ACS Synthetic Biology

Synthetic biology is as an excellent vehicle for education, as it enables creative combination of engineering and molecular biology approaches for quantitative characterisations of the assembled constructs. However, there is a limited number of resources available for such applications in the educational context, where straightforward setup, easily measurable phenotypes and extensibility are of particular importance. To expand the availability of education-friendly resources to teach synthetic biology and genetic engineering, we developed Unigems, a set of 10 plasmids that enable out-of-the-box investigations of principles of gene expression control, as well as more complex designs a biological logic gate. The system uses a common high-copy plasmid backbone and a common set of primers to enable Gibson-assembly of PCR-generated or synthesised parts into a target vector. It currently has two reporter genes with either two constitutive (high- or low-level) or two inducible (lactose- or arabinose-) promoters, as well as a single-plasmid implementation of an AND logic gate. The Unigems system has already been employed in undergraduate teaching settings, during outreach events and for training of iGEM teams. All plasmids have been deposited in Addgene.

Characterizing and cataloging genetic parts are critical to the design of useful genetic circuits. Having well-characterized parts allows for the fine-tuning of genetic circuits, such that their function results in predictable outcomes. With the growth of synthetic biology as a field, there has been an explosion of genetic circuits that have been implemented in microbes to execute functions pertaining to sensing, metabolic alteration, and cellular computing. Here, we show a cost-effective and rapid method for characterizing genetic parts. Our method utilizes cell-free lysate, prepared in-house, as a medium to evaluate parts via the expression of a reporter protein. Template DNA is prepared by PCR-amplification using inexpensive primers to add variant parts to the reporter gene, and the template is added to the reaction as linear DNA without cloning. Parts that can be added in this way include promoters, operators, ribosome binding sites, insulators, and terminators. This approach, combined with the incorporation of an acoustic liquid handler and 384-well plates, allows the user to carry out high-throughput evaluations of genetic parts in a single day. By comparison, cell-based screening approaches require time-consuming cloning and have longer testing times due to overnight culture and culture density normalization steps. Further, working in cell-free lysate allows the user to exact tighter control over the expression conditions through the addition of exogenous components, or by titrating DNA concentrations rather than relying on limited plasmid copy numbers. Because this method retains a cell-like environment, the function of the genetic part will typically mimic its function in whole cells. SUMMARYWell-characterized genetic parts are necessary for the design of novel genetic circuits. Here we describe a cost-effective, high-throughput method for rapidly characterizing genetic parts. Our method reduces cost and time by combining cell-free lysates, linear DNA to avoid cloning, and acoustic liquid handling to increase throughput and reduce reaction volumes.

Quorum sensing-based genetic circuits are gaining traction in synthetic biology as they link population-level behaviour to individual cell responses. However, tuning these circuits remains challenging due to complex dynamics, particularly during the Learn phase of the Design-Build-Test-Learn (DBTL) cycle. To accelerate this process, we developed a mathematical model to predict how varying expression levels of the transcription factor and synthase affect the response of the EsaI/EsaR quorum sensing system. A strain library was constructed, and experimental data were used to optimize the model. The final model could successfully differentiate between the effects of these expression levels on the response of the bidirectional promoter. It allowed visualization of all potential system outcomes and emphasized the transcription factors critical role in tuning the circuit. This model offers a valuable tool for fine-tuning EsaI/EsaR-based systems for synthetic biology applications. Moreover, given the homology within the LuxR-family quorum sensing systems, this modelling approach may serve as a foundation for model-based tuning of other quorum sensing systems.

Azospirillum brasilense is an important plant-growth promoting bacterium found in the rhizosphere and used extensively in commercial agriculture. It is an attractive candidate for use in the development of novel crop interventions, such as the targeted delivery of plant hormones and similar compounds to the roots, shoots or leaves, or the introduction of smart bacterial sensors and actuators in the rhizosphere. In order to be able to engineer A. brasilense with these novel and complex behaviours, we require a collection of predictable and reliable genetic parts, consisting of constitutive and inducible promoters, terminators, and a stable expression plasmid. To date, such a genetic toolkit does not exist for the genus. In this work, for the first time we have designed and tested a synthetic constitutive promoter library with a wide-range of transcriptional strengths, a tightly-regulated inducible promoter with a non-metabolisable inducer, strong Rho-independent transcriptional terminators and synthetic small RNAs for post-transcriptional regulation.. We have adapted them for use in a one-pot assembly method and demonstrated their utility in the overproduction of the important plant hormone, indole 3-acetic acid (IAA).

