotec.2013.12.011
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    Swingle, B.; Bao, Z.; Markel, E.; Chambers, A.; Cartinhour, S. Recombineering using recTE from Pseudomonas syringae. Appl. Environ. Microbiol. 2010, 76 (15), 4960– 4968,  DOI: 10.1128/AEM.00911-10
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    Recombineering using RecTE from Pseudomonas syringae
    Swingle, Bryan; Bao, Zhongmeng; Markel, Eric; Chambers, Alan; Cartinhour, Samuel
    Applied and Environmental Microbiology (2010), 76 (15), 4960-4968CODEN: AEMIDF; ISSN:0099-2240. (American Society for Microbiology)
    In this report, we describe the identification of functions that promote genomic recombination of linear DNA introduced into Pseudomonas cells by electroporation. The genes encoding these functions were identified in Pseudomonas syringae pv. syringae B728a based on similarity to the lambda Red Exo/Beta and RecET proteins encoded by the lambda and Rac bacteriophages of Escherichia coli. The ability of the pseudomonad-encoded proteins to promote recombination was tested in P. syringae pv. tomato DC3000 using a quant. assay based on recombination frequency. The results show that the Pseudomonas RecT homolog is sufficient to promote recombination of single-stranded DNA oligonucleotides and that efficient recombination of double-stranded DNA requires the expression of both the RecT and RecE homologs. Addnl., we illustrate the utility of this recombineering system to make targeted gene disruptions in the P. syringae chromosome.
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    Arndt, D.; Grant, J. R.; Marcu, A. PHASTER: a better, faster version of the PHAST phage search tool. Nucleic Acids Res. 2016, 44 (W1), W16– W21,  DOI: 10.1093/nar/gkw387
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    PHASTER: a better, faster version of the PHAST phage search tool
    Arndt, David; Grant, Jason R.; Marcu, Ana; Sajed, Tanvir; Pon, Allison; Liang, Yongjie; Wishart, David S.
    Nucleic Acids Research (2016), 44 (W1), W16-W21CODEN: NARHAD; ISSN:0305-1048. (Oxford University Press)
    PHASTER (PHAge Search Tool - Enhanced Release)is a significant upgrade to the popular PHAST web server for the rapid identification and annotation of prophage sequences within bacterial genomes and plasmids. Although the steps in the phage identification pipeline in PHASTER remain largely the same as in the original PHAST, numerous software improvements and significant hardware enhancements have now made PHASTER faster, more efficient, more visually appealing and much more user friendly. In particular, PHASTER is now 4.3× faster than PHAST when analyzing a typical bacterial genome. Mores pecifically, software optimizations have made the backend of PHASTER 2.7X faster than PHAST, while the addn. of 80 CPUs to the PHASTER compute cluster are responsible for the remaining speed up. PHASTER can now process a typical bacterial genome in 3 min from the raw sequence alone, or in1.5 min when given a pre-annotated GenBank file. Anumber of other optimizations have also been implemented, including automated algorithms to reduce the size and redundancy of PHASTER's databases, improvements in handling multiple (meta genomic)queries and higher user traffic, along with the ability to perform automated look-ups against 14000 previously PHAST/PHASTER annotated bacterial genomes (which can lead to complete phage annotations in seconds as opposed to minutes). PHASTER's web interface has also been entirely rewritten. A new graphical genome browser has been added, gene/genome visualization tools have been improved, and the graphical interface is now more modern, robust and user-friendly.
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    Canchaya, C.; Proux, C.; Fournous, G.; Bruttin, A.; Brüssow, H. Prophage Genomics. Microbiol. Mol. Biol. Rev. 2003, 67 (3), 473,  DOI: 10.1128/MMBR.67.3.473.2003
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    Dong, H.; Tao, W.; Zhang, Y.; Li, Y. Development of an anhydrotetracycline-inducible gene expression system for solvent-producing Clostridium acetobutylicum: A useful tool for strain engineering. Metab. Eng. 2012, 14 (1), 59– 67,  DOI: 10.1016/j.ymben.2011.10.004
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    Development of an anhydrotetracycline-inducible gene expression system for solvent-producing Clostridium acetobutylicum: A useful tool for strain engineering
    Dong, Hongjun; Tao, Wenwen; Zhang, Yanping; Li, Yin
    Metabolic Engineering (2012), 14 (1), 59-67CODEN: MEENFM; ISSN:1096-7176. (Elsevier B. V.)
    Clostridium acetobutylicum is an important solvent (acetone-butanol-ethanol) producing bacterium. However, a stringent, effective, and convenient-to-use inducible gene expression system that can be used for regulating the gene expression strength in C. acetobutylicum is currently not available. Here, we report an anhydrotetracycline-inducible gene expression system for solvent-producing bacterium C. acetobutylicum. This system consists of a functional chloramphenicol acetyltransferase gene promoter contg. tet operators (tetO), Pthl promoter (thiolase gene promoter from C. acetobutylicum) controlling TetR repressor expression cassette, and the chem. inducer anhydrotetracycline (aTc). The optimized system, designated as pGusA2-2tetO1, allows gene regulation in an inducer aTc concn.-dependent way, with an inducibility of over two orders of magnitude. The stringency of TetR repression supports the introduction of the genes encoding counterselective marker into C. acetobutylicum, which can be used to increase the mutant screening efficiency. This aTc-inducible gene expression system will thus increase the genetic manipulation capability for engineering C. acetobutylicum.
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    Aparicio, T.; de Lorenzo, V.; Martínez-García, E. CRISPR/Cas9-enhanced ssDNA recombineering for Pseudomonas putida. Microb. Biotechnol. 2019, 12 (5), 1076– 1089,  DOI: 10.1111/1751-7915.13453
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    CRISPR/Cas9-enhanced ssDNA recombineering for Pseudomonas putida
    Aparicio, Tomas; de Lorenzo, Victor; Martinez-Garcia, Esteban
    Microbial Biotechnology (2019), 12 (5), 1076-1089CODEN: MBIIB2; ISSN:1751-7915. (Wiley-Blackwell)
    Summary : Implementation of single-stranded DNA (ssDNA) recombineering in Pseudomonas putida has widened the range of genetic manipulations applicable to this biotechnol. relevant bacterium. Yet, the relatively low efficiency of the technol. hampers identification of mutated clones lacking conspicuous phenotypes. Fortunately, the use of CRISPR/Cas9 as a device for counterselection of wild-type sequences helps to overcome this limitation. Merging ssDNA recombineering with CRISPR/Cas9 thus enables a suite of genomic edits with a straightforward approach: a CRISPR plasmid provides the spacer DNA sequence that directs the Cas9 nuclease ribonucleoprotein complex to cleave the genome at the wild-type sequences that have not undergone the change entered by the mutagenic ssDNA oligonucleotide(s). This protocol describes a complete workflow of the method optimized for P. putida, although it could in principle be applicable to many other pseudomonads. As an example, we show the deletion of the edd gene that encodes one key enzyme that operates the EDEMP cycle for glucose metab. in P. putidaEM42. By combining two incompatible CRISPR plasmids with different antibiotic selection markers, we show that the procedure can be cycled to implement consecutive deletions in the same strain, e.g. deletion of the pyrF gene following that of the edd mutant. This approach adds to the wealth of genetic technologies available for P. putida and strengthens its status as a chassis of choice for a suite of biotechnol. applications.
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    Wyman, C.; Ristic, D.; Kanaar, R. Homologous recombination-mediated double-strand break repair. DNA Repair 2004, 3 (8–9), 827– 833,  DOI: 10.1016/j.dnarep.2004.03.037
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    Sawitzke, J. A.; Costantino, N.; Li, X. t. Probing cellular processes with oligo-mediated recombination and using the knowledge gained to optimize recombineering. J. Mol. Biol. 2011, 407 (1), 45– 59,  DOI: 10.1016/j.jmb.2011.01.030
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    Inhibition of deoxyribonucleases by phosphorothioate groups in oligodeoxyribonucleotides
    Spitzer, Stephan; Eckstein, Fritz
    Nucleic Acids Research (1988), 16 (24), 11691-704CODEN: NARHAD; ISSN:0305-1048.
    The Rp- and Sp-diastereomers of the phosphorothiolate-contg. oligonucleotide d[ApAp(S)ApA] (I) were synthesized. They and the tetramer d[ApApApA] were tested as substrates for staphylococcal nuclease, DNase II, and spleen phosphodiesterase. For digestions with DNase I these oligonucleotides were converted to the 5'-phosphorylated derivs. The reactions with the nucleases were analyzed by HPLC. The phosphorothioate groups of both diastereomers were resistant to the action of staphylococcal nuclease, DNase I, and DNase II. While the phosphorothioate group of the Rp-diastereomer was resistant to the action of spleen phosphodiesterase, the Sp-diastereomer was hydrolyzed at an estd. rate 1/100 the rate of cleavage of the unmodified tetramer. The presence of the phosphorothiolate group in the center of the mol. affected the rate of hydrolysis of neighboring phosphate groups for some enzymes. In particular, very slow release of 3'-dAMP from the Rp-diastereomer occurred on incubation with staphylococcal nuclease but the Sp-diastereomer was completely resistant. DNase II produced 3'-dAMP quite rapidly from both diastereomers of I and DNase I released 5'-dAMP from both diastereomers of I only slowly.
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    Jiang, Y.; Chen, B.; Duan, C.; Sun, B.; Yang, J.; Yang, S. Multigene editing in the Escherichia coli genome via the CRISPR-Cas9 system. Appl. Environ. Microbiol. 2015, 81 (7), 2506– 2514,  DOI: 10.1128/AEM.04023-14
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    Multigene editing in the Escherichia coli genome via the CRISPR-Cas9 system
    Jiang, Yu; Chen, Biao; Duan, Chunlan; Sun, Bingbing; Yang, Junjie; Yang, Sheng
    Applied and Environmental Microbiology (2015), 81 (7), 2506-2514CODEN: AEMIDF; ISSN:1098-5336. (American Society for Microbiology)
    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%.
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  • Abstract

