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Bacterial microcompartments (MCPs) are protein-based organelles that encapsulate metabolic pathways. Metabolic engineers have recently sought to repurpose MCPs to encapsulate heterologous pathways to increase flux through pathways of interest. As MCP engineering becomes more common, standardized methods for analyzing changes to MCPs and interpreting results across studies will become increasingly important. In this study, we demonstrate that different imaging techniques yield variations in the apparent size of purified MCPs from Salmonella enterica serovar Typhimurium LT2, likely due to variations in sample preparation methods. We provide guidelines for preparing samples for MCP imaging and outline expected variations in apparent size and morphology between methods. With this report we aim to establish an aid for comparing results across studies., Competing Interests: The authors have declared that no competing interests exist.

Cell-free biology is the activation of biological processes without the use of intact living cells. It has been used for more than 50 years across the life sciences as a foundational research tool, but a recent technical renaissance has facilitated high-yielding (grams of protein per litre), cell-free gene expression systems from model bacteria, the development of cell-free platforms from non-model organisms and multiplexed strategies for rapidly assessing biological design. These advances provide exciting opportunities to profoundly transform synthetic biology by enabling new approaches to the model-driven design of synthetic gene networks, the fast and portable sensing of compounds, on-demand biomanufacturing, building cells from the bottom up, and next-generation educational kits.

Directed evolution of the ribosome for expanded substrate incorporation and novel functions is challenging because the requirement of cell viability limits the mutations that can be made. Here we address this challenge by combining cell-free synthesis and assembly of translationally competent ribosomes with ribosome display to develop a fully in vitro methodology for ribosome synthesis and evolution (called RISE). We validate the RISE method by selecting active genotypes from a ~1.7 × 10 7 member library of ribosomal RNA (rRNA) variants, as well as identifying mutant ribosomes resistant to the antibiotic clindamycin from a library of ~4 × 10 3 rRNA variants. We further demonstrate the prevalence of positive epistasis in resistant genotypes, highlighting the importance of such interactions in selecting for new function. We anticipate that RISE will facilitate understanding of molecular translation and enable selection of ribosomes with altered properties.

Protein glycosylation is a common post-translational modification that influences the functions and properties of proteins. Despite advances in methods to produce defined glycoproteins by chemoenzymatic elaboration of monosaccharides, the understanding and engineering of glycoproteins remain challenging, in part, due to the difficulty of site-specifically controlling glycosylation at each of several positions within a protein. Here, we address this limitation by discovering and exploiting the unique, conditionally orthogonal peptide acceptor specificities of N -glycosyltransferases (NGTs). We used cell-free protein synthesis and mass spectrometry of self-assembled monolayers to rapidly screen 41 putative NGTs and rigorously characterize the unique acceptor sequence preferences of four NGT variants using 1254 acceptor peptides and 8306 reaction conditions. We then used the optimized NGT-acceptor sequence pairs to sequentially install monosaccharides at four sites within one target protein. This strategy to site-specifically control the installation of N -linked monosaccharides for elaboration to a variety of functional N -glycans overcomes a major limitation in synthesizing defined glycoproteins for research and therapeutic applications., Competing Interests: The authors declare no competing financial interest., (Copyright © 2020 American Chemical Society.)

Engineering the process of molecular translation, or protein biosynthesis, has emerged as a major opportunity in synthetic and chemical biology to generate novel biological insights and enable new applications (e.g. designer protein therapeutics). Here, we review methods for engineering the process of translation in vitro. We discuss the advantages and drawbacks of the two major strategies-purified and extract-based systems-and how they may be used to manipulate and study translation. Techniques to engineer each component of the translation machinery are covered in turn, including transfer RNAs, translation factors, and the ribosome. Finally, future directions and enabling technological advances for the field are discussed., (© The Author(s) 2019. Published by Oxford University Press on behalf of Nucleic Acids Research.)

High-throughput quantification of the post-translational modification of many individual protein samples is challenging with current label-based methods. This paper demonstrates an efficient method that addresses this gap by combining Escherichia coli -based cell-free protein synthesis (CFPS) and self-assembled monolayers for matrix-assisted laser desorption/ionization mass spectrometry (SAMDI-MS) to analyze intact proteins. This high-throughput approach begins with polyhistidine-tagged protein substrates expressed from linear DNA templates by CFPS. Here, we synthesized an 87-member library of the E. coli Immunity Protein 7 (Im7) containing an acceptor sequence optimized for glycosylation by the Actinobacillus pleuropneumoniae N -glycosyltransferase (NGT) at every possible position along the protein backbone. These protein substrates were individually treated with NGT and then selectively immobilized to self-assembled monolayers presenting nickel-nitrilotriacetic acid (Ni-NTA) complexes before final analysis by SAMDI-MS to quantify the conversion of substrate to glycoprotein. This method offers new opportunities for rapid synthesis and quantitative evaluation of intact glycoproteins.

Rapid molecular biosensing is an emerging application area for synthetic biology. Here, we engineer a portable biosensor for cyanuric acid (CYA), an analyte of interest for human and environmental health, using a LysR-type transcription regulator (LTTR) from Pseudomonas within the context of Escherichia coli gene expression machinery. To overcome cross-host portability challenges of LTTRs, we rationally engineered hybrid Pseudomonas-E. coli promoters by integrating DNA elements required for transcriptional activity and ligand-dependent regulation from both hosts, which enabled E. coli to function as a whole-cell biosensor for CYA. To alleviate challenges of whole-cell biosensing, we adapted these promoter designs to function within a freeze-dried E. coli cell-free system to sense CYA. This portable, on-demand system robustly detects CYA within an hour from laboratory and real-world samples and works with both fluorescent and colorimetric reporters. This work elucidates general principles to facilitate the engineering of a wider array of LTTR-based environmental sensors.

Advances in biosensor engineering have enabled the design of programmable molecular systems to detect a range of pathogens, nucleic acids, and chemicals. Here, we engineer and field-test a biosensor for fluoride, a major groundwater contaminant of global concern. The sensor consists of a cell-free system containing a DNA template that encodes a fluoride-responsive riboswitch regulating genes that produce a fluorescent or colorimetric output. Individual reactions can be lyophilized for long-term storage and detect fluoride at levels above 2 ppm, the Environmental Protection Agency's most stringent regulatory standard, in both laboratory and field conditions. Through onsite detection of fluoride in a real-world water source, this work provides a critical proof-of-principle for the future engineering of riboswitches and other biosensors to address challenges for global health and the environment.

