CRISPR interference-guided balancing of a biosynthetic mevalonate pathway increases terpenoid production

New strategies to study in depth the metabolic mechanism of astaxanthin biosynthesis in Phaffia rhodozyma

Astaxanthin, a ketone carotenoid known for its high antioxidant activity, holds significant potential for application in nutraceuticals, aquaculture, and cosmetics. The increasing market demand necessitates a higher production of astaxanthin using Phaffia rhodozyma. Despite extensive research efforts focused on optimizing fermentation conditions, employing mutagenesis treatments, and utilizing genetic engineering technologies to enhance astaxanthin yield in P. rhodozyma, progress in this area remains limited. This review provides a comprehensive summary of the current understanding of rough metabolic pathways, regulatory mechanisms, and preliminary strategies for enhancing astaxanthin yield. However, further investigation is required to fully comprehend the intricate and essential metabolic regulation mechanism underlying astaxanthin synthesis. Specifically, the specific functions of key genes, such as crtYB, crtS, and crtI, need to be explored in detail. Additionally, a thorough understanding of the action mechanism of bifunctional enzymes and alternative splicing products is imperative. Lastly, the regulation of metabolic flux must be thoroughly investigated to reveal the complete pathway of astaxanthin synthesis. To obtain an in-depth mechanism and improve the yield of astaxanthin, this review proposes some frontier methods, including: omics, genome editing, protein structure-activity analysis, and synthetic biology. Moreover, it further elucidates the feasibility of new strategies using these advanced methods in various effectively combined ways to resolve these problems mentioned above. This review provides theory and method for studying the metabolic pathway of astaxanthin in P. rhodozyma and the industrial improvement of astaxanthin, and provides new insights into the flexible combined use of multiple modern advanced biotechnologies.

Metabolic engineering for enhanced terpenoid production: Leveraging new horizons with an old technique

Terpenoids are a vast class of plant specialized metabolites (PSMs) manufactured by plants and are involved in their interactions with environment. In addition, they add health benefits to human nutrition and are widely used as pharmaceutically active compounds. However, native plants produce a limited amount of terpenes restricting metabolite yield of terpene-related metabolites. Exponential growth in the plant metabolome data and the requirement of alternative approaches for producing the desired amount of terpenoids, has redirected plant biotechnology research to plant metabolic engineering, which requires in-depth knowledge and precise expertise about dynamic plant metabolic pathways and cellular physiology. Metabolic engineering is an assuring tool for enhancing the concentration of terpenes by adopting specific strategies such as overexpression of the key genes associated with the biosynthesis of targeted metabolites, controlling the modulation of transcription factors, downregulation of competitive pathways (RNAi), co-expression of the biosynthetic pathway genes in heterologous system and other combinatorial approaches. Microorganisms, fast-growing host plants (such as Nicotiana benthamiana), and cell suspension/callus cultures have provided better means for producing valuable terpenoids. Manipulation in the biosynthetic pathways responsible for synthesis of terpenoids can provide opportunities to enhance the content of desired terpenoids and open up new avenues to enhance their production. This review deliberates the worth of metabolic engineering in medicinal plants to resolve issues associated with terpenoid production at a commercial scale. However, to bring the revolution through metabolic engineering, further implementation of genome editing, elucidation of metabolic pathways using omics approaches, system biology approaches, and synthetic biology tactics are essentially needed.

The non-mevalonate pathway requires a delicate balance of intermediates to maximize terpene production

