Engineering an Escherichia coli strain for production of long single-stranded DNA

Long single-stranded DNA (ssDNA) is a versatile molecular reagent with applications including RNA-guided genome engineering and DNA nanotechnology, yet its production is typically resource-intensive. We introduce a novel method utilizing an engineered Escherichia coli 'helper' strain and phagemid system that simplifies long ssDNA generation to a straightforward transformation and purification procedure. Our method obviates the need for helper plasmids and their associated contamination by integrating M13mp18 genes directly into the E. coli chromosome. We achieved ssDNA lengths ranging from 504 to 20 724 nt with titers up to 250 μg/l following alkaline lysis purification. The efficacy of our system was confirmed through its application in primary T-cell genome modifications and DNA origami folding. The reliability, scalability and ease of our approach promise to unlock new experimental applications requiring large quantities of long ssDNA.

Engineering an Escherichia coli strain for production of long single-stranded DNA

Long single-stranded DNA (ssDNA) is a versatile molecular reagent with applications including RNA-guided genome engineering and DNA nanotechnology, yet its production is typically resource-intensive. We introduce a novel method utilizing an engineered E. coli "helper" strain and phagemid system that simplifies long ssDNA generation to a straightforward transformation and purification procedure. Our method obviates the need for helper plasmids and their associated contamination by integrating M13mp18 genes directly into the E. coli chromosome. We achieved ssDNA lengths ranging from 504 to 20,724 nucleotides with titers up to 250 μg/L following alkaline-lysis purification. The efficacy of our system was confirmed through its application in primary T cell genome modifications and DNA origami folding. The reliability, scalability, and ease of our approach promises to unlock new experimental applications requiring large quantities of long ssDNA.

Peroxisome compartmentalization of a toxic enzyme improves alkaloid production

Eukaryotic cells compartmentalize metabolic pathways in organelles to achieve optimal reaction conditions and avoid crosstalk with cytosolic factors. We found that cytosolic expression of norcoclaurine synthase (NCS), the enzyme that catalyzes the first committed reaction in benzylisoquinoline alkaloid biosynthesis, is toxic in Saccharomyces cerevisiae and, consequently, restricts (S)-reticuline production. We developed a compartmentalization strategy that alleviates NCS toxicity while promoting increased (S)-reticuline titer. This strategy is achieved through efficient targeting of toxic NCS to the peroxisome while, crucially, taking advantage of the free flow of metabolite substrates and products across the peroxisome membrane. We demonstrate that expression of engineered transcription factors can mimic the oleate response for larger peroxisomes, further increasing benzylisoquinoline alkaloid titer without the requirement for peroxisome induction with fatty acids. This work specifically addresses the challenges associated with toxic NCS expression and, more broadly, highlights the potential for engineering organelles with desired characteristics for metabolic engineering.

Exploration of Acetylation as a Base-Labile Protecting Group in Escherichia coli for an Indigo Precursor

Biochemical protecting groups are observed in natural metabolic pathways to control reactivity and properties of chemical intermediates; similarly, they hold promise as a tool for metabolic engineers to achieve the same goals. Protecting groups come with costs: lower yields from carbon, metabolic load to the production host, deprotection catalyst costs and kinetics limitations, and wastewater treatment of the group. Compared to glycosyl biochemical protection, such as glucosyl groups, acetylation can mitigate each of these costs. As an example application where these benefits could be valuable, we explored acetylation protection of indoxyl, the reactive precursor to the clothing dye, indigo. First, we demonstrated denim dyeing with chemically sourced indoxyl acetate by deprotection with base, showing results comparable to industry-standard denim dyeing. Second, we modified an Escherichia coli production host for improved indoxyl acetate stability by the knockout of 14 endogenous hydrolases. Cumulatively, these knockouts yielded a 67% reduction in the indoxyl acetate hydrolysis rate from 0.22 mmol/g DCW/h to 0.07 mmol/g DCW/h. To biosynthesize indoxyl acetate, we identified three promiscuous acetyltransferases which acetylate indoxyl in vivo. Indoxyl acetate titer, while low, was improved 50%, from 43 μM to 67 μM, in the hydrolase knockout strain compared to wild-type E. coli. Unfortunately, low millimolar concentrations of indoxyl acetate proved to be toxic to the E. coli production host; however, the principle of acetylation as a readily cleavable and low impact biochemical protecting group and the engineered hydrolase knockout production host should prove useful for other metabolic products.

