Automated characterization and analysis of expression compatibility between regulatory sequences and metabolic genes in Escherichia coli.

Utilizing standardized artificial regulatory sequences to fine-tuning the expression of multiple metabolic pathways/genes is a key strategy in the creation of efficient microbial cell factories. However, when regulatory sequence expression strengths are characterized using only a few reporter genes, they may not be applicable across diverse genes. This introduces great uncertainty into the precise regulation of multiple genes at multiple expression levels. To address this, our study adopted a fluorescent protein fusion strategy for a more accurate assessment of target protein expression levels. We combined 41 commonly-used metabolic genes with 15 regulatory sequences, yielding an expression dataset encompassing 520 unique combinations. This dataset highlighted substantial variation in protein expression level under identical regulatory sequences, with relative expression levels ranging from 2.8 to 176-fold. It also demonstrated that improving the strength of regulatory sequences does not necessarily lead to significant improvements in the expression levels of target proteins. Utilizing this dataset, we have developed various machine learning models and discovered that the integration of promoter regions, ribosome binding sites, and coding sequences significantly improves the accuracy of predicting protein expression levels, with a Spearman correlation coefficient of 0.72, where the promoter sequence exerts a predominant influence. Our study aims not only to provide a detailed guide for fine-tuning gene expression in the metabolic engineering of Escherichia coli but also to deepen our understanding of the compatibility issues between regulatory sequences and target genes.

Repurposing conformational changes in ANL superfamily enzymes to rapidly generate biosensors for organic and amino acids.

Biosensors are powerful tools for detecting, real-time imaging, and quantifying molecules, but rapidly constructing diverse genetically encoded biosensors remains challenging. Here, we report a method to rapidly convert enzymes into genetically encoded circularly permuted fluorescent protein-based indicators to detect organic acids (GECFINDER). ANL superfamily enzymes undergo hinge-mediated ligand-coupling domain movement during catalysis. We introduce a circularly permuted fluorescent protein into enzymes hinges, converting ligand-induced conformational changes into significant fluorescence signal changes. We obtain 11 GECFINDERs for detecting phenylalanine, glutamic acid and other acids. GECFINDER-Phe3 and GECFINDER-Glu can efficiently and accurately quantify target molecules in biological samples in vitro. This method simplifies amino acid quantification without requiring complex equipment, potentially serving as point-of-care testing tools for clinical applications in low-resource environments. We also develop a GECFINDER-enabled droplet-based microfluidic high-throughput screening method for obtaining high-yield industrial strains. Our method provides a foundation for using enzymes as untapped blueprint resources for biosensor design, creation, and application.

A Programmable CRISPR/Cas9 Toolkit Improves Lycopene Production in Bacillus subtilis.

Bacillus subtilis has been widely used and generally recognized as a safe host for the production of recombinant proteins, high-value chemicals, and pharmaceuticals. Thus, its metabolic engineering attracts significant attention. Nevertheless, the limited availability of selective markers makes this process difficult and time-consuming, especially in the case of multistep biosynthetic pathways. Here, we employ CRISPR/Cas9 technology to build an easy cloning toolkit that addresses commonly encountered obstacles in the metabolic engineering of B. subtilis, including the chromosomal integration locus, promoter, terminator, and guide RNA (gRNA) target. Six promoters were characterized, and the promoter strengths ranged from 0.9- to 23-fold that of the commonly used strong promoter P43. We characterized seven terminators in B. subtilis, and the termination efficiencies (TEs) of the seven terminators are all more than 90%. Six gRNA targets were designed upstream of the promoter and downstream of the terminator. Using a green fluorescent protein (GFP) reporter, we confirmed integration efficiency with the single-locus integration site is up to 100%. We demonstrated the applicability of this toolkit by optimizing the expression of a challenging but industrially important product, lycopene. By heterologous expression of the essential genes for lycopene synthesis on the B. subtilis genome, a total of 13 key genes involved in the lycopene biosynthetic pathway were manipulated. Moreover, our findings showed that the gene cluster ispG-idi-dxs-ispD could positively affect the production of lycopene, while the cluster dxr-ispE-ispF-ispH had a negative effect on lycopene production. Hence, our multilocus integration strategy can facilitate the pathway assembly for production of complex chemicals and pharmaceuticals in B. subtilis. IMPORTANCE We present a toolkit that allows for rapid cloning procedures and one-step subcloning to move from plasmid-based expression to stable chromosome integration and expression in a production strain in less than a week. The utility of the customized tool was demonstrated by integrating the MEP (2C-methyl-d-erythritol-4-phosphate) pathway, part of the pentose phosphate pathway (PPP), and the hetero-lycopene biosynthesis genes by stable expression in the genome. The tool could be useful to engineer B. subtilis strains through diverse recombination events and ultimately improve its potential and scope of industrial application as biological chassis.

AutoESD: a web tool for automatic editing sequence design for genetic manipulation of microorganisms.