Tracking progress towards Target 6.1 of the United Nations Sustainable Development Goals, "achieving universal and equitable access to safe and affordable drinking water for all", necessitates the development of simple, inexpensive tools to monitor water quality. The rapidly growing field of synthetic biology has the potential to address this need by taking DNA-encoded sensing elements from nature and reassembling them to create field-deployable 'biosensors' that can detect pathogenic or chemical water contaminants. Here we describe water quality monitoring strategies enabled by synthetic biology and compare them to previous approaches used to detect three priority water contaminants: fecal pathogens, arsenic, and fluoride in order to explain the potential for engineered biosensors to simplify and decentralize water quality monitoring. We also briefly discuss expanding biosensors to detect emerging contaminants including metals and pharmaceuticals. We conclude with an outlook on the future of biosensor development, in which we discuss adaptability to emerging contaminants, outline current limitations, and propose steps to overcome the field's outstanding challenges to facilitate global water quality monitoring., Competing Interests: Competing Interests Statement K.K.A. and J.B.L. are founders and have a financial interest in Stemloop, Inc. These latter interests were reviewed and managed by Northwestern University in accordance with their conflict of interest policies. All other authors declare no conflicts of interest.

Organism engineering requires the selection of an appropriate chassis, editing its genome, combining traits from different source species, and controlling genes with synthetic circuits. When a strain is needed for a new target objective, for example, to produce a chemical-of-need, the best strains, genes, techniques, software, and expertise may be distributed across laboratories. Here, we report a project where we were assigned phloroglucinol (PG) as a target, and then combined unique capabilities across the United States Army, Navy, and Air Force service laboratories with the shared goal of designing an organism to produce this molecule. In addition to the laboratory strain Escherichia coli , organisms were screened from soil and seawater. Putative PG-producing enzymes were mined from a strain bank of bacteria isolated from aircraft and fuel depots. The best enzyme was introduced into the ocean strain Marinobacter atlanticus CP1 with its genome edited to redirect carbon flux from natural fatty acid ester (FAE) production. PG production was also attempted in Bacillus subtilis and Clostridium acetobutylicum . A genetic circuit was constructed in E. coli that responds to PG accumulation, which was then ported to an in vitro paper-based system that could serve as a platform for future low-cost strain screening or for in-field sensing. Collectively, these efforts show how distributed biotechnology laboratories with domain-specific expertise can be marshalled to quickly provide a solution for a targeted organism engineering project, and highlights data and material sharing protocols needed to accelerate future efforts.

The site-specific incorporation of non-canonical amino acids (ncAAs) into proteins via amber suppression provides access to novel protein properties, structures, and functions. Historically, poor protein expression yields resulting from release factor 1 (RF1) competition has limited this technology. To address this limitation, we develop a high-yield, one-pot cell-free platform for synthesizing proteins bearing ncAAs based on genomically recoded Escherichia coli lacking RF1. A key feature of this platform is the independence on the addition of purified T7 DNA-directed RNA polymerase (T7RNAP) to catalyze transcription. Extracts derived from our final strain demonstrate high productivity, synthesizing 2.67 ± 0.06 g/L superfolder GFP in batch mode without supplementation of purified T7RNAP. Using an optimized one-pot platform, we demonstrate multi-site incorporation of the ncAA p-acetyl-L-phenylalanine into an elastin-like polypeptide with high accuracy of incorporation and yield. Our work has implications for chemical and synthetic biology., (Copyright © 2019 Elsevier Ltd. All rights reserved.)

Glycosylation plays important roles in cellular function and endows protein therapeutics with beneficial properties. However, constructing biosynthetic pathways to study and engineer precise glycan structures on proteins remains a bottleneck. Here, we report a modular, versatile cell-free platform for glycosylation pathway assembly by rapid in vitro mixing and expression (GlycoPRIME). In GlycoPRIME, glycosylation pathways are assembled by mixing-and-matching cell-free synthesized glycosyltransferases that can elaborate a glucose primer installed onto protein targets by an N-glycosyltransferase. We demonstrate GlycoPRIME by constructing 37 putative protein glycosylation pathways, creating 23 unique glycan motifs, 18 of which have not yet been synthesized on proteins. We use selected pathways to synthesize a protein vaccine candidate with an α-galactose adjuvant motif in a one-pot cell-free system and human antibody constant regions with minimal sialic acid motifs in glycoengineered Escherichia coli. We anticipate that these methods and pathways will facilitate glycoscience and make possible new glycoengineering applications.

The need for sustainable, low-cost production of bioenergy and commodity chemicals is increasing. Unfortunately, the engineering potential of whole-cell catalysts to address this need can be hampered by cellular toxicity. When such bottlenecks limit the commercial feasibility of whole-cell fermentation, cell-free, or in vitro , based approaches may offer an alternative. Here, we assess the impact of three classes of growth toxic compounds on crude extract-based, cell-free chemical conversions. As a model system, we test a metabolic pathway for conversion of glucose to 2,3-butanediol (2,3-BDO) in lysates of Escherichia coli . First, we characterized 2,3-BDO production with different classes of antibiotics and found, as expected, that the system is uninhibited by compounds that prevent cell growth by means of cell wall replication and DNA, RNA, and protein synthesis. Second, we considered the impact of polar solvent addition ( e.g., methanol, n-butanol) . We observed that volumetric productivities (g/L/h) were slowed with increasing hydrophobicity of added alcohols. Finally, we investigated the effects of using pretreated biomass hydrolysate as a feed stock, observing a 25% reduction in 2,3-BDO production as a result of coumaroyl and feruloyl amides. Overall, we find the cell-free system to be robust to working concentrations of antibiotics and other compounds that are toxic to cell growth, but do not denature or inhibit relevant enzymes., Competing Interests: None., (© 2019 The Authors.)