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    Figure 1

    Figure 1. Screening a recombinase library reveals a highly active RecT protein from the E. limosum SA11 prophage. (a) Scheme for testing recombinase library by activating a mutated ermB gene containing early stop codons. (b) Number of clarithromycin-resistant colonies obtained by transforming recombinase library or no-library control with 120 bp ssDNA recombinogenic oligos targeting ermB′. Error bars indicate standard deviation of biological triplicate transformations. (c) Proportion of clarithromycin resistant colonies, from a sample of 100 colonies each, encoding any given recombinase from conditions treated with either leading or lagging strand oligos for ermB′ correction.

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    Figure 2

    Figure 2. Oligo and process modifications affect recombineering efficiency. (a) Effect of oligo size and concentration on recombineering efficiency (% viable cells recombined). (b) Effect of oligo modifications on recombineering efficiency. “S-oligo” refers to oligos with phosphonothioate bases in the first and final three positions of the oligo, “wobble” refers to oligos encoding consecutive wobble-base codon mutations in addition to the original targeted mutation, and “generic” refers to oligos without either of these alterations. (c) Effect of recovery time on recombineering efficiency. In all cases error bars represent standard deviation of biological triplicate transformations. ND: not detected.

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    Figure 3

    Figure 3. Scheme for improved recombineering efficiency using Cas9 counterselection. Plasmid encoding RecT and two sgRNAs-one targeting a PAM site near the desired mutation and one targeting the RecT plasmid-is transformed into E. limosum expressing BgaB for a black phenotype. That strain is again transformed with Cas9 plasmid and recombinogenic oligos targeting inactivation of BgaB and selected on tetracycline. Expression of Cas9 kills unrecombined colonies and destroys the recombinase plasmid. Finally, surviving colonies are subcultured to cure the Cas9 plasmid resulting in a strain with a markerless and scarless genomic point mutation to inactivate BgaB, leaving cells with a white phenotype.

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    Figure 4

    Figure 4. Cas9 counterselection boosts proportion of viable recombined cells to 100%. Error bars represent standard deviation of biological triplicates. (a) Recombination efficiency as determined by colony color, (b) representative 96-well plates for colonies picked from condition treated with RecT and recombinogenic oligos but lacking Cas9 and sgRNA (left) and condition with Cas9 and sgRNA (right). Arrows on the left plate indicate recombined strains. Every colony on the right plate lacks BgaB activity. (c) Number of colonies obtained on tetracycline and nonselective agar plates showing that on the order of hundreds of recombined colonies may be obtained from a transformation. (d) Colony PCR results showing removal of pEL11-BgaB (plasmid encoding RecT and sgRNAs) and pRec-Cas2 (Plasmid encoding Cas9) following counterselection and curing. ND: not detected.