Targeted Diversification in the S. cerevisiae Genome with CRISPR-Guided DNA Polymerase I

New technologies to target nucleotide diversification in vivo are promising enabling strategies to perform directed evolution for engineering applications and forward genetics for addressing biological questions. Recently, we reported EvolvR-a system that employs CRISPR-guided Cas9 nickases fused to nick-translating, error-prone DNA polymerases to diversify targeted genomic loci-in E. coli. As CRISPR-Cas9 has shown activity across diverse cell types, EvolvR has the potential to be ported into other organisms, including eukaryotes, if nick-translating polymerases can be active across species. Here, we implement and characterize EvolvR's function in Saccharomyces cerevisiae, representing a key first step to enable EvolvR-mediated mutagenesis in eukaryotes. This advance will be useful for mutagenesis of user-defined loci in the yeast chromosomes for both engineering and basic research applications, and it furthermore provides a platform to develop the EvolvR technology for performance in higher eukaryotes.

A yeast platform for high-level synthesis of tetrahydroisoquinoline alkaloids

The tetrahydroisoquinoline (THIQ) moiety is a privileged substructure of many bioactive natural products and semi-synthetic analogs. Plants manufacture more than 3,000 THIQ alkaloids, including the opioids morphine and codeine. While microbial species have been engineered to synthesize a few compounds from the benzylisoquinoline alkaloid (BIA) family of THIQs, low product titers impede industrial viability and limit access to the full chemical space. Here we report a yeast THIQ platform by increasing production of the central BIA intermediate (S)-reticuline to 4.6 g L⁻¹, a 57,000-fold improvement over our first-generation strain. We show that gains in BIA output coincide with the formation of several substituted THIQs derived from amino acid catabolism. We use these insights to repurpose the Ehrlich pathway and synthesize an array of THIQ structures. This work provides a blueprint for building diverse alkaloid scaffolds and enables the targeted overproduction of thousands of THIQ products, including natural and semi-synthetic opioids.

Plants synthesize more than 3000 tetrahydroisoquinoline (THIQ) alkaloids, but only a few of them have been produced by engineered microbes and titers are very low. Here, the authors increase (S)-reticuline titer to 4.6 g/L and repurpose the yeast Ehrlich pathway to synthesize a diverse array of THIQ scaffolds.

Publisher Correction: Modular and tunable biological feedback control using a de novo protein switch

An amendment to this paper has been published and can be accessed via a link at the top of the paper.

Iterative screening methodology enables isolation of strains with improved properties for a FACS-based screen and increased L-DOPA production

Optimizing microbial hosts for the large-scale production of valuable metabolites often requires multiple mutations and modifications to the host's genome. We describe a three-round screen for increased L-DOPA production in S. cerevisiae using FACS enrichment of an enzyme-coupled biosensor for L-DOPA. Multiple rounds of screening were enabled by a single build of a barcoded in vitro transposon-mediated disruption library. New background strains for screening were built for each iteration using results from previous iterations. The same in vitro transposon-mediated disruption library was integrated by homologous recombination into new background strains in each round of screening. Compared with creating new transposon insertions in each round, this method takes less time and saves the cost of additional sequencing to characterize transposon insertion sites. In the first two rounds of screening, we identified deletions that improved biosensor compartmentalization and, consequently, improved our ability to screen for L-DOPA production. In a final round, we discovered that deletion of heme oxygenase (HMX1) increases total heme concentration and increases L-DOPA production, using dopamine measurement as a proxy. We further demonstrated that deleting HMX1 may represent a general strategy for P450 function improvement by improving activity of a second P450 enzyme, BM3, which performs a distinct reaction.

CRISPR-guided DNA polymerases enable diversification of all nucleotides in a tunable window

The capacity to diversify genetic codes advances our ability to understand and engineer biological systems^1,2. A method for continuously diversifying user-defined regions of a genome would enable forward genetic approaches in systems that are not amenable to efficient homology-directed oligonucleotide integration. It would also facilitate the rapid evolution of biotechnologically useful phenotypes through accelerated and parallelized rounds of mutagenesis and selection, as well as cell-lineage tracking through barcode mutagenesis. Here we present EvolvR, a system that can continuously diversify all nucleotides within a tunable window length at user-defined loci. This is achieved by directly generating mutations using engineered DNA polymerases targeted to loci via CRISPR-guided nickases. We identified nickase and polymerase variants that offer a range of targeted mutation rates that are up to 7,770,000-fold greater than rates seen in wild-type cells, and editing windows with lengths of up to 350 nucleotides. We used EvolvR to identify novel ribosomal mutations that confer resistance to the antibiotic spectinomycin. Our results demonstrate that CRISPR-guided DNA polymerases enable multiplexed and continuous diversification of user-defined genomic loci, which will be useful for a broad range of basic and biotechnological applications.

