CRISPR/Cas9 recombineering-mediated deep mutational scanning of essential genes in Escherichia coli.

Deep mutational scanning can provide significant insights into the function of essential genes in bacteria. Here, we developed a high-throughput method for mutating essential genes of Escherichia coli in their native genetic context. We used Cas9-mediated recombineering to introduce a library of mutations, created by error-prone PCR, within a gene fragment on the genome using a single gRNA pre-validated for high efficiency. Tracking mutation frequency through deep sequencing revealed biases in the position and the number of the introduced mutations. We overcame these biases by increasing the homology arm length and blocking mismatch repair to achieve a mutation efficiency of 85% for non-essential genes and 55% for essential genes. These experiments also improved our understanding of poorly characterized recombineering process using dsDNA donors with single nucleotide changes. Finally, we applied our technology to target rpoB, the beta subunit of RNA polymerase, to study resistance against rifampicin. In a single experiment, we validate multiple biochemical and clinical observations made in the previous decades and provide insights into resistance compensation with the study of double mutants.

Building a genome engineering toolbox in nonmodel prokaryotic microbes.

The realization of a sustainable bioeconomy requires our ability to understand and engineer complex design principles for the development of platform organisms capable of efficient conversion of cheap and sustainable feedstocks (e.g., sunlight, CO2 , and nonfood biomass) into biofuels and bioproducts at sufficient titers and costs. For model microbes, such as Escherichia coli, advances in DNA reading and writing technologies are driving the adoption of new paradigms for engineering biological systems. Unfortunately, microbes with properties of interest for the utilization of cheap and renewable feedstocks, such as photosynthesis, autotrophic growth, and cellulose degradation, have very few, if any, genetic tools for metabolic engineering. Therefore, it is important to develop "design rules" for building a genetic toolbox for novel microbes. Here, we present an overview of our current understanding of these rules for the genetic manipulation of prokaryotic microbes and the available genetic tools to expand our ability to genetically engineer nonmodel systems.

Engineering Saccharomyces cerevisiae fatty acid composition for increased tolerance to octanoic acid.

Biorenewable chemicals such as short and medium chain fatty acids enable functional or direct substitution of petroleum-derived building blocks, allowing reduction of anthropogenic greenhouse gases while meeting market needs of high-demand products like aliphatic alcohols and alpha olefins. However, producing these fatty acids in microorganisms can be challenging due to toxicity issues. Octanoic acid (C8) can disrupt the integrity of the cell membrane in yeast, and exogenous supplementation of oleic acid has been shown to help alleviate this. We recently engineered the Saccharomyces cerevisiae enzyme acetyl-CoA carboxylase by replacing serine residue 1157 with alanine to prevent deactivation by phosphorylation. Expression of Acc1S1157A in S. cerevisiae resulted in an increase in total fatty acid production, with the largest increase for oleic acid. In this study, we evaluated the effect of this modified lipid profile on C8 toxicity to the yeast. Expression of Acc1S1157A in S. cerevisiae BY4741 increased the percentage of oleic acid 3.1- and 1.6-fold in the absence and presence of octanoic acid challenge, respectively. Following exposure to 0.9 mM of C8 for 24 h, the engineered yeast had a 10-fold higher cell density relative to the baseline strain. Moreover, overexpressing Acc1S1157A allowed survival at C8 concentrations that were lethal for the baseline strain. This marked reduction of toxicity was shown to be due to higher membrane integrity as an 11-fold decrease in leakage of intracellular magnesium was observed. Due to the increase in oleic acid, this approach has the potential to reduce toxicity of other valuable bioproducts such as shorter chain aliphatic acids and alcohols and other membrane stressors. In an initial screen, increased resistance to n-butanol, 2-propanol, and hexanoic acid was demonstrated with cell densities 3.2-, 1.8-, and 29-fold higher than the baseline strain, respectively. Biotechnol. Bioeng. 2017;114: 1531-1538. © 2017 Wiley Periodicals, Inc.

Dynamic and single cell characterization of a CRISPR-interference toolset in Pseudomonas putida KT2440 for ß-ketoadipate production from p-coumarate.

Pseudomonas putida KT2440 is a well-studied bacterium for the conversion of lignin-derived aromatic compounds to bioproducts. The development of advanced genetic tools in P. putida has reduced the turnaround time for hypothesis testing and enabled the construction of strains capable of producing various products of interest. Here, we evaluate an inducible CRISPR-interference (CRISPRi) toolset on fluorescent, essential, and metabolic targets. Nuclease-deficient Cas9 (dCas9) expressed with the arabinose (8K)-inducible promoter was shown to be tightly regulated across various media conditions and when targeting essential genes. In addition to bulk growth data, single cell time lapse microscopy was conducted, which revealed intrinsic heterogeneity in knockdown rate within an isoclonal population. The dynamics of knockdown were studied across genomic targets in exponentially-growing cells, revealing a universal 1.75 ± 0.38 h quiescent phase after induction where 1.5 ± 0.35 doublings occur before a phenotypic response is observed. To demonstrate application of this CRISPRi toolset, ß-ketoadipate, a monomer for performance-advantaged nylon, was produced at a 4.39 ± 0.5 g/L and yield of 0.76 ± 0.10 mol/mol from p-coumarate, a hydroxycinnamic acid that can be derived from grasses. These cultivation metrics were achieved by using the higher strength IPTG (1K)-inducible promoter to knockdown the pcaIJ operon in the ßKA pathway during early exponential phase. This allowed the majority of the carbon to be shunted into the desired product while eliminating the need for a supplemental carbon and energy source to support growth and maintenance.

High-Throughput Functional Genomics for Energy Production.

Functional genomics remains a foundational field for establishing genotype-phenotype relationships that enable strain engineering. High-throughput (HTP) methods accelerate the Design-Build-Test-Learn cycle that currently drives synthetic biology towards a forward engineering future. Trackable mutagenesis techniques including transposon insertion sequencing and CRISPR-Cas-mediated genome editing allow for rapid fitness profiling of a collection, or library, of mutants to discover beneficial mutations. Due to the relative speed of these experiments compared to adaptive evolution experiments, iterative rounds of mutagenesis can be implemented for next-generation metabolic engineering efforts to design complex production and tolerance phenotypes. Additionally, the expansion of these mutagenesis techniques to novel bacteria are opening up industrial microbes that show promise for establishing a bio-based economy.