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    Figure 5

    Figure 5. Recombination for biofilm abolition with Cas9 counterselection. (a) Scheme for eps gene cluster inactivation. (b) Colony PCR results for amplification across eps gene cluster region +150 bp on either side. Bands show full length gene cluster of ∼5 kb for WT strain and removal of operon for engineered strain leaving just the ∼150 bp on either side. (c) Centrifugation analysis demonstrating Δeps strain pellets at low speed (3500g). Both strains are fully suspended before centrifugation (left), but only the Δeps strain pellets after centrifugation (right).

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      There is no corresponding record for this reference.
    18. 18
      Sanford, P. A.; Miller, K. G.; Hoyt, K. O.; Woolston, B. M. Deletion of biofilm synthesis in Eubacterium limosum ATCC 8486 improves handling and transformation efficiency. FEMS Microbiol. Lett. 2023, 370, fnad030  DOI: 10.1093/femsle/fnad030
      There is no corresponding record for this reference.
    19. 19
      Vees, C. A.; Neuendorf, C. S.; Pflügl, S. Towards continuous industrial bioprocessing with solventogenic and acetogenic clostridia: challenges, progress and perspectives. J. Ind. Microbiol. Biotechnol. 2020, 47 (9–10), 753– 787,  DOI: 10.1007/s10295-020-02296-2
      There is no corresponding record for this reference.
    20. 20
      Charubin, K.; Bennett, R. K.; Fast, A. G.; Papoutsakis, E. T. Engineering Clostridium organisms as microbial cell-factories: challenges & opportunities. Metab. Eng. 2018, 50, 173– 191,  DOI: 10.1016/j.ymben.2018.07.012
      20
      Engineering Clostridium organisms as microbial cell-factories: challenges & opportunities
      Charubin, Kamil; Bennett, R. Kyle; Fast, Alan G.; Papoutsakis, Eleftherios T.
      Metabolic Engineering (2018), 50 (), 173-191CODEN: MEENFM; ISSN:1096-7176. (Elsevier B.V.)
      Clostridium organisms are of major importance in the development of technologies to produce biofuels and chems. They are uniquely capable of utilizing virtually all biomass-derived carbohydrates, as well as waste gases, waste materials, and C1 compds., and they possess diverse biosynthetic capabilities for producing a broad spectrum of metabolites, including those of C4-C8 chain length. They can also be readily used in synthetic, syntrophic, and other microbial consortia to broaden the biosynthetic repertoire of individual organisms, thus enabling the development of novel biotechnol. processes. Engineering Clostridium organisms at the mol. and population level is hampered by genetic engineering, genome engineering, and microbial-population engineering tools. We discuss these challenges, and the promise that derives from their resoln. aiming to usher in an era of broader use of Clostridium organisms as biotechnol. platforms.
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    21. 21
      Sanford, P. A.; Woolston, B. M. Synthetic or natural? Metabolic engineering for assimilation and valorization of methanol. Curr. Opin. Biotechnol. 2022, 74, 171– 179,  DOI: 10.1016/j.copbio.2021.12.001
      21
      Synthetic or natural? Metabolic engineering for assimilation and valorization of methanol
      Sanford, Patrick A.; Woolston, Benjamin M.
      Current Opinion in Biotechnology (2022), 74 (), 171-179CODEN: CUOBE3; ISSN:0958-1669. (Elsevier B.V.)
      Single carbon (C1) substrates such as methanol are gaining increasing attention as cost-effective and environmentally friendly microbial feedstocks. Recent impressive metabolic engineering efforts to import C1 catabolic pathways into the non-methylotrophic bacterium Escherichia coli have led to synthetic strains growing on methanol as the sole carbon source. However, the growth rate and product yield in these strains remain inferior to native methylotrophs. Meanwhile, an ever-expanding genetic engineering toolbox is increasing the tractability of native C1 utilizers, raising the question of whether it is best to use an engineered strain or a native host for the microbial assimilation of C1 substrates. Here we provide perspective on this debate, using recent work in E. coli and the methylotrophic acetogen Eubacterium limosum as case studies.
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    22. 22
      Pines, G.; Freed, E. F.; Winkler, J. D.; Gill, R. T. Bacterial Recombineering: Genome Engineering via Phage-Based Homologous Recombination. ACS Synth. Biol. 2015, 4 (11), 1176– 1185,  DOI: 10.1021/acssynbio.5b00009
      22
      Bacterial recombineering: Genome engineering via phage-based homologous recombination
      Pines, Gur; Freed, Emily F.; Winkler, James D.; Gill, Ryan T.
      ACS Synthetic Biology (2015), 4 (11), 1176-1185CODEN: ASBCD6; ISSN:2161-5063. (American Chemical Society)
      A review. The ability to specifically modify bacterial genomes in a precise and efficient manner is highly desired in various fields, ranging from mol. genetics to metabolic engineering and synthetic biol. Much has changed from the initial realization that phage-derived genes may be employed for such tasks to today, where recombineering enables complex genetic edits within a genome or a population. Here, we review the major developments leading to recombineering becoming the method of choice for in situ bacterial genome editing while highlighting the various applications of recombineering in pushing the boundaries of synthetic biol. We also present the current understanding of the mechanism of recombineering. Finally, we discuss in detail issues surrounding recombineering efficiency and future directions for recombineering-based genome editing.
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    23. 23
      Murphy, K. C.; Campellone, K. G. Lambda Red-mediated recombinogenic engineering of enterohemorrhagic and enteropathogenic E. coli. BMC Mol. Biol. 2003, 4, 1– 12,  DOI: 10.1186/1471-2199-4-11
      There is no corresponding record for this reference.
    24. 24
      Wang, H. H.; Isaacs, F. J.; Carr, P. A. Programming cells by multiplex genome engineering and accelerated evolution. Nature 2009, 460 (7257), 894– 898,  DOI: 10.1038/nature08187
      24
      Programming cells by multiplex genome engineering and accelerated evolution
      Wang, Harris H.; Isaacs, Farren J.; Carr, Peter A.; Sun, Zachary Z.; Xu, George; Forest, Craig R.; Church, George M.
      Nature (London, United Kingdom) (2009), 460 (7257), 894-898CODEN: NATUAS; ISSN:0028-0836. (Nature Publishing Group)