Employing a biochemical protecting group for a sustainable indigo dyeing strategy

Indigo is an ancient dye uniquely capable of producing the signature tones in blue denim; however, the dyeing process requires chemical steps that are environmentally damaging. We describe a sustainable dyeing strategy that not only circumvents the use of toxic reagents for indigo chemical synthesis but also removes the need for a reducing agent for dye solubilization. This strategy utilizes a glucose moiety as a biochemical protecting group to stabilize the reactive indigo precursor indoxyl to form indican, preventing spontaneous oxidation to crystalline indigo during microbial fermentation. Application of a β-glucosidase removes the protecting group from indican, resulting in indigo crystal formation in the cotton fibers. We identified the gene coding for the glucosyltransferase PtUGT1 from the indigo plant Polygonum tinctorium and solved the structure of PtUGT1. Heterologous expression of PtUGT1 in Escherichia coli supported high indican conversion, and biosynthesized indican was used to dye cotton swatches and a garment.

Bioproduction of a betalain color palette in Saccharomyces cerevisiae

Betalains are a family of natural pigments found exclusively in the plant order Caryophyllales. All members of this chemical family are biosynthesized through the common intermediate betalamic acid, which is capable of spontaneously condensing with various primary and secondary amines to produce betalains. Of particular interest is the red-violet betanin, most commonly obtained from Beta vulgaris (beet) as a natural food dye. We demonstrate the first complete microbial production of betanin in Saccharomyces cerevisiae from glucose, an early step towards a fermentation process enabling rapid, on-demand production of this natural dye. A titer of 17mg/L was achieved, corresponding to a color intensity obtained from 10g/L of beetroot extract. Further, we expanded the spectrum of betalain colors by condensing betalamic acid with various amines fed to an engineered strain of S. cerevisiae. Our work establishes a platform for microbial production of betalains of various colors as a potential alternative to land- and resource-intensive agricultural production.

Correction to: Identification and characterization of a galacturonic acid transporter from Neurospora crassa and its application for Saccharomyces cerevisiae fermentation processes

[This corrects the article DOI: 10.1186/1754-6834-7-20.].

Iterative optimization of xylose catabolism in Saccharomyces cerevisiae using combinatorial expression tuning

A common challenge in metabolic engineering is rapidly identifying rate-controlling enzymes in heterologous pathways for subsequent production improvement. We demonstrate a workflow to address this challenge and apply it to improving xylose utilization in Saccharomyces cerevisiae. For eight reactions required for conversion of xylose to ethanol, we screened enzymes for functional expression in S. cerevisiae, followed by a combinatorial expression analysis to achieve pathway flux balancing and identification of limiting enzymatic activities. In the next round of strain engineering, we increased the copy number of these limiting enzymes and again tested the eight-enzyme combinatorial expression library in this new background. This workflow yielded a strain that has a ∼70% increase in biomass yield and ∼240% increase in xylose utilization. Finally, we chromosomally integrated the expression library. This library enriched for strains with multiple integrations of the pathway, which likely were the result of tandem integrations mediated by promoter homology. Biotechnol. Bioeng. 2017;114: 1301-1309. © 2017 Wiley Periodicals, Inc.

Towards repurposing the yeast peroxisome for compartmentalizing heterologous metabolic pathways

Compartmentalization of enzymes into organelles is a promising strategy for limiting metabolic crosstalk and improving pathway efficiency, but improved tools and design rules are needed to make this strategy available to more engineered pathways. Here we focus on the Saccharomyces cerevisiae peroxisome and develop a sensitive high-throughput assay for peroxisomal cargo import. We identify an enhanced peroxisomal targeting signal type 1 (PTS1) for rapidly sequestering non-native cargo proteins. Additionally, we perform the first systematic in vivo measurements of nonspecific metabolite permeability across the peroxisomal membrane using a polymer exclusion assay. Finally, we apply these new insights to compartmentalize a two-enzyme pathway in the peroxisome and characterize the expression regimes where compartmentalization leads to improved product titre. This work builds a foundation for using the peroxisome as a synthetic organelle, highlighting both promise and future challenges on the way to realizing this goal.

