Independent and tight regulation of transcriptional units in Escherichia coli via the LacR/O, the TetR/O and AraC/I1-I2 regulatory elements

Soluble versions of outer membrane cytochromes function as exporters for heterologously produced cargo proteins

This study reveals that it is possible to secrete truncated versions of outer membrane cytochromes into the culture supernatant and that these proteins can provide a basis for the export of heterologously produced proteins. Different soluble and truncated versions of the outer membrane cytochrome MtrF were analyzed for their suitability to be secreted. A protein version with a very short truncation of the N-terminus to remove the recognition sequence for the addition of a lipid anchor is secreted efficiently to the culture supernatant, and moreover this protein could be further truncated by a deletion of 160 amino acid and still is detectable in the supernatant. By coupling a cellulase to this soluble outer membrane cytochrome, the export efficiency was measured by means of relative cellulase activity. We conclude that outer membrane cytochromes of S. oneidensis can be applied as transporters for the export of target proteins into the medium using the type II secretion pathway.

Design, Fabrication, and Device Chemistry of a 3-Input-3-Output Synthetic Genetic Combinatorial Logic Circuit with a 3-Input AND Gate in a Single Bacterial Cell

Advancement of in-cell molecular computation requires multi-input-multi-output genetic logic devices. However, increased physical size, a higher number of molecular interactions, cross-talk, and complex systems level device chemistry limited the realization of such multi-input-multi-output devices in a single bacterial cell. Here, by adapting a circuit minimization and conjugated promoter engineering approach, we created the first 3-input-3-output logic function in a single bacterial cell. The circuit integrated three extracellular chemical signals as inputs and produced three different fluorescent proteins as outputs following the truth table of the circuit. First, we created a noncascaded 1-gate-3-input synthetic genetic AND gate in bacteria. We showed that the 3-input AND gate was digital in nature and mathematically predictable, two important characteristics, which were not reported for previous 3-input AND gates in bacteria. Our design consists of a 128 bp DNA scaffold, which conjugated various protein-binding sites in a single piece of DNA and worked as a hybrid promoter. The scaffold was a few times smaller than the similar 3-input synthetic genetic AND gate promoter reported. Integrating this AND gate with a new 2-input-2-output integrated circuit, which was also digital-like and predictive, we created a 3-input-3-output combinatorial logic circuit. This work demonstrated the integration of a 3-input AND gate in a larger circuit and a 3-input-3-output synthetic genetic circuit, both for the first time. The work has significance in molecular computation, biorobotics, DNA nanotechnology, and synthetic biology.

Engineering DNA recognition and allosteric response properties of TetR family proteins by using a module-swapping strategy

The development of synthetic biological systems requires modular biomolecular components to flexibly alter response pathways. In previous studies, we have established a module-swapping design principle to engineer allosteric response and DNA recognition properties among regulators in the LacI family, in which the engineered regulators served as effective components for implementing new cellular behavior. Here we introduced this protein engineering strategy to two regulators in the TetR family: TetR (UniProt Accession ID: P04483) and MphR (Q9EVJ6). The TetR DNA-binding module and the MphR ligand-binding module were used to create the TetR-MphR. This resulting hybrid regulator possesses DNA-binding properties of TetR and ligand response properties of MphR, which is able to control gene expression in response to a molecular signal in cells. Furthermore, we studied molecular interactions between the TetR DNA-binding module and MphR ligand-binding module by using mutant analysis. Together, we demonstrated that TetR family regulators contain discrete and functional modules that can be used to build biological components with novel properties. This work highlights the utility of rational design as a means of creating modular parts for cell engineering and introduces new possibilities in rewiring cellular response pathways.

Engineering repressors with coevolutionary cues facilitates toggle switches with a master reset

Engineering allosteric transcriptional repressors containing an environmental sensing module (ESM) and a DNA recognition module (DRM) has the potential to unlock a combinatorial set of rationally designed biological responses. We demonstrated that constructing hybrid repressors by fusing distinct ESMs and DRMs provides a means to flexibly rewire genetic networks for complex signal processing. We have used coevolutionary traits among LacI homologs to develop a model for predicting compatibility between ESMs and DRMs. Our predictions accurately agree with the performance of 40 engineered repressors. We have harnessed this framework to develop a system of multiple toggle switches with a master OFF signal that produces a unique behavior: each engineered biological activity is switched to a stable ON state by different chemicals and returned to OFF in response to a common signal. One promising application of this design is to develop living diagnostics for monitoring multiple parameters in complex physiological environments and it represents one of many circuit topologies that can be explored with modular repressors designed with coevolutionary information.

Control of ϕC31 integrase-mediated site-specific recombination by protein trans-splicing

Serine integrases are emerging as core tools in synthetic biology and have applications in biotechnology and genome engineering. We have designed a split-intein serine integrase-based system with potential for regulation of site-specific recombination events at the protein level in vivo. The ϕC31 integrase was split into two extein domains, and intein sequences (Npu DnaEN and Ssp DnaEC) were attached to the two termini to be fused. Expression of these two components followed by post-translational protein trans-splicing in Escherichia coli generated a fully functional ϕC31 integrase. We showed that protein splicing is necessary for recombination activity; deletion of intein domains or mutation of key intein residues inactivated recombination. We used an invertible promoter reporter system to demonstrate a potential application of the split intein-regulated site-specific recombination system in building reversible genetic switches. We used the same split inteins to control the reconstitution of a split Integrase-Recombination Directionality Factor fusion (Integrase-RDF) that efficiently catalysed the reverse attR x attL recombination. This demonstrates the potential for split-intein regulation of the forward and reverse reactions using the integrase and the integrase-RDF fusion, respectively. The split-intein integrase is a potentially versatile, regulatable component for building synthetic genetic circuits and devices.

CRIMoClo plasmids for modular assembly and orthogonal chromosomal integration of synthetic circuits in Escherichia coli

Background

Synthetic biology heavily depends on rapid and simple techniques for DNA engineering, such as Ligase Cycling Reaction (LCR), Gibson assembly and Golden Gate assembly, all of which allow for fast, multi-fragment DNA assembly. A major enhancement of Golden Gate assembly is represented by the Modular Cloning (MoClo) system that allows for simple library propagation and combinatorial construction of genetic circuits from reusable parts. Yet, one limitation of the MoClo system is that all circuits are assembled in low- and medium copy plasmids, while a rapid route to chromosomal integration is lacking. To overcome this bottleneck, here we took advantage of the conditional-replication, integration, and modular (CRIM) plasmids, which can be integrated in single copies into the chromosome of Escherichia coli and related bacteria by site-specific recombination at different phage attachment (att) sites.

Results

By combining the modularity of the MoClo system with the CRIM plasmids features we created a set of 32 novel CRIMoClo plasmids and benchmarked their suitability for synthetic biology applications. Using CRIMoClo plasmids we assembled and integrated a given genetic circuit into four selected phage attachment sites. Analyzing the behavior of these circuits we found essentially identical expression levels, indicating orthogonality of the loci. Using CRIMoClo plasmids and four different reporter systems, we illustrated a framework that allows for a fast and reliable sequential integration at the four selected att sites. Taking advantage of four resistance cassettes the procedure did not require recombination events between each round of integration. Finally, we assembled and genomically integrated synthetic ECF σ factor/anti-σ switches with high efficiency, showing that the growth defects observed for circuits encoded on medium-copy plasmids were alleviated.

