The Cas9 nuclease has become central to modern methods and technologies in synthetic biology, largely due to the ease with which it can be targeted to specific DNA loci via guide RNAs (gRNAs). Reports vary widely on the actual specificity of this targeting, with some studies observing 60% of gRNAs possessing no activity against the genome, yet an assumption persists within the E. coli community that inactive gRNAs are rare. To resolve these contradictions, we evaluated the activity of 463 000 unique gRNAs in the E. coli K12 MG1655 genome. We show that the overwhelming majority (at least 93%) of unique gRNAs are functional while only 0.3% are nonfunctional. These nonfunctional gRNAs exhibit strong spacer self-interaction, which can either be excluded using a simple design rule or "repaired" during library design. Finally, this work provides the greater microbial synthetic biology community both a set of nearly half a million empirically evaluated E. coli gRNAs as well as a thoroughly evaluated experimental procedure, complete with appropriate controls for Cas9 activity, for conducting Cas9 assays in E. coli specifically and bacteria more generally. Lastly, we have produced a webapp to allow users to easily browse and extract gRNA sequences from the E. coli genome, which can be accessed at https://grna.ornl.gov.

Cas9-based genome editing technologies can rapidly generate mutations to probe a diverse array of mutant genotypes. However, aberrant Cas9 nuclease translation and activity can occur despite the use of inducible promoters to control expression, leading to extensive cell death. This background killing caused by promoter leakiness severely limits the application of Cas9 for generating mutant libraries because of the potential for population skew. We demonstrate the utility of temperature sensitive RNA elements as a layer of post-transcriptional regulation to reduce the impact of promoter leak. We observe significant temperature-dependent increases in cell survival when certain RNA thermometers (RNATs) are placed upstream of the cas9 coding sequence. We also show that the most highly repressing RNAT, hsp17rep, significantly reduces population skew with a library of characterized guide RNAs in Escherichia coli. This strategy should be applicable to all bacterial Cas9-based methods and technologies.

RNA plays a central role in plants, governing various cellular and physiological processes. Monitoring its dynamic abundance provides a discerning understanding of molecular mechanisms underlying plant responses to internal (developmental) and external (environmental) stimuli, paving the way for advances in plant biotechnology to engineer crops with improved resilience, quality and productivity. In general, traditional methods for analysis of RNA abundance in plants require destructive, labour-intensive and time-consuming assays. To overcome these limitations, we developed a transformative innovation for in vivo RNA imaging in plants. Specifically, we established a synthetic split ribozyme system that converts various RNA signals to orthogonal protein outputs, enabling in vivo visualisation of various RNA signals in plants. We demonstrated the utility of this system in transient expression experiments (i.e., leaf infiltration in Nicotiana benthamiana) to detect RNAs derived from transgenes and tobacco rattle virus, respectively. Also, we successfully engineered a split ribozyme-based biosensor in Arabidopsis thaliana for in vivo visualisation of endogenous gene expression at the cellular level, demonstrating the feasibility of multi-scale (e.g., cellular and tissue level) RNA imaging in plants. Furthermore, we developed a platform for easy incorporation of different protein outputs, allowing for flexible choice of reporters to optimise the detection of target RNAs.

Cyanobacteria are promising microbial platforms for a diverse set of biotechnology applications, from living materials to photosynthetic chemical production, but are less well characterized than commonly engineered microbes such as Escherichia coli. This study facilitates genetic engineering in Synechococcus sp. PCC 7002, a fast-growing, halotolerant, and naturally competent strain, by identifying ten native methylation motifs and designing shuttle strains that mimic the native methylation state by expressing a subset of heterologous methyltransferases. DNA methylation in E. coli with as few as two active methyltransferases increased transformation efficiency up to 30-fold across four distinct integration sites in PCC 7002. This work provides an experimental framework to bypass native restriction-modification systems for efficient genome editing and metabolic engineering in nonmodel bacteria.

The field of synthetic biology is essential to the continued development of a bio-based economy, creating mechanisms to supply carbon needed in the economy by both converting existing end-of-life wastes as well as by creating novel, purpose-grown and sustainable feedstocks. Here, we first discuss the near- and long-term resources available for use as feedstocks for bioconversion as well as the output molecules needed for building the foundation of an expanded bio-based economy. We then outline the organisms and phenotypic traits that are needed for the performance-advantaged chassis organisms of the future. Furthermore, we detail the advances, challenges, and opportunities in both microbial and plant synthetic biology relevant to expanding the bio-based economy. Finally, we explore technologies that have and will further enable advances in synthetic biology and the greater bio-based economy.