Plant synthetic biology is a fast-evolving field that employs engineering principles to empower research and bioproduction in plant systems. Nevertheless, in the whole synthetic biology landscape, plant systems lag compared to microbial and mammalian systems. When it comes to multigene delivery to plants, the predictability of the outcome is decreased since it depends on three different chassis: E. coli, Agrobacterium, and the plant species. Here we aimed to develop standardised and streamlined tools for genetic engineering in plant synthetic biology. We have devised Mobius Assembly for Plant Systems (MAPS), a user-friendly Golden Gate Assembly system for fast and easy generation of complex DNA constructs. MAPS is based on a new group of small plant binary vectors (pMAPs) that contains an origin of replication from a cryptic plasmid of Paracoccus pantotrophus. The functionality of the pMAP vectors was confirmed by transforming the MM1 cell culture, demonstrating for the first time that plant transformation is dependent on the Agrobacterium strains and plasmids; plasmid stability was highly dependent on the plasmid and bacterial strain. We made a library of new short promoters and terminators and characterised them using a high-throughput protoplast expression assay. Our results underscored the strong influence of terminators in gene expression, and they altered the strength of promoters in some combinations and indicated the presence of synergistic interactions between promoters and terminators. Overall this work will further facilitate plant synthetic biology and contribute to improving its predictability, which is challenged by combinatorial interactions among the genetic parts, vectors, and chassis.

Escherichia coli Nissle 1917 (EcN) is a probiotic microbe that has the potential to be developed as a promising chassis for synthetic biology applications. However, the molecular tools and techniques for utilizing EcN have not been fully explored. To address this opportunity, we systematically expanded the EcN-based toolbox, enabling EcN as a powerful platform for more applications. First, two EcN cryptic plasmids and other compatible plasmids were genetically engineered to enrich the manipulable plasmid toolbox for multiple gene coexpression. Next, we developed two EcN-based enabling technologies, including the conjugation strategy for DNA transfer, and quantification of protein expression capability. Finally, we expanded the EcN-based applications by developing EcN native integrase-mediated genetic engineering capabilities and establishing an in vitro cell-free protein synthesis (CFPS) system. Overall, this study expanded the toolbox for manipulating EcN as a commonly used probiotic chassis, providing several simplified, dependable, and predictable strategies for researchers working in synthetic biology fields. For Table of Contents Use Only O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=133 SRC="FIGDIR/small/543671v1_ufig1.gif" ALT="Figure 1"> View larger version (27K): org.highwire.dtl.DTLVardef@1f8e91forg.highwire.dtl.DTLVardef@90fb70org.highwire.dtl.DTLVardef@6badbforg.highwire.dtl.DTLVardef@15be840_HPS_FORMAT_FIGEXP M_FIG C_FIG

Synthetic developmental biology uses engineering approaches to understand multicellularity with goals ranging from recapitulating development to building synthetic organisms. Current approaches include engineering multicellular patterning, controlling differentiation, and implementing cooperative cellular behaviors in model systems. Synthetic biology tools enable these pursuits with genetic circuits that drive customized responses to arbitrary stimuli, synthetic receptors that enable orthogonal signaling channels, and light- or drug-inducible systems that enable precise spatial and temporal control of cell function. Mouse embryonic stem cells (mESCs) offer a well-studied and genetically tractable pluripotent chassis for pursuing synthetic development questions however, there is minimal characterization of existing synthetic biology tools in mESCs and we lack genetic toolkits for rapid iterative engineering of synthetic development workflows. Here, we began to address this challenge by characterizing small molecule and cell contact-inducible systems for gene expression in and differentiation of mESCs. We show that small molecule and cell-contact inducible systems work reliably and efficiently for controlling expression of arbitrary genetic payloads. Furthermore, we show that these systems can drive direct differentiation of mESCs into neurons. Each of these systems can readily be used on their own or in combination, opening many possibilities for studying developmental principles with high precision.