Terpenes are valuable industrial chemicals whose demands are increasingly being met by bioengineering microbes such as E. coli. Although the bioengineering efforts commonly involve installing the mevalonate (MVA) pathway in E. coli for terpene production, the less studied methylerythritol phosphate (MEP) pathway is a more attractive target due to its higher energy efficiency and theoretical yield, despite its tight regulation. In this study, we integrated an additional copy of the entire MEP pathway into the E. coli genome for stable, marker-free terpene production. The genomically integrated strain produced more monoterpene geraniol than a plasmid-based system. The pathway genes' transcription was modulated using different promoters to produce geraniol as the reporter of the pathway flux. Pathway genes, including dxs, idi, and ispDF, expressed from a medium-strength promoter, led to the highest geraniol production. Quantifying the MEP pathway intermediates revealed that the highest geraniol producers had high levels of isopentenyl pyrophosphate (IPP) and dimethylallyl pyrophosphate (DMAPP), but moderate levels of the pathway intermediates upstream of these two building blocks. A principal component analysis demonstrated that 1-deoxy-D-xylulose 5-phosphate (DXP), the product of the first enzyme of the pathway, was critical for determining the geraniol titer, whereas MEP, the product of DXP reductoisomerase (Dxr or IspC), was the least essential. This work shows that an intricate balance of the MEP pathway intermediates determines the terpene yield in engineered E. coli. The genetically stable and intermediate-balanced strains created in this study will serve as a chassis for producing various terpenes. KEY POINTS: • Genome-integrated MEP pathway afforded higher strain stability • Genome-integrated MEP pathway produced more terpene than the plasmid-based system • High monoterpene production requires a fine balance of MEP pathway intermediates.

The HpSGNi system: A compact approach for genetic suppression without sequence limitation in Escherichia coli

Targeted gene regulation is indispensable for exploring gene functions in microbes and the development of microbial cell factories. While most loci can be regulated by CRISPRi, it cannot be used for targets lacking protospacer adjacent motifs (PAM) or protospacer flanking sequences (PFS). Here, we characterized a genetic suppression approach named the hpDNA-assisted structure-guided nuclease mediating interference system (HpSGNi). It was composed of a flap endonuclease 1 (FEN1) and mis-hairpin DNA probes (mis-hpDNA) to suppress the expression of target genes simply and efficiently in microbe without sequence restrictions. By inhibiting the initiation and elongation of the transcription, HpSGNi successfully silenced the transcription of exogenous fluorescent protein genes, ampicillin resistance gene and endogenous folP/sulA genes in Escherichia coli BL21(DE3) and K-12 MG1655. Meanwhile, aiming at optimizing the mis-hpDNA, we displayed the characteristics by detecting the tolerance to the single base mismatch and length of the guide sequence. This DNA-guided recognition platform provides a simple approach for selectively inhibiting gene expression.

Taming CRISPRi: Dynamic range tuning through guide RNA diversion

CRISPRi is a powerful technique to repress gene expression in a targeted and highly efficient manner. However, this potency is a double-edged sword in inducible systems, as even leaky expression of guide RNA results in a repression phenotype, complicating applications such as dynamic metabolic engineering. We evaluated three methods to enhance the controllability of CRISPRi by modulating the level of free and DNA-bound guide RNA complexes. Overall repression can be attenuated through rationally designed mismatches in the reversibility determining region of the guide RNA sequence; decoy target sites can selectively modulate repression at low levels of induction; and the implementation of feedback control not only enhances the linearity of induction, but broadens the dynamic range of the output as well. Furthermore, feedback control significantly enhances the recovery rate after induction is removed. Used in combination, these techniques enable the fine-tuning of CRISPRi to meet restrictions imposed by the target and match the input signal required for induction.

Enhancing isoprenol production by systematically tuning metabolic pathways using CRISPR interference in E. coli

Regulation of metabolic gene expression is crucial for maximizing bioproduction titers. Recent engineering tools including CRISPR/Cas9, CRISPR interference (CRISPRi), and CRISPR activation (CRISPRa) have enabled effective knock-out, knock-down, and overexpression of endogenous pathway genes, respectively, for advanced strain engineering. CRISPRi in particular has emerged as a powerful tool for gene repression through the use of a deactivated Cas9 (dCas9) protein and target guide RNA (gRNA). By constructing gRNA arrays, CRISPRi has the capacity for multiplexed gene downregulation across multiple orthogonal pathways for enhanced bioproduction titers. In this study, we harnessed CRISPRi to downregulate 32 essential and non-essential genes in E. coli strains heterologously expressing either the original mevalonate pathway or isopentenyl diphosphate (IPP) bypass pathway for isoprenol biosynthesis. Isoprenol remains a candidate bioproduct both as a drop-in blend additive and as a precursor for the high-performance sustainable aviation fuel, 1,4-dimethylcyclooctane (DMCO). Of the 32 gRNAs targeting genes associated with isoprenol biosynthesis, a subset was found to vastly improve product titers. Construction of a multiplexed gRNA library based on single guide RNA (sgRNA) performance enabled simultaneous gene repression, yielding a 3 to 4.5-fold increase in isoprenol titer (1.82 ± 0.19 g/L) on M9-MOPS minimal medium. We then scaled the best performing CRISPRi strain to 2-L fed-batch cultivation and demonstrated translatable titer improvements, ultimately obtaining 12.4 ± 1.3 g/L isoprenol. Our strategy further establishes CRISPRi as a powerful tool for tuning metabolic flux in production hosts and that titer improvements are readily scalable with potential for applications in industrial bioproduction.