Cyanobacteria are the oldest form of photosynthetic life on Earth and contribute to primary production in nearly every habitat, from permafrost to hot springs. Despite longstanding interest in the acclimation of these microbes, it remains poorly understood and challenging to rewire. This study uses a high-density, genome-wide CRISPR interference screen to examine the influence of gene-specific transcriptional variation on the growth of Synechococcus sp. PCC 7002 under environmental extremes. Surprisingly, many partial knockdowns enhanced fitness under cold monochromatic conditions. Transcriptional repression of genes for core subunits of the NDH-1 complex, which are important for photosynthesis and carbon uptake, improved growth rates under both red and blue light but at distinct, color-specific optima. Most genes with fitness-improving knockdowns were distinct to each light color, and dual-target transcriptional repression produced nonadditive effects. Findings reveal diverse routes to improved acclimation in cyanobacteria (e.g., attenuation of genes involved in CO2 uptake, light harvesting, translation, and purine metabolism) and provide an approach for using gradients in sgRNA activity to pinpoint biochemically influential transcriptional changes in cells.

The emerging bioeconomy presents a promising solution to both economic and environmental challenges. Within the bioeconomy, plants serve as a renewable, sustainable, and cost-effective source of foods, fuels, chemicals, and materials. However, traditional breeding and single-gene engineering approaches fall short in addressing complex traits (e.g., drought tolerance, disease resistance, yield, nutrient use efficiency) which are controlled by multiple genes. The complexity of plant biology often necessitates the use of multigene engineering (MGE), which involves simultaneous ectopic expression, up/down-regulation, or editing of multiple genes, to enhance plant traits relevant to the bioeconomy. These genes may be associated with distinct traits or function as components of specific metabolic and regulatory pathways. This review summarizes current technologies for MGE within the synthetic biology-driven Design-Build-Test-Learn (DBTL) framework, detailing its four key stages: Design - gene construct development; Build - DNA assembly and plant transformation; Test - the molecular, biochemical, and physiological characterization of engineered plants; and Learn - computational modeling to refine, multiplex and iterate the process. Despite good progress in the applications of MGE in biofortification, metabolic engineering, and stress resilience, challenges remain in construct stability, coordinated gene expression, and regulatory predictability. We identified optimization paths and future directions to accelerate MGE deployment in sustainable agriculture, with possible societal benefits including reduced production costs, increased yield, and improved food and nutritional security.

Cas9-based genome editing technologies can rapidly generate mutations to probe a diverse array of mutant genotypes. However, aberrant Cas9 nuclease translation and activity can occur despite the use of inducible promoters to control expression, leading to extensive cell death. This background killing caused by promoter leakiness severely limits the application of Cas9 for generating mutant libraries because of the potential for population skew. We demonstrate the utility of temperature sensitive RNA elements as a layer of post-transcriptional regulation to reduce the impact of promoter leak. We observe significant temperature-dependent increases in cell survival when certain RNA thermometers (RNATs) are placed upstream of the cas9 coding sequence. We show that the most highly repressing RNAT, hsp17rep , significantly reduces population skew with a library of characterized guide RNAs (gRNAs). This information should be applicable to all Cas9-based methods and technologies. Notice This manuscript has been authored by UT-Battelle, LLC, under contract DE-AC05-00OR22725 with the US Department of Energy (DOE). The US government retains and the publisher, by accepting the article for publication, acknowledges that the US government retains a nonexclusive, paid-up, irrevocable, worldwide license to publish or reproduce the published form of this manuscript, or allow others to do so, for US government purposes. DOE will provide public access to these results of federally sponsored research in accordance with the DOE Public Access Plan (). ![Figure][1] Graphical Abstract ### Competing Interest Statement The authors have declared no competing interest. United States Department of Energy, https://ror.org/01bj3aw27, ERKP886 [1]: pending:yes