Inducible gene expression systems can be used to control the expression of a gene of interest by means of a small-molecule. One of the most common designs involves engineering a small-molecule responsive transcription factor (TF) and its cognate promoter, which often results in a compromise between minimal uninduced background expression (leakiness) and maximal induced expression. Here, we focussed on an alternative strategy using quantitative synthetic biology to mitigate leakiness while maintaining high expression, without modifying neither the TF nor the promoter. Through mathematical modelling and experimental validations, we designed the CASwitch, a mammalian synthetic gene circuit based on combining two well-known network motifs: the Coherent Feed-Forward Loop (CFFL) and the Mutual Inhibition (MI). The CASwitch combines the CRISPR-Cas endoribonuclease CasRx with the state-of-the-art Tet-On3G inducible gene system to achieve high performances. To demonstrate the potentialities of the CASwitch, we applied it to three different scenarios: enhancing a whole-cell biosensor, controlling expression of a toxic gene and inducible production of Adeno-Associated Virus (AAV) vectors.

Whole-cell biosensors are emerging as promising tools for monitoring environmental pollutants such as heavy metals. These sensors constitute a genetic circuit comprising a sensing module and an output module, such that a detectable signal is produced in the presence of the desired analyte. The MerR family of metal-responsive regulators offers great potential for the construction of metal sensing circuits, due to their high sensitivity, tight transcription control and large diversity in metal-specificity. However, the sensing diversity is broadest in Gram-negative systems, while chassis organisms are often selected from Gram-positive species, particularly sporulating bacilli. This can be problematic, because Gram-negative biological parts, such as promoters, are frequently observed to be non-functional in Gram-positive hosts. Herein, we combined construction of synthetic genetic circuits and chimeric MerR regulators, supported by structure-guided design, to generate metal-sensitive biosensor modules that are functional in the biotechnological work-horse species Bacillus subtilis. These chimeras consist of a constant Gram-positive derived DNA-binding domain fused to variable metal binding domains of Gram-negative origins. To improve the specificity of the whole-cell biosensor, we developed a modular AND gate logic system based on the B. subtilis natively split {sigma}-factor, SigO-RsoA, designed to maximise future use for synthetic biology applications in B. subtilis. This work provides insights into the use of modular regulators, such as the MerR family, in the design of synthetic circuits for the detection of heavy metals, with potential wider applicability of the approach to other systems and genetic backgrounds.

Environmental isolates are promising candidates for new chassis of synthetic biology because of their inherent conversion capabilities and resilience to environmental stresses; however, many remain genetically intractable and unamenable to established genetic tools tailored for model bacteria. Acinetobacter sp. Tol 5 possesses intriguing properties for use in synthetic biology applications. However, genetic manipulation via electroporation is hindered by its low transformation efficiency. This study demonstrated the genetic refinement of the Tol 5 strain, achieving efficient transformation via electroporation. We deleted two genes encoding restriction enzymes. The resulting mutant strain not only exhibited marked efficiency of electrotransformation but also proved receptive to both in vitro and in vivo DNA assembly technologies, thereby facilitating the construction of recombinant DNA. In addition, we successfully adapted a CRISPR-Cas9-based base-editing platform developed for other Acinetobacter species. Our genetic modification strategy allows for the domestication of non-model bacteria, streamlining their utilization in synthetic biology applications.

A genomically recoded Escherichia coli strain that lacks all amber codons and release factor 1 (C321.{Delta}A) enables efficient genetic encoding of chemically diverse, non-canonical amino acids (ncAAs) into proteins. While C321.{Delta}A has opened new opportunities in chemical and synthetic biology, this strain has not been optimized for protein production, limiting its utility in widespread industrial and academic applications. To address this limitation, we describe the construction of a series of genomically recoded organisms that are optimized for cellular protein production. We demonstrate that the functional deactivation of nucleases (e.g., rne, endA) and proteases (e.g., lon) increases production of wild-type superfolder green fluorescent protein (sfGFP) and sfGFP containing two ncAAs up to [~]5-fold. Additionally, we introduce a genomic IPTG-inducible T7 RNA polymerase (T7RNAP) cassette into these strains. Using an optimized platform, we demonstrated the ability to introduce 2 identical N6-(propargyloxycarbonyl)-L-Lysine residues site specifically into sfGFP with a 17-fold improvement in production relative to the parent. We envision that our library of organisms will provide the community with multiple options for increased expression of proteins with new and diverse chemistries.