The site-specific incorporation of noncanonical monomers into polypeptides through genetic code reprogramming permits synthesis of bio-based products that extend beyond natural limits. To better enable such efforts, flexizymes (transfer RNA (tRNA) synthetase-like ribozymes that recognize synthetic leaving groups) have been used to expand the scope of chemical substrates for ribosome-directed polymerization. The development of design rules for flexizyme-catalyzed acylation should allow scalable and rational expansion of genetic code reprogramming. Here we report the systematic synthesis of 37 substrates based on 4 chemically diverse scaffolds (phenylalanine, benzoic acid, heteroaromatic, and aliphatic monomers) with different electronic and steric factors. Of these substrates, 32 were acylated onto tRNA and incorporated into peptides by in vitro translation. Based on the design rules derived from this expanded alphabet, we successfully predicted the acylation of 6 additional monomers that could uniquely be incorporated into peptides and direct N-terminal incorporation of an aldehyde group for orthogonal bioconjugation reactions.

Easy-to-perform, relatively inexpensive blood diagnostics have transformed at-home healthcare for some patients, but they require analytical equipment and are not easily adapted to measuring other biomarkers. The requirement for reliable quantification in complex sample types (such as blood) has been a critical roadblock in developing and deploying inexpensive, minimal-equipment diagnostics. Here, we developed a platform for inexpensive, easy-to-use diagnostics that uses cell-free expression to generate colored readouts that are visible to the naked eye, yet quantitative and robust to the interference effects seen in complex samples. We achieved this via a parallelized calibration scheme that uses the patient sample to generate custom reference curves. We used this approach to quantify a clinically relevant micronutrient and to quantify nucleic acids, demonstrating a generalizable platform for low-cost quantitative diagnostics., (Copyright © 2019 The Authors, some rights reserved; exclusive licensee American Association for the Advancement of Science. No claim to original U.S. Government Works. Distributed under a Creative Commons Attribution NonCommercial License 4.0 (CC BY-NC).)

Ribo-T is a ribosome with covalently tethered subunits where core 16S and 23S ribosomal RNAs form a single chimeric molecule. Ribo-T makes possible a functionally orthogonal ribosome-mRNA system in cells. Unfortunately, use of Ribo-T has been limited because of low activity of its original version. Here, to overcome this limitation, we use an evolutionary approach to select new tether designs that are capable of supporting faster cell growth and increased protein expression. Further, we evolve new orthogonal Ribo-T/mRNA pairs that function in parallel with, but independent of, natural ribosomes and mRNAs, increasing the efficiency of orthogonal protein expression. The Ribo-T with optimized designs is able to synthesize a diverse set of proteins, and can also incorporate multiple non-canonical amino acids into synthesized polypeptides. The enhanced Ribo-T designs should be useful for exploring poorly understood functions of the ribosome and engineering ribosomes with altered catalytic properties.

RNA nanotechnology seeks to create nanoscale machines by repurposing natural RNA modules. The field is slowed by the current need for human intuition during three-dimensional structural design. Here, we demonstrate that three distinct problems in RNA nanotechnology can be reduced to a pathfinding problem and automatically solved through an algorithm called RNAMake. First, RNAMake discovers highly stable single-chain solutions to the classic problem of aligning a tetraloop and its sequence-distal receptor, with experimental validation from chemical mapping, gel electrophoresis, solution X-ray scattering and crystallography with 2.55 Å resolution. Second, RNAMake automatically generates structured tethers that integrate 16S and 23S ribosomal RNAs into single-chain ribosomal RNAs that remain uncleaved by ribonucleases and assemble onto messenger RNA. Third, RNAMake enables the automated stabilization of small-molecule binding RNAs, with designed tertiary contacts that improve the binding affinity of the ATP aptamer and improve the fluorescence and stability of the Spinach RNA in cell extracts and in living Escherichia coli cells.

Metabolic engineering uses enzymes to produce small molecules with industrial, pharmaceutical, and energy applications. However, efforts to optimize enzymatic pathways for commercial production are limited by the throughput of assays for quantifying metabolic intermediates and end products. We developed a multiplexed method for profiling CoA-dependent pathways that uses a cysteine-terminated peptide to covalently capture CoA-bound metabolites. Captured metabolites are then rapidly separated from the complex mixture by immobilization onto arrays of self-assembled monolayers and directly quantified by SAMDI mass spectrometry. We demonstrate the throughput of the assay by characterizing the cell-free synthesis of HMG-CoA, a key intermediate in the biosynthesis of isoprenoids, collecting over 10,000 individual spectra to map more than 800 unique reaction conditions. We anticipate that our rapid and robust analytical method will accelerate efforts to engineer metabolic pathways.

Recent advances in synthetic biology have resulted in biological technologies with the potential to reshape the way we understand and treat human disease. Educating students about the biology and ethics underpinning these technologies is critical to empower them to make informed future policy decisions regarding their use and to inspire the next generation of synthetic biologists. However, hands-on, educational activities that convey emerging synthetic biology topics can be difficult to implement due to the expensive equipment and expertise required to grow living cells. We present BioBits Health, an educational kit containing lab activities and supporting curricula for teaching antibiotic resistance mechanisms and CRISPR-Cas9 gene editing in high school classrooms. This kit links complex biological concepts to visual, fluorescent readouts in user-friendly freeze-dried cell-free reactions. BioBits Health represents a set of educational resources that promises to encourage teaching of cutting-edge, health-related synthetic biology topics in classrooms and other nonlaboratory settings.

Over the last 50 years, Cell-Free Protein Synthesis (CFPS) has emerged as a powerful technology to harness the transcriptional and translational capacity of cells within a test tube. By obviating the need to maintain the viability of the cell, and by eliminating the cellular barrier, CFPS has been foundational to emerging applications in biomanufacturing of traditionally challenging proteins, as well as applications in rapid prototyping for metabolic engineering, and functional genomics. Our methods for implementing an E. coli-based CFPS platform allow new users to access many of these applications. Here, we describe methods to prepare extract through the use of enriched media, baffled flasks, and a reproducible method of tunable sonication-based cell lysis. This extract can then be used for protein expression capable of producing 900 µg/mL or more of super folder green fluorescent protein (sfGFP) in just 5 h from experimental setup to data analysis, given that appropriate reagent stocks have been prepared beforehand. The estimated startup cost of obtaining reagents is $4,500 which will sustain thousands of reactions at an estimated cost of $0.021 per µg of protein produced or $0.019 per µL of reaction. Additionally, the protein expression methods mirror the ease of the reaction setup seen in commercially available systems due to optimization of reagent pre-mixes, at a fraction of the cost. In order to enable the user to leverage the flexible nature of the CFPS platform for broad applications, we have identified a variety of aspects of the platform that can be tuned and optimized depending on the resources available and the protein expression outcomes desired.