With the expansion of domesticated microbes producing biomaterials and chemicals to support a growing circular bioeconomy, the variety of waste and sustainable substrates that can support microbial growth and production will also continue to expand. The diversity of these microbes also requires a range of compatible genetic tools to engineer improved robustness and economic viability. As we still do not fully understand the function of many genes in even highly studied model microbes, engineering improved microbial performance requires introducing genome-scale genetic modifications followed by screening or selecting mutants that enhance growth under prohibitive conditions encountered during production. These approaches include adaptive laboratory evolution, random or directed mutagenesis, transposon-mediated gene disruption, or CRISPR interference (CRISPRi). Although any of these approaches may be applicable for identifying engineering targets, here we focus on using CRISPRi to reduce the time required to engineer more robust microbes for industrial applications.One-Sentence Summary The development of genome scale CRISPR-based libraries in new microbes enables discovery of genetic factors linked to desired traits for engineering more robust microbial systems. Graphical Abstract

Cyanobacteria are promising microbial platforms for a myriad of biotechnological applications, from sustainable biomaterials to photosynthetic chemical production, but still lack the breadth of genetic tools available for more commonly engineered microbes such as Escherichia coli . This study develops genetic tools to enhance the transformation efficiency and heterologous gene expression in Picosynechococcus sp. PCC 7002, a fast-growing, halotolerant, and naturally competent strain. Integration of fluorescent reporter cassettes across the genome revealed an integration site that yields a fourfold improvement in gene expression relative to previously reported sites. Protocol optimization and engineered DNA methylation in E. coli increased transformation efficiency by over tenfold. This work provides an experimental framework for efficient genome editing and metabolic engineering in the model cyanobacterium PCC 7002. ### Competing Interest Statement The authors have declared no competing interest.

ABSTRACT Cyanobacteria are the oldest form of photosynthetic life on Earth and contribute to primary production in nearly every habitat, from permafrost to hot springs. Despite longstanding interest in the biochemical basis of environmental adaptation in these microbes, it remains poorly understood and challenging to re-wire. This study uses a high-density, genome-wide CRISPR interference (CRISPRi) screen to examine the influence of gene-specific transcriptional variation on the growth of Synechococcus sp. PCC 7002 under environmental extrema. Surprisingly, many partial knockdowns enhanced fitness under cold monochromatic conditions. Notably, transcriptional repression of a gene for a core subunit of the NDH-1 complex, which is important for photosynthesis and carbon uptake, improved growth rates under both red and blue light but at distinct, color-specific optima. In general, most genes with fitness-improving knockdowns were distinct to each light color, evidencing unique stress responses and alleviation mechanisms. Multi-target transcriptional repression produced nonadditive effects. Findings reveal diverse mechanisms of environmental adaptation in cyanobacteria and provide a new approach for using gradients in sgRNA activity to pinpoint biochemically influential transcriptional changes in cells. SIGNIFICANCE STATEMENT Cyanobacteria are the most abundant photosynthetic organisms on Earth, where they endure a striking variety of environmental fluctuations. This study examines the biochemical basis of environmental adaptation in Synechococcus sp. PCC 7002, an important model strain, by modulating the expression of every gene in its genome. Results show that partial, but not complete, reduction in the expression of a subset of influential genes can improve growth under cold monochromatic conditions. Optimal expression levels differ between red and blue light and shift with multi-gene adjustments. Findings show how minor transcriptional adjustments can yield major improvements in growth under environmental extrema and provide a powerful systems-level approach for studying—and fine-tuning—the adaptive capacity of microbes.

Bacterial DNA methylation is involved in diverse cellular functions, including modulation of gene expression, DNA repair, and restriction-modification systems for defense against viruses and other foreign DNA. Restriction systems hinder efforts to engineer organisms to produce fuels and chemicals from waste and renewable feedstocks by degrading DNA during transformation. Methylome analysis allows identification of motifs within a bacterial chromosome that may be targeted by native restriction enzymes. Further expression of the corresponding methyltransferases in Escherichia coli allows plasmid DNA to be protected from restriction in the target organism, thereby drastically enhancing transformation efficiency. Nanopore sequencing can detect methylated bases, but software is needed to transform modified base coordinates into methylated motifs. Here, we develop MIJAMP (MIJAMP Is Just A  MethylBED Parser), a software package that was developed to discover methylated motifs from the output of ONT's Modkit or other data in the methylBED format. MIJAMP employs a human-driven refinement strategy that empirically validates all motifs against genome-wide methylation data, thus eliminating incorrect motifs. MIJAMP also reports methylation data on specific, user-defined motifs. Using MIJAMP, we determined the methylated motifs both in a control strain (wild-type E. coli) and in Synecococcus sp. strain PCC7002, laying the foundation for improved transformation in this organism. MIJAMP is available at https://code.ornl.gov/alexander-public/mijamp/. One Sentence Summary: Here we describe software written to discover DNA methylation motifs from nanopore sequencing data.