Compartmentalization of enzymes into cellular organelles is a promising strategy for improving pathway efficiency. Here, the authors use a high-throughput assay to identify enhanced peroxisomal targeting signals in yeast, and study the effects of peroxisomal compartmentalization on the performance of a model pathway.

A Barcoding Strategy Enabling Higher-Throughput Library Screening by Microscopy

Dramatic progress has been made in the design and build phases of the design-build-test cycle for engineering cells. However, the test phase usually limits throughput, as many outputs of interest are not amenable to rapid analytical measurements. For example, phenotypes such as motility, morphology, and subcellular localization can be readily measured by microscopy, but analysis of these phenotypes is notoriously slow. To increase throughput, we developed microscopy-readable barcodes (MiCodes) composed of fluorescent proteins targeted to discernible organelles. In this system, a unique barcode can be genetically linked to each library member, making possible the parallel analysis of phenotypes of interest via microscopy. As a first demonstration, we MiCoded a set of synthetic coiled-coil leucine zipper proteins to allow an 8 × 8 matrix to be tested for specific interactions in micrographs consisting of mixed populations of cells. A novel microscopy-readable two-hybrid fluorescence localization assay for probing candidate interactions in the cytosol was also developed using a bait protein targeted to the peroxisome and a prey protein tagged with a fluorescent protein. This work introduces a generalizable, scalable platform for making microscopy amenable to higher-throughput library screening experiments, thereby coupling the power of imaging with the utility of combinatorial search paradigms.

A Highly Characterized Yeast Toolkit for Modular, Multipart Assembly

Saccharomyces cerevisiae is an increasingly attractive host for synthetic biology because of its long history in industrial fermentations. However, until recently, most synthetic biology systems have focused on bacteria. While there is a wealth of resources and literature about the biology of yeast, it can be daunting to navigate and extract the tools needed for engineering applications. Here we present a versatile engineering platform for yeast, which contains both a rapid, modular assembly method and a basic set of characterized parts. This platform provides a framework in which to create new designs, as well as data on promoters, terminators, degradation tags, and copy number to inform those designs. Additionally, we describe genome-editing tools for making modifications directly to the yeast chromosomes, which we find preferable to plasmids due to reduced variability in expression. With this toolkit, we strive to simplify the process of engineering yeast by standardizing the physical manipulations and suggesting best practices that together will enable more straightforward translation of materials and data from one group to another. Additionally, by relieving researchers of the burden of technical details, they can focus on higher-level aspects of experimental design.

Avoidance of truncated proteins from unintended ribosome binding sites within heterologous protein coding sequences

Genetic sequences ported into non-native hosts for synthetic biology applications can gain unexpected properties. In this study, we explored sequences functioning as ribosome binding sites (RBSs) within protein coding DNA sequences (CDSs) that cause internal translation, resulting in truncated proteins. Genome-wide prediction of bacterial RBSs, based on biophysical calculations employed by the RBS calculator, suggests a selection against internal RBSs within CDSs in Escherichia coli, but not those in Saccharomyces cerevisiae. Based on these calculations, silent mutations aimed at removing internal RBSs can effectively reduce truncation products from internal translation. However, a solution for complete elimination of internal translation initiation is not always feasible due to constraints of available coding sequences. Fluorescence assays and Western blot analysis showed that in genes with internal RBSs, increasing the strength of the intended upstream RBS had little influence on the internal translation strength. Another strategy to minimize truncated products from an internal RBS is to increase the relative strength of the upstream RBS with a concomitant reduction in promoter strength to achieve the same protein expression level. Unfortunately, lower transcription levels result in increased noise at the single cell level due to stochasticity in gene expression. At the low expression regimes desired for many synthetic biology applications, this problem becomes particularly pronounced. We found that balancing promoter strengths and upstream RBS strengths to intermediate levels can achieve the target protein concentration while avoiding both excessive noise and truncated protein.