      The breadth of genomic diversity found among organisms in nature allows populations to adapt to diverse environments. However, genomic diversity is difficult to generate in the lab. and new phenotypes do not easily arise on practical timescales. Although in vitro and directed evolution methods have created genetic variants with usefully altered phenotypes, these methods are limited to laborious and serial manipulation of single genes and are not used for parallel and continuous directed evolution of gene networks or genomes. Here, we describe multiplex automated genome engineering (MAGE) for large-scale programming and evolution of cells. MAGE simultaneously targets many locations on the chromosome for modification in a single cell or across a population of cells, thus producing combinatorial genomic diversity. Because the process is cyclical and scalable, we constructed prototype devices that automate the MAGE technol. to facilitate rapid and continuous generation of a diverse set of genetic changes (mismatches, insertions, deletions). We applied MAGE to optimize the 1-deoxy-d-xylulose-5-phosphate (DXP) biosynthesis pathway in Escherichia coli to overproduce the industrially important isoprenoid lycopene. Twenty-four genetic components in the DXP pathway were modified simultaneously using a complex pool of synthetic DNA, creating over 4.3 billion combinatorial genomic variants per day. We isolated variants with more than fivefold increase in lycopene prodn. within 3 days, a significant improvement over existing metabolic engineering techniques. Our multiplex approach embraces engineering in the context of evolution by expediting the design and evolution of organisms with new and improved properties.
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    25. 25
      Kolodner, R.; Hall, S. D.; Luisi-DeLuca, C. Homologous pairing proteins encoded by the Escherichia coli recE and recT genes. Mol. Microbiol. 1994, 11 (1), 23– 30,  DOI: 10.1111/j.1365-2958.1994.tb00286.x
      There is no corresponding record for this reference.
    26. 26
      Pyne, M. E.; Moo-Young, M.; Chung, D. A.; Chou, C. P. Coupling the CRISPR/Cas9 system with lambda red recombineering enables simplified chromosomal gene replacement in Escherichia coli. Appl. Environ. Microbiol. 2015, 81 (15), 5103– 5114,  DOI: 10.1128/AEM.01248-15
      26
      Coupling the CRISPR/Cas9 system with lambda Red recombineering enables simplified chromosomal gene replacement in Escherichia coli
      Pyne, Michael E.; Moo-Young, Murray; Chung, Duane A.; Chou, C. Perry
      Applied and Environmental Microbiology (2015), 81 (15), 5103-5114CODEN: AEMIDF; ISSN:1098-5336. (American Society for Microbiology)
      To date, most genetic engineering approaches coupling the type II Streptococcus pyogenes clustered regularly interspaced short palindromic repeat (CRISPR)/Cas9 system to lambda Red recombineering have involved minor single nucleotide mutations. Here we show that procedures for carrying out more complex chromosomal gene replacements in Escherichia coli can be substantially enhanced through implementation of CRISPR/Cas9 genome editing. We developed a three-plasmid approach that allows not only highly efficient recombination of short single-stranded oligonucleotides but also replacement of multigene chromosomal stretches of DNA with large PCR products. By systematically challenging the proposed system with respect to the magnitude of chromosomal deletion and size of DNA insertion, we demonstrated DNA deletions of up to 19.4 kb, encompassing 19 nonessential chromosomal genes, and insertion of up to 3 kb of heterologous DNA with recombination efficiencies permitting mutant detection by colony PCR screening. Since CRISPR/Cas9-coupled recombineering does not rely on the use of chromosome- encoded antibiotic resistance, or flippase recombination for antibiotic marker recycling, our approach is simpler, less labor-intensive, and allows efficient prodn. of gene replacement mutants that are both markerless and "scar"-less.
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    27. 27
      Reisch, C. R.; Prather, K. L. J. The no-SCAR (Scarless Cas9 Assisted Recombineering) system for genome editing in Escherichia coli. Sci. Rep. 2015, 5, 15096  DOI: 10.1038/srep15096
      27
      The no-SCAR (Scarless Cas9 Assisted Recombineering) system for genome editing in Escherichia coli
      Reisch, Chris R.; Prather, Kristala L. J.
      Scientific Reports (2015), 5 (), 15096CODEN: SRCEC3; ISSN:2045-2322. (Nature Publishing Group)
      Genome engineering methods in E. coli allow for easy to perform manipulations of the chromosome in vivo with the assistance of the λ-Red recombinase system. These methods generally rely on the insertion of an antibiotic resistance cassette followed by removal of the same cassette, resulting in a two-step procedure for genomic manipulations. Here we describe a method and plasmid system that can edit the genome of E. coli without chromosomal markers. This system, known as Scarless Cas9 Assisted Recombineering (no-SCAR), uses λ-Red to facilitate genomic integration of donor DNA and double stranded DNA cleavage by Cas9 to counter select against wild-type cells. We show that point mutations, gene deletions, and short sequence insertions were efficiently performed in several genomic loci in a single-step with regards to the chromosome and did not leave behind scar sites. The single-guide RNA encoding plasmid can be easily cured due to its temp. sensitive origin of replication, allowing for iterative chromosomal manipulations of the same strain, as is often required in metabolic engineering. In addn., we demonstrate the ability to efficiently cure the second plasmid in the system by targeting with Cas9, leaving the cells plasmid-free.
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    28. 28
      Ellis, H. M.; Yu, D.; DiTizio, T.; Court, D. L. High efficiency mutagenesis, repair, and engineering of chromosomal DNA using single-stranded oligonucleotides. Proc. Natl. Acad. Sci. U.S.A. 2001, 98 (12), 6742– 6746,  DOI: 10.1073/pnas.121164898
      28
      High efficiency mutagenesis, repair, and engineering of chromosomal DNA using single-stranded oligonucleotides
      Ellis, Hilary M.; Yu, Daiguan; DiTizio, Tina; Court, Donald L.
      Proceedings of the National Academy of Sciences of the United States of America (2001), 98 (12), 6742-6746CODEN: PNASA6; ISSN:0027-8424. (National Academy of Sciences)
