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.

ML-enhanced peroxisome capacity enables compartmentalization of multienzyme pathway.

Repurposing an organelle for specialized metabolism provides an avenue for fermentable, unicellular organisms such as Saccharomyces cerevisiae to mimic compartmentalization of metabolic pathways within different plant tissues. Peroxisomes are attractive organelles for repurposing as they are not required for yeast viability when grown on glucose and can efficiently compartmentalize heterologous enzymes to enable physical separation of cytosolic native metabolism and peroxisomal engineered metabolism. However, when not required, peroxisomes are repressed, leading to low functional capacities for heterologous proteins. Here we engineer peroxisomes with enhanced functional capacities, with the goal of compartmentalizing up to eight metabolic enzymes to enhance titers. We implement a machine learning pipeline that allows the identification of factors to overexpress, culminating in a 137% increase in peroxisome functional capacity compared to a wild-type strain. Improved pathway compartmentalization enables an 80% increase in the biosynthesis titers of the monoterpene geraniol, up to 9.5 g L-1.

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.

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.

Modular and tunable biological feedback control using a de novo protein switch.

De novo-designed proteins1-3 hold great promise as building blocks for synthetic circuits, and can complement the use of engineered variants of natural proteins4-7. One such designer protein-degronLOCKR, which is based on 'latching orthogonal cage-key proteins' (LOCKR) technology8-is a switch that degrades a protein of interest in vivo upon induction by a genetically encoded small peptide. Here we leverage the plug-and-play nature of degronLOCKR to implement feedback control of endogenous signalling pathways and synthetic gene circuits. We first generate synthetic negative and positive feedback in the yeast mating pathway by fusing degronLOCKR to endogenous signalling molecules, illustrating the ease with which this strategy can be used to rewire complex endogenous pathways. We next evaluate feedback control mediated by degronLOCKR on a synthetic gene circuit9, to quantify the feedback capabilities and operational range of the feedback control circuit. The designed nature of degronLOCKR proteins enables simple and rational modifications to tune feedback behaviour in both the synthetic circuit and the mating pathway. The ability to engineer feedback control into living cells represents an important milestone in achieving the full potential of synthetic biology10,11,12. More broadly, this work demonstrates the large and untapped potential of de novo design of proteins for generating tools that implement complex synthetic functionalities in cells for biotechnological and therapeutic applications.

De novo design of bioactive protein switches.

Allosteric regulation of protein function is widespread in biology, but is challenging for de novo protein design as it requires the explicit design of multiple states with comparable free energies. Here we explore the possibility of designing switchable protein systems de novo, through the modulation of competing inter- and intramolecular interactions. We design a static, five-helix 'cage' with a single interface that can interact either intramolecularly with a terminal 'latch' helix or intermolecularly with a peptide 'key'. Encoded on the latch are functional motifs for binding, degradation or nuclear export that function only when the key displaces the latch from the cage. We describe orthogonal cage-key systems that function in vitro, in yeast and in mammalian cells with up to 40-fold activation of function by key. The ability to design switchable protein functions that are controlled by induced conformational change is a milestone for de novo protein design, and opens up new avenues for synthetic biology and cell engineering.

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 systems1,2. 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.

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.

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 ß-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.

An enzyme-coupled biosensor enables (S)-reticuline production in yeast from glucose.

Benzylisoquinoline alkaloids (BIAs) are a diverse family of plant-specialized metabolites that include the pharmaceuticals codeine and morphine and their derivatives. Microbial synthesis of BIAs holds promise as an alternative to traditional crop-based manufacturing. Here we demonstrate the production of the key BIA intermediate (S)-reticuline from glucose in Saccharomyces cerevisiae. To aid in this effort, we developed an enzyme-coupled biosensor for the upstream intermediate L-3,4-dihydroxyphenylalanine (L-DOPA). Using this sensor, we identified an active tyrosine hydroxylase and improved its L-DOPA yields by 2.8-fold via PCR mutagenesis. Coexpression of DOPA decarboxylase enabled what is to our knowledge the first demonstration of dopamine production from glucose in yeast, with a 7.4-fold improvement in titer obtained for our best mutant enzyme. We extended this pathway to fully reconstitute the seven-enzyme pathway from L-tyrosine to (S)-reticuline. Future work to improve titers and connect these steps with downstream pathway branches, already demonstrated in S. cerevisiae, will enable low-cost production of many high-value BIAs.