Design and implementation of a biomolecular concentration tracker

As a field, synthetic biology strives to engineer increasingly complex artificial systems in living cells. Active feedback in closed loop systems offers a dynamic and adaptive way to ensure constant relative activity independent of intrinsic and extrinsic noise. In this work, we use synthetic protein scaffolds as a modular and tunable mechanism for concentration tracking through negative feedback. Input to the circuit initiates scaffold production, leading to colocalization of a two-component system and resulting in the production of an inhibitory antiscaffold protein. Using a combination of modeling and experimental work, we show that the biomolecular concentration tracker circuit achieves dynamic protein concentration tracking in Escherichia coli and that steady state outputs can be tuned.

Engineering complex synthetic transcriptional programs with CRISPR RNA scaffolds

Eukaryotic cells execute complex transcriptional programs in which specific loci throughout the genome are regulated in distinct ways by targeted regulatory assemblies. We have applied this principle to generate synthetic CRISPR-based transcriptional programs in yeast and human cells. By extending guide RNAs to include effector protein recruitment sites, we construct modular scaffold RNAs that encode both target locus and regulatory action. Sets of scaffold RNAs can be used to generate synthetic multigene transcriptional programs in which some genes are activated and others are repressed. We apply this approach to flexibly redirect flux through a complex branched metabolic pathway in yeast. Moreover, these programs can be executed by inducing expression of the dCas9 protein, which acts as a single master regulatory control point. CRISPR-associated RNA scaffolds provide a powerful way to construct synthetic gene expression programs for a wide range of applications, including rewiring cell fates or engineering metabolic pathways.

Selection of chromosomal DNA libraries using a multiplex CRISPR system

The directed evolution of biomolecules to improve or change their activity is central to many engineering and synthetic biology efforts. However, selecting improved variants from gene libraries in living cells requires plasmid expression systems that suffer from variable copy number effects, or the use of complex marker-dependent chromosomal integration strategies. We developed quantitative gene assembly and DNA library insertion into the Saccharomyces cerevisiae genome by optimizing an efficient single-step and marker-free genome editing system using CRISPR-Cas9. With this Multiplex CRISPR (CRISPRm) system, we selected an improved cellobiose utilization pathway in diploid yeast in a single round of mutagenesis and selection, which increased cellobiose fermentation rates by over 10-fold. Mutations recovered in the best cellodextrin transporters reveal synergy between substrate binding and transporter dynamics, and demonstrate the power of CRISPRm to accelerate selection experiments and discoveries of the molecular determinants that enhance biomolecule function.

DOI:http://dx.doi.org/10.7554/eLife.03703.001

Over the course of billions of years, natural evolution has produced new proteins and adapted existing ones so that they work better. Scientists have learned how to use the principles that underlie evolution to similarly engineer proteins in the laboratory. This process, known as directed evolution, is a powerful tool for improving how proteins function. Directed evolution normally involves mutating the gene that encodes the protein of interest, selecting the genes that produce the most promising proteins for another round of mutation, and repeating the process until the desired protein function is achieved.

In the first step of directed evolution, a gene is usually mutated randomly in order to create a large ‘library’ of different forms of the gene. These are joined to circular pieces of DNA known as plasmids that can replicate themselves inside cells. However, the number of plasmids than can be taken up differs from cell to cell. This complicates experiments, and the ideal directed evolution experiment would have the same number of plasmids, or target genes, being delivered into each cell.

Ryan et al. have developed a new method for performing directed evolution experiments that uses a recently developed technique called the CRISPR-Cas9 system. This can make direct changes to a DNA strand such as inserting or deleting specific sequences that code for proteins. Ryan et al. used the CRISPR-Cas9 system to create multiple DNA breaks simultaneously across the genome of yeast cells, and joined ‘barcoded’ DNA or DNA for intact genes to these breaks. This avoids the need to use plasmids to introduce foreign DNA into cells. Ryan et al. have named this method the Multiplex CRISPR (or CRISPRm) system.

Having established CRISPRm, Ryan et al. tested whether it could be used to engineer improved proteins by attempting to modify a transporter protein called CDT-1. This protein transports the sugar cellobiose into yeast cells, where it can be converted into alcohol by fermentation. This is important for making biofuel from plants. After just one round of directed evolution using CRISPRm, Ryan et al. successfully isolated a form of the CDT-1 protein that increased the rate of fermentation over 10-fold; hence this CDT-1 variant could be used to increase biofuel production.