      Homologous DNA recombination is a fundamental, regenerative process within living organisms. However, in most organisms, homologous recombination is a rare event, requiring a complex set of reactions and extensive homol. We demonstrate in this paper that Beta protein of phage λ generates recombinants in chromosomal DNA by using synthetic single-stranded DNAs (ssDNA) as short as 30 bases long. This ssDNA recombination can be used to mutagenize or repair the chromosome with efficiencies that generate up to 6% recombinants among treated cells. Mechanistically, it appears that Beta protein, a Rad52-like protein, binds and anneals the ssDNA donor to a complementary single-strand near the DNA replication fork to generate the recombinant. This type of homologous recombination with ssDNA provides new avenues for studying and modifying genomes ranging from bacterial pathogens to eukaryotes. Beta protein and ssDNA may prove generally applicable for repairing DNA in many organisms.
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    29. 29
      Wannier, T. M.; Nyerges, A.; Kuchwara, H. M. Improved bacterial recombineering by parallelized protein discovery. Proc. Natl. Acad. Sci. U.S.A. 2020, 117 (24), 13689– 13698,  DOI: 10.1073/pnas.2001588117
      29
      Improved bacterial recombineering by parallelized protein discovery
      Wannier, Timothy M.; Nyerges, Akos; Kuchwara, Helene M.; Czikkely, Marton; Balogh, David; Filsinger, Gabriel T.; Borders, Nathaniel C.; Gregg, Christopher J.; Lajoie, Marc J.; Rios, Xavier; Pal, Csaba; Church, George M.
      Proceedings of the National Academy of Sciences of the United States of America (2020), 117 (24), 13689-13698CODEN: PNASA6; ISSN:0027-8424. (National Academy of Sciences)
      Exploiting bacteriophage-derived homologous recombination processes has enabled precise, multiplex editing of microbial genomes and the construction of billions of customized genetic variants in a single day. The techniques that enable this, multiplex automated genome engineering (MAGE) and directed evolution with random genomic mutations (DIvERGE), are however, currently limited to a handful of microorganisms for which single-stranded DNA-annealing proteins (SSAPs) that promote efficient recombineering have been identified. Thus, to enable genome-scale engineering in new hosts, efficient SSAPs must first be found. Here the authors introduce a high-throughput method for SSAP discovery that the authors call 'serial enrichment for efficient recombineering' (SEER). By performing SEER in Escherichia coli to screen hundreds of putative SSAPs, the authors identify highly active variants PapRecT and CspRecT. CspRecT increases the efficiency of single-locus editing to as high as 50% and improves multiplex editing by 5-10-fold in E. coli, while PapRecT enables efficient recombineering in Pseudomonas aeruginosa, a concerning human pathogen. CspRecT and PapRecT are also active in other, clin. and biotechnol. relevant enterobacteria. The authors envision that the deployment of SEER in new species will pave the way toward pooled interrogation of genotype-to-phenotype relations in previously intractable bacteria.
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    30. 30
      Datta, S.; Costantino, N.; Zhou, X.; Court, D. L. Identification and analysis of recombineering functions from Gram-negative and Gram-positive bacteria and their phages. Proc. Natl. Acad. Sci. U.S.A. 2008, 105 (5), 1626– 1631,  DOI: 10.1073/pnas.0709089105
      30
      Identification and analysis of recombineering functions from Gram-negative and Gram-positive bacteria and their phages
      Datta, Simanti; Costantino, Nina; Zhou, Xiaomei; Court, Donald L.
      Proceedings of the National Academy of Sciences of the United States of America (2008), 105 (5), 1626-1631CODEN: PNASA6; ISSN:0027-8424. (National Academy of Sciences)
      We report the identification and functional anal. of nine genes from Gram-pos. and Gram-neg. bacteria and their phages that are similar to lambda (λ) bet or Escherichia coli recT. Beta and RecT are single-strand DNA annealing proteins, referred to here as recombinases. Each of the nine other genes when expressed in E. col carries out oligonucleotide-mediated recombination. To our knowledge, this is the first study showing single-strand recombinase activity from diverse bacteria. Similar to bet and recT, most of these other recombinases were found to be assocd. with putative exonuclease genes. Beta and RecT in conjunction with their cognate exonucleases carry out recombination of linear double-strand DNA. Among four of these foreign recombinase/exonuclease pairs tested for recombination with double-strand DNA, three had activity, albeit barely detectable. Thus, although these recombinases can function in E. col to catalyze oligonucleotide recombination, the double-strand DNA recombination activities with their exonuclease partners were inefficient. This study also demonstrated that Gam, by inhibiting host RecBCD nuclease activity, helps to improve the efficiency of λ Red-mediated recombination with linear double-strand DNA, but Gam is not absolutely essential. Thus, in other bacterial species where Gam analogs have not been identified, double-strand DNA recombination may still work in the absence of a Gam-like function. We anticipate that at least some of the recombineering systems studied here will potentiate oligonucleotide and double-strand DNA-mediated recombineering in their native or related bacteria.
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    31. 31
      Noirot, P.; Kolodner, R. D. DNA strand invasion promoted by Escherichia coli RecT protein. J. Biol. Chem. 1998, 273 (20), 12274– 12280,  DOI: 10.1074/jbc.273.20.12274
      There is no corresponding record for this reference.
    32. 32
      Roca, A.; Cox, M.; Brenner, S. The RecA Protein: Structure and Function. Crit. Rev. Biochem. Mol. Biol. 1990, 25 (6), 415– 456,  DOI: 10.3109/10409239009090617
      There is no corresponding record for this reference.
    33. 33
      Dong, H.; Tao, W.; Gong, F.; Li, Y.; Zhang, Y. A functional recT gene for recombineering of Clostridium. J. Biotechnol. 2014, 173 (1), 65– 67,  DOI: 10.1016/j.jbiotec.2013.12.011
      There is no corresponding record for this reference.
    34. 34
      Swingle, B.; Bao, Z.; Markel, E.; Chambers, A.; Cartinhour, S. Recombineering using recTE from Pseudomonas syringae. Appl. Environ. Microbiol. 2010, 76 (15), 4960– 4968,  DOI: 10.1128/AEM.00911-10
      34