Conclusions

The CRIMoClo system enables the generation of genetic circuits from reusable, MoClo-compatible parts and their integration into 4 orthogonal att sites into the genome of E. coli. Utilizing four different resistance modules the CRIMoClo system allows for easy, fast, and reliable multiple integrations. Moreover, utilizing CRIMoClo plasmids and MoClo reusable parts, we efficiently integrated and alleviated the toxicity of plasmid-borne circuits. Finally, since CRIMoClo framework allows for high flexibility, it is possible to utilize plasmid-borne and chromosomally integrated circuits simultaneously. This increases our ability to permute multiple genetic modules and allows for an easier design of complex synthetic metabolic pathways in E. coli.

Directed evolution of the PcaV allosteric transcription factor to generate a biosensor for aromatic aldehydes

Background

Transcription factor-based biosensors are useful tools for the detection of metabolites and industrially valuable molecules, and present many potential applications in biotechnology and biomedicine. However, the most common approach to develop biosensors relies on employing a limited set of naturally occurring allosteric transcription factors (aTFs). Therefore, altering the ligand specificity of aTFs towards the detection of new effectors is an important goal.

Results

Here, the PcaV repressor, a member of the MarR aTF family, was used to develop a biosensor for the detection of hydroxyl-substituted benzoic acids, including protocatechuic acid (PCA). The PCA biosensor was further subjected to directed evolution to alter its ligand specificity towards vanillin and other closely related aromatic aldehydes, to generate the Van2 biosensor. Ligand recognition of Van2 was explored in vitro using a range of biochemical and biophysical analyses, and extensive in vivo genetic-phenotypic analysis was performed to determine the role of each amino acid change upon biosensor performance.

Conclusions

This is the first study to report directed evolution of a member of the MarR aTF family, and demonstrates the plasticity of the PCA biosensor by altering its ligand specificity to generate a biosensor for aromatic aldehydes.

Design and Control of Extrachromosomal Elements in Methylorubrum extorquens AM1

Genetic tools are a prerequisite to engineer cellular factories for synthetic biology and biotechnology. Methylorubrum extorquens AM1 is an important platform organism of a future C₁-bioeconomy. However, its application is currently limited by the availability of genetic tools. Here we systematically tested repABC regions to maintain extrachromosomal DNA in M. extorquens. We used three elements to construct mini-chromosomes that are stably inherited at single copy number and can be shuttled between Escherichia coli and M. extorquens. These mini-chromosomes are compatible among each other and with high-copy number plasmids of M. extorquens. We also developed a set of inducible promoters of wide expression range, reaching levels exceeding those currently available, notably the P_mxaF-promoter. In summary, we provide a set of tools to control the dynamic expression and copy number of genetic elements in M. extorquens, which opens new ways to unleash the metabolic and biotechnological potential of this organism for future applications.

The Biogenesis of SRP RNA Is Modulated by an RNA Folding Intermediate Attained during Transcription

The signal recognition particle (SRP), responsible for co-translational protein targeting and delivery to cellular membranes, depends on the native long-hairpin fold of its RNA to confer functionality. Since RNA initiates folding during its synthesis, we used high-resolution optical tweezers to follow in real time the co-transcriptional folding of SRP RNA. Surprisingly, SRP RNA folding is robust to transcription rate changes and the presence or absence of its 5'-precursor sequence. The folding pathway also reveals the obligatory attainment of a non-native hairpin intermediate (H1) that eventually rearranges into the native fold. Furthermore, H1 provides a structural platform alternative to the native fold for RNase P to bind and mature SRP RNA co-transcriptionally. Delays in attaining the final native fold are detrimental to the cell, altogether showing that a co-transcriptional folding pathway underpins the proper biogenesis of function-essential SRP RNA.

The serine transporter SdaC prevents cell lysis upon glucose depletion in Escherichia coli

The amino acid serine plays diverse metabolic roles, yet bacteria actively degrade exogenously provided serine via deamination to pyruvate. Serine deamination is thought to be a detoxification mechanism due to the ability of serine to inhibit several biosynthetic reactions, but this pathway remains highly active even in nutrient-replete conditions. While investigating the physiological roles of serine deamination in different growth conditions, we discovered that Escherichia coli cells lacking the sdaCB operon, which encodes the serine transporter SdaC and the serine deaminase SdaB, lyse upon glucose depletion in a medium containing no exogenous serine but all other amino acids and nucleobases. Unexpectedly, this lysis phenotype can be recapitulated by deleting sdaC alone and can be rescued by heterologous expression of SdaC. Lysis of ΔsdaC cells can be prevented by omitting glycine from the medium, inhibiting the glycine cleavage system, or by increasing alanine availability. Together, our results reveal that the serine transporter SdaC plays a critical role in maintaining amino acid homeostasis during shifts in nutrient availability in E. coli.

Stimulus response-based fine-tuning of polyhydroxyalkanoate pathway in Halomonas

Optimization of intracellular biosynthesis process involving regulation of multiple gene expressions is dependent on the efficient and accurate expression of each expression unit independently. However, challenges of analyzing intermediate products seriously hinder the application of high throughput assays. This study aimed to develop an engineering approach for unsterile production of poly(3-hydroxybutyrate-co-4-hydroxybutyrate) or (P3HB4HB) by recombinant Halomonas bluephagenesis (H. bluephagenesis) constructed via coupling the design of GFP-mediated transcriptional mapping and high-resolution control of gene expressions (HRCGE), which consists of two inducible systems with high- and low-dynamic ranges employed to search the exquisite transcription level of each expression module in the presence of γ-butyrolactone, the intermediate for 4-hydroxybutyrate (4HB) synthesis. It has been successful to generate a recombinant H. bluephagenesis, namely TD68-194, able to produce over 36 g/L P3HB4HB consisting of 16 mol% 4HB during a 7-L lab-scale fed-batch growth process, of which cell dry weight and PHA content reached up to 48.22 g/L and 74.67%, respectively, in 36 h cultivation. HRCGE has been found useful for metabolic pathway construction.

BldD, a master developmental repressor, activates antibiotic production in two Streptomyces species

BldD generally functions as a repressor controlling morphological development of Streptomyces. In this work, evidences that BldD also activates antibiotic production are provided. In Streptomyces roseosporus (which produces daptomycin widely used for treatment of human infections), deletion of bldD notably reduced daptomycin production, but enhanced sporulation. BldD stimulated daptomycin production by directly activating transcription of dpt structural genes and dptR3 (which encodes an indirect activator of daptomycin production), and repressed its own gene. BldD-binding sites on promoter regions of dptE, dptR3, and bldD were all found to contain BldD box-like sequences, facilitating prediction of new BldD targets. Two Streptomyces global regulatory genes, adpA and afsR, were confirmed to be directly activated by BldD. The protein AfsR was shown to act as an activator of daptomycin production, but a repressor of development. BldD directly represses nine key developmental genes. In Streptomyces avermitilis (which produces effective anthelmintic agents avermectins), BldD homolog (BldDsav) directly activates avermectin production through ave structural genes and cluster-situated activator gene aveR. This is the first report that BldD activates antibiotic biosynthesis both directly and via a cascade mechanism. BldD homologs are widely distributed among Streptomyces, our findings suggest that BldD may activate antibiotic production in other Streptomyces species.