In the future, it will be important to implement multiple selection rounds with CRISPRm, and to test how large the DNA libraries can be for directed evolution. In time, CRISPRm could find use in evolving and engineering different combinations of genes, metabolic pathways, and possibly entire genomes.

DOI:http://dx.doi.org/10.7554/eLife.03703.002

Identification and characterization of a galacturonic acid transporter from Neurospora crassa and its application for Saccharomyces cerevisiae fermentation processes

BACKGROUND

Pectin-rich agricultural wastes potentially represent favorable feedstocks for the sustainable production of alternative energy and bio-products. Their efficient utilization requires the conversion of all major constituent sugars. The current inability of the popular fermentation host Saccharomyces cerevisiae to metabolize the major pectic monosaccharide D-galacturonic acid (D-GalA) significantly hampers these efforts. While it has been reasoned that the optimization of cellular D-GalA uptake will be critical for the engineering of D-GalA utilization in yeast, no dedicated eukaryotic transport protein has been biochemically described. Here we report for the first time such a eukaryotic D-GalA transporter and characterize its functionality in S. cerevisiae.

RESULTS

We identified and characterized the D-GalA transporter GAT-1 out of a group of candidate genes obtained from co-expression analysis in N. crassa. The N. crassa Δgat-1 deletion strain is substantially affected in growth on pectic substrates, unable to take up D-GalA, and impaired in D-GalA-mediated signaling events. Moreover, expression of a gat-1 construct in yeast conferred the ability for strong high-affinity D-GalA accumulation rates, providing evidence for GAT-1 being a bona fide D-GalA transport protein. By recombinantly co-expressing D-galacturonate reductase or uronate dehydrogenase in yeast we furthermore demonstrated a transporter-dependent conversion of D-GalA towards more reduced (L-galactonate) or oxidized (meso-galactaric acid) downstream products, respectively, over a broad concentration range.

CONCLUSIONS

By utilizing the novel D-GalA transporter GAT-1 in S. cerevisiae we successfully generated a transporter-dependent uptake and catalysis system for D-GalA into two products with high potential for utilization as platform chemicals. Our data thereby provide a considerable first step towards a more complete utilization of biomass for biofuel and value-added chemicals production.

Expression-level optimization of a multi-enzyme pathway in the absence of a high-throughput assay

Engineered metabolic pathways often suffer from flux imbalances that can overburden the cell and accumulate intermediate metabolites, resulting in reduced product titers. One way to alleviate such imbalances is to adjust the expression levels of the constituent enzymes using a combinatorial expression library. Typically, this approach requires high-throughput assays, which are unfortunately unavailable for the vast majority of desirable target compounds. To address this, we applied regression modeling to enable expression optimization using only a small number of measurements. We characterized a set of constitutive promoters in Saccharomyces cerevisiae that spanned a wide range of expression and maintained their relative strengths irrespective of the coding sequence. We used a standardized assembly strategy to construct a combinatorial library and express for the first time in yeast the five-enzyme violacein biosynthetic pathway. We trained a regression model on a random sample comprising 3% of the total library, and then used that model to predict genotypes that would preferentially produce each of the products in this highly branched pathway. This generalizable method should prove useful in engineering new pathways for the sustainable production of small molecules.

DNA-guided assembly of biosynthetic pathways promotes improved catalytic efficiency

Synthetic scaffolds that permit spatial and temporal organization of enzymes in living cells are a promising post-translational strategy for controlling the flow of information in both metabolic and signaling pathways. Here, we describe the use of plasmid DNA as a stable, robust and configurable scaffold for arranging biosynthetic enzymes in the cytoplasm of Escherichia coli. This involved conversion of individual enzymes into custom DNA-binding proteins by genetic fusion to zinc-finger domains that specifically bind unique DNA sequences. When expressed in cells that carried a rationally designed DNA scaffold comprising corresponding zinc finger binding sites, the titers of diverse metabolic products, including resveratrol, 1,2-propanediol and mevalonate were increased as a function of the scaffold architecture. These results highlight the utility of DNA scaffolds for assembling biosynthetic enzymes into functional metabolic structures. Beyond metabolism, we anticipate that DNA scaffolds may be useful in sequestering different types of enzymes for specifying the output of biological signaling pathways or for coordinating other assembly-line processes such as protein folding, degradation and post-translational modifications.