Efficient Myrcene Production Using Linalool Dehydratase Isomerase and Rational Biochemical Process in Escherichia coli

Microbial synthesis of plant-based myrcene is of great interest because of its high demand, however, achieving high biosynthetic titers remains a great challenge. Previous strategies adopted for microbial myrcene production have relied on the recruitment of a multi-step biosynthetic pathway which requires complex metabolic regulation or high activity of myrcene synthase, hindering its application. Here, we present an effective one-step biotransformation system for myrcene biosynthesis from geraniol, using a linalool dehydratase isomerase (LDI) to overcome these limitations. The truncated LDI possesses nominal activity that catalyzes the isomerization of geraniol to linalool and the subsequent dehydration to myrcene in anaerobic environment. In order to improve the robustness of engineered strains for the efficient conversion of geraniol to myrcene, rational enzyme modification and a series of biochemical process engineering were employed to maintain and improve the anaerobic catalytic activity of LDI. Finally, by introducing the optimized myrcene biosynthetic capability in the existing geraniol-production strain, we achieve de novo biosynthesis of myrcene at 1.25g/L from glycerol during 84h aerobic-anaerobic two-stage fermentation, which is much higher than previously reported myrcene levels. This work highlights the value of dehydratase isomerase-based biocatalytic in establishing novel biosynthetic pathways and lays a reliable foundation for the microbial synthesis of myrcene.

Overproduction of Patchoulol in Metabolically Engineered Komagataella phaffii

Patchoulol, a plant-derived sesquiterpene compound, is widely used in perfumes, cosmetics, and pharmaceuticals. Microbial production provides a promising alternative approach for the efficient and sustainable production of patchoulol. However, there are no systematic engineering studies on Komagataella phaffii aimed at achieving high-yield patchoulol production. Herein, by fusion overexpression of FPP synthase and patchoulol synthase (ERG20LPTS), increasing the precursor supply, adjusting the copy number of ERG20LPTS and PTS, and combined with adding auxiliary carbon source and methanol concentration optimization, we constructed a high-yield patchoulol-producing strain P6H53, which produced 149.64 mg/L patchoulol in shake-flask fermentation with methanol as the substrate. In fed-batch fermentation, strain P6H53 achieved the highest production (2.47 g/L, 21.48 mg/g DCW, and 283.25 mg/L/d) to date in a 5 L fermenter. This study will lay a good foundation for the development of K. phaffii as a promising chassis microbial cell for the synthesis of patchoulol and other sesquiterpenes with methanol as the carbon source.

Recent Advances in CRISPR-Cas Technologies for Synthetic Biology

With developments in synthetic biology, "engineering biology" has emerged through standardization and platformization based on hierarchical, orthogonal, and modularized biological systems. Genome engineering is necessary to manufacture and design synthetic cells with desired functions by using bioparts obtained from sequence databases. Among various tools, the CRISPR-Cas system is modularly composed of guide RNA and Cas nuclease; therefore, it is convenient for editing the genome freely. Recently, various strategies have been developed to accurately edit the genome at a single nucleotide level. Furthermore, CRISPR-Cas technology has been extended to molecular diagnostics for nucleic acids and detection of pathogens, including disease-causing viruses. Moreover, CRISPR technology, which can precisely control the expression of specific genes in cells, is evolving to find the target of metabolic biotechnology. In this review, we summarize the status of various CRISPR technologies that can be applied to synthetic biology and discuss the development of synthetic biology combined with CRISPR technology in microbiology.