Construction of Two-Input Logic Gates Using Transcriptional Interference

Transcriptional interference (TI) has been shown to regulate gene expression at the DNA level via different molecular mechanisms. The obstacles present on the DNA that a transcribing RNA polymerase might encounter, for example, a DNA-bound protein or another RNA polymerase, can result in TI causing termination of transcription, thus reducing gene expression. However, the potential of TI as a new strategy to engineer complex gene expression modules has not been fully explored yet. Here we created a series of two-input genetic devices that use the presence of a roadblocking protein to control gene expression and analyzed their behaviors using both experimental and mathematical modeling approaches. We explored how multiple characteristics affect the response of genetic devices engineered to act like either AND, OR, or single input logic gates. We show that the dissociation constant of the roadblocking protein, inducer activation of promoter and operator sites, and distance between tandem promoters tune gate behavior. This work highlights the potential of rationally creating different types of genetic responses using the same transcription factors in subtly different genetic architectures.

Exploring the Potential of Cell-Free Protein Synthesis for Extending the Abilities of Biological Systems

Cell-free protein synthesis (CFPS) system is a simple, rapid, and sensitive tool that is devoid of membrane-bound barriers, yet contains all the mandatory substrates, biomolecules, and machineries required for the synthesis of the desired proteins. It has the potential to overcome loopholes in the current in vivo production systems and is a promising tool in both basic and applied scientific research. It facilitates a simplified organization of desired experiments with a variety of reaction conditions, making CFPS a powerful tool in biological research. It has been used for the expansion of genetic code, assembly of viruses, and in metabolic engineering for production of toxic and complex proteins. Subsequently, CFPS systems have emerged as potent technology for high-throughput production of membrane proteins, enzymes, and therapeutics. The present review highlights the recent advances and uses of CFPS systems in biomedical, therapeutic, and biotechnological applications. Additionally, we highlight possible solutions to the potential biosafety issues that may be encountered while using CFPS technology.

Adaption of an Episomal Antisense Silencing Approach for Investigation of the Phenotype Switch of Staphylococcus aureus Small-Colony Variants

Staphylococcus aureus small-colony variants (SCVs) are associated with chronic, persistent, and relapsing courses of infection and are characterized by slow growth combined with other phenotypic and molecular traits. Although certain mechanisms have been described, the genetic basis of clinical SCVs remains often unknown. Hence, we adapted an episomal tool for rapid identification and investigation of putative SCV phenotype-associated genes via antisense gene silencing based on previously described Tnl0-encoded tet-regulatory elements. Targeting the SCV phenotype-inducing enoyl-acyl-carrier-protein reductase gene (fabI), plasmid pSN1-AS‘fabI’ was generated leading to antisense silencing, which was proven by pronounced growth retardation in liquid cultures, phenotype switch on solid medium, and 200-fold increase of antisense ‘fabI’ expression. A crucial role of TetR repression in effective regulation of the system was demonstrated. Based on the use of anhydrotetracycline as effector, an easy-to-handle one-plasmid setup was set that may be applicable to different S. aureus backgrounds and cell culture studies. However, selection of the appropriate antisense fragment of the target gene remains a critical factor for effectiveness of silencing. This inducible gene expression system may help to identify SCV phenotype-inducing genes, which is prerequisite for the development of new antistaphylococcal agents and future alternative strategies to improve treatment of therapy-refractory SCV-related infections by iatrogenically induced phenotypic switch. Moreover, it can be used as controllable phenotype switcher to examine important aspects of SCV biology in cell culture as well as in vivo.

Development of a Vanillate Biosensor for the Vanillin Biosynthesis Pathway in E. coli

The engineered de novo vanillin biosynthesis pathway constructed in Escherichia coli is industrially relevant but limited by the reaction catalyzed by catechol O-methyltransferase, which is intended to catalyze the conversion of protocatechuate to vanillate. To identify alternative O-methyltransferases, we constructed a vanillate sensor based on the Caulobacter crescentus VanR-VanO system. Using an E. coli promoter library, we achieved greater than 14-fold dynamic range in our best rationally constructed sensor. We found that this construct and an evolved variant demonstrate remarkable substrate selectivity, exhibiting no detectable response to the regioisomer byproduct isovanillate and minimal response to structurally similar pathway intermediates. We then harnessed the evolved biosensor to conduct rapid bioprospecting of natural catechol O-methyltransferases and identified three previously uncharacterized but active O-methyltransferases. Collectively, these efforts enrich our knowledge of how biosensing can aid metabolic engineering and constitute the foundation for future improvements in vanillin pathway productivity.

Functional metagenomics-guided discovery of potent Cas9 inhibitors in the human microbiome

CRISPR-Cas systems protect bacteria and archaea from phages and other mobile genetic elements, which use small anti-CRISPR (Acr) proteins to overcome CRISPR-Cas immunity. Because Acrs are challenging to identify, their natural diversity and impact on microbial ecosystems are underappreciated. To overcome this discovery bottleneck, we developed a high-throughput functional selection to isolate ten DNA fragments from human oral and fecal metagenomes that inhibit Streptococcus pyogenes Cas9 (SpyCas9) in Escherichia coli. The most potent Acr from this set, AcrIIA11, was recovered from a Lachnospiraceae phage. We found that AcrIIA11 inhibits SpyCas9 in bacteria and in human cells. AcrIIA11 homologs are distributed across diverse bacteria; many distantly-related homologs inhibit both SpyCas9 and a divergent Cas9 from Treponema denticola. We find that AcrIIA11 antagonizes SpyCas9 using a different mechanism than other previously characterized Type II-A Acrs. Our study highlights the power of functional selection to uncover widespread Cas9 inhibitors within diverse microbiomes.

Gene networks that compensate for crosstalk with crosstalk

Crosstalk is a major challenge to engineering sophisticated synthetic gene networks. A common approach is to insulate signal-transduction pathways by minimizing molecular-level crosstalk between endogenous and synthetic genetic components, but this strategy can be difficult to apply in the context of complex, natural gene networks and unknown interactions. Here, we show that synthetic gene networks can be engineered to compensate for crosstalk by integrating pathway signals, rather than by pathway insulation. We demonstrate this principle using reactive oxygen species (ROS)-responsive gene circuits in Escherichia coli that exhibit concentration-dependent crosstalk with non-cognate ROS. We quantitatively map the degree of crosstalk and design gene circuits that introduce compensatory crosstalk at the gene network level. The resulting gene network exhibits reduced crosstalk in the sensing of the two different ROS. Our results suggest that simple network motifs that compensate for pathway crosstalk can be used by biological networks to accurately interpret environmental signals.

Crosstalk between genetic circuits is a major challenge for engineering sophisticated networks. Here the authors design networks that compensate for crosstalk by integrating, not insulating, pathways.