Synthetic biology offers powerful tools to engineer biological systems for diverse applications. However, key challenges persists before achieving real-world applications like environmental bioremediation or therapeutic microrobots for targeted drug delivery. This study aimed to precisely control bacterial movement by modulating gene expression using engineered promoters in Escherichia coli. We focused on Escherichia coli, a model organism, and manipulated its motility by engineering the expression of flagellin, a crucial protein for bacterial movement. To achieve this, specific genetic promoters were employed to regulate the production of flagellin, thereby dictating the movement capabilities of these bacteria. The promoters enabled targeted adjustments to flagellin expression, which in turn allowed for the enhancement or suppression of bacterial locomotion. Interestingly, the relationship between promoter design parameters and gene expression levels was non-linear, highlighting complex underlying dynamics. Optimal bacterial motility occurred at 30{degrees}C, illustrating the influence of environmental factors. Our findings demonstrate the ability to effectively regulate complex microbial phenotypes like motility using genetic engineering strategies. The results not only extend our understanding of bacterial gene regulation but also highlight the transformative potential of synthetic biology in creating functional and adaptable microbial phenotypes for diverse biotechnological applications. O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=99 SRC="FIGDIR/small/635866v1_ufig1.gif" ALT="Figure 1"> View larger version (53K): org.highwire.dtl.DTLVardef@1fca66dorg.highwire.dtl.DTLVardef@12529dorg.highwire.dtl.DTLVardef@ede03aorg.highwire.dtl.DTLVardef@11c2c6c_HPS_FORMAT_FIGEXP M_FIG C_FIG

DNA modifying enzymes are ubiquitous reagents in synthetic biology. Producing these enzymes often requires large culture volumes, purified nucleases and chromatographic separations to make enzymes of necessary quality. We sought to leverage synthetic biology tools to develop engineered strains allowing for not only the production but rapid purification of these reagents. Toward this goal, we report an E. coli strain enabling the rapid production and purification of Taq polymerase. The method relies on 1) autoinducible expression achieving high protein titers, 2) autolysis and auto DNA/RNA hydrolysis via lysozyme and a mutant benzonase, and 3) heat denaturation under reducing conditions to precipitate contaminating proteins including the mutant benzonase. Taq polymerase is obtained at high purities (>95% pure by SDS-PAGE) and is readily usable in standard reactions. The method takes less than 1 hour of hands-on time, does not require special equipment, expensive reagents or affinity purification. We expect this simple methodology and approach will improve access not only to Taq polymerase but to numerous additional commonly utilized reagent proteins. HighlightsO_LIProtein titers ~ 1g/L achieved in 20 mL shakeflasks. C_LIO_LI4 mg of purified Taq (corresponding to 5,000 Units, or 4,000 PCR reactions) per 20 mL shake flask. C_LIO_LIInstant Taq is equivalent to commercial preparations in routine PCR C_LI

The development of CRISPR-Cas-based engineering technologies has revolutionized the microbial biotechnology field. Over the years, the Class II Type II CRISPR-Cas9 system has become the gold standard for genome editing in many bacterial hosts. However, the Cas9 system does not allow efficient genomic integration in Pseudomonas putida, an emerging Synthetic Biology host, without the assistance of lambda-Red recombineering. In this work, we utilize the alternative Class I Type I-C CRISPR-Cas3 system from Pseudomonas aeruginosa as a highly-efficient genome editing tool for P. putida and P. aeruginosa. This system consists of two vectors, one encoding the Cas genes, CRISPR array and targeting spacer, and a second SEVA-vector, containing the homologous repair template. Both vectors are Golden Gate compatible for rapid cloning and are available with multiple antibiotic markers, for application in various Gram-negative hosts and different designs. By employing this Cas3 system, we successfully integrated an 820-bp cassette in the genome of P. putida and performed several genomic deletions in P. aeruginosa within four days, with an efficiency of >83% for both hosts. Moreover, by introducing a universal self-targeting spacer, the Cas3 system rapidly cures all helper vectors, including itself, from the host strain in a matter of days. As such, this system constitutes a valuable engineering tool for Pseudomonas, to complement the existing range of Cas9-based editing methods and facilitates genomic engineering efforts of this important genus. ImportanceThe CRISPR-Cas3 editing system as presented here facilitates the creation of genomic alterations in P. putida and P. aeruginosa in a straightforward manner. By providing the Cas3 system as a vector set with Golden Gate compatibility and different antibiotic markers, as well as by employing the established SEVA vector set to provide the homology repair template, this system is flexible and can readily be ported to a multitude of Gram-negative hosts. Besides genome editing, the Cas3 system can also be used as an effective and universal tool for vector curing. This is achieved by introducing a spacer that targets the oriT, present on the majority of established (SEVA) vectors. Based on this, the Cas3 system efficiently removes up to three vectors in only a few days. As such, this curing approach may also benefit other genomic engineering methods or remove naturally-occurring plasmids from bacteria.