      Recombineering using RecTE from Pseudomonas syringae
      Swingle, Bryan; Bao, Zhongmeng; Markel, Eric; Chambers, Alan; Cartinhour, Samuel
      Applied and Environmental Microbiology (2010), 76 (15), 4960-4968CODEN: AEMIDF; ISSN:0099-2240. (American Society for Microbiology)
      In this report, we describe the identification of functions that promote genomic recombination of linear DNA introduced into Pseudomonas cells by electroporation. The genes encoding these functions were identified in Pseudomonas syringae pv. syringae B728a based on similarity to the lambda Red Exo/Beta and RecET proteins encoded by the lambda and Rac bacteriophages of Escherichia coli. The ability of the pseudomonad-encoded proteins to promote recombination was tested in P. syringae pv. tomato DC3000 using a quant. assay based on recombination frequency. The results show that the Pseudomonas RecT homolog is sufficient to promote recombination of single-stranded DNA oligonucleotides and that efficient recombination of double-stranded DNA requires the expression of both the RecT and RecE homologs. Addnl., we illustrate the utility of this recombineering system to make targeted gene disruptions in the P. syringae chromosome.
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    35. 35
      Arndt, D.; Grant, J. R.; Marcu, A. PHASTER: a better, faster version of the PHAST phage search tool. Nucleic Acids Res. 2016, 44 (W1), W16– W21,  DOI: 10.1093/nar/gkw387
      35
      PHASTER: a better, faster version of the PHAST phage search tool
      Arndt, David; Grant, Jason R.; Marcu, Ana; Sajed, Tanvir; Pon, Allison; Liang, Yongjie; Wishart, David S.
      Nucleic Acids Research (2016), 44 (W1), W16-W21CODEN: NARHAD; ISSN:0305-1048. (Oxford University Press)
      PHASTER (PHAge Search Tool - Enhanced Release)is a significant upgrade to the popular PHAST web server for the rapid identification and annotation of prophage sequences within bacterial genomes and plasmids. Although the steps in the phage identification pipeline in PHASTER remain largely the same as in the original PHAST, numerous software improvements and significant hardware enhancements have now made PHASTER faster, more efficient, more visually appealing and much more user friendly. In particular, PHASTER is now 4.3× faster than PHAST when analyzing a typical bacterial genome. Mores pecifically, software optimizations have made the backend of PHASTER 2.7X faster than PHAST, while the addn. of 80 CPUs to the PHASTER compute cluster are responsible for the remaining speed up. PHASTER can now process a typical bacterial genome in 3 min from the raw sequence alone, or in1.5 min when given a pre-annotated GenBank file. Anumber of other optimizations have also been implemented, including automated algorithms to reduce the size and redundancy of PHASTER's databases, improvements in handling multiple (meta genomic)queries and higher user traffic, along with the ability to perform automated look-ups against 14000 previously PHAST/PHASTER annotated bacterial genomes (which can lead to complete phage annotations in seconds as opposed to minutes). PHASTER's web interface has also been entirely rewritten. A new graphical genome browser has been added, gene/genome visualization tools have been improved, and the graphical interface is now more modern, robust and user-friendly.
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    36. 36
      Canchaya, C.; Proux, C.; Fournous, G.; Bruttin, A.; Brüssow, H. Prophage Genomics. Microbiol. Mol. Biol. Rev. 2003, 67 (3), 473,  DOI: 10.1128/MMBR.67.3.473.2003
      There is no corresponding record for this reference.
    37. 37
      Dong, H.; Tao, W.; Zhang, Y.; Li, Y. Development of an anhydrotetracycline-inducible gene expression system for solvent-producing Clostridium acetobutylicum: A useful tool for strain engineering. Metab. Eng. 2012, 14 (1), 59– 67,  DOI: 10.1016/j.ymben.2011.10.004
      37
      Development of an anhydrotetracycline-inducible gene expression system for solvent-producing Clostridium acetobutylicum: A useful tool for strain engineering
      Dong, Hongjun; Tao, Wenwen; Zhang, Yanping; Li, Yin
      Metabolic Engineering (2012), 14 (1), 59-67CODEN: MEENFM; ISSN:1096-7176. (Elsevier B. V.)
      Clostridium acetobutylicum is an important solvent (acetone-butanol-ethanol) producing bacterium. However, a stringent, effective, and convenient-to-use inducible gene expression system that can be used for regulating the gene expression strength in C. acetobutylicum is currently not available. Here, we report an anhydrotetracycline-inducible gene expression system for solvent-producing bacterium C. acetobutylicum. This system consists of a functional chloramphenicol acetyltransferase gene promoter contg. tet operators (tetO), Pthl promoter (thiolase gene promoter from C. acetobutylicum) controlling TetR repressor expression cassette, and the chem. inducer anhydrotetracycline (aTc). The optimized system, designated as pGusA2-2tetO1, allows gene regulation in an inducer aTc concn.-dependent way, with an inducibility of over two orders of magnitude. The stringency of TetR repression supports the introduction of the genes encoding counterselective marker into C. acetobutylicum, which can be used to increase the mutant screening efficiency. This aTc-inducible gene expression system will thus increase the genetic manipulation capability for engineering C. acetobutylicum.
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    38. 38
      Aparicio, T.; de Lorenzo, V.; Martínez-García, E. CRISPR/Cas9-enhanced ssDNA recombineering for Pseudomonas putida. Microb. Biotechnol. 2019, 12 (5), 1076– 1089,  DOI: 10.1111/1751-7915.13453
      38
      CRISPR/Cas9-enhanced ssDNA recombineering for Pseudomonas putida
      Aparicio, Tomas; de Lorenzo, Victor; Martinez-Garcia, Esteban
      Microbial Biotechnology (2019), 12 (5), 1076-1089CODEN: MBIIB2; ISSN:1751-7915. (Wiley-Blackwell)