CRISPRi-based programmable logic inverter cascade for antibiotic-free selection and maintenance of multiple plasmids

Antibiotics have been widely used for plasmid-mediated cell engineering. However, continued use of antibiotics increases the metabolic burden, horizontal gene transfer risks, and biomanufacturing costs. There are limited approaches to maintaining multiple plasmids without antibiotics. Herein, we developed an inverter cascade using CRISPRi by building a plasmid containing a single guide RNA (sgRNA) landing pad (pSLiP); this inhibited host cell growth by repressing an essential cellular gene. Anti-sgRNAs on separate plasmids restored cell growth by blocking the expression of growth-inhibitory sgRNAs in pSLiP. We maintained three plasmids in Escherichia coli with a single antibiotic selective marker. To completely avoid antibiotic use and maintain the CRISPRi-based logic inverter cascade, we created a novel d-glutamate auxotrophic E. coli. This enabled the stable maintenance of the plasmid without antibiotics, enhanced the production of the terpenoid, (-)-α-bisabolol, and generation of an antibiotic-resistance gene-free plasmid. CRISPRi is therefore widely applicable in genetic circuits and may allow for antibiotic-free biomanufacturing.

Dynamic and single cell characterization of a CRISPR-interference toolset in Pseudomonas putida KT2440 for β-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, β-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 βKA 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.

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Developed an inducible dCas9-based CRISPR interference toolset in Pseudomonas putida KT2440.

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Characterized single-cell dynamics of fluorescent and essential gene knockdown.

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Applied the toolset for glucose-free production of β-ketoadipate from p-coumarate.

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Produced β-ketoadipate at titer of 4.39 ± 0.5 g/L and 0.76 ± 0.10 mol/mol yield.

Two strategies to improve the supply of PKS extender units for ansamitocin P-3 biosynthesis by CRISPR-Cas9

Ansamitocin P-3 (AP-3) produced by Actinosynnema pretiosum is a potent antitumor agent. However, lack of efficient genome editing tools greatly hinders the AP-3 overproduction in A. pretiosum. To solve this problem, a tailor-made pCRISPR-Cas9apre system was developed from pCRISPR-Cas9 for increasing the accessibility of A. pretiosum to genetic engineering, by optimizing cas9 for the host codon preference and replacing pSG5 with pIJ101 replicon. Using pCRISPR-Cas9apre, five large-size gene clusters for putative competition pathway were individually deleted with homology-directed repair (HDR) and their effects on AP-3 yield were investigated. Especially, inactivation of T1PKS-15 increased AP-3 production by 27%, which was most likely due to the improved intracellular triacylglycerol (TAG) pool for essential precursor supply of AP-3 biosynthesis. To enhance a "glycolate" extender unit, two combined bidirectional promoters (BDPs) ermEp-kasOp and j23119p-kasOp were knocked into asm12-asm13 spacer in the center region of gene cluster, respectively, by pCRISPR-Cas9apre. It is shown that in the two engineered strains BDP-ek and BDP-jk, the gene transcription levels of asm13-17 were significantly upregulated to improve the methoxymalonyl-acyl carrier protein (MM-ACP) biosynthetic pathway and part of the post-PKS pathway. The AP-3 yields of BDP-ek and BDP-jk were finally increased by 30% and 50% compared to the parent strain L40. Both CRISPR-Cas9-mediated engineering strategies employed in this study contributed to the availability of AP-3 PKS extender units and paved the way for further metabolic engineering of ansamitocin overproduction.