Ribo-T is an engineered ribosome whose small and large subunits are tethered together by linking 16S rRNA and 23S rRNA in a single molecule. Although Ribo-T can support cell proliferation in the absence of wild type ribosomes, Ribo-T cells grow slower than those with wild type ribosomes. Here, we show that cell growth defect is likely explained primarily by slow Ribo-T assembly rather than its imperfect functionality. Ribo-T maturation is stalled at a late assembly stage. Several post-transcriptional rRNA modifications and some ribosomal proteins are underrepresented in the accumulated assembly intermediates and rRNA ends are incompletely trimmed. Ribosome profiling of Ribo-T cells shows no defects in translation elongation but reveals somewhat higher occupancy by Ribo-T of the start codons and to a lesser extent stop codons, suggesting that subunit tethering mildly affects the initiation and termination stages of translation. Understanding limitations of Ribo-T system offers ways for its future development.

Recent advances in cell-free gene expression (CFE) systems have enabled their use for a host of synthetic biology applications, particularly for rapid prototyping of genetic circuits and biosensors. Despite the proliferation of cell-free protein synthesis platforms, the large number of currently existing protocols for making CFE extracts muddles the collective understanding of how the extract preparation method affects its functionality. A key aspect of extract performance relevant to many applications is the activity of the native host transcriptional machinery that can mediate protein synthesis. However, protein yields from genes transcribed in vitro by the native Escherichia coli RNA polymerase are variable for different extract preparation techniques, and specifically low in some conventional crude extracts originally optimized for expression by the bacteriophage transcriptional machinery. Here, we show that cell-free expression of genes under bacterial σ 70 promoters is constrained by the rate of transcription in crude extracts, and that processing the extract with a ribosomal runoff reaction and subsequent dialysis alleviates this constraint. Surprisingly, these processing steps only enhance protein synthesis in genes under native regulation, indicating that the translation rate is unaffected. We further investigate the role of other common extract preparation process variants on extract performance and demonstrate that bacterial transcription is inhibited by including glucose in the growth culture but is unaffected by flash-freezing the cell pellet prior to lysis. Our final streamlined and detailed protocol for preparing extract by sonication generates extract that facilitates expression from a diverse set of sensing modalities including protein and RNA regulators. We anticipate that this work will clarify the methodology for generating CFE extracts that are active for biosensing using native transcriptional machinery and will encourage the further proliferation of cell-free gene expression technology for new applications.

Isoprenoids are an attractive class of metabolites for enzymatic synthesis from renewable substrates. However, metabolic engineering of microorganisms for monoterpenoid production is limited by the need for time-consuming, and often non-intuitive, combinatorial tuning of biosynthetic pathway variations to meet design criteria. Towards alleviating this limitation, the goal of this work was to build a modular, cell-free platform for construction and testing of monoterpenoid pathways, using the fragrance and flavoring molecule limonene as a model. In this platform, multiple Escherichia coli lysates, each enriched with a single overexpressed pathway enzyme, are mixed to construct the full biosynthetic pathway. First, we show the ability to synthesize limonene from six enriched lysates with mevalonate substrate, an adenosine triphosphate (ATP) source, and cofactors. Next, we extend the pathway to use glucose as a substrate, which relies on native metabolism in the extract to convert glucose to acetyl-CoA along with three additional enzymes to convert acetyl-CoA to mevalonate. We find that the native E. coli farnesyl diphosphate synthase (IspA) is active in the lysate and diverts flux from the pathway intermediate geranyl pyrophospahte to farnesyl pyrophsophate and the byproduct farnesol. By adjusting the relative levels of cofactors NAD + , ATP and CoA, the system can synthesize 0.66 mM (90.2 mg l -1 ) limonene over 24 h, a productivity of 3.8 mg l -1 h -1 . Our results highlight the flexibility of crude lysates to sustain complex metabolism and, by activating a glucose-to-limonene pathway with 9 heterologous enzymes encompassing 20 biosynthetic steps, expands an approach of using enzyme-enriched lysates for constructing, characterizing and prototyping enzymatic pathways.

Building variant ribosomes offers opportunities to reveal fundamental principles underlying ribosome biogenesis and to make ribosomes with altered properties. However, cell viability limits mutations that can be made to the ribosome. To address this limitation, the in vitro integrated synthesis, assembly and translation (iSAT) method for ribosome construction from the bottom up was recently developed. Unfortunately, iSAT is complex, costly, and laborious to researchers, partially due to the high cost of reaction buffer containing over 20 components. In this study, we develop iSAT in Escherichia coli BL21Rosetta2 cell lysates, a commonly used bacterial strain, with a cost-effective poly sugar and nucleotide monophosphate-based metabolic scheme. We achieved a 10-fold increase in protein yield over our base case with an evolutionary design of experiments approach, screening 490 reaction conditions to optimize the reaction buffer. The computationally guided, cell-free, high-throughput technology presented here augments the way we approach multicomponent synthetic biology projects and efforts to repurpose ribosomes.

Neisseria gonorrhoeae mounts a substantial transcriptional program in response to hydrogen peroxide (HP), a prominent reactive oxygen species (ROS) encountered during infection. We tested which strain FA1090 genes show differential transcript abundance in response to sublethal amounts of HP to differentiate HP-responsive signaling from widespread cellular death and dysregulation. RNA sequencing (RNA-Seq) revealed that 150 genes were significantly upregulated and 143 genes downregulated following HP exposure. We annotated HP-responsive operons and all transcriptional start sites (TSSs) and identified which TSSs responded to HP treatment. We compared the HP responses and other previously reported genes and found only partial overlapping of other regulatory networks, indicating that the response to HP involves multiple biological functions. Using a representative subset of responsive genes, we validated the RNA-Seq results and found that the HP transcriptome was similar to that of sublethal organic peroxide. None of the genes in the representative subset, however, responded to sublethal levels of HOCl or O 2 - . These results support the idea that N. gonorrhoeae may use variations in HP levels as a signal for different stages of infection. IMPORTANCE The strict human pathogen Neisseria gonorrhoeae is the only causative agent of the sexually transmitted disease gonorrhea. This bacterium encounters hydrogen peroxide produced from host cells during infection, but the organism survives in the presence of this antimicrobial agent. This work shows that the bacterium responds to hydrogen peroxide by regulating the expression of many genes involved in multiple processes.