      Summary : Implementation of single-stranded DNA (ssDNA) recombineering in Pseudomonas putida has widened the range of genetic manipulations applicable to this biotechnol. relevant bacterium. Yet, the relatively low efficiency of the technol. hampers identification of mutated clones lacking conspicuous phenotypes. Fortunately, the use of CRISPR/Cas9 as a device for counterselection of wild-type sequences helps to overcome this limitation. Merging ssDNA recombineering with CRISPR/Cas9 thus enables a suite of genomic edits with a straightforward approach: a CRISPR plasmid provides the spacer DNA sequence that directs the Cas9 nuclease ribonucleoprotein complex to cleave the genome at the wild-type sequences that have not undergone the change entered by the mutagenic ssDNA oligonucleotide(s). This protocol describes a complete workflow of the method optimized for P. putida, although it could in principle be applicable to many other pseudomonads. As an example, we show the deletion of the edd gene that encodes one key enzyme that operates the EDEMP cycle for glucose metab. in P. putidaEM42. By combining two incompatible CRISPR plasmids with different antibiotic selection markers, we show that the procedure can be cycled to implement consecutive deletions in the same strain, e.g. deletion of the pyrF gene following that of the edd mutant. This approach adds to the wealth of genetic technologies available for P. putida and strengthens its status as a chassis of choice for a suite of biotechnol. applications.
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    39. 39
      Wyman, C.; Ristic, D.; Kanaar, R. Homologous recombination-mediated double-strand break repair. DNA Repair 2004, 3 (8–9), 827– 833,  DOI: 10.1016/j.dnarep.2004.03.037
      There is no corresponding record for this reference.
    40. 40
      Sawitzke, J. A.; Costantino, N.; Li, X. t. Probing cellular processes with oligo-mediated recombination and using the knowledge gained to optimize recombineering. J. Mol. Biol. 2011, 407 (1), 45– 59,  DOI: 10.1016/j.jmb.2011.01.030
      There is no corresponding record for this reference.
    41. 41
      Nakayama, M.; Ohara, O. Improvement of Recombination Efficiency by Mutation of Red Proteins. Biotechniques 2005, 38 (6), 917– 924,  DOI: 10.2144/05386RR02
      There is no corresponding record for this reference.
    42. 42
      Ding, Y.; Wang, K. F.; Wang, W. J. Increasing the homologous recombination efficiency of eukaryotic microorganisms for enhanced genome engineering. Appl. Microbiol. Biotechnol. 2019, 103 (11), 4313– 4324,  DOI: 10.1007/s00253-019-09802-2
      There is no corresponding record for this reference.
    43. 43
      Ren, J.; Karna, S.; Lee, H. M.; Yoo, S. M.; Na, D. Artificial transformation methodologies for improving the efficiency of plasmid DNA transformation and simplifying its use. Appl. Microbiol. Biotechnol. 2019, 103 (23–24), 9205– 9215,  DOI: 10.1007/s00253-019-10173-x
      There is no corresponding record for this reference.
    44. 44
      Dutra, B. E.; Sutera, V. A.; Lovett, S. T. RecA-independent recombination is efficient but limited by exonucleases. Proc. Natl. Acad. Sci. U.S.A. 2007, 104 (1), 216– 221,  DOI: 10.1073/pnas.0608293104
      There is no corresponding record for this reference.
    45. 45
      Spitzer, S.; Eckstein, F. Inhibition of deoxyribonucleases by phosphorothioate groups in oligodeoxyribonucleotides. Nucleic Acids Res. 1988, 16 (24), 11691– 11704,  DOI: 10.1093/nar/16.24.11691
      45
      Inhibition of deoxyribonucleases by phosphorothioate groups in oligodeoxyribonucleotides
      Spitzer, Stephan; Eckstein, Fritz
      Nucleic Acids Research (1988), 16 (24), 11691-704CODEN: NARHAD; ISSN:0305-1048.
      The Rp- and Sp-diastereomers of the phosphorothiolate-contg. oligonucleotide d[ApAp(S)ApA] (I) were synthesized. They and the tetramer d[ApApApA] were tested as substrates for staphylococcal nuclease, DNase II, and spleen phosphodiesterase. For digestions with DNase I these oligonucleotides were converted to the 5'-phosphorylated derivs. The reactions with the nucleases were analyzed by HPLC. The phosphorothioate groups of both diastereomers were resistant to the action of staphylococcal nuclease, DNase I, and DNase II. While the phosphorothioate group of the Rp-diastereomer was resistant to the action of spleen phosphodiesterase, the Sp-diastereomer was hydrolyzed at an estd. rate 1/100 the rate of cleavage of the unmodified tetramer. The presence of the phosphorothiolate group in the center of the mol. affected the rate of hydrolysis of neighboring phosphate groups for some enzymes. In particular, very slow release of 3'-dAMP from the Rp-diastereomer occurred on incubation with staphylococcal nuclease but the Sp-diastereomer was completely resistant. DNase II produced 3'-dAMP quite rapidly from both diastereomers of I and DNase I released 5'-dAMP from both diastereomers of I only slowly.
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    46. 46
      Jiang, Y.; Chen, B.; Duan, C.; Sun, B.; Yang, J.; Yang, S. Multigene editing in the Escherichia coli genome via the CRISPR-Cas9 system. Appl. Environ. Microbiol. 2015, 81 (7), 2506– 2514,  DOI: 10.1128/AEM.04023-14
      46
      Multigene editing in the Escherichia coli genome via the CRISPR-Cas9 system
      Jiang, Yu; Chen, Biao; Duan, Chunlan; Sun, Bingbing; Yang, Junjie; Yang, Sheng
      Applied and Environmental Microbiology (2015), 81 (7), 2506-2514CODEN: AEMIDF; ISSN:1098-5336. (American Society for Microbiology)
      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%.
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    47. 47
      Thomason, L. C.; Costantino, N.; Li, X.; Court, D. L. Recombineering: Genetic Engineering in Escherichia coli Using Homologous Recombination. Curr. Protoc. 2023, 3, e656  DOI: 10.1002/cpz1.656
      There is no corresponding record for this reference.
    48. 48
      Yuan, J.; Martinez-Bilbao, M.; Huber, R. E. Substitutions for Glu-537 of β-galactosidase from Escherichia coli cause large decreases in catalytic activity. Biochem. J. 1994, 299 (2), 527– 531,  DOI: 10.1042/bj2990527
      48
      Substitutions for Glu-537 of β-galactosidase from Escherichia coli cause large decreases in catalytic activity
      Yuan, Jingming; Martinez-Bilbao, Mercedes; Huber, Reuben E.
      Biochemical Journal (1994), 299 (2), 527-31CODEN: BIJOAK; ISSN:0264-6021.
