Search

Systems metabolic engineering faces the formidable task of rewiring microbial metabolism to cost-effectively generate high-value molecules from a variety of inexpensive feedstocks for many different applications. Because these cellular systems are still too complex to model accurately, vast collections of engineered organism variants must be systematically created and evaluated through an enormous trial-and-error process in order to identify a manufacturing-ready strain. The high-throughput screening of strains to optimize their scalable manufacturing potential requires execution of many carefully controlled, parallel, miniature fermentations, followed by high-precision analysis of the resulting complex mixtures. This review discusses strategies for the design of high-throughput, small-scale fermentation models to predict improved strain performance at large commercial scale. Established and promising approaches from industrial and academic groups are presented for both cell culture and analysis, with primary focus on microplate- and microfluidics-based screening systems.

The ability to engineer natural proteins is pivotal toafuture, 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.

Metabolic engineering offers the potential to renewably produce important classes of chemicals, particularly biofuels, at an industrial scale. DNA synthesis and editing techniques can generate large pathway libraries, yet identifying the best variants is slow and cumbersome. Traditionally, analytical methods like chromatography and mass spectrometry have been used to evaluate pathway variants, but such techniques cannot be performed with high throughput. Biosensors - genetically encoded components that actuate a cellular output in response to a change in metabolite concentration - are therefore a promising tool for rapid and high-throughput evaluation of candidate pathway variants. Applying biosensors can also dynamically tune pathways in response to metabolic changes, improving balance and productivity. Here, we describe the major classes of biosensors and briefly highlight recent progress in applying them to biofuel-related metabolic pathway engineering.

The clustered, regularly interspaced, short palindromic repeats (CRISPR)-associated protein Cas9 from Streptococcus pyogenes is an RNA-guided DNA endonuclease with widespread utility for genome modification. However, the structural constraints limiting the engineering of Cas9 have not been determined. Here we experimentally profile Cas9 using randomized insertional mutagenesis and delineate hotspots in the structure capable of tolerating insertions of a PDZ domain without disruption of the enzyme's binding and cleavage functions. Orthogonal domains or combinations of domains can be inserted into the identified sites with minimal functional consequence. To illustrate the utility of the identified sites, we construct an allosterically regulated Cas9 by insertion of the estrogen receptor-α ligand-binding domain. This protein showed robust, ligand-dependent activation in prokaryotic and eukaryotic cells, establishing a versatile one-component system for inducible and reversible Cas9 activation. Thus, domain insertion profiling facilitates the rapid generation of new Cas9 functionalities and provides useful data for future engineering of Cas9.

Single-fluorescent protein biosensors (SFPBs) are an important class of probes that enable the single-cell quantification of analytes in vivo. Despite advantages over other detection technologies, their use has been limited by the inherent challenges of their construction. Specifically, the rational design of green fluorescent protein (GFP) insertion into a ligand-binding domain, generating the requisite allosteric coupling, remains a rate-limiting step. Here, we describe an unbiased approach, termed domain-insertion profiling with DNA sequencing (DIP-seq), that combines the rapid creation of diverse libraries of potential SFPBs and high-throughput activity assays to identify functional biosensors. As a proof of concept, we construct an SFPB for the important regulatory sugar trehalose. DIP-seq analysis of a trehalose-binding-protein reveals allosteric hotspots for GFP insertion and results in high-dynamic range biosensors that function robustly in vivo. Taken together, DIP-seq simultaneously accelerates metabolite biosensor construction and provides a novel tool for interrogating protein allostery., In the construction of single fluorescent protein biosensors, selection of the insertion point of a fluorescent protein into a ligand-binding domain is a rate-limiting step. Here, the authors develop an unbiased, high-throughput approach, called domain insertion profiling with DNA sequencing (DIP-seq), to generate a novel trehalose biosensor.

CRISPR/Cas systems act to protect the cell from invading nucleic acids in many bacteria and archaea. The bacterial immune protein Cas9 is a component of one of these CRISPR/Cas systems and has recently been adapted as a tool for genome editing. Cas9 is easily targeted to bind and cleave a DNA sequence via a complimentary RNA; this straightforward programmability has gained Cas9 rapid acceptance in the field of genetic engineering. While this technology has developed quickly, a number of challenges regarding Cas9 specificity, efficiency, fusion protein function, and spatiotemporal control within the cell remain. In this work, we develop a platform for constructing novel proteins to address these open questions. We demonstrate methods to either screen or select active Cas9 mutants and use the screening technique to isolate functional Cas9 variants with a heterologous PDZ domain inserted directly into the protein. As a proof of concept, these methods lay the groundwork for the future construction of diverse Cas9 proteins. Straightforward and accessible techniques for genetic editing are helping to elucidate biology in new and exciting ways; a platform to engineer new functionalities into Cas9 will help forge the next generation of genome modifying tools.

Despite extensive studies on microbial and enzymatic lignocellulose degradation, relatively few Archaea are known to deconstruct crystalline cellulose. Here we describe a consortium of three hyperthermophilic archaea enriched from a continental geothermal source by growth at 90 °C on crystalline cellulose, representing the first instance of Archaea able to deconstruct lignocellulose optimally above 90 °C. Following metagenomic studies on the consortium, a 90 kDa, multidomain cellulase, annotated as a member of the TIM barrel glycosyl hydrolase superfamily, was characterized. The multidomain architecture of this protein is uncommon for hyperthermophilic endoglucanases, and two of the four domains of the enzyme have no characterized homologues. The recombinant enzyme has optimal activity at 109 °C, a half-life of 5 h at 100 °C, and resists denaturation in strong detergents, high-salt concentrations, and ionic liquids. Cellulases active above 100 °C may assist in biofuel production from lignocellulosic feedstocks by hydrolysing cellulose under conditions typically employed in biomass pretreatment.

Improving the catalytic activity of cellulases requires screening variants against solid substrates. Expressing cellulases in microbial hosts is time-consuming, can be cellulase specific, and often leads to inactive forms and/or low yields. These limitations have been obstacles for improving cellulases in a high-throughput manner. We have developed a cell-free expression system and used it to express 54 chimeric bacterial and archaeal endoglucanases (EGs), with and without cellulose binding modules (CBMs) at either the N- or C-terminus, in active enzyme yields of 100-350 µg/mL. The platform was employed to systematically study the role of CBMs in cellulose hydrolysis toward a variety of natural and pretreated solid substrates, including ionic-liquid pretreated Miscanthus and AFEX-pretreated corn stover. Adding a CBM generally increased activity against crystalline Avicel, whereas for pretreated substrates the effect of CBM addition depended on the source of cellulase. The cell-free expression platform can thus provide insights into cellulase structure-function relationships for any substrate, and constitutes a powerful discovery tool for evaluating or engineering cellulolytic enzymes for biofuels production.