Engineering cell morphology by CRISPR interference in Acinetobacter baylyi ADP1

Microbial production of intracellular compounds can be engineered by redirecting the carbon flux towards products and increasing the cell size. Potential engineering strategies include exploiting clustered regularly interspaced short palindromic repeats interference (CRISPRi)-based tools for controlling gene expression. Here, we applied CRISPRi for engineering Acinetobacter baylyi ADP1, a model bacterium for synthesizing intracellular storage lipids, namely wax esters. We first established an inducible CRISPRi system for strain ADP1, which enables tightly controlled repression of target genes. We then targeted the glyoxylate shunt to redirect carbon flow towards wax esters. Second, we successfully employed CRISPRi for modifying cell morphology by repressing ftsZ, an essential gene required for cell division, in combination with targeted knock-outs to generate significantly enlarged filamentous or spherical cells respectively. The engineered cells sustained increased wax ester production metrics, demonstrating the potential of cell morphology engineering in the production of intracellular lipids.

CRISPR/dCas9-based metabolic pathway engineering for the systematic optimization of exopolysaccharide biosynthesis in Streptococcus thermophilus

Streptococcus thermophilus is used extensively in the dairy industry and has shown great promise as a chassis cell for the biosynthesis of high-value metabolites. However, metabolic engineering in S. thermophilus lacks effective genetic modification tools to modulate gene expression to relieve metabolic burden and maximize the production of desired compounds. Here, we developed a clustered regularly interspaced short palindromic repeats interference (CRISPRi) system for efficient gene transcriptional modulation in S. thermophilus. Our CRISPRi system typically achieved 66 to 98% knockdown of single or multiple gene expression. We used CRISPRi for the biosynthesis of a new exopolysaccharide (EPS) as a paradigm model. Repression of galK at module of uridine diphosphate glucose sugar metabolism and overexpression of epsA and epsE at EPS synthesis module resulted in an approximately 2-fold increase in EPS titer (277 mg/L) when compared with a control strain. This study demonstrated the effectiveness of CRISPRi as a powerful metabolic engineering tool and synthetic biology strategy for S. thermophilus.

Activating natural product synthesis using CRISPR interference and activation systems in Streptomyces

The rise of antibiotic-resistant bacteria represents a major threat to global health, creating an urgent need to discover new antibiotics. Natural products derived from the genus Streptomyces represent a rich and diverse repertoire of chemical molecules from which new antibiotics are likely to be found. However, a major challenge is that the biosynthetic gene clusters (BGCs) responsible for natural product synthesis are often poorly expressed under laboratory culturing conditions, thus preventing the isolation and screening of novel chemicals. To address this, we describe a novel approach to activate silent BGCs through rewiring endogenous regulation using synthetic gene regulators based upon CRISPR-Cas. First, we refine CRISPR interference (CRISPRi) and create CRISPR activation (CRISPRa) systems that allow for highly programmable and effective gene repression and activation in Streptomyces. We then harness these tools to activate a silent BGC by perturbing its endogenous regulatory network. Together, this work advances the synthetic regulatory toolbox for Streptomyces and facilitates the programmable activation of silent BGCs for novel chemical discovery.

Development of a gRNA Expression and Processing Platform for Efficient CRISPR-Cas9-Based Gene Editing and Gene Silencing in Candida tropicalis