Regulation of bistability in the std fimbrial operon of Salmonella enterica by DNA adenine methylation and transcription factors HdfR, StdE and StdF

Bistable expression of the Salmonella enterica std operon is controlled by an AND logic gate involving three transcriptional activators: the LysR-type factor HdfR and the StdE and StdF regulators encoded by the std operon itself. StdE activates transcription of the hdfR gene, and StdF activates std transcription together with HdfR. Binding of HdfR upstream of the std promoter is hindered by methylation of GATC sites located within the upstream activating sequence (UAS). Epigenetic control by Dam methylation thus antagonizes formation of the StdE-StdF-HdfR loop and tilts the std switch toward the StdOFF state. In turn, HdfR binding hinders methylation of the UAS, permitting activation of the StdE-StdF-HdfR loop and concomitant formation of StdON cells. Bistability is thus the outcome of competition between DNA adenine methylation and the StdE-StdF-HdfR activator loop.

Start-Stop Assembly: a functionally scarless DNA assembly system optimized for metabolic engineering

DNA assembly allows individual DNA constructs or libraries to be assembled quickly and reliably. Most methods are either: (i) Modular, easily scalable and suitable for combinatorial assembly, but leave undesirable 'scar' sequences; or (ii) bespoke (non-modular), scarless but less suitable for construction of combinatorial libraries. Both have limitations for metabolic engineering. To overcome this trade-off we devised Start-Stop Assembly, a multi-part, modular DNA assembly method which is both functionally scarless and suitable for combinatorial assembly. Crucially, 3 bp overhangs corresponding to start and stop codons are used to assemble coding sequences into expression units, avoiding scars at sensitive coding sequence boundaries. Building on this concept, a complete DNA assembly framework was designed and implemented, allowing assembly of up to 15 genes from up to 60 parts (or mixtures); monocistronic, operon-based or hybrid configurations; and a new streamlined assembly hierarchy minimizing the number of vectors. Only one destination vector is required per organism, reflecting our optimization of the system for metabolic engineering in diverse organisms. Metabolic engineering using Start-Stop Assembly was demonstrated by combinatorial assembly of carotenoid pathways in Escherichia coli resulting in a wide range of carotenoid production and colony size phenotypes indicating the intended exploration of design space.

Recent advances in plasmid-based tools for establishing novel microbial chassis

A key challenge for domesticating alternative cultivable microorganisms with biotechnological potential lies in the development of innovative technologies. Within this framework, a myriad of genetic tools has flourished, allowing the design and manipulation of complex synthetic circuits and genomes to become the general rule in many laboratories rather than the exception. More recently, with the development of novel technologies such as DNA automated synthesis/sequencing and powerful computational tools, molecular biology has entered the synthetic biology era. In the beginning, most of these technologies were established in traditional microbial models (known as chassis in the synthetic biology framework) such as Escherichia coli and Saccharomyces cerevisiae, enabling fast advances in the field and the validation of fundamental proofs of concept. However, it soon became clear that these organisms, although extremely useful for prototyping many genetic tools, were not ideal for a wide range of biotechnological tasks due to intrinsic limitations in their molecular/physiological properties. Over the last decade, researchers have been facing the great challenge of shifting from these model systems to non-conventional chassis with endogenous capacities for dealing with specific tasks. The key to address these issues includes the generation of narrow and broad host plasmid-based molecular tools and the development of novel methods for engineering genomes through homologous recombination systems, CRISPR/Cas9 and other alternative methods. Here, we address the most recent advances in plasmid-based tools for the construction of novel cell factories, including a guide for helping with "build-your-own" microbial host.

An Adaptive Synthetic Cell Based on Mechanosensing, Biosensing, and Inducible Gene Circuits

The bottom-up assembly of synthetic cell systems capable of recapitulating biological functions has become a means to understand living matter by construction. The integration of biomolecular components into active, cell-sized, genetically programmed compartments remains, however, a major bottleneck for building synthetic cells. A primary feature of real cells is their ability to actively interact with their surroundings, particularly in stressed conditions. Here, we construct a synthetic cell equipped with an inducible genetic circuit that responds to changes in osmotic pressure through the mechanosensitive channel MscL. Liposomes loaded with an E. coli cell-free transcription-translation (TXTL) system are induced with IPTG when exposed to hypo-osmotic solution, resulting in the expression of a bacterial cytoskeletal protein MreB. MreB associates with the membrane to generate a cortex-like structure. Our work provides the first example of molecular integration that couples mechanosensitivity, gene expression, and self-assembly at the inner membrane of synthetic cells.

Targeting 16S rDNA for Stable Recombinant Gene Expression in Pseudomonas

Ribosomal RNA (rRNA) operons have recently been identified as promising sites for chromosomal integration of genetic elements in Pseudomonas putida, a bacterium that has gained considerable popularity as a microbial cell factory. We have developed a tool for targeted integration of recombinant genes into the rRNA operons of various Pseudomonas strains, where the native context of the rRNA clusters enables effective transcription of heterologous genes. However, a sufficient translation of foreign mRNA  transcriptionally fused to rRNA required optimization of RNA secondary structures, which was achieved utilizing synthetic ribozymes and a bicistronic design. The generated tool further enabled the characterization of the six rRNA promoter units of P. putida S12 under different growth conditions. The presence of multiple, almost identical rRNA operons in Pseudomonas also allowed the integration of multiple copies of heterologous genetic elements. The integration of two expression cassettes and the resulting disruption of rRNA units only moderately affects growth rates, and the constructs were highly stable over more than 160 generations.

Developing a pyruvate-driven metabolic scenario for growth-coupled microbial production

Microbial-based chemical synthesis serves as a promising approach for sustainable production of industrially important products. However, limited production performance caused by metabolic burden or genetic variations poses one of the major challenges in achieving an economically viable biomanufacturing process. To address this issue, one superior strategy is to couple the product synthesis with cellular growth, which renders production obligatory for cell survival. Here we create a pyruvate-driven metabolic scenario in engineered Escherichia coli for growth-coupled bioproduction, with which we demonstrate its application in boosting production of anthranilate and its derivatives. Deletion of a minimal set of endogenous pyruvate-releasing pathways engenders anthranilate synthesis as the salvage route for pyruvate generation to support cell growth, concomitant with simultaneous anthranilate production. Further introduction of native and non-native downstream pathways affords production enhancement of two anthranilate-derived high-value products including L-tryptophan and cis, cis-muconic acid from different carbon sources. The work reported here presents a new growth-coupled strategy with demonstrated feasibility for promoting microbial production.

An Engineered B. subtilis Inducible Promoter System with over 10 000-Fold Dynamic Range

Bacillus subtilis is the leading model Gram-positive bacterium, and a widely used chassis for industrial protein production. However, B. subtilis research is limited by a lack of inducible promoter systems with low leakiness and high dynamic range. Here, we engineer an inducible promoter system based on the T7 RNA Polymerase (T7 RNAP), the lactose repressor LacI, and the chimeric promoter P_T7lac, integrated as a single copy in the B. subtilis genome. In the absence of IPTG, LacI strongly represses T7 RNAP and P_T7lac and minimizes leakiness. Addition of IPTG derepresses P_T7lac and simultaneously induces expression of T7RNAP, which results in very high output expression. Using green fluorescent and β-galactosidase reporter proteins, we estimate that this LacI-T7 system can regulate expression with a dynamic range of over 10 000, by far the largest reported for an inducible B. subtilis promoter system. Furthermore, LacI-T7 responds to similar IPTG concentrations and with similar kinetics as the widely used P_hy-spank IPTG-inducible system, which we show has a dynamic range of at most 300 in a similar genetic context. Due to its superior performance, our LacI-T7 system should have broad applications in fundamental B. subtilis biology studies and biotechnology.