Advances in genetic manipulation and genome engineering techniques have enabled on-demand targeted deletion, insertion, and substitution of DNA sequences. One important step in these techniques is the design of editing sequences (e.g. primers, homologous arms) to precisely target and manipulate DNA sequences of interest. Experimental biologists can employ multiple tools in a stepwise manner to assist editing sequence design (ESD), but this requires various software involving non-standardized data exchange and input/output formats. Moreover, necessary quality control steps might be overlooked by non-expert users. This approach is low-throughput and can be error-prone, which illustrates the need for an automated ESD system. In this paper, we introduce AutoESD (https://autoesd.biodesign.ac.cn/), which designs editing sequences for all steps of genetic manipulation of many common homologous-recombination techniques based on screening-markers. Notably, multiple types of manipulations for different targets (CDS or intergenic region) can be processed in one submission. Moreover, AutoESD has an entirely cloud-based serverless architecture, offering high reliability, robustness and scalability which is capable of parallelly processing hundreds of design tasks each having thousands of targets in minutes. To our knowledge, AutoESD is the first cloud platform enabling precise, automated, and high-throughput ESD across species, at any genomic locus for all manipulation types.

[Advances in automated high-throughput editing and screening of engineered strains].

One of the revolutionary features of synthetic biology is that the standardization and modularization of biological experimental objects, methods, technologies and processes can be combined with various software and hardware to forge into an automated high-throughput synthetic biology biofoundry. Disrupting the conventional labor-intensive research paradigm, biofoundry represents a novel research paradigm with highly enhanced technical iteration capabilities, and remarkably promotes the development and industrial applications of synthetic biology. On the occasion of the 10th anniversary of the founding of Tianjin Institute of Industrial Biotechnology, Chinese Academy of Sciences, this review summarized a series of important achievements in the field of automated high-throughput editing and screening of industrial strains. These achievements range from automated editing technologies such as gene cloning, genome editing, and editing sequence design, to high-throughput screening technologies such as fluorescence activated cell sorting, fluorescence activated droplet sorting, and genome-scale gene perturb sequencing. Moreover, we prospected future development of this field, hoping to provide overall support for intelligent, automated and full chain integrated creation of excellent industrial strains with intellectual property rights.

A CRISPR/dCas9-assisted system to clone toxic genes in Escherichia coli.

BACKGROUND: The cloning of toxic genes in E. coli requires strict regulation of the target genes' leaky expression. Many methods facilitating successful gene cloning of toxic genes are commonly exploited, but the applicability is severely limited. METHODS: A CRISPR/dCas9-assisted system was used to clone toxic genes in E. coli. The plasmid-based and genome-integrated systems were designed in this study. And the green fluorescent protein characterization system was used to test the repression efficiency of the two systems. RESULTS: We optimized the plasmid-based CRISPR/dCas9-assisted repression system via testing different sgRNAs targeting the Ptrc promoter and achieved inhibition efficiency up to 64.8%. The genome-integrated system represented 35.9% decreased GFP expression and was successfully employed to cloned four toxic genes from Corynebacterium glutamicum in E. coli. CONCLUSIONS: Using this method, we successfully cloned four C. glutamicum-derived toxic genes that had been failed to clone in conventional ways. The CRISPR/dCas9-assisted gene cloning method was a promising tool to facilitate precise gene cloning of different origins in E. coli. GENERAL SIGNIFICANCE: This system will be useful for cloning toxic genes from different origins in E. coli, and can accelerate the related research of gene characterization and heterologous expression in the metagenomic era.

In-situ generation of large numbers of genetic combinations for metabolic reprogramming via CRISPR-guided base editing.

Reprogramming complex cellular metabolism requires simultaneous regulation of multigene expression. Ex-situ cloning-based methods are commonly used, but the target gene number and combinatorial library size are severely limited by cloning and transformation efficiencies. In-situ methods such as multiplex automated genome engineering (MAGE) depends on high-efficiency transformation and incorporation of heterologous DNA donors, which are limited to few microorganisms. Here, we describe a Base Editor-Targeted and Template-free Expression Regulation (BETTER) method for simultaneously diversifying multigene expression. BETTER repurposes CRISPR-guided base editors and in-situ generates large numbers of genetic combinations of diverse ribosome binding sites, 5' untranslated regions, or promoters, without library construction, transformation, and incorporation of DNA donors. We apply BETTER to simultaneously regulate expression of up to ten genes in industrial and model microorganisms Corynebacterium glutamicum and Bacillus subtilis. Variants with improved xylose catabolism, glycerol catabolism, or lycopene biosynthesis are respectively obtained. This technology will be useful for large-scale fine-tuning of multigene expression in both genetically tractable and intractable microorganisms.

Expanding targeting scope, editing window, and base transition capability of base editing in Corynebacterium glutamicum.