Clostridium thermocellum is an anaerobic thermophile capable of producing ethanol and other commodity chemicals from lignocellulosic biomass. The insertion of heterologous DNA into the C. thermocellum chromosome is currently achieved via a time-consuming homologous recombination process, where a single stable insertion can take 2-4 weeks or more to construct. In this work, we developed a thermostable version of the Serine recombinase Assisted Genome Engineering (tSAGE) approach for gene insertion in C. thermocellum utilizing a site-specific recombinase from Geobacillus sp. Y412MC61, enabling quick and easy insertion of DNA into the chromosome for accelerated genetic tool screening and heterologous gene expression. Using tSAGE, chromosomal insertion of plasmid DNA occurred at a maximum transformation efficiency of 5 × 103 CFU/µg, which is comparable to the transformation efficiency of a replicating control plasmid in C. thermocellum. Using tSAGE, we chromosomally integrated and characterized 17 reporter genes, 15 homologous and 31 heterologous constitutive promoters of varying strengths, 4 inducible promoters, and 5 riboswitches in C. thermocellum. We also determined that a 6-7 nucleotide gap between the ribosome binding site (RBS) and the start codon is optimal for high expression by employing a library of superfolder green fluorescent protein expression constructs driven by our strongest tested promoter (Pclo1313_1194) with different distances between the RBS and start codon. The tools developed here will aid in accelerating C. thermocellum strain engineering for producing sustainable fuels and chemicals directly from plant biomass. One-Sentence Summary: A highly efficient site-specific recombination system was created for Clostridium thermocellum, which enabled the rapid characterization of a large collection of genetic parts for controlled gene expression.

Detection of gene expression in plants is critical for understanding the molecular basis of complex plant biosystems and plant responses to environmental stresses. Here, we report the development of a split ribozyme-based biosensor that enables in vivo visualization of gene expression in plants. We demonstrated the utility of this biosensor in transient expression experiments (i.e., leaf infiltration in Nicotiana benthamiana) to detect RNAs derived from transgenes and tobacco rattle virus, respectively. Furthermore, we successfully engineered a split ribozyme-based biosensor in Arabidopsis thaliana for in vivo visualization of endogenous gene expression at the cellular level. In addition, we developed a platform for easy incorporation of different reporters into the RNA biosensor. ### Competing Interest Statement The authors have declared no competing interest.

CRISPR-Cas9 tools have transformed genetic manipulation capabilities in the laboratory. Empirical rules-of-thumb have been developed for only a narrow range of model organisms, and mechanistic underpinnings for sgRNA efficiency remain poorly understood. This work establishes a novel feature set and new public resource, produced with quantum chemical tensors, for interpreting and predicting sgRNA efficiency. Feature engineering for sgRNA efficiency is performed using an explainable-artificial intelligence model: iterative Random Forest (iRF). By encoding quantitative attributes of position-specific sequences for Escherichia coli sgRNAs, we identify important traits for sgRNA design in bacterial species. Additionally, we show that expanding positional encoding to quantum descriptors of base-pair, dimer, trimer, and tetramer sequences captures intricate interactions in local and neighboring nucleotides of the target DNA. These features highlight variation in CRISPR-Cas9 sgRNA dynamics between E. coli and H. sapiens genomes. These novel encodings of sgRNAs enhance our understanding of the elaborate quantum biological processes involved in CRISPR-Cas9 machinery.