Synthetic symbiosis combining plasmid displacement enables rapid construction of phenotype-stable strains

Plasmid-based microbial systems have been a major workhorse for chemical and pharmaceutical production. The biosafety issues and elevated industrial cost of antibiotic usage have led to the development of alternative strategies for plasmid selection and maintenance. Such strategies, including auxotrophy complementation, post-segregational killing, operator-repressor and RNA-based interactions often require extensive engineering of various elements and may result in extra metabolic burden in the cells. Herein, we report a design of synthetic symbiosis combining plasmid displacement to construct a phenotype-stable microbial system. By sequestrating an endogenous essential gene folP, cells obtained long-term plasmid maintenance with minimum cost. The phenotype performance was also inherited for up to 80 generations demonstrated by the production of salicylic acid in Escherichia coli. Meanwhile, the temperature-induced curing method of the intermediate plasmids enables rapid engineering. This design can lead to broad applications as a reliable and convenient plasmid-based expression system.

The emergence of the two cell fates and their associated switching for a negative auto-regulating gene

Background

Decisions in the cell that lead to its ultimate fate are important for fundamental cellular functions such as proliferation, growth, differentiation, development, and death. These cell fate decisions can be influenced by both the gene regulatory network and also environmental factors and can be modeled using simple gene feedback circuits. Negative auto-regulation is a common feedback motif in the gene circuits. It can act to reduce gene expression noise or induce oscillatory expression and is thought to lead to only one cell fate. Here, we present experimental and modeling data to suggest that a self-repressor circuit can lead to two cell fates under specific conditions.

Results

We show that the introduction of inducers capable of binding and unbinding to a self-repressing gene product (protein), thus regulating the associated gene, can lead to the emergence of two cell states. We suggest that the inducers can alter the effective regulatory binding and unbinding speed of the self-repressor regulatory protein to its destination DNA without changing the gene itself. The corresponding simulation results are consistent with the experimental findings. We propose physical and quantitative explanations for the origin of the two phenotypic cell fates.

Conclusions

Our results suggest a mechanism for the emergence of multiple cell fates. This may explain the heterogeneity often observed among cell states, while illustrating that altering gene regulation strength can influence cell fates and their decision-making processes without genetic changes.

Electronic supplementary material

The online version of this article (10.1186/s12915-019-0666-0) contains supplementary material, which is available to authorized users.

A bacterial secreted translocator hijacks riboregulators to control type III secretion in response to host cell contact

Numerous Gram-negative pathogens use a Type III Secretion System (T3SS) to promote virulence by injecting effector proteins into targeted host cells, which subvert host cell processes. Expression of T3SS and the effectors is triggered upon host cell contact, but the underlying mechanism is poorly understood. Here, we report a novel strategy of Yersinia pseudotuberculosis in which this pathogen uses a secreted T3SS translocator protein (YopD) to control global RNA regulators. Secretion of the YopD translocator upon host cell contact increases the ratio of post-transcriptional regulator CsrA to its antagonistic small RNAs CsrB and CsrC and reduces the degradosome components PNPase and RNase E levels. This substantially elevates the amount of the common transcriptional activator (LcrF) of T3SS/Yop effector genes and triggers the synthesis of associated virulence-relevant traits. The observed hijacking of global riboregulators allows the pathogen to coordinate virulence factor expression and also readjusts its physiological response upon host cell contact.

Reversible Gene Expression Control in Yersinia pestis by Using an Optimized CRISPR Interference System

Many genes in the bacterial pathogen Yersinia pestis, the causative agent of three plague pandemics, remain uncharacterized, greatly hampering the development of measures for plague prevention and control. Clustered regularly interspaced short palindromic repeat interference (CRISPRi) has been shown to be an effective tool for gene knockdown in model bacteria. In this system, a catalytically dead Cas9 (dCas9) and a small guide RNA (sgRNA) form a complex, binding to the specific DNA target through base pairing, thereby impeding RNA polymerase binding and causing target gene repression. Here, we introduce an optimized CRISPRi system using Streptococcus pyogenes Cas9-derived dCas9 for gene knockdown in Y. pestis Multiple genes harbored on either the chromosome or plasmids of Y. pestis were efficiently knocked down (up to 380-fold) in a strictly anhydrotetracycline-inducible manner using this CRISPRi approach. Knockdown of hmsH (responsible for biofilm formation) or cspB (encoding a cold shock protein) resulted in greatly decreased biofilm formation or impaired cold tolerance in in vitro phenotypic assays. Furthermore, silencing of the virulence-associated genes yscB or ail using this CRISPRi system resulted in attenuation of virulence in HeLa cells and mice similar to that previously reported for yscB and ail null mutants. Taken together, our results confirm that this optimized CRISPRi system can reversibly and efficiently repress the expression of target genes in Y. pestis, providing an alternative to conventional gene knockdown techniques, as well as a strategy for high-throughput phenotypic screening of Y. pestis genes with unknown functions.IMPORTANCE Yersinia pestis is a lethal pathogen responsible for millions of human deaths in history. It has also attracted much attention for potential uses as a bioweapon or bioterrorism agent, against which new vaccines are desperately needed. However, many Y. pestis genes remain uncharacterized, greatly hampering the development of measures for plague prevention and control. Clustered regularly interspaced short palindromic repeat interference (CRISPRi) has been successfully used in a variety of bacteria in functional genomic studies, but no such genetic tool has been reported in Y. pestis Here, we systematically optimized the CRISPRi approach for use in Y. pestis, which ultimately repressed target gene expression with high efficiency in a reversible manner. Knockdown of functional genes using this method produced phenotypes that were readily detected by in vitro assays, cell infection assays, and mouse infection experiments. This is a report of a CRISPRi approach in Y. pestis and highlights the potential use of this approach in high-throughput functional genomics studies of this pathogen.

Plasmids for Independently Tunable, Low-Noise Expression of Two Genes

Microbiologists often express foreign proteins in bacteria in order study them or to use bacteria as a microbial factory. Usually, this requires controlling the number of foreign proteins expressed in each cell, but for many common protein expression systems, it is difficult to “tune” protein expression without large cell-to-cell variation in expression levels (called “noise” in protein expression). This work describes two protein expression systems that can be combined in the same cell, with tunable expression levels and very low protein expression noise. One new system was used to detect single mRNA molecules by fluorescence microscopy, and the two systems were shown to be independent of each other. These protein expression systems may be useful in any experiment or biotechnology application that can be improved with low protein expression noise.