CRISPR/Cas9-guided cytidine deaminase enables C:G to T:A base editing in bacterial genome without introduction of lethal double-stranded DNA break, supplement of foreign DNA template, or dependence on inefficient homologous recombination. However, limited by genome-targeting scope, editing window, and base transition capability, the application of base editing in metabolic engineering has not been explored. Herein, four Cas9 variants accepting different protospacer adjacent motif (PAM) sequences were used to increase the genome-targeting scope of bacterial base editing. After a comprehensive evaluation, we demonstrated that PAM requirement of bacterial base editing can be relaxed from NGG to NG using the Cas9 variants, providing 3.9-fold more target loci for gene inactivation in Corynebacterium glutamicum. Truncated or extended guide RNAs were employed to expand the canonical 5-bp editing window to 7-bp. Bacterial adenine base editing was also achieved with Cas9 fused to adenosine deaminase. With these updates, base editing can serve as an enabling tool for fast metabolic engineering. To demonstrate its potential, base editing was used to deregulate feedback inhibition of aspartokinase via amino acid substitution for lysine overproduction. Finally, a user-friendly online tool named gBIG was provided for designing guide RNAs for base editing-mediated inactivation of given genes in any given sequenced genome (www.ibiodesign.net/gBIG).

Co-expression of L-glutamate oxidase and catalase in Escherichia coli to produce α-ketoglutaric acid by whole-cell biocatalyst.

OBJECTIVES: To improve the production of α-ketoglutaric acid (α-KG) from L-glutamate by whole-cell biocatalysis. RESULTS: A novel and highly active L-glutamate oxidase, SmlGOX, from Streptomyces mobaraensis was overexpressed and purified. The recombinant SmlGOX was approx. 64 kDa by SDS-PAGE. SmlGOX had a maximal activity of 125 ± 2.7 U mg-1 at pH 6.0, 35 oC. The apparent Km and Vmax values of SmlGOX were 9.3 ± 0.5 mM and 159 ± 3 U mg-1, respectively. Subsequently, a co-expression plasmid containing the SmlGOX and KatE genes was constructed to remove H2O2, and the protein levels of SmlGOX were improved by codon optimization. Finally, by optimizing the whole-cell transformation conditions, the production of α-KG reached 77.4 g l-1 with a conversion rate from L-glutamate of 98.5% after 12 h. CONCLUSIONS: An efficient method for the production of α-KG was established in the recombinant Escherichia coli, and it has a potential prospect in industrial application.

Expression, characterization and mutagenesis of a novel glutamate decarboxylase from Bacillus megaterium.

OBJECTIVES: To search for a novel glutamate decarboxylase (GAD) with an optimum pH towards near-neutrality in order to improve production of gamma-aminobutyric acid (GABA) in recombinant hosts. RESULTS: A novel glutamate decarboxylase, BmGAD, from Bacillus megaterium was overexpressed and purified. BmGAD was approximately 53 kDa by SDS-PAGE analysis. Its optimum activity was at pH 5 and 50 °C. BmGAD had a specific activity of 59 ± 5.2 U mg(-1) at pH 6, which is the highest value reported so far. The apparent Km and Vmax values of BmGAD were 8 ± 0.5 mM and 150 ± 4.7 U mg(-1), respectively. Through site-directed mutagenesis, two BmGAD mutants (E294R and H467A) showed higher Vmax values than that of wild-type, with the values of 210 ± 6.9 and 180 ± 4.1 U mg(-1) at pH 5 and 50 °C, respectively. CONCLUSIONS: The unusual high activity of BmGAD at pH 6 makes it an attractive GABA-producing candidate in industrial application.

In vitro functional characterization of the Na+/H+ antiporters in Corynebacterium glutamicum.

Corynebacterium glutamicum, typically used as industrial workhorse for amino acid production, is a moderately salt-alkali-tolerant microorganism with optimal growth at pH 7-9. However, little is known about the mechanisms of salt-alkali tolerance in C. glutamicum. Here, the catalytic capacity of three putative Na(+)/H(+) antiporters from C. glutamicum (designated as Cg-Mrp1, Cg-Mrp2 and Cg-NhaP) were characterized in an antiporter-deficient Escherichia coli KNabc strain. Only Cg-Mrp1 was able to effectively complement the Na(+)-sensitive of E. coli KNabc. Cg-Mrp1 exhibited obvious Na(+)(Li(+))/H(+) antiport activities with low apparent Km values of 1.08 mM and 1.41 mM for Na(+) and Li(+), respectively. The Na(+)/H(+) antiport activity of Cg-Mrp1 was optimal in the alkaline pH range. All three antiporters showed detectable K(+)/H(+) antiport activitiy. Cg-NhaP also exhibited Na(+)(Li(+),Rb(+))/H(+) antiport activities but at lower levels of activity. Interestingly, overexpression of Cg-Mrp2 exhibited clear Na(+)(K(+))/H(+) antiport activities. These results suggest that C. glutamicum Na(+)(K(+))/H(+) antiporters may have overlapping roles in coping with salt-alkali and perhaps high-osmolarity stress.