Candida tropicalis, a nonmodel diploid microbe, has been applied in industry as a chassis cell. Metabolic engineering of C. tropicalis is challenging due to a lack of gene editing and regulation tools. Here, we report a tRNA:guide RNA (gRNA) platform for boosting gene editing and silencing efficiency in C. tropicalis. As the endogenous tRNA-processing system enables autocleavage for producing a large number of mature gRNAs, a tRNAGly sequence from the genome of C. tropicalis ATCC 20336 was selected for constructing the tRNA:gRNA platform. In the CRISPR-Cas9 system, the tRNA:gRNA platform proved to be efficient in single-gene and multi-gene editing. Furthermore, based on the tRNA:gRNA platform, a CRISPR interference (CRISPRi) system was developed to construct an efficient dCas9-mediated gene expression regulation system for C. tropicalis. The CRISPRi system was employed to regulate the expression of the exogenous gene GFP3 (green fluorescent protein) and the endogenous gene ADE2 (phosphoribosylaminoimidazole carboxylase). Different regions of GFP3 and ADE2 were targeted with the gRNAs processed by the tRNAGly, and the transcription levels of GFP3 and ADE2 were successfully downregulated to 23.9% ± 4.1% and 38.0% ± 7.4%, respectively. The effects of the target regions on gene regulation were also investigated. Additionally, the regulation system was applied to silence ERG9 (squalene synthase) to enhance β-carotene biosynthesis in a metabolically modified C. tropicalis strain. The results suggest that the endogenous tRNAGly and the CRISPRi system have great potential for metabolic engineering of C. tropicalis. IMPORTANCE In the nonmodel yeast Candida tropicalis, a lack of available RNA polymerase type III (Pol III) promoters hindered the development of guide RNA (gRNA) expression platforms for the establishment of CRISPR-Cas-mediated genome editing and silencing strategies. Here, a tRNA:gRNA platform was constructed. We show that this platform allows efficient and precise expression and processing of different gRNAs from a single polycistronic gene capable of mediating multi-gene editing in combination with CRISPR-Cas9. Furthermore, in combination with dCas9, the tRNA:gRNA platform was efficiently used for silencing of exogenous and endogenous genes, representing the first CRISPR interference tool (CRISPRi) in C. tropicalis. Importantly, the established CRISPRi-tRNA:gRNA tool was also used for metabolic engineering by regulating β-carotene biosynthesis in C. tropicalis. The results suggest that the tRNA:gRNA platform and the CRISPRi system will further advance the application of the CRISPR-Cas-based editing and CRISPRi systems for metabolic engineering in C. tropicalis.

Transcriptional Tuning of Mevalonate Pathway Enzymes to Identify the Impact on Limonene Production in Escherichia coli

Heterologous production of limonene in microorganisms through the mevalonate (MVA) pathway has traditionally imposed metabolic burden and reduced cell fitness, where imbalanced stoichiometries among sequential enzymes result in the accumulation of toxic intermediates. Although prior studies have shown that changes to mRNA stability, RBS strength, and protein homology can be effective strategies for balancing enzyme levels in the MVA pathway, testing different variations of these parameters often requires distinct genetic constructs, which can exponentially increase assembly costs as pathways increase in size. Here, we developed a multi-input transcriptional circuit to regulate the MVA pathway, where four chemical inducers, l-arabinose (Ara), choline chloride (Cho), cuminic acid (Cuma), and isopropyl β-d-1-thiogalactopyranoside (IPTG), each regulate one of four orthogonal promoters. We tested modular transcriptional regulation of the MVA pathway by placing this circuit in an engineered Escherichia coli "marionette" strain, which enabled systematic and independent tuning of the first three enzymes (AtoB, HMGS, and HMGR) in the MVA pathway. By systematically testing combinations of chemical inducers as inputs, we investigated relationships between the expressions of different MVA pathway submodules, finding that limonene yields are sensitive to the coordinated transcriptional regulation of HMGS and HMGR.

CRISPR-Cas system in microbial hosts for terpenoid production

Terpenoids represent the largest group of secondary metabolites with variable structures and functions. Terpenoids are well known for their beneficial application in human life, such as pharmaceutical products, vitamins, hormones, anticancer drugs, cosmetics, flavors and fragrances, foods, agriculture, and biofuels. Recently, engineering microbial cells have been provided with a sustainable approach to produce terpenoids with high yields. Noticeably, the clustered regularly interspaced short palindromic repeats (CRISPR)-associated (Cas) system has emerged as one of the most efficient genome-editing technologies to engineer microorganisms for improving terpenoid production. In this review, we summarize the application of the CRISPR-Cas system for the production of terpenoids in microbial hosts such as Escherichia coli, Saccharomyces cerevisiae, Corynebacterium glutamicum, and Pseudomonas putida. CRISPR-Cas9 deactivated Cas9 (dCas9)-based CRISPR (CRISPRi), and the dCas9-based activator (CRISPRa) have been used in either individual or combinatorial systems to control the metabolic flux for enhancing the production of terpenoids. Finally, the prospects of using the CRISPR-Cas system in terpenoid production are also discussed.