Diatoms produce 20% of the worlds fixed organic carbon annually, making them vital to global carbon fixation and climate change mitigation. Their potential as cell factories for biofuels, proteins, and other high value chemicals remains underutilized due to a lack of genetic engineering tools, with DNA delivery being one of the biggest challenges. Here, we present optimized electroporation and polyethylene glycol transformation methods for delivering DNA and ribonucleoprotein complexes to Phaeodactylum tricornutum, a model diatom species and emerging chassis for algal biotechnology. It was possible to recover transformants with as little as 1 ng of DNA, and to transform linear or circular episomes as large as 55.6 kb. With the optimized electroporation protocol, episomes can be assembled in the algal cell de novo through diatom in vivo assembly (DIVA), forgoing the need for time-consuming traditional cloning steps in Escherichia coli and Saccharomyces cerevisiae. It was also possible to electroporate a Cas9 ribonucleoprotein complex in P. tricornutum, providing an alternative to biolistics for DNA free genome engineering. We have demonstrated that the PEG approach can be adapted to successfully transform Thalassiosira pseudonana, demonstrating the applicability of our methods for engineering other diatom species. These tools can be used to accelerate diatom synthetic biology projects and, therefore, the development of sustainable technologies.

The discovery of new genes regulating essential biological processes has become increasingly important, and CRISPRi has emerged as a powerful tool for achieving this goal. This method has been used in many model organisms to decrease the expression of specific genes and assess their impact on phenotype. Pooled CRISPRi libraries in bacteria have been particularly useful in discovering new regulators of growth, division, and other biological processes. However, these libraries rely on the induction of dCas9 via an inducible promoter, which can be problematic due to promoter leakiness. This is a widespread phenomenon of any inducible promoter that can result in the unwanted downregulation of genes and the emergence of genetic suppressors when essential genes are knocked down. To overcome this issue, we have developed a novel strategy that eliminates dCas9 leakiness and enables reversible knockdown control using the rapamycin-dependent degron system in Bacillus subtilis. This degron system causes rapid degradation of dCas9, resulting in an almost instant reset of the system. Our results demonstrate that it is possible to achieve zero CRISPRi activity in the uninduced state and full activity in the induced state. This improved CRISPRi system will enable researchers to investigate phenotypic changes more effectively while reducing the undesirable effects of leaky expression and noise in their phenotypic data. Moreover, a rapid degradation system could serve as a tool for dynamic perturbation before compensation mechanisms or stress responses kick in. Finally, this approach can be adapted to other organisms and other promoter-inducible systems, potentially opening up strategies for tighter control of gene expression.

A universal biochemical signal for bacterial cell-cell communication could facilitate programming dynamic responses in diverse bacterial consortia. However, the classical quorum sensing paradigm is that gram-negative and gram-positive bacteria generally communicate via homoserine lactones (HSL) or oligopeptide molecular signals, respectively, to elicit population responses. Here, we create synthetic HSL sensors for gram-positive Bacillus subtilis 168 using allosteric LuxR-type regulators (RpaR, LuxR, RhlR, and CinR) and synthetic promoters. Promoters were combinatorially designed from different sequence elements (-35, -16, -10, and transcriptional start regions). We quantified the effects of these combinatorial promoters on sensor activity and determined how regulator expression affects its activation, achieving up to 293-fold activation. Using statistical design of experiments, we identified significant effects of promoter regions and pairwise interactions on sensor activity, which helped to understand the sequence-function relationships for synthetic promoter design. We present the first known set of functional HSL sensors ([≥] 20-fold dynamic range) in B. subtilis for four different HSL chemical signals: p-coumaroyl-HSL, 3-oxohexanoyl-HSL, n-butyryl-HSL, and n-(3-hydroxytetradecanoyl)-HSL. This set of synthetic HSL sensors for a gram-positive bacterium can pave the way for designable interspecies communication within microbial consortia.

