Bacteriophages (phages) are critical players in the dynamics and function of microbial communities and drive processes as diverse as global biogeochemical cycles and human health. Phages tend to be predators finely tuned to attack specific hosts, even down to the strain level, which in turn defend themselves using an array of mechanisms. However, to date, efforts to rapidly and comprehensively identify bacterial host factors important in phage infection and resistance have yet to be fully realized. Here, we globally map the host genetic determinants involved in resistance to 14 phylogenetically diverse double-stranded DNA phages using two model Escherichia coli strains (K-12 and BL21) with known sequence divergence to demonstrate strain-specific differences. Using genome-wide lossof-function and gain-of-function genetic technologies, we are able to confirm previously described phage receptors as well as uncover a number of previously unknown host factors that confer resistance to one or more of these phages. We uncover differences in resistance factors that strongly align with the susceptibility of K-12 and BL21 to specific phage. We also identify both phage-specific mechanisms, such as the unexpected role of cyclic-di-GMP in host sensitivity to phage N4, and more generic defenses, such as the overproduction of colanic acid capsular polysaccharide that defends against a wide array of phages. Our results indicate that host responses to phages can occur via diverse cellular mechanisms. Our systematic and high-throughput genetic workflow to characterize phage-host interaction determinants can be extended to diverse bacteria to generate datasets that allow predictive models of how phage-mediated selection will shape bacterial phenotype and evolution. The results of this study and future efforts to map the phage resistance landscape will lead to new insights into the coevolution of hosts and their phage, which can

The ability to engineer natural proteins is pivotal to a future, pragmatic biology. CRISPR proteins have revolutionized genome modification, yet the CRISPR-Cas9 scaffold is not ideal for fusions or activation by cellular triggers. Here, we show that a topological rearrangement of Cas9 using circular permutation provides an advanced platform for RNA-guided genome modification and protection. Through systematic interrogation, we find that protein termini can be positioned adjacent to bound DNA, offering a straightforward mechanism for strategically fusing functional domains. Additionally, circular permutation enabled protease-sensing Cas9s (ProCas9s), a unique class of single-molecule effectors possessing programmable inputs and outputs. ProCas9s can sense a wide range of proteases, and we demonstrate that ProCas9 can orchestrate a cellular response to pathogen-associated protease activity. Together, these results provide a toolkit of safer and more efficient genome-modifying enzymes and molecular recorders for the advancement of precision genome engineering in research, agriculture, and biomedicine.

Bacteriophages (phages) are critical players in the dynamics and function of microbial communities and drive processes as diverse as global biogeochemical cycles and human health. Phages tend to be predators finely tuned to attack specific hosts, even down to the strain level, which in turn defend themselves using an array of mechanisms. However, to date, efforts to rapidly and comprehensively identify bacterial host factors important in phage infection and resistance have yet to be fully realized. Here, we globally map the host genetic determinants involved in resistance to 14 phylogenetically diverse double-stranded DNA phages using two model Escherichia coli strains (K-12 and BL21) with known sequence divergence to demonstrate strain-specific differences. Using genome-wide loss-of-function and gain-of-function genetic technologies, we are able to confirm previously described phage receptors as well as uncover a number of previously unknown host factors that confer resistance to one or more of these phages. We uncover differences in resistance factors that strongly align with the susceptibility of K-12 and BL21 to specific phage. We also identify both phage specific mechanisms, such as the unexpected role of cyclic-di-GMP in host sensitivity to phage N4, and more generic defenses, such as the overproduction of colanic acid capsular polysaccharide that defends against a wide array of phages. Our results indicate that host responses to phages can occur via diverse cellular mechanisms. Our systematic and highthroughput genetic workflow to characterize phage-host interaction determinants can be extended to diverse bacteria to generate datasets that allow predictive models of how phagemediated selection will shape bacterial phenotype and evolution. The results of this study and future efforts to map the phage resistance landscape will lead to new insights into the coevolution of hosts and their phage, which can ultimately be used to design better phage therapeutic treatments and tools for precision microbiome engineering.3 Introduction:

22Genome-wide repression screens using CRISPR interference (CRISPRi) have enabled the high-throughput 23 identification of essential genes in bacteria. However, there is a lack of functional studies leveraging 24 CRISPRi to systematically explore targeting of both the coding and non-coding genome in bacteria. Here 25 we perform CRISPRi screens in Escherichia coli MG1655 K-12 targeting ~13,000 genomic features, 26including nearly all protein-coding genes, non-coding RNAs, promoters, and transcription factor binding 27 sites (TFBSs) using a ~33,000-member sgRNA library, which represents the most compact and 28 comprehensive genome-wide CRISPRi library in E. coli to date. Our data reveal insights into the conditional 29 essentiality of the genome with key refinements to screen design and profiling. First, we demonstrate that 30 strong fitness defects associated with essential cellular processes can be resolved using inducible time-31 series measurements. We show that knockdowns of different classes of genes exhibit distinct, transient 32 responses that are correlated to gene function with genes involved in translation exhibiting the strongest 33 responses. We also query feature essentiality across several biochemical conditions and show that several 34 genes, sRNAs, and operons exhibit conditional phenotypes not reported by previous high-throughput 35 efforts. Second, we evaluate systematically targeting non-genic features (promoters and TFBSs) in the E. 36 coli genome. We show that promoter-targeting guides can be used to add phenotypic confidence to 37 promoter annotations and verify computationally predicted promoters. In contrast to prior studies, we find 38 that promoter knockdowns exhibit a strong targeting orientation dependency where targeting the non-39 template strand of the promoter closest to the target gene is more effective in knocking down gene 40 expression than other promoter targeting orientations. Unlike eukaryotic genomes, we note that interpreting 41 the effects of TFBS targeting is particularly challenging due to the small size of such features and their 42 proximity to and overlap with other genomic features. Together, this work reveals novel conditionally 43 essential gene phenotypes, provides a characterized set of sgRNAs for future E. coli CRISPRi screens, 44 and highlights considerations for CRISPRi library design and screening for microbial genome 45 characterization. 46 47 48 50 51 The nuclease deactivated dCas9 protein has been developed as a powerful tool for programmable gene 52 repression 1 , and the ability to induce genetic perturbation at a user-defined time -a feature not available 53 in conventional gene disruption or deletion techniques -has enabled the CRISPRi-mediated 54 characterization of essential genes in a number of bacteria 2-7 . The programmability of CRISPRi targeting 55 also enables the interrogation of smaller non-coding DNA (ncDNA) features such as non-coding RNA 56 (ncRNA) genes, promoters, and transcription factor binding sites (TFBSs). ncDNA features, which 57 r...

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

Precision genome editing accelerates the discovery of the genetic determinants of phenotype and the engineering of novel behaviors in organisms. Advances in DNA synthesis and recombineering have enabled high-throughput engineering of genetic circuits and biosynthetic pathways via directed mutagenesis of bacterial chromosomes. However, the highest recombination efficiencies have to date been reported in persistent mutator strains, which suffer from reduced genomic fidelity. The absence of inducible transcriptional regulators in these strains also prevents concurrent control of genome engineering tools and engineered functions. Here, we introduce a new recombineering platform strain, BioDesignER, which incorporates (i) a refactored λ-Red recombination system that reduces toxicity and accelerates multi-cycle recombination, (ii) genetic modifications that boost recombination efficiency, and (iii) four independent inducible regulators to control engineered functions. These modifications resulted in single-cycle recombineering efficiencies of up to 25% with a 7-fold increase in recombineering fidelity compared to the widely used recombineering strain EcNR2. To facilitate genome engineering in BioDesignER, we have curated eight context­-neutral genomic loci, termed Safe Sites, for stable gene expression and consistent recombination efficiency. BioDesignER is a platform to develop and optimize engineered cellular functions and can serve as a model to implement comparable recombination and regulatory systems in other bacteria.

