Synthetic genomes unveil the effects of synonymous recoding.

Engineering the genetic code of an organism provides the basis for (i) making any organism safely resistant to natural viruses and (ii) preventing genetic information flow into and out of genetically modified organisms while (iii) allowing the biosynthesis of genetically encoded unnatural polymers 1-4 . Achieving these three goals requires the reassignment of multiple of the 64 codons nature uses to encode proteins. However, synonymous codon replacement-recoding-is frequently lethal, and how recoding impacts fitness remains poorly explored. Here, we explore these effects using whole-genome synthesis, multiplexed directed evolution, and genome-transcriptome-translatome-proteome co-profiling on multiple recoded genomes. Using this information, we assemble a synthetic Escherichia coli genome in seven sections using only 57 codons to encode proteins. By discovering the rules responsible for the lethality of synonymous recoding and developing a data-driven multi-omics-based genome construction workflow that troubleshoots synthetic genomes, we overcome the lethal effects of 62,007 synonymous codon swaps and 11,108 additional genomic edits. We show that synonymous recoding induces transcriptional noise including new antisense RNAs, leading to drastic transcriptome and proteome perturbation. As the elimination of select codons from an organism's genetic code results in the widespread appearance of cryptic promoters, we show that synonymous codon choice may naturally evolve to minimize transcriptional noise. Our work provides the first genome-scale description of how synonymous codon changes influence organismal fitness and paves the way for the construction of functional genomes that provide genetic firewalls from natural ecosystems and safely produce biopolymers, drugs, and enzymes with an expanded chemistry.

Characterization of ColE1 Production for Robust tolC Plate Dual-Selection in E. coli.

Bacterial selection is an indispensable tool for E. coli genetic engineering. Marker genes allow for mutant isolation even at low editing efficiencies. TolC is an especially useful E. coli marker: its presence can be selected for with sodium dodecyl sulfate, while its absence can be selected for with the bactericidal protein ColE1. However, utilization of this selection system is greatly limited by the lack of commercially available ColE1 protein. Here, we provide a simple, plate-based, ColE1 negative-selection protocol that does not require purification of ColE1. Using agar plates containing a nonpurified lysate from a ColE1-production strain, we achieved a stringent negative selection with an escape rate of 10-7. Using this powerful negative-selection assay, we then performed the scarless deletion of multiple, large genomic loci (>10 kb), screening only 12 colonies each. We hope this accessible protocol for ColE1 production will lower the barrier of entry for any lab that wishes to harness tolC's dual selection for genetic engineering.

Functional genomics of the rapidly replicating bacterium Vibrio natriegens by CRISPRi.

The fast-growing Gram-negative bacterium Vibrio natriegens is an attractive microbial system for molecular biology and biotechnology due to its remarkably short generation time1,2 and metabolic prowess3,4. However, efforts to uncover and utilize the mechanisms underlying its rapid growth are hampered by the scarcity of functional genomic data. Here, we develop a pooled genome-wide clustered regularly interspaced short palindromic repeats (CRISPR) interference (CRISPRi) screen to identify a minimal set of genes required for rapid wild-type growth. Targeting 4,565 (99.7%) of predicted protein-coding genes, our screen uncovered core genes comprising putative essential and growth-supporting genes that are enriched for respiratory pathways. We found that 96% of core genes were located on the larger chromosome 1, with growth-neutral duplicates of core genes located primarily on chromosome 2. Our screen also refines metabolic pathway annotations by distinguishing functional biosynthetic enzymes from those predicted on the basis of comparative genomics. Taken together, this work provides a broadly applicable platform for high-throughput functional genomics to accelerate biological studies and engineering of V. natriegens.

Cell-free Protein Expression Using the Rapidly Growing Bacterium Vibrio natriegens.

The marine bacterium Vibrio natriegens has garnered considerable attention as an emerging microbial host for biotechnology due to its fast growth rate. A general protocol is described for the preparation of V. natriegens crude cell extracts using common laboratory equipment. This high yielding protocol has been specifically optimized for user accessibility and reduced cost. Cell-free protein synthesis (CFPS) can be carried out in small scale 10 µL batch reactions in either a 96- or 384-well format and reproducibly yields concentrations of > 260 µg/mL super folder GFP (sfGFP) within 3 h. Overall, crude cell extract preparation and CFPS can be achieved in 1-2 full days by a single user. This protocol can be easily integrated into existing protein synthesis pipelines to facilitate advances in bio-production and synthetic biology applications.

Establishing a Cell-Free Vibrio natriegens Expression System.

The fast growing bacterium Vibrio natriegens is an emerging microbial host for biotechnology. Harnessing its productive cellular components may offer a compelling platform for rapid protein production and prototyping of metabolic pathways or genetic circuits. Here, we report the development of a V. natriegens cell-free expression system. We devised a simplified crude extract preparation protocol and achieved >260 µg/mL of superfolder GFP in a small-scale batch reaction after 3 h. Culturing conditions, including growth media and cell density, significantly affect translation kinetics and protein yield of extracts. We observed maximal protein yield at incubation temperatures of 26 or 30 °C, and show improved yield by tuning ions crucial for ribosomal stability. This work establishes an initial V. natriegens cell-free expression system, enables probing of V. natriegens biology, and will serve as a platform to accelerate metabolic engineering and synthetic biology applications.