The Shine-Dalgarno (SD) sequence motif facilitates translation initiation and is frequently found upstream of bacterial start codons. However, thousands of instances of this motif occur throughout the middle of protein coding genes in a typical bacterial genome. Here, we use comparative evolutionary analysis to test whether SD sequences located within genes are functionally constrained. We measure the conservation of SD sequences across Enterobacteriales, and find that they are significantly less conserved than expected. Further, the strongest SD sequences are the least conserved whereas we find evidence of conservation for the weakest possible SD sequences given amino acid constraints. Our findings indicate that most SD sequences within genes are likely to be deleterious and removed via selection. To illustrate the origin of these deleterious costs, we show that ATG start codons are significantly depleted downstream of SD sequences within genes, highlighting the constraint that these sequences impose on the surrounding nucleotides to minimize the potential for erroneous translation initiation.

A new wave of interest in cell-free protein synthesis (CFPS) systems has shown their utility for producing proteins at high titers, establishing genetic regulatory element libraries ( e.g., promoters, ribosome binding sites) in nonmodel organisms, optimizing biosynthetic pathways before implementation in cells, and sensing biomarkers for diagnostic applications. Unfortunately, most previous efforts have focused on a select few model systems, such as Escherichia coli. Broadening the spectrum of organisms used for CFPS promises to better mimic host cell processes in prototyping applications and open up new areas of research. Here, we describe the development and characterization of a facile CFPS platform based on lysates derived from the fast-growing bacterium Vibrio natriegens, which is an emerging host organism for biotechnology. We demonstrate robust preparation of highly active extracts using sonication, without specialized and costly equipment. After optimizing the extract preparation procedure and cell-free reaction conditions, we show synthesis of 1.6 ± 0.05 g/L of superfolder green fluorescent protein in batch mode CFPS, making it competitive with existing E. coli CFPS platforms. To showcase the flexibility of the system, we demonstrate that it can be lyophilized and retain biosynthesis capability, that it is capable of producing antimicrobial peptides, and that our extract preparation procedure can be coupled with the recently described Vmax Express strain in a one-pot system. Finally, to further increase system productivity, we explore a knockout library in which putative negative effectors of CFPS are genetically removed from the source strain. Our V. natriegens-derived CFPS platform is versatile and simple to prepare and use. We expect it will facilitate expansion of CFPS systems into new laboratories and fields for compelling applications in synthetic biology.

The original version of this Article contained an error in Figure 2, wherein the bottom right western blot panel in Figure 2a was blank. This has now been corrected in both the PDF and HTML versions of the Article.

Synthetic biology offers opportunities for experiential educational activities at the intersection of the life sciences, engineering, and design. However, implementation of hands-on biology activities in classrooms is challenging because of the need for specialized equipment and expertise to grow living cells. We present BioBits™ Bright, a shelf-stable, just-add-water synthetic biology education kit with easy visual outputs enabled by expression of fluorescent proteins in freeze-dried, cell-free reactions. We introduce activities and supporting curricula for teaching the central dogma, tunable protein expression, and design-build-test cycles and report data generated by K-12 teachers and students. We also develop inexpensive incubators and imagers, resulting in a comprehensive kit costing

Hands-on demonstrations greatly enhance the teaching of science, technology, engineering, and mathematics (STEM) concepts and foster engagement and exploration in the sciences. While numerous chemistry and physics classroom demonstrations exist, few biology demonstrations are practical and accessible due to the challenges and concerns of growing living cells in classrooms. We introduce BioBits™ Explorer, a synthetic biology educational kit based on shelf-stable, freeze-dried, cell-free (FD-CF) reactions, which are activated by simply adding water. The FD-CF reactions engage the senses of sight, smell, and touch with outputs that produce fluorescence, fragrances, and hydrogels, respectively. We introduce components that can teach tunable protein expression, enzymatic reactions, biomaterial formation, and biosensors using RNA switches, some of which represent original FD-CF outputs that expand the toolbox of cell-free synthetic biology. The BioBits™ Explorer kit enables hands-on demonstrations of cutting-edge science that are inexpensive and easy to use, circumventing many current barriers for implementing exploratory biology experiments in classrooms.

The emerging discipline of bacterial glycoengineering has made it possible to produce designer glycans and glycoconjugates for use as vaccines and therapeutics. Unfortunately, cell-based production of homogeneous glycoproteins remains a significant challenge due to cell viability constraints and the inability to control glycosylation components at precise ratios in vivo. To address these challenges, we describe a novel cell-free glycoprotein synthesis (CFGpS) technology that seamlessly integrates protein biosynthesis with asparagine-linked protein glycosylation. This technology leverages a glyco-optimized Escherichia coli strain to source cell extracts that are selectively enriched with glycosylation components, including oligosaccharyltransferases (OSTs) and lipid-linked oligosaccharides (LLOs). The resulting extracts enable a one-pot reaction scheme for efficient and site-specific glycosylation of target proteins. The CFGpS platform is highly modular, allowing the use of multiple distinct OSTs and structurally diverse LLOs. As such, we anticipate CFGpS will facilitate fundamental understanding in glycoscience and make possible applications in on demand biomanufacturing of glycoproteins.

The ribosome is the cell's factory for protein synthesis. With protein synthesis rates of up to 20 amino acids per second and at an accuracy of 99.99%, the extraordinary catalytic capacity of the bacterial translation machinery has attracted extensive efforts to engineer, reconstruct, and repurpose it for biochemical studies and novel functions. Despite these efforts, the potential for harnessing the translation apparatus to manufacture bio-based products beyond natural limits remains underexploited, and fundamental constraints on the chemistry that the ribosome's RNA-based active site can carry out are unknown. This review aims to cover the past and present advances in ribosome design and engineering to understand the fundamental biology of the ribosome to facilitate the construction of synthetic manufacturing machines. The prospects for the development of engineered, or designer, ribosomes for novel polymer synthesis are reviewed, future challenges are considered, and promising advances in a variety of applications are discussed.