21Precision genome editing accelerates the discovery of the genetic determinants of phenotype and the 22 engineering of novel behaviors in organisms. Advances in DNA synthesis and recombineering have 23 enabled high-throughput engineering of genetic circuits and biosynthetic pathways via directed 24 mutagenesis of bacterial chromosomes. However, the highest recombination efficiencies have to date 25 been reported in persistent mutator strains, which suffer from reduced genomic fidelity. The absence of 26 inducible transcriptional regulators in these strains also prevents concurrent control of genome 27 engineering tools and engineered functions. Here, we introduce a new recombineering platform strain, 28BioDesignER, which incorporates (1) a refactored λ-Red recombination system that reduces toxicity 29 and accelerates multi-cycle recombination, (2) genetic modifications that boost recombination 30 efficiency, and (3) four independent inducible regulators to control engineered functions. These 31 modifications resulted in single-cycle recombineering efficiencies of up to 25% with a seven-fold 32 increase in recombineering fidelity compared to the widely used recombineering strain EcNR2. To 33 facilitate genome engineering in BioDesignER, we have curated eight context-neutral genomic loci,34 termed Safe Sites, for stable gene expression and consistent recombination efficiency. BioDesignER is 35 a platform to develop and optimize engineered cellular functions and can serve as a model to implement 36 comparable recombination and regulatory systems in other bacteria. 37 INTRODUCTION 38The design-build-test (DBT) cycle is a common paradigm used in engineering disciplines. Within the 39 context of synthetic biology it is employed to engineer user-defined cellular functions for applications 40 such as metabolic engineering, biosensing, and therapeutics (1, 2). The rapid prototyping of engineered 41 functions has been facilitated by advances in in vitro DNA assembly, and plasmids have traditionally 42 been used to implement designs in vivo given their ease-of-assembly and portability. However, for 43 deployment in contexts beyond the laboratory such as large-scale industrial bioprocesses or among 44 complex microbial communities, plasmid-based circuits suffer from multiple limitations: high intercellular 45 variation in gene expression, genetic instability from random partitioning of plasmids during cell division, 46 and plasmid loss in environments for which antibiotic use could disrupt native microbial communities or 47 is economically infeasible (3, 4). These shortcomings can be ameliorated once a design is transferred 48 from a plasmid to the host genome, which offers improved genetic stability and lower expression 49 variation (5) along with reduced metabolic load (6). However, behaviors optimized for plasmid contexts 50 often do not map predictably to the genome. As such, building and testing designs directly on the 51 genome can reduce the DBT cycle time and facilitate engineering cellular programs for complex 52 e...

Our knowledge of the relationship between the gut microbiome and health has rapidly expanded in recent years. Diet has been shown to have causative effects on microbiome composition, which can have subsequent implications on health. Soylent 2.0 is a liquid meal replacement drink that satisfies nearly 20% of all recommended daily intakes per serving. This study aims to characterize the changes in gut microbiota composition resulting from a short-term Soylent diet. Fourteen participants were separated into two groups: 5 in the regular diet group and 9 in the Soylent diet group. The regular diet group maintained a diet closely resembling self-reported regular diets. The Soylent diet group underwent three dietary phases: A) a regular diet for 2 days, B) a Soylent-only diet (five servings of Soylent daily and water as needed) for 4 days, and C) a regular diet for 4 days. Daily logs self-reporting diet, Bristol stool ratings, and any abdominal discomfort were electronically submitted. Eight fecal samples per participant were collected using fecal sampling kits, which were subsequently sent to uBiome, Inc. for sample processing and V4 16S rDNA sequencing. Reads were clustered into operational taxonomic units (OTUs) and taxonomically identified against the GreenGenes 16S database. We find that an individual’s alpha-diversity is not significantly altered during a Soylent-only diet. In addition, principal coordinate analysis using the unweighted UniFrac distance metric shows samples cluster strongly by individual and not by dietary phase. Among Soylent dieters, we find a significant increase in the ratio of Bacteroidetes to Firmicutes abundance, which is associated with several positive health outcomes, including reduced risks of obesity and intestinal inflammation.