A modular yeast biosensor for low-cost point-of-care pathogen detection.

The availability of simple, specific, and inexpensive on-site detection methods is of key importance for deployment of pathogen surveillance networks. We developed a nontechnical and highly specific colorimetric assay for detection of pathogen-derived peptides based on Saccharomyces cerevisiae-a genetically tractable model organism and household product. Integrating G protein-coupled receptors with a visible, reagent-free lycopene readout, we demonstrate differential detection of major human, plant, and food fungal pathogens with nanomolar sensitivity. We further optimized a one-step rapid dipstick prototype that can be used in complex samples, including blood, urine, and soil. This modular biosensor can be economically produced at large scale, is not reliant on cold-chain storage, can be detected without additional equipment, and is thus a compelling platform scalable to global surveillance of pathogens.

Design, synthesis, and testing toward a 57-codon genome.

Recoding--the repurposing of genetic codons--is a powerful strategy for enhancing genomes with functions not commonly found in nature. Here, we report computational design, synthesis, and progress toward assembly of a 3.97-megabase, 57-codon Escherichia coli genome in which all 62,214 instances of seven codons were replaced with synonymous alternatives across all protein-coding genes. We have validated 63% of recoded genes by individually testing 55 segments of 50 kilobases each. We observed that 91% of tested essential genes retained functionality with limited fitness effect. We demonstrate identification and correction of lethal design exceptions, only 13 of which were found in 2229 genes. This work underscores the feasibility of rewriting genomes and establishes a framework for large-scale design, assembly, troubleshooting, and phenotypic analysis of synthetic organisms.

FtsZ Cytoskeletal Filaments as a Template for Metallic Nanowire Fabrication.

Supramolecular protein assemblies can serve as templates for the fabrication of inorganic nanowires due to their morphological reproducibility and innate proclivity to form well-ordered structures. Amongst the variety of naturally occurring nano-scale assemblies, cytoskeletal fibers from diverse biological sources represent a unique family of scaffolds for biomimetics as they efficiently self-assemble in vitro in a controllable manner to form stable filaments. Here, we harness the bacterial FtsZ filament system as a scaffold for protein-based metal nanowires, and further demonstrate the control of wire alignment with the use of an external magnetic field. Due to the ease at which the bacterial FtsZ is overexpressed and purified, as well as the extensive studies of its ultrastructural properties and physiological significance, FtsZ filaments are an ideal substrate for large-scale production and chemical manipulation. Using a biologically compatible electroless metal deposition technique initiated by adsorption of platinum as a surface catalyst, we demonstrate the coating of assembled FtsZ filaments with iron, nickel, gold, and copper to fabricate continuous nanowires with diameters ranging from 10-50 nm. Organic-inorganic hybrid wires were analyzed using high-resolution field-emission-gun transmission and scanning electron microscopy, and confirmed by energy-dispersive elemental analysis. We also achieved alignment of ferrofluid-coated FtsZ filaments using an external magnetic field. Overall, we provide evidence for the robustness of the FtsZ filament system as a molecular scaffold, and offer an efficient, biocompatible procedure for facile bottom-up assembly of metallic wires on biological templates. We believe that bottom-up fabrication methods as reported herein significantly contribute to the expanding toolkit available for the incorporation of biological materials in nano-scale devices for electronic and electromechanical applications.

Gene assembly and combinatorial libraries in S. cerevisiae via reiterative recombination.

While mutagenesis of single genes is now common practice in molecular biology, engineering multiple target genes still requires complex cloning techniques and thus is limited to expert laboratories. Here, we describe "Reiterative Recombination," a user-friendly DNA assembly technique in Saccharomyces cerevisiae for the integration of an indefinite number of DNA fragments sequentially into the yeast genome. The high efficiency of chromosomal integration can further be utilized for the assembly of large combinatorial libraries for metabolic engineering.

PCRless library mutagenesis via oligonucleotide recombination in yeast.

The directed evolution of biomolecules with new functions is largely performed in vitro, with PCR mutagenesis followed by high-throughput assays for desired activities. As synthetic biology creates impetus for generating biomolecules that function in living cells, new technologies are needed for performing mutagenesis and selection for directed evolution in vivo. Homologous recombination, routinely exploited for targeted gene alteration, is an attractive tool for in vivo library mutagenesis, yet surprisingly is not routinely used for this purpose. Here, we report the design and characterization of a yeast-based system for library mutagenesis of protein loops via oligonucleotide recombination. In this system, a linear vector is co-transformed with single-stranded mutagenic oligonucleotides. Using repair of nonsense codons engineered in three different active-site loops in the selectable marker TRP1 as a model system, we first optimized the recombination efficiency. Single-loop recombination was highly efficient, averaging 5%, or 4.0×10(5) recombinants. Multiple loops could be simultaneously mutagenized, although the efficiencies dropped to 0.2%, or 6.0×10(3) recombinants, for two loops and 0.01% efficiency, or 1.5×10(2) recombinants, for three loops. Finally, the utility of this system for directed evolution was tested explicitly by selecting functional variants from a mock library of 1:10(6) wild-type:nonsense codons. Sequencing showed that oligonucleotide recombination readily covered this large library, mutating not only the target codon but also encoded silent mutations on either side of the library cassette. Together these results establish oligonucleotide recombination as a simple and powerful library mutagenesis technique and advance efforts to engineer the cell for fully in vivo directed evolution.