Glycosylation is an abundant post-translational modification that is important in disease and biotechnology. Current methods to understand and engineer glycosylation cannot sufficiently explore the vast experimental landscapes required to accurately predict and design glycosylation sites modified by glycosyltransferases. Here we describe a systematic platform for glycosylation sequence characterization and optimization by rapid expression and screening (GlycoSCORES), which combines cell-free protein synthesis and mass spectrometry of self-assembled monolayers. We produced six N- and O-linked polypeptide-modifying glycosyltransferases from bacteria and humans in vitro and rigorously determined their substrate specificities using 3,480 unique peptides and 13,903 unique reaction conditions. We then used GlycoSCORES to optimize and design small glycosylation sequence motifs that directed efficient N-linked glycosylation in vitro and in the Escherichia coli cytoplasm for three heterologous proteins, including the human immunoglobulin Fc domain. We find that GlycoSCORES is a broadly applicable method to facilitate fundamental understanding of glycosyltransferases and engineer synthetic glycoproteins.

Cell-free protein synthesis (CFPS) systems enable the production of protein without the use of living, intact cells. An emerging area of interest is to use CFPS systems to characterize individual elements for genetic programs [e.g. promoters, ribosome binding sites (RBS)]. To enable this research area, robust CFPS systems must be developed from new chassis organisms. One such chassis is the Gram-negative Pseudomonas bacteria, which have been studied extensively for their diverse metabolism with promises in the field of bioremediation and biosynthesis. Here, we report the development and optimization of a high-yielding (198 ± 5.9 µg/ml) batch CFPS system from Pseudomonas putida ATCC 12633. Importantly, both circular and linear DNA templates can be applied directly to the CFPS reaction to program protein synthesis. Therefore, it is possible to prepare hundreds or even thousands of DNA templates without time-consuming cloning work. This opens the possibility to rapidly assess and validate genetic part performance in vitro before performing experiments in cells. To validate the P. putida CFPS system as a platform for prototyping genetic parts, we designed and constructed a library consisting of 15 different RBSs upstream of the reporter protein sfGFP, which covered an order of magnitude range in expression. Looking forward, our P. putida CFPS platform will not only expand the protein synthesis toolkit for synthetic biology but also serve as a platform in expediting the screening and prototyping of gene regulatory elements., (© The Author(s) 2018. Published by Oxford University Press.)

Lantibiotics are ribosomally synthesized and post-translationally modified peptide natural products that contain thioether cross-links formed by lanthionine and methyllanthionine residues. They exert potent antimicrobial activity against Gram-positive bacteria. We herein report production of analogues of two lantibiotics, lacticin 481 and nisin, that contain nonproteinogenic amino acids using two different strategies involving amber stop codon suppression technology. These methods complement recent alternative approaches to incorporate nonproteinogenic amino acids into lantibiotics.

Centralized facilities for genetic engineering, or "biofoundries", offer the potential to design organisms to address emerging needs in medicine, agriculture, industry, and defense. The field has seen rapid advances in technology, but it is difficult to gauge current capabilities or identify gaps across projects. To this end, our foundry was assessed via a timed "pressure test", in which 3 months were given to build organisms to produce 10 molecules unknown to us in advance. By applying a diversity of new approaches, we produced the desired molecule or a closely related one for six out of 10 targets during the performance period and made advances toward production of the others as well. Specifically, we increased the titers of 1-hexadecanol, pyrrolnitrin, and pacidamycin D, found novel routes to the enediyne warhead underlying powerful antimicrobials, established a cell-free system for monoterpene production, produced an intermediate toward vincristine biosynthesis, and encoded 7802 individually retrievable pathways to 540 bisindoles in a DNA pool. Pathways to tetrahydrofuran and barbamide were designed and constructed, but toxicity or analytical tools inhibited further progress. In sum, we constructed 1.2 Mb DNA, built 215 strains spanning five species ( Saccharomyces cerevisiae, Escherichia coli, Streptomyces albidoflavus, Streptomyces coelicolor, and Streptomyces albovinaceus), established two cell-free systems, and performed 690 assays developed in-house for the molecules.

Cell-free protein synthesis has emerged as a powerful approach for expanding the range of genetically encoded chemistry into proteins. Unfortunately, efforts to site-specifically incorporate multiple non-canonical amino acids into proteins using crude extract-based cell-free systems have been limited by release factor 1 competition. Here we address this limitation by establishing a bacterial cell-free protein synthesis platform based on genomically recoded Escherichia coli lacking release factor 1. This platform was developed by exploiting multiplex genome engineering to enhance extract performance by functionally inactivating negative effectors. Our most productive cell extracts enabled synthesis of 1,780 ± 30 mg/L superfolder green fluorescent protein. Using an optimized platform, we demonstrated the ability to introduce 40 identical p-acetyl-L-phenylalanine residues site specifically into an elastin-like polypeptide with high accuracy of incorporation ( ≥ 98%) and yield (96 ± 3 mg/L). We expect this cell-free platform to facilitate fundamental understanding and enable manufacturing paradigms for proteins with new and diverse chemistries.

The Shine-Dalgarno (SD) sequence motif is frequently found upstream of protein coding genes and is thought to be the dominant mechanism of translation initiation used by bacteria. Experimental studies have shown that the SD sequence facilitates start codon recognition and enhances translation initiation by directly interacting with the highly conserved anti-SD sequence on the 30S ribosomal subunit. However, the proportion of SD-led genes within a genome varies across species and the factors governing this variation in translation initiation mechanisms remain largely unknown. Here, we conduct a phylogenetically informed analysis and find that species capable of rapid growth contain a higher proportion of SD-led genes throughout their genomes. We show that SD sequence utilization covaries with a suite of genomic features that are important for efficient translation initiation and elongation. In addition to these endogenous genomic factors, we further show that exogenous environmental factors may influence the evolution of translation initiation mechanisms by finding that thermophilic species contain significantly more SD-led genes than mesophiles. Our results demonstrate that variation in translation initiation mechanisms across bacterial species is predictable and is a consequence of differential life-history strategies related to maximum growth rate and environmental-specific constraints., (© The Author 2017. Published by Oxford University Press on behalf of the Society for Molecular Biology and Evolution.)