Some microbiology experiments and biotechnology applications can be improved if it is possible to tune the expression of two different genes at the same time with cell-to-cell variation at or below the level of genes constitutively expressed from the chromosome (the “extrinsic noise limit”). This was recently achieved for a single gene by exploiting negative autoregulation by the tetracycline repressor (TetR) and bicistronic gene expression to reduce gene expression noise. We report new plasmids that use the same principles to achieve simultaneous, low-noise expression for two genes in Escherichia coli. The TetR system was moved to a compatible plasmid backbone, and a system based on the lac repressor (LacI) was found to also exhibit gene expression noise below the extrinsic noise limit. We characterized gene expression mean and noise across the range of induction levels for these plasmids, applied the LacI system to tune expression for single-molecule mRNA detection under two different growth conditions, and showed that two plasmids can be cotransformed to independently tune expression of two different genes.

IMPORTANCE Microbiologists often express foreign proteins in bacteria in order study them or to use bacteria as a microbial factory. Usually, this requires controlling the number of foreign proteins expressed in each cell, but for many common protein expression systems, it is difficult to “tune” protein expression without large cell-to-cell variation in expression levels (called “noise” in protein expression). This work describes two protein expression systems that can be combined in the same cell, with tunable expression levels and very low protein expression noise. One new system was used to detect single mRNA molecules by fluorescence microscopy, and the two systems were shown to be independent of each other. These protein expression systems may be useful in any experiment or biotechnology application that can be improved with low protein expression noise.

A single-input binary counting module based on serine integrase site-specific recombination

A device that counts and records the number of events experienced by an individual cell could have many uses in experimental biology and biotechnology. Here, we report a DNA-based 'latch' that switches between two states upon each exposure to a repeated stimulus. The key component of the latch is a DNA segment whose orientation is inverted by the actions of ϕC31 integrase and its recombination directionality factor (RDF). Integrase expression is regulated by an external input, while RDF expression is controlled by the state of the latch, such that the orientation of the invertible segment switches efficiently each time the device receives an input pulse. Recombination occurs over a time scale of minutes after initiation of integrase expression. The latch requires a delay circuit, implemented with a transcriptional repressor expressed in only one state, to ensure that each input pulse results in only one inversion of the DNA segment. Development and optimization of the latch in living cells was driven by mathematical modelling of the recombination reactions and gene expression regulated by the switch. We discuss how N latches built with orthogonal site-specific recombination systems could be chained together to form a binary ripple counter that could count to 2N - 1.

A simple and dual expression plasmid system in prokaryotic (E. coli) and mammalian cells

We introduce a simple and universal cloning plasmid system for gene expression in prokaryotic (Escherichia coli) and mammalian cells. This novel system has two expression modes: the (subcloning) prokaryotic and mammalian modes. This system streamlines the process of producing mammalian gene expression plasmids with desired genes. The plasmid (prokaryotic mode) has an efficient selection system for DNA insertion, multiple component genes with rare restriction sites at both ends (termed “units”), and a simple transformation to mammalian expression mode utilizing rare restriction enzymes and re-ligation (deletion step). The new plasmid contains the lac promoter and operator followed by a blunt-end EcoRV recognition site, and a DNA topoisomerase II toxin-originated gene for effective selection with isopropyl-β-D-thiogalactoside (IPTG) induction. This system is highly efficient for the subcloning of blunt-end fragments, including PCR products. After the insertion of the desired gene, protein encoded by the desired gene can be detected in E. coli with IPTG induction. Then, the lac promoter and operator are readily deleted with 8-nucleotide rare-cutter blunt-end enzymes (deletion step). Following re-ligation and transformation, the plasmid is ready for mammalian expression analysis (mammalian mode). This idea (conversion from prokaryotic to mammalian mode) can be widely adapted. The pgMAX system overwhelmingly simplifies prokaryotic and mammalian gene expression analyses.

Construction and validation of the Tn5-PLtetO-1-msfGFP transposon as a tool to probe protein expression and localization

In this study we report the design, construction and validation of a novel transposon aimed to systematically screen for protein localization and expression patterns in prokaryotes using fluorescence microscopy. Upon random insertion in an open reading frame in the proper frame and orientation, the transposon creates an N-terminal fluorescent protein fusion to the msfGFP reporter. Moreover, in order to examine the localization of fusion proteins whose native expression might be too low or absent, the transposon was fitted with a P_LtetO-1 promoter that makes the expression of the generated fluorescent protein fusions controllable by anhydrotetracycline. Importantly, upon flipping out the P_LtetO-1 promoter and neighboring antibiotic resistance marker, an in-frame "sandwich" msfGFP fusion is created in which the N- and C-terminal portions of the targeted protein are again controlled by its native promoter.

Discovery and Characterization of a Nitroreductase Capable of Conferring Bacterial Resistance to Chloramphenicol

Widespread antibiotic resistance has led to the reappraisal of abandoned antibiotics including chloramphenicol. However, enzyme(s) underlying one form of chloramphenicol resistance, nitroreduction, have eluded identification. Here we demonstrate that expression of the Haemophilus influenzae nitroreductase gene nfsB confers chloramphenicol resistance in Escherichia coli. We characterized the enzymatic product of H. influenzae NfsB acting on chloramphenicol and found it to be amino-chloramphenicol. Kinetic analysis revealed reduction of diverse substrates including the incomplete reduction of 5-nitro antibiotics metronidazole and nitrofurantoin, likely resulting in activation of these antibiotic pro-drugs to their cytotoxic forms. We observed that expression of the H. influenzae nfsB gene in E. coli results in significantly increased susceptibility to metronidazole. Finally, we found that in this strain metronidazole attenuates chloramphenicol resistance synergistically, and in vitro metronidazole weakly inhibits chloramphenicol reduction by NfsB. Our findings reveal the underpinnings of a chloramphenicol resistance mechanism nearly 70 years after its description.

Adaptive Strategies of the Candidate Probiotic E. coli Nissle in the Mammalian Gut

Probiotics are living microorganisms that are increasingly used as gastrointestinal therapeutics by virtue of their innate or engineered genetic function. Unlike abiotic therapeutics, probiotics can replicate in their intended site, subjecting their genomes and therapeutic properties to natural selection. We exposed the candidate probiotic E. coli Nissle (EcN) to the mouse gastrointestinal tract over several weeks, systematically altering the diet and background microbiota complexity. In-transit EcN accumulates genetic mutations that modulate carbohydrate utilization, stress response, and adhesion to gain competitive fitness, while previous exposure to antibiotics reveals an acquisition of resistance. We then leveraged these insights to generate an EcN strain that shows therapeutic efficacy in a mouse model of phenylketonuria and found that it was genetically stable over 1 week, thereby validating EcN's utility as a chassis for engineering. Collectively, we demonstrate a generalizable pipeline that can be applied to other probiotics to better understand their safety and engineering potential.