Toward improved terpenoids biosynthesis: strategies to enhance the capabilities of cell factories

Terpenoids form the most diversified class of natural products, which have gained application in the pharmaceutical, food, transportation, and fine and bulk chemical industries. Extraction from naturally occurring sources does not meet industrial demands, whereas chemical synthesis is often associated with poor enantio-selectivity, harsh working conditions, and environmental pollutions. Microbial cell factories come as a suitable replacement. However, designing efficient microbial platforms for isoprenoid synthesis is often a challenging task. This has to do with the cytotoxic effects of pathway intermediates and some end products, instability of expressed pathways, as well as high enzyme promiscuity. Also, the low enzymatic activity of some terpene synthases and prenyltransferases, and the lack of an efficient throughput system to screen improved high-performing strains are bottlenecks in strain development. Metabolic engineering and synthetic biology seek to overcome these issues through the provision of effective synthetic tools. This review sought to provide an in-depth description of novel strategies for improving cell factory performance. We focused on improving transcriptional and translational efficiencies through static and dynamic regulatory elements, enzyme engineering and high-throughput screening strategies, cellular function enhancement through chromosomal integration, metabolite tolerance, and modularization of pathways.

OUP accepted manuscript

Optimization of metabolism to maximize production of bio-based chemicals must consistently balance cellular resources for biocatalyst growth and desired compound synthesis. This mini-review discusses synthetic biology strategies for dynamically controlling expression of genes to enable dual-phase fermentations in which growth and production are separated into dedicated phases. Emphasis is placed on practical examples which can be reliably scaled to commercial production with the current state of technology. Recent case studies are presented, and recommendations are provided for environmental signals and genetic control circuits.

Diversifying Isoprenoid Platforms via Atypical Carbon Substrates and Non-model Microorganisms

Isoprenoid compounds are biologically ubiquitous, and their characteristic modularity has afforded products ranging from pharmaceuticals to biofuels. Isoprenoid production has been largely successful in Escherichia coli and Saccharomyces cerevisiae with metabolic engineering of the mevalonate (MVA) and methylerythritol phosphate (MEP) pathways coupled with the expression of heterologous terpene synthases. Yet conventional microbial chassis pose several major obstacles to successful commercialization including the affordability of sugar substrates at scale, precursor flux limitations, and intermediate feedback-inhibition. Now, recent studies have challenged typical isoprenoid paradigms by expanding the boundaries of terpene biosynthesis and using non-model organisms including those capable of metabolizing atypical C1 substrates. Conversely, investigations of non-model organisms have historically informed optimization in conventional microbes by tuning heterologous gene expression. Here, we review advances in isoprenoid biosynthesis with specific focus on the synergy between model and non-model organisms that may elevate the commercial viability of isoprenoid platforms by addressing the dichotomy between high titer production and inexpensive substrates.

CRISPR/Cas Technologies and Their Applications in Escherichia coli

The clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated protein (Cas) systems have revolutionized genome editing and greatly promoted the development of biotechnology. However, these systems unfortunately have not been developed and applied in bacteria as extensively as in eukaryotic organism. Here, the research progress on the most widely used CRISPR/Cas tools and their applications in Escherichia coli is summarized. Genome editing based on homologous recombination, non-homologous DNA end-joining, transposons, and base editors are discussed. Finally, the state of the art of transcriptional regulation using CRISPRi is briefly reviewed. This review provides a useful reference for the application of CRISPR/Cas systems in other bacterial species.

Synthetic Biology Advanced Natural Product Discovery

A wide variety of bacteria, fungi and plants can produce bioactive secondary metabolites, which are often referred to as natural products. With the rapid development of DNA sequencing technology and bioinformatics, a large number of putative biosynthetic gene clusters have been reported. However, only a limited number of natural products have been discovered, as most biosynthetic gene clusters are not expressed or are expressed at extremely low levels under conventional laboratory conditions. With the rapid development of synthetic biology, advanced genome mining and engineering strategies have been reported and they provide new opportunities for discovery of natural products. This review discusses advances in recent years that can accelerate the design, build, test, and learn (DBTL) cycle of natural product discovery, and prospects trends and key challenges for future research directions.