      Glu-537 of β-galactosidase (EC 3.2.1.23) was replaced by Asp, Gln, and Val using synthetic oligonucleotides. The kcat values of the purified enzyme mixts. were reduced by ∼100-fold for the Asp mutant, 30,000-60,000-fold for the Val mutant, and 160,000-300,000-fold for the Gln mutant. The greatest differences in properties from the wild-type enzyme were found for the Asp-substituted enzyme: the Km values increased (from 0.12 to 0.42 mM for o-nitrophenyl β-D-galactopyranoside and from 0.04 to 0.37 mM for p-nitrophenyl β-D-galactopyranoside); the Ki for iso-Pr β-D-galactopyranoside increased (from 0.11 to 0.30 mM), the stability to heat decreased, and MeOH did not act as an acceptor. The enzymes with the other 2 substitutions had properties similar to those of the wild-type enzyme. For all 3 substituted enzymes, the inhibitory effects of the transition-state analogs (2-deoxy-2-amino-D-galactose and L-ribose) and the Mg2+ effects were similar to those of the normal enzyme. As all of the properties (except the kcat values) of the Gln- and Val-substituted enzyme prepns. were similar to those of the wild-type enzyme, the activities in those prepns. were probably due to the presence of a few wild-type enzyme mols. (formed from misreads) among the substituted enzymes. The enzymes with Gln and Val substitutions appeared to be totally inactive. The results obtained supported a recent suggestion that Glu-537 is an important catalytic residue of β-galactosidase.
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    49. 49
      Morrison, K. L.; Weiss, G. A. Combinatorial alanine-scanning. Curr. Opin. Chem. Biol. 2001, 5 (3), 302– 307,  DOI: 10.1016/S1367-5931(00)00206-4
      49
      Combinatorial alanine-scanning
      Morrison, Kim L.; Weiss, Gregory A.
      Current Opinion in Chemical Biology (2001), 5 (3), 302-307CODEN: COCBF4; ISSN:1367-5931. (Elsevier Science Ltd.)
      A review with 37 refs. Combinatorial libraries of alanine-substituted proteins can be used to rapidly identify residues important for protein function, stability and shape. Each alanine substitution examines the contribution of an individual amino acid side chain to the functionality of the protein. The recently described method of shotgun scanning uses phage-displayed libraries of alanine-substituted proteins for high-throughput anal.
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    50. 50
      Loubiere, P.; Gros, E.; Paquet, V.; Lindley, N. D. Kinetics and physiological implications of the growth behaviour of Eubacterium limosum on glucose/methanol mixtures. J. Gen. Microbiol. 1992, 138 (5), 979– 985,  DOI: 10.1099/00221287-138-5-979
      There is no corresponding record for this reference.
    51. 51
      Heap, J. T.; Pennington, O. J.; Cartman, S. T.; Minton, N. P. A modular system for Clostridium shuttle plasmids. J. Microbiol. Methods 2009, 78 (1), 79– 85,  DOI: 10.1016/j.mimet.2009.05.004
      51
      A modular system for Clostridium shuttle plasmids
      Heap, John T.; Pennington, Oliver J.; Cartman, Stephen T.; Minton, Nigel P.
      Journal of Microbiological Methods (2009), 78 (1), 79-85CODEN: JMIMDQ; ISSN:0167-7012. (Elsevier B.V.)
      Despite their medical and industrial importance, our basic understanding of the biol. of the genus Clostridium is rudimentary in comparison to their aerobic counterparts in the genus Bacillus. A major contributing factor has been the comparative lack of sophistication in the gene tools available to the clostridial mol. biologist, which are immature, and in clear need of development. The transfer and maintenance of recombinant, replicative plasmids into various species of Clostridium has been reported, and several elements suitable as shuttle plasmid components are known. However, these components have to-date only been available in disparate plasmid contexts, and their use has not been broadly explored. Here we describe the specification, design and construction of a standardized modular system for Clostridium-Escherichia coli shuttle plasmids. Existing replicons and selectable markers were incorporated, along with a novel clostridial replicon. The properties of these components were compared, and the data allow researchers to identify combinations of components potentially suitable for particular hosts and applications. The system has been extensively tested in our lab., where it is utilized in all ongoing recombinant work. We propose that adoption of this modular system as a std. would be of substantial benefit to the Clostridium research community, whom we invite to use and contribute to the system.
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    52. 52
      Waterhouse, A.; Bertoni, M.; Bienert, S. SWISS-MODEL: Homology modelling of protein structures and complexes. Nucleic Acids Res. 2018, 46 (W1), W296– W303,  DOI: 10.1093/nar/gky427
      52
      SWISS-MODEL: homology modelling of protein structures and complexes
      Waterhouse, Andrew; Bertoni, Martino; Bienert, Stefan; Studer, Gabriel; Tauriello, Gerardo; Gumienny, Rafal; Heer, Florian T.; de Beer, Tjaart A. P.; Rempfer, Christine; Bordoli, Lorenza; Lepore, Rosalba; Schwede, Torsten
      Nucleic Acids Research (2018), 46 (W1), W296-W303CODEN: NARHAD; ISSN:1362-4962. (Oxford University Press)
      Homol. modeling has matured into an important technique in structural biol., significantly contributing to narrowing the gap between known protein sequences and exptl. detd. structures. Fully automated workflows and servers simplify and streamline the homol. modeling process, also allowing users without a specific computational expertise to generate reliable protein models and have easy access to modeling results, their visualization and interpretation. Here, we present an update to the SWISS-MODEL server, which pioneered the field of automated modeling 25 years ago and been continuously further developed. Recently, its functionality has been extended to the modeling of homo- and heteromeric complexes. Starting from the amino acid sequences of the interacting proteins, both the stoichiometry and the overall structure of the complex are inferred by homol. modeling. Other major improvements include the implementation of a new modeling engine, ProMod3 and the introduction a new local model quality estn. method, QMEANDisCo.
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    53. 53
      Viborg, A. H.; Fredslund, F.; Katayama, T. A β1–6/β1–3 galactosidase from Bifidobacterium animalis subsp. lactisBl-04 gives insight into sub-specificities of β-galactoside catabolism within Bifidobacterium. Mol. Microbiol. 2014, 94 (5), 1024– 1040,  DOI: 10.1111/mmi.12815
      There is no corresponding record for this reference.

  • Supporting Information

    Supporting Information

    The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acssynbio.4c00253.

    • Tables of the recombinases, plasmids, oligos, and guide RNAs used in the study, along with enzyme assay data, sequencing, plating, and colony PCR data to further validate results presented in the main text (PDF)

    • Nucleotide sequences for the recombinase library members (XLSX)


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