RNA polymerase pausing at a protein roadblock can enhance transcriptional interference by promoter occlusion

Convergent promoters exert transcriptional interference (TI) by several mechanisms including promoter occlusion, where elongating RNA polymerases (RNAPs) block access to a promoter. Here, we tested whether pausing of RNAPs by obstructive DNA‐bound proteins can enhance TI by promoter occlusion. Using the Lac repressor as a ‘roadblock’ to induce pausing over a target promoter, we found only a small increase in TI, with mathematical modelling suggesting that rapid termination of the stalled RNAP was limiting the occlusion effect. As predicted, the roadblock‐enhanced occlusion was significantly increased in the absence of the Mfd terminator protein. Thus, protein roadblocking of RNAP may cause pause‐enhanced occlusion throughout genomes, and the removal of stalled RNAP may be needed to minimize unwanted TI.

Comparison between Effects of Retroactivity and Resource Competition upon Change in Downstream Reporter Genes of Synthetic Genetic Circuits

Reporter genes have contributed to advancements in molecular biology. Binding of an upstream regulatory protein to a downstream reporter promoter allows quantification of the activity of the upstream protein produced from the corresponding gene. In studies of synthetic biology, analyses of reporter gene activities ensure control of the cell with synthetic genetic circuits, as achieved using a combination of in silico and in vivo experiments. However, unexpected effects of downstream reporter genes on upstream regulatory genes may interfere with in vivo observations. This phenomenon is termed as retroactivity. Using in silico and in vivo experiments, we found that a different copy number of regulatory protein-binding sites in a downstream gene altered the upstream dynamics, suggesting retroactivity of reporters in this synthetic genetic oscillator. Furthermore, by separating the two sources of retroactivity (titration of the component and competition for degradation), we showed that, in the dual-feedback oscillator, the level of the fluorescent protein reporter competing for degradation with the circuits' components is important for the stability of the oscillations. Altogether, our results indicate that the selection of reporter promoters using a combination of in silico and in vivo experiments is essential for the advanced design of genetic circuits.

Cell-free gene-regulatory network engineering with synthetic transcription factors

Gene-regulatory networks are ubiquitous in nature and critical for bottom-up engineering of synthetic networks. Transcriptional repression is a fundamental function that can be tuned at the level of DNA, protein, and cooperative protein-protein interactions, necessitating high-throughput experimental approaches for in-depth characterization. Here, we used a cell-free system in combination with a high-throughput microfluidic device to comprehensively study the different tuning mechanisms of a synthetic zinc-finger repressor library, whose affinity and cooperativity can be rationally engineered. The device is integrated into a comprehensive workflow that includes determination of transcription-factor binding-energy landscapes and mechanistic modeling, enabling us to generate a library of well-characterized synthetic transcription factors and corresponding promoters, which we then used to build gene-regulatory networks de novo. The well-characterized synthetic parts and insights gained should be useful for rationally engineering gene-regulatory networks and for studying the biophysics of transcriptional regulation.

CRISPR-interference-based modulation of mobile genetic elements in bacteria

Spontaneous mutagenesis of synthetic genetic constructs by mobile genetic elements frequently results in the rapid loss of engineered functions. Previous efforts to minimize such mutations required the exceedingly time-consuming manipulation of bacterial chromosomes and the complete removal of insertional sequences (ISes). To this aim, we developed a single plasmid-based system (pCRIS) that applies CRISPR-interference to inhibit the transposition of bacterial ISes. pCRIS expresses multiple guide RNAs to direct inactivated Cas9 (dCas9) to simultaneously silence IS1, IS3, IS5 and IS150 at up to 38 chromosomal loci in Escherichia coli, in vivo. As a result, the transposition rate of all four targeted ISes dropped to negligible levels at both chromosomal and episomal targets. Most notably, pCRIS, while requiring only a single plasmid delivery performed within a single day, provided a reduction of IS-mobility comparable to that seen in genome-scale chromosome engineering projects. The fitness cost of multiple IS-knockdown, detectable in flask-and-shaker systems was readily outweighed by the less frequent inactivation of the transgene, as observed in green fluorescent protein (GFP)-overexpression experiments. In addition, global transcriptomics analysis revealed only minute alterations in the expression of untargeted genes. Finally, the transposition-silencing effect of pCRIS was easily transferable across multiple E. coli strains. The plasticity and robustness of our IS-silencing system make it a promising tool to stabilize bacterial genomes for synthetic biology and industrial biotechnology applications.

A frame-shifted gene, which rescued its function by non-natural start codons and its application in constructing synthetic gene circuits

Background

Frame-shifted genes results in non-functional peptides. Because of this complete loss of function, frame-shifted genes have never been used in constructing synthetic gene circuits.

Results

Here we report that the function of gene circuits is rescued by a frame-shifted gene, which functions by translating from a non-natural start codon. We report a single nucleotide deletion mutation that developed in the λ-repressor cI within a synthetic genetic NOT gate in Escherichia coli during growth and through this mutation, a non-functional synthetic gene circuit became functional. This mutation resulted in a frame-shifted cI, which showed effective functionality among genetic NOT-gates in Escherichia coli with high regulatory ranges (> 300) and Hill coefficient (> 6.5). The cI worked over a large range of relative copy numbers between the frame-shifted gene and its target promoter. These properties make this frame-shifted gene an excellent candidate for building synthetic gene circuits. We hypothesized a new operating mechanism and showed evidence that frame-shifted cI was translated from non-natural start codon. We have engineered and tested a series of NOT gates made from a library of cI genes, each of which starts from a different codon within the first several amino acids of the frame-shifted cI. It is found that one form with start codon ACA, starting from the 3rd codon had similar repression behavior as the whole frame-shifted gene. We demonstrated synthetic genetic NAND and NOR logic-gates with frame-shifted cI. This is the first report of synthetic-gene-circuits made from a frame-shifted gene.

Conclusions

This study inspires a new view on frame-shifted gene and may serve as a novel way of building and optimizing synthetic-gene-circuits. This work may also have significance in the understanding of non-directed evolution of synthetic genetic circuits.

Electronic supplementary material

The online version of this article (10.1186/s13036-019-0151-x) contains supplementary material, which is available to authorized users.

Synthetic Switches and Regulatory Circuits in Plants

Synthetic biology is an established but ever-growing interdisciplinary field of research currently revolutionizing biomedicine studies and the biotech industry. The engineering of synthetic circuitry in bacterial, yeast, and animal systems prompted considerable advances for the understanding and manipulation of genetic and metabolic networks; however, their implementation in the plant field lags behind. Here, we review theoretical-experimental approaches to the engineering of synthetic chemical- and light-regulated (optogenetic) switches for the targeted interrogation and control of cellular processes, including existing applications in the plant field. We highlight the strategies for the modular assembly of genetic parts into synthetic circuits of different complexity, ranging from Boolean logic gates and oscillatory devices up to semi- and fully synthetic open- and closed-loop molecular and cellular circuits. Finally, we explore potential applications of these approaches for the engineering of novel functionalities in plants, including understanding complex signaling networks, improving crop productivity, and the production of biopharmaceuticals.

Riboregulated toehold-gated gRNA for programmable CRISPR-Cas9 function

Predictable control over gene expression is essential to elicit desired synthetic cellular phenotypes. Although CRISPR-Cas9 offers a simple RNA-guided method for targeted transcriptional control, it lacks the ability to integrate endogenous cellular information for efficient signal processing. Here, we present a new class of riboregulators termed toehold-gated gRNA (thgRNA) by integrating toehold riboswitches into sgRNA scaffolds, and demonstrate their programmability for multiplexed regulation in Escherichia coli with minimal cross-talks.