CRISPR-Cas9 is a versatile genome editing system widely used since 2013 to introduce site-specific modifications into the genomes of model and non-model species. This technology is used in various applications, from gene knock-outs, knock-ins, and over-expressions to more precise changes, such as the introduction of nucleotides at a targeted locus. CRISPR-Cas9 has been demonstrated to be easy to establish in new species and highly efficient and specific compared to previous gene editing strategies such as Zinc finger nucleases and transcription activator-like effector nucleases. Grand challenges for emerging CRISPR-Cas9 tools in filamentous fungi are developing efficient transformation methods for non-model organisms. In this paper, we have leveraged the establishment of CRISPR-Cas9 genome editing tool that relies on Cas9/sgRNA ribonucleoprotein complexes (RNPs) in the model species Trichoderma reesei and developed the first protocol to efficiently transform the non-model species, Sphaerulina musiva. This fungal pathogen constitutes a real threat to the genus Populus, a foundational bioenergy crop used for biofuel production. Herein, we highlight the general considerations to design sgRNAs and their computational validation. We also describe the use of isolated protoplasts to deliver the CRISPR-Cas9 RNP components in both species and the screening for targeted genome editing events. The development of engineering tools in S. musiva can be used for studying genes involved in diverse processes such as secondary metabolism, establishment, and pathogenicity, among many others, but also for developing genetic mitigation approaches. The approach described here provides guidance for potential development of transformation systems in other non-model spore-bearing ascomycetes.

Conversion of CO2 to value-added products presents an opportunity to reduce GHG emissions while generating revenue. Formate, which can be generated by the electrochemical reduction of CO2, has been proposed as a promising intermediate compound for microbial upgrading. Here we present progress towards improving the soil bacterium Cupriavidus necator H16, which is capable of growing on formate as its sole source of carbon and energy using the Calvin–Benson–Bassham (CBB) cycle, as a host for formate utilization. Using adaptive laboratory evolution, we generated several isolates that exhibited faster growth rates on formate. The genomes of these isolates were sequenced, and resulting mutations were systematically reintroduced by metabolic engineering, to identify those that improved growth. The metabolic impact of several mutations was investigated further using RNA-seq transcriptomics. We found that deletion of a transcriptional regulator implicated in quorum sensing, PhcA, reduced expression of several operons and led to improved growth on formate. Growth was also improved by deleting large genomic regions present on the extrachromosomal megaplasmid pHG1, particularly two hydrogenase operons and the megaplasmid CBB operon, one of two copies present in the genome. Based on these findings, we generated a rationally engineered ΔphcA and megaplasmid-deficient strain that exhibited a 24% faster maximum growth rate on formate. Moreover, this strain achieved a 7% growth rate improvement on succinate and a 19% increase on fructose, demonstrating the broad utility of microbial genome reduction. This strain has the potential to serve as an improved microbial chassis for biological conversion of formate to value-added products.

Precise modification of plant genomes, such as seamless insertion, deletion, or replacement of DNA sequences at a predefined site, is a challenging task. Gene targeting (GT) and prime editing are currently the best approaches for this purpose. However, these techniques are inefficient in plants, which limits their applications for crop breeding programs. Recently, substantial developments have been made to improve the efficiency of these techniques in plants. Several strategies, such as RNA donor templating, chemically modified donor DNA template, and tandem-repeat homology-directed repair, are aimed at improving GT. Additionally, improved prime editing gRNA design, use of engineered reverse transcriptase enzymes, and splitting prime editing components have improved the efficacy of prime editing in plants. These emerging strategies and existing technologies are reviewed along with various perspectives on their future improvement and the development of robust precision genome editing technologies for plants.

Protein engineering has been used successfully in fields ranging from medicine to food science to biofuels. Applications of protein engineering include developing antiviral peptides or other protein therapeutics, antibody engineering, designing protein-based logic circuits, engineering enzymes to be more specific or to function under industrially relevant conditions such as at higher temperatures or high/low pH, modifying cell signaling or regulatory functions, and so on. Advances in recombinant DNA, "omics," and CRISPR-Cas (clustered regularly interspaced short palindromic repeats and its associated proteins) technologies, combined with high-throughput screening facilities, will lead to improved methods for protein engineering, enabling easy modification of more proteins/enzymes for new specific applications. New methods for rational design, directed evolution, and computer-aided protein design will further accelerate the speed of protein evolution and expand the scope for protein engineering. In this chapter we discuss general protein engineering strategies and advances in engineering proteins with desired functions, focusing on the "design" and "build" part of the design-build-test-learn cycle.