Protein glycosylation, or the attachment of sugar moieties (glycans) to proteins, is important for protein stability, activity, and immunogenicity. However, understanding the roles and regulations of site-specific glycosylation events remains a significant challenge due to several technological limitations. These limitations include a lack of available tools for biochemical characterization of enzymes involved in glycosylation. A particular challenge is the synthesis of oligosaccharyltransferases (OSTs), which catalyze the attachment of glycans to specific amino acid residues in target proteins. The difficulty arises from the fact that canonical OSTs are large (>70 kDa) and possess multiple transmembrane helices, making them difficult to overexpress in living cells. Here, we address this challenge by establishing a bacterial cell-free protein synthesis platform that enables rapid production of a variety of OSTs in their active conformations. Specifically, by using lipid nanodiscs as cellular membrane mimics, we obtained yields of up to 420 μg/ml for the single-subunit OST enzyme, "Protein glycosylation B" (PglB) from Campylobacter jejuni, as well as for three additional PglB homologs from Campylobacter coli, Campylobacter lari, and Desulfovibrio gigas. Importantly, all of these enzymes catalyzed N-glycosylation reactions in vitro with no purification or processing needed. Furthermore, we demonstrate the ability of cell-free synthesized OSTs to glycosylate multiple target proteins with varying N-glycosylation acceptor sequons. We anticipate that this broadly applicable production method will advance glycoengineering efforts by enabling preparative expression of membrane-embedded OSTs from all kingdoms of life., (© 2017 Wiley Periodicals, Inc.)

Despite decades of accumulated knowledge about proteins and their post-translational modifications (PTMs), numerous questions remain regarding their molecular composition and biological function. One of the most fundamental queries is the extent to which the combinations of DNA-, RNA- and PTM-level variations explode the complexity of the human proteome. Here, we outline what we know from current databases and measurement strategies including mass spectrometry-based proteomics. In doing so, we examine prevailing notions about the number of modifications displayed on human proteins and how they combine to generate the protein diversity underlying health and disease. We frame central issues regarding determination of protein-level variation and PTMs, including some paradoxes present in the field today. We use this framework to assess existing data and to ask the question, "How many distinct primary structures of proteins (proteoforms) are created from the 20,300 human genes?" We also explore prospects for improving measurements to better regularize protein-level biology and efficiently associate PTMs to function and phenotype.

Building biosynthetic pathways and engineering metabolic reactions in cells can be time-consuming due to complexities in cellular metabolism. These complexities often convolute the combinatorial testing of biosynthetic pathway designs needed to define an optimal biosynthetic system. To simplify the optimization of biosynthetic systems, we recently reported a new cell-free framework for pathway construction and testing. In this framework, multiple crude-cell extracts are selectively enriched with individual pathway enzymes, which are then mixed to construct full biosynthetic pathways on the time scale of a day. This rapid approach to building pathways aids in the study of metabolic pathway performance by providing a unique freedom of design to modify and control biological systems for both fundamental and applied biotechnology. The goal of this work was to demonstrate the ability to probe biosynthetic pathway performance in our cell-free framework by perturbing physiochemical conditions, using n-butanol synthesis as a model. We carried out three unique case studies. First, we demonstrated the power of our cell-free approach to maximize biosynthesis yields by mapping physiochemical landscapes using a robotic liquid-handler. This allowed us to determine that NAD and CoA are the most important factors that govern cell-free n-butanol metabolism. Second, we compared metabolic profile differences between two different approaches for building pathways from enriched lysates, heterologous expression and cell-free protein synthesis. We discover that phosphate from PEP utilization, along with other physiochemical reagents, during cell-free protein synthesis-coupled, crude-lysate metabolic system operation inhibits optimal cell-free n-butanol metabolism. Third, we show that non-phosphorylated secondary energy substrates can be used to fuel cell-free protein synthesis and n-butanol biosynthesis. Taken together, our work highlights the ease of using cell-free systems to explore physiochemical perturbations and suggests the need for a more controllable, multi-step, separated cell-free framework for future pathway prototyping and enzyme discovery efforts., (Copyright © 2017 International Metabolic Engineering Society. Published by Elsevier Inc. All rights reserved.)

The unequal utilization of synonymous codons affects numerous cellular processes including translation rates, protein folding and mRNA degradation. In order to understand the biological impact of variable codon usage bias (CUB) between genes and genomes, it is crucial to be able to accurately measure CUB for a given sequence. A large number of metrics have been developed for this purpose, but there is currently no way of systematically testing the accuracy of individual metrics or knowing whether metrics provide consistent results. This lack of standardization can result in false-positive and false-negative findings if underpowered or inaccurate metrics are applied as tools for discovery. Here, we show that the choice of CUB metric impacts both the significance and measured effect sizes in numerous empirical datasets, raising questions about the generality of findings in published research. To bring about standardization, we developed a novel method to create synthetic protein-coding DNA sequences according to different models of codon usage. We use these benchmark sequences to identify the most accurate and robust metrics with regard to sequence length, GC content and amino acid heterogeneity. Finally, we show how our benchmark can aid the development of new metrics by providing feedback on its performance compared to the state of the art., (© 2018 The Author(s).)

Engineering biological systems for the production of biofuels and bioproducts holds great potential to transform the bioeconomy, but often requires laborious, time-consuming design-build-test cycles. For decades cell-free systems have offered quick and facile approaches to study enzymes with hopes of informing cellular processes, mainly in the form of purified single-enzyme activity assays. Over the past 20 years, cell-free systems have grown to include multienzymatic systems, both purified and crude. By decoupling cellular growth objectives from enzyme pathway engineering objectives, cell-free systems provide a controllable environment to direct substrates toward a single, desired product. Cell-free approaches are being developed for prototyping and for biomanufacturing. In prototyping applications, the idea is to use cell-free systems to test and optimize biosynthetic pathways before implementation in live cells and scale-up. We present a detailed method for the generation of crude lysates for cell-free pathway prototyping, mix-and-match cell-free metabolic engineering using preenriched lysates, and cell-free protein synthesis driven cell-free metabolic engineering. The cell-free synthetic biology methods described herein are generalizable to any biosynthetic pathway of interest and provide a powerful approach to building pathways in crude lysates for the purpose of prototyping. The foundational principle of the presented approach is that we can construct discrete metabolic pathways through modular assembly of cell-free lysates containing enzyme components produced by overexpression in the lysate chassis strain or by cell-free protein synthesis (in vitro production). Overall, the modular and cell-free nature of our pathway prototyping framework is poised to facilitate multiplexed, automated study of biosynthetic pathways to inform systems-level cellular design., (© 2018 Elsevier Inc. All rights reserved.)