Engineering a New Class of Anti-LacI Transcription Factors with Alternate DNA Recognition

The lactose repressor, LacI (I⁺_YQR), is an archetypal transcription factor that has been a workhorse in many synthetic genetic networks. LacI represses gene expression (apo ligand) and is induced upon binding of the ligand isopropyl β-d-1-thiogalactopyranoside (IPTG). Recently, laboratory evolution was used to confer inverted function in the native LacI topology resulting in anti-LacI (antilac) function (I^A_YQR), where IPTG binding results in gene suppression. Here we engineered 46 antilacs with alternate DNA binding function (I^A_ADR). Phenotypically, I^A_ADR transcription factors are the inverse of wild-type I⁺_YQR function and possess alternate DNA recognition (ADR). This collection of bespoke I^A_ADR bind orthogonally to disparate non-natural operator DNA sequences and suppress gene expression in the presence of IPTG. This new class of I^A_ADR gene regulators were designed modularly via the systematic pairing of nine alternate allosteric regulatory cores with six alternate DNA binding domains that interact with complementary synthetic operator DNA sequences. The 46 I^A_ADR identified in this study are also orthogonal to the naturally occurring operator O¹. Finally, a demonstration of full orthogonality was achieved via the construction of synthetic genetic toggle switches composed of two nonsynonymous unit pair operations that control two distinct fluorescent outputs. This new class of I^A_ADR transcription factors will facilitate the expansion of the computational capacity of engineered gene circuits, via the scalable increase in the control over the number of gene outputs by way of the expansion of the number of unique transcription factors (or systems of transcription factors) that can simultaneously regulate one or more promoter(s).

Incorporation of a Synthetic Amino Acid into dCas9 Improves Control of Gene Silencing

The CRISPR-Cas9 nuclease has been repurposed as a tool for gene repression (CRISPRi). This catalytically dead Cas9 (dCas9) variant inhibits transcription by blocking either initiation or elongation by the RNA polymerase complex. Conditional control of dCas9-mediated repression has been achieved with inducible promoters that regulate the expression of the dcas9 gene. However, as dCas9-mediated gene silencing is very efficient, even slightly leaky dcas9 expression leads to significant background levels of repression of the target gene. In this study, we report on the development of optimized control of dCas9-mediated silencing through additional regulation at the translation level. We have introduced the TAG stop codon in the dcas9 gene in order to insert a synthetic amino acid, l-biphenylalanine (BipA), at a permissive site in the dCas9 protein. In the absence of BipA, a nonfunctional, truncated dCas9 is produced, but when BipA is present, the TAG codon is translated resulting in a functional, full-length dCas9 protein. This synthetic, BipA-containing dCas9 variant (dCas9-BipA) could still fully repress gene transcription. Comparison of silencing mediated by dCas9 to dCas9-BipA revealed a 14-fold reduction in background repression by the latter system. The here developed proof-of-principle system thus reduces unwanted background levels of gene silencing, allowing for tight and timed control of target gene expression.

Temporal order and precision of complex stress responses in individual bacteria

Sudden stress often triggers diverse, temporally structured gene expression responses in microbes, but it is largely unknown how variable in time such responses are and if genes respond in the same temporal order in every single cell. Here, we quantified timing variability of individual promoters responding to sublethal antibiotic stress using fluorescent reporters, microfluidics, and time-lapse microscopy. We identified lower and upper bounds that put definite constraints on timing variability, which varies strongly among promoters and conditions. Timing variability can be interpreted using results from statistical kinetics, which enable us to estimate the number of rate-limiting molecular steps underlying different responses. We found that just a few critical steps control some responses while others rely on dozens of steps. To probe connections between different stress responses, we then tracked the temporal order and response time correlations of promoter pairs in individual cells. Our results support that, when bacteria are exposed to the antibiotic nitrofurantoin, the ensuing oxidative stress and SOS responses are part of the same causal chain of molecular events. In contrast, under trimethoprim, the acid stress response and the SOS response are part of different chains of events running in parallel. Our approach reveals fundamental constraints on gene expression timing and provides new insights into the molecular events that underlie the timing of stress responses.

Discovery and Characterization of Cas9 Inhibitors Disseminated across Seven Bacterial Phyla

CRISPR-Cas systems in bacteria and archaea provide immunity against bacteriophages and plasmids. To overcome CRISPR immunity, phages have acquired anti-CRISPR genes that reduce CRISPR-Cas activity. Using a synthetic genetic circuit, we developed a high-throughput approach to discover anti-CRISPR genes from metagenomic libraries based on their functional activity rather than sequence homology or genetic context. We identified 11 DNA fragments from soil, animal, and human metagenomes that circumvent Streptococcus pyogenes Cas9 activity in our selection strain. Further in vivo and in vitro characterization of a subset of these hits validated the activity of four anti-CRISPRs. Notably, homologs of some of these anti-CRISPRs were detected in seven different phyla, namely Firmicutes, Proteobacteria, Bacteroidetes, Actinobacteria, Cyanobacteria, Spirochaetes, and Balneolaeota, and have high sequence identity suggesting recent horizontal gene transfer. Thus, anti-CRISPRs against type II-A CRISPR-Cas systems are widely distributed across bacterial phyla, suggesting a more complex ecological role than previously appreciated.

Mapping DNA sequence to transcription factor binding energy in vivo

Despite the central importance of transcriptional regulation in biology, it has proven difficult to determine the regulatory mechanisms of individual genes, let alone entire gene networks. It is particularly difficult to decipher the biophysical mechanisms of transcriptional regulation in living cells and determine the energetic properties of binding sites for transcription factors and RNA polymerase. In this work, we present a strategy for dissecting transcriptional regulatory sequences using in vivo methods (massively parallel reporter assays) to formulate quantitative models that map a transcription factor binding site’s DNA sequence to transcription factor-DNA binding energy. We use these models to predict the binding energies of transcription factor binding sites to within 1 k_BT of their measured values. We further explore how such a sequence-energy mapping relates to the mechanisms of trancriptional regulation in various promoter contexts. Specifically, we show that our models can be used to design specific induction responses, analyze the effects of amino acid mutations on DNA sequence preference, and determine how regulatory context affects a transcription factor’s sequence specificity.

It has been said that we live in the “genomic era,” a time where we can readily sequence full genomes at will. However, it remains difficult to interpret much of the information within a genome. This is especially true of non-coding sequences such as promoters, which contain a number of features such as transcription factor binding sites that determine how genes are regulated. There is no straightforward regulatory “code” that tells us how transcription factor binding sites are organized within a promoter. In this work we examine how DNA sequence determines one of the most important features of a promoter, the strength with which a transcription factor binds to its DNA binding site. We discuss an approach to modeling DNA sequence-specific transcription factor binding energies in vivo using a massively parellel reporter assay. We develop models that allow us to predict the binding energy between a transcription factor and a mutated version of its binding site. We then show that this modeling technique can be used to address a number of scientific and design questions, such as engineering the behavior of genetic circuit elements or examining how transcription factors and their binding sites co-evolve.