BglBricks: A flexible standard for biological part assembly

BACKGROUND

Standard biological parts, such as BioBricks parts, provide the foundation for a new engineering discipline that enables the design and construction of synthetic biological systems with a variety of applications in bioenergy, new materials, therapeutics, and environmental remediation. Although the original BioBricks assembly standard has found widespread use, it has several shortcomings that limit its range of potential applications. In particular, the system is not suitable for the construction of protein fusions due to an unfavorable scar sequence that encodes an in-frame stop codon.

RESULTS

Here, we present a similar but new composition standard, called BglBricks, that addresses the scar translation issue associated with the original standard. The new system employs BglII and BamHI restriction enzymes, robust cutters with an extensive history of use, and results in a 6-nucleotide scar sequence encoding glycine-serine, an innocuous peptide linker in most protein fusion applications. We demonstrate the utility of the new standard in three distinct applications, including the construction of constitutively active gene expression devices with a wide range of expression profiles, the construction of chimeric, multi-domain protein fusions, and the targeted integration of functional DNA sequences into specific loci of the E. coli genome.

CONCLUSIONS

The BglBrick standard provides a new, more flexible platform from which to generate standard biological parts and automate DNA assembly. Work on BglBrick assembly reactions, as well as on the development of automation and bioinformatics tools, is currently underway. These tools will provide a foundation from which to transform genetic engineering from a technically intensive art into a purely design-based discipline.

The pathogen protein EspF(U) hijacks actin polymerization using mimicry and multivalency

Enterohaemorrhagic Escherichia coli attaches to the intestine through actin pedestals that are formed when the bacterium injects its protein EspF(U) (also known as TccP) into host cells. EspF(U) potently activates the host WASP (Wiskott-Aldrich syndrome protein) family of actin-nucleating factors, which are normally activated by the GTPase CDC42, among other signalling molecules. Apart from its amino-terminal type III secretion signal, EspF(U) consists of five-and-a-half 47-amino-acid repeats. Here we show that a 17-residue motif within this EspF(U) repeat is sufficient for interaction with N-WASP (also known as WASL). Unlike most pathogen proteins that interface with the cytoskeletal machinery, this motif does not mimic natural upstream activators: instead of mimicking an activated state of CDC42, EspF(U) mimics an autoinhibitory element found within N-WASP. Thus, EspF(U) activates N-WASP by competitively disrupting the autoinhibited state. By mimicking an internal regulatory element and not the natural activator, EspF(U) selectively activates only a precise subset of CDC42-activated processes. Although one repeat is able to stimulate actin polymerization, we show that multiple-repeat fragments have notably increased potency. The activities of these EspF(U) fragments correlate with their ability to coordinate activation of at least two N-WASP proteins. Thus, this pathogen has used a simple autoinhibitory fragment as a component to build a highly effective actin polymerization machine.

Engineering synthetic signaling proteins with ultrasensitive input/output control

Many signaling proteins are built from simple, modular components, yet display highly complex signal-processing behavior. Here we explore how modular domains can be used to build an ultrasensitive switch--a nonlinear input/output function that is central to many complex biological behaviors. By systematically altering the number and affinity of modular autoinhibitory interactions, we show that we can predictably convert a simple linear signaling protein into an ultrasensitive switch.

Rewiring cell signaling: the logic and plasticity of eukaryotic protein circuitry

Living cells rival computers in their ability to process external information and make complex behavioral decisions. Many of these decisions are made by networks of interacting signaling proteins. Ongoing structural, biochemical and cell-based studies have begun to reveal several common principles by which protein components are used to specifically transmit and process information. Recent engineering studies demonstrate that these relatively simple principles can be used to rewire signaling behavior in a process that mimics the evolution of new phenotypic responses.

Reprogramming Control of an Allosteric Signaling Switch Through Modular Recombination

Many eukaryotic signaling proteins are composed of simple modular binding domains, yet they can display sophisticated behaviors such as allosteric gating and multi-input signal integration, properties essential for complex cellular circuits. To understand how such behavior can emerge from combinations of simple domains, we engineered variants of the actin regulatory protein N-WASP (neuronal Wiskott-Aldrich syndrome protein) in which the "output" domain of N-WASP was recombined with heterologous autoinhibitory "input" domains. Synthetic switch proteins were created with diverse gating behaviors in response to nonphysiological inputs. Thus, this type of modular framework can facilitate the evolution or engineering of cellular signaling circuits.