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.

Application of a Palladium-Catalyzed C-H Functionalization/Indolization Method to Syntheses of cis-Trikentrin†A and Herbindole†B.

We describe herein formal syntheses of the indole alkaloids cis-trikentrin A and herbindole B from a common meso-hydroquinone intermediate prepared by a ruthenium-catalyzed [2+2+1+1] cycloaddition that has not been used previously in natural product synthesis. Key steps include a sterically demanding Buchwald-Hartwig amination as well as a unique C(sp(3) )-H amination/indole formation. Studies toward a selective desymmetrization of the meso-hydroquinone are also reported.

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.

Engineering robust control of two-component system phosphotransfer using modular scaffolds.

Synthetic biology applies engineering principles to facilitate the predictable design of biological systems. Biological systems composed of modular parts with clearly defined interactions are generally easier to manipulate than complex systems exhibiting a large number of subtle interactions. However, recreating the function of a naturally complex system with simple modular parts can increase fragility. Here, inspired by scaffold-directed signaling in higher organisms, we modularize prokaryotic signal transduction to allow programmable redirection of phosphate flux from a histidine kinase to response regulators based on targeting by eukaryotic protein-protein interaction domains. Although scaffold-directed colocalization alone was sufficient to direct signaling between components, this minimal system suffered from high sensitivity to changing expression levels of each component. To address this fragility, we demonstrate how to engineer autoinhibition into the kinase so that phosphotransfer is possible only upon binding to the scaffold. This system, in which scaffold performs the dual functions of activating this autoinhibited kinase and directing flux to the cotargeted response regulator, was significantly more robust to varying component concentrations. Thus, we demonstrate that design principles inspired by the complex signal-transduction pathways of eukaryotes may be generalized, abstracted, and applied to prokaryotes using well-characterized parts.

Employing a combinatorial expression approach to characterize xylose utilization in Saccharomyces cerevisiae.

Fermentation of xylose, a major constituent of lignocellulose, will be important for expanding sustainable biofuel production. We sought to better understand the effects of intrinsic (genotypic) and extrinsic (growth conditions) variables on optimal gene expression of the Scheffersomyces stipitis xylose utilization pathway in Saccharomyces cerevisiae by using a set of five promoters to simultaneously regulate each gene. Three-gene (xylose reductase, xylitol dehydrogenase (XDH), and xylulokinase) and eight-gene (expanded with non-oxidative pentose phosphate pathway enzymes and pyruvate kinase) promoter libraries were enriched under aerobic and anaerobic conditions or with a mutant XDH with altered cofactor usage. Through characterization of enriched strains, we observed (1) differences in promoter enrichment for the three-gene library depending on whether the pentose phosphate pathway genes were included during the aerobic enrichment; (2) the importance of selection conditions, where some aerobically-enriched strains underperform in anaerobic conditions compared to anaerobically-enriched strains; (3) improved growth rather than improved fermentation product yields for optimized strains carrying the mutant XDH compared to the wild-type XDH.

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.

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.

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.

Ten Years of the Synthetic Biology Summer Course at Cold Spring Harbor Laboratory.

The Cold Spring Harbor Laboratory (CSHL) Summer Course on Synthetic Biology, established in 2013, has emerged as a premier platform for immersive education and research in this dynamic field. Rooted in CSHL's rich legacy of biological discovery, the course offers a comprehensive exploration of synthetic biology's fundamentals and applications. Led by a consortium of faculty from diverse institutions, the course structure seamlessly integrates practical laboratory sessions, exploratory research rotations, and enriching seminars by leaders in the field. Over the years, the curriculum has evolved to cover essential topics such as cell-free transcription-translation, DNA construction, computational modeling of gene circuits, engineered gene regulation, and CRISPR technologies. In this review, we describe the history, development, and structure of the course, and discuss how elements of the course might inform the development of other short courses in synthetic biology. We also demonstrate the course's impact beyond the lab with a summary of alumni contributions to research, education, and entrepreneurship. Through these efforts, the CSHL Summer Course on Synthetic Biology remains at the forefront of shaping the next generation of synthetic biologists.