Recently, a new subset of fluorescent proteins has been identified from the Aequorea species of jellyfish. These fluorescent proteins were characterized in vivo; however, there has not been validation of these proteins within cell-free systems. Cell-free systems and technology development is a rapidly expanding field, encompassing foundational research, synthetic cells, bioengineering, biomanufacturing and drug development. Cell-free systems rely heavily on fluorescent proteins as reporters. Here we characterize and validate this new set of Aequorea proteins for use in a variety of cell-free and synthetic cell expression platforms. O_FIG O_LINKSMALLFIG WIDTH=144 HEIGHT=200 SRC="FIGDIR/small/519681v1_ufig1.gif" ALT="Figure 1"> View larger version (41K): org.highwire.dtl.DTLVardef@3a3100org.highwire.dtl.DTLVardef@672233org.highwire.dtl.DTLVardef@f65f32org.highwire.dtl.DTLVardef@e40aaf_HPS_FORMAT_FIGEXP M_FIG C_FIG

The engineering of genetic circuits to perform predefined computations is central to synthetic biology, enabling living cells with new functionalities applicable across various domains. However, these circuits are often specifically tailored to particular cellular hosts, with Escherichia coli being the most popular. Consequently, their intended functions may not translate well to other organisms, limiting their scope. Understanding circuit dynamics in less familiar organisms is crucial, especially for niche-specific applications requiring cellular chassis different from model organisms generally used in synthetic biology. Here, we develop a combined experimental and theoretical pipeline to evaluate the performance of NOT logic circuits, also called inverters, in the soil bacterium Pseudomonas protegens Pf-5--a host renowned for its unique environmental functions and a newcomer to genetic circuitry. Inverters were experimentally tested to characterize input-output functionality, and mathematical modelling was used to infer the dynamic principles of circuit modules. The model quantified the individual impacts of key parameters--such as translation efficiency, repressor performance, and promoter activity--on output levels, enabling predictions about inter-host circuit portability. This parameter calibration revealed unique properties of the chassis, including faster transitions between on and off circuit states compared to the synthetic biology workhorse Pseudomonas putida. These characteristics may reflect adaptations to the fluctuating conditions of the plant rhizosphere, where this bacteria thrives. As a result, our work provides DNA parts, circuits and mathematical characterizations to establish P. protegens Pf-5 as a viable chassis for environmental synthetic biology.

Transposons have been instrumental tools in microbiology enabling random mutagenesis, with transposons like Tn5 and Mariner, and site-specific DNA integrations with Tn7. However, programmable targeting of transposons was impossible until CRISPR-associated transposase (CasTn) systems were described. Like other CRISPR-derived systems, CasTn can be programmed with a short DNA encoded sequence that is transcribed into a guide-RNA. Here we describe a broad-host-range CasTn system and demonstrate its function in bacteria from three classes of the Proteobacteria. The CasTn genes are expressed from a broad-host-range replicative plasmid, while the guide-RNA and transposon are provided on a high-copy pUC plasmid that is suicidal in most bacteria outside of E. coli. Using our CasTn system, single-gene disruptions were performed with on-target efficiencies approaching 100% in the Beta- and Gammaproteobacteria, Burkholderia thailandensis, and Pseudomonas putida, respectively. The results were more modest in the Alphaproteobacterium Agrobacterium fabrum, with a peak efficiency of 45%, though for routine single-gene disruptions, this efficiency is adequate. In B. thailandensis, the system allowed simultaneous co-integration of transposons at two different target sites. The CasTn system is also capable of high-efficiency large transposon insertion totaling over 11 kbp in P. putida. Given the iterative capabilities and large payload size, this system will be helpful for genome engineering experiments across several fields of research. SignificanceThe genetic modification of bacteria to disrupt native genes and integrate recombinant genes is necessary for basic and applied research. Traditional methods for targeted disruptions and insertions are often cumbersome and inefficient, limiting experiments' scale and throughput. This work developed a system for targeted transposon mutagenesis that is easy to use, iterative, and efficient. We demonstrate that the system functions across three different classes of the Proteobacteria in species widely used in research and biotechnology. Moreover, the framework of the system and accompanying plasmids that we developed will facilitate porting the system to other bacteria. Our system provides a fast and efficient protocol to genetically modify these bacteria by inserting desired genetic cargo into specific genomic targets.