The translation system is the cell's factory for protein biosynthesis, stitching together hundreds to thousands of amino acids into proteins, which are required for the structure, function, and regulation of living systems. The extraordinary synthetic capability of this system, which includes the ribosome and its associated factors required for polymerization, has driven extensive efforts to harness it for societal use in areas as diverse as energy, materials, and medicine. A powerful example is recombinant protein production, which has impacted the lives of patients through the synthesis of biopharmaceuticals such as insulin. In nature, however, only limited sets of monomers are utilized, thereby resulting in limited sets of biopolymers (i.e., proteins). Expanding nature's repertoire of ribosomal monomers could yield new classes of enzymes, therapeutics, materials, and chemicals with diverse, genetically encoded chemistry. Here, we discuss recent progress towards engineering ribosomes both in vivo and in vitro. These fundamental and technical breakthroughs open doors for advanced applications in biotechnology and synthetic biology., (Copyright © 2017 Elsevier Ltd. All rights reserved.)

Chinese Hamster Ovary (CHO) cells are routinely optimized to stably express monoclonal antibodies (mAbs) at high titers. At the early stages of lead isolation and optimization, hundreds of sequences for the target protein of interest are screened. Typically, cell-based transient expression technology platforms are used for expression screening, but these can be time- and resource-intensive. Here, we have developed a cell-free protein synthesis (CFPS) platform utilizing a commercially available CHO extract for the rapid in vitro synthesis of active, aglycosylated mAbs. Specifically, we optimized reaction conditions to maximize protein yields, established an oxidizing environment to enable disulfide bond formation, and demonstrated the importance of temporal addition of heavy chain and light chain plasmids for intact mAb production. Using our optimized platform, we demonstrate for the first time to our knowledge the cell-free synthesis of biologically active, intact mAb at >100 mg/L using a eukaryotic-based extract. We then explored the utility of our system as a tool for ranking yields of candidate antibodies. Unlike stable or transient transfection-based screening, which requires a minimum of 7 days for setup and execution, results using our CHO-based CFPS platform are attained within 2 days and it is well-suited for automation. Further development would provide a tool for rapid, high-throughput prediction of mAb expression ranking to accelerate design-build-test cycles required for antibody expression and engineering. Looking forward, the CHO-based CFPS platform could facilitate the synthesis of toxic proteins as well.

To build replicating systems with new functions, the engineering of existing biological machineries requires a sensible strategy. Protein synthesis Using Recombinant Elements (PURE) system consists of the desired components for transcription, translation, aminoacylation and energy regeneration. PURE might be the basis for a radically alterable, lifelike system after optimization. Here, we regenerated 54 E. coli ribosomal (r-) proteins individually from DNA templates in the PURE system. We show that using stable isotope labeling with amino acids, mass spectrometry based quantitative proteomics could detect 26 of the 33 50S and 20 of the 21 30S subunit r-proteins when coexpressed in batch format PURE system. By optimizing DNA template concentrations and adapting a miniaturized Fluid Array Device with optimized feeding solution, we were able to cogenerate and detect at least 29 of the 33 50S and all of the 21 30S subunit r-proteins in one pot. The boost on yield of a single r-protein in coexpression pool varied from ∼1.5 to 5-fold compared to the batch mode, with up to ∼2.4 μM yield for a single r-protein. Reconstituted ribosomes under physiological condition from PURE system synthesized 30S r-proteins and native 16S rRNA showed ∼13% activity of native 70S ribosomes, which increased to 21% when supplemented with GroEL/ES. This work also points to what is still needed to obtain self-replicating synthetic ribosomes in situ in the PURE system.

Cell-free protein synthesis (CFPS) has emerged as a powerful platform for applied biotechnology and synthetic biology, with a range of applications in synthesizing proteins, evolving proteins, and prototyping genetic circuits. To expand the current CFPS repertoire, we report here the development and optimization of a Streptomyces-based CFPS system for the expression of GC-rich genes. By developing a streamlined crude extract preparation protocol and optimizing reaction conditions, we were able to achieve active enhanced green fluorescent protein (EGFP) yields of greater than 50 μg/mL with batch reactions lasting up to 3 h. By adopting a semi-continuous reaction format, the EGFP yield could be increased to 282 ± 8 μg/mL and the reaction time was extended to 48 h. Notably, our extract preparation procedures were robust to multiple Streptomyces lividans and Streptomyces coelicolor strains, although expression yields varied. We show that our optimized Streptomyces lividans system provides benefits when compared to an Escherichia coli-based CFPS system for increasing percent soluble protein expression for four Streptomyces-originated high GC-content genes that are involved in biosynthesis of the nonribosomal peptides tambromycin and valinomycin. Looking forward, we believe that our Streptomyces-based CFPS system will contribute significantly towards efforts to express complex natural product gene clusters (e.g., nonribosomal peptides and polyketides), providing a new avenue for obtaining and studying natural product biosynthesis pathways. Biotechnol. Bioeng. 2017;114: 1343-1353. © 2017 Wiley Periodicals, Inc., (© 2017 Wiley Periodicals, Inc.)

The ability to site-specifically incorporate non-canonical amino acids (ncAAs) into proteins has made possible the study of protein structure and function in fundamentally new ways, as well as the bio synthesis of unnatural polymers. However, the task of site-specifically incorporating multiple ncAAs into proteins with high purity and yield continues to present a challenge. At the heart of this challenge lies the lower efficiency of engineered orthogonal translation system components compared to their natural counterparts (e.g., translation elements that specifically use a ncAA and do not interact with the cell's natural translation apparatus). Here, we show that evolving and tuning expression levels of multiple components of an engineered translation system together as a whole enhances ncAA incorporation efficiency. Specifically, we increase protein yield when incorporating multiple p-azido-phenylalanine(pAzF) residues into proteins by (i) evolving the Methanocaldococcus jannaschii p-azido-phenylalanyl-tRNA synthetase anti-codon binding domain, (ii) evolving the elongation factor Tu amino acid-binding pocket, and (iii) tuning the expression of evolved translation machinery components in a single vector. Use of the evolved translation machinery in a genomically recoded organism lacking release factor one enabled enhanced multi-site ncAA incorporation into proteins. We anticipate that our approach to orthogonal translation system development will accelerate and expand our ability to site-specifically incorporate multiple ncAAs into proteins and biopolymers, advancing new horizons for synthetic and chemical biotechnology. Biotechnol. Bioeng. 2017;114: 1074-1086. © 2016 Wiley Periodicals, Inc., (© 2016 Wiley Periodicals, Inc.)