Competitive dCas9 binding as a mechanism for transcriptional control

Catalytically dead Cas9 (dCas9) is a programmable transcription factor that can be targeted to promoters through the design of small guide RNAs (sgRNAs), where it can function as an activator or repressor. Natural promoters use overlapping binding sites as a mechanism for signal integration, where the binding of one can block, displace, or augment the activity of the other. Here, we implemented this strategy in Escherichia coli using pairs of sgRNAs designed to repress and then derepress transcription through competitive binding. When designed to target a promoter, this led to 27-fold repression and complete derepression. This system was also capable of ratiometric input comparison over two orders of magnitude. Additionally, we used this mechanism for promoter sequence-independent control by adopting it for elongation control, achieving 8-fold repression and 4-fold derepression. This work demonstrates a new genetic control mechanism that could be used to build analog circuit or implement cis-regulatory logic on CRISPRi-targeted native genes.

CRISPR/dCas9-Based Systems: Mechanisms and Applications in Plant Sciences

RNA-guided genomic transcriptional regulation tools, namely clustered regularly interspaced short palindromic repeats interference (CRISPRi) and CRISPR-mediated gene activation (CRISPRa), are a powerful technology for gene functional studies. Deriving from the CRISPR/Cas9 system, both systems consist of a catalytically dead Cas9 (dCas9), a transcriptional effector and a single guide RNA (sgRNA). This type of dCas9 is incapable to cleave DNA but retains its ability to specifically bind to DNA. The binding of the dCas9/sgRNA complex to a target gene results in transcriptional interference. The CRISPR/dCas9 system has been explored as a tool for transcriptional modulation and genome imaging. Despite its potential applications and benefits, the challenges and limitations faced by the CRISPR/dCas9 system include the off-target effects, protospacer adjacent motif (PAM) sequence requirements, efficient delivery methods and the CRISPR/dCas9-interfered crops being labeled as genetically modified organisms in several countries. This review highlights the progression of CRISPR/dCas9 technology as well as its applications and potential challenges in crop improvement.

Rational engineering of a modular bacterial CRISPR-Cas activation platform with expanded target range

CRISPR-Cas activator (CRISPRa) systems that selectively turn on transcription of a target gene are a potentially transformative technology for programming cellular function. While in eukaryotes versatile CRISPRa systems exist, in bacteria these systems suffer from a limited ability to activate different genes due to strict distance-dependent requirements of functional target binding sites, and require greater customization to optimize performance in different genetic and cellular contexts. To address this, we apply a rational protein engineering approach to create a new CRISPRa platform that is highly modular to allow for easy customization and has increased targeting flexibility through harnessing engineered Cas proteins. We first demonstrate that transcription activation domains can be recruited by CRISPR-Cas through noncovalent protein-protein interactions, which allows each component to be encoded on separate and easily interchangeable plasmid elements. We then exploit this modularity to rapidly screen a library of different activation domains, creating new systems with distinct regulatory properties. Furthermore, we demonstrate that by harnessing a library of circularly permuted Cas proteins, we can create CRISPRa systems that have different target binding site requirements, which together, allow for expanded target range.

A plasmid toolset for CRISPR-mediated genome editing and CRISPRi gene regulation in Escherichia coli

CRISPR technologies have become standard laboratory tools for genetic manipulations across all kingdoms of life. Despite their origins in bacteria, the development of CRISPR tools for engineering bacteria has been slower than for eukaryotes; nevertheless, their function and application for genome engineering and gene regulation via CRISPR interference (CRISPRi) has been demonstrated in various bacteria, and adoption has become more widespread. Here, we provide simple plasmid-based systems for genome editing (gene knockouts/knock-ins, and genome integration of large DNA fragments) and CRISPRi in E. coli using a CRISPR-Cas12a system. The described genome engineering protocols allow markerless deletion or genome integration in just seven working days with high efficiency (> 80% and 50%, respectively), and the CRISPRi protocols allow robust transcriptional repression of target genes (> 90%) with a single cloning step. The presented minimized plasmids and their associated design and experimental protocols provide efficient and effective CRISPR-Cas12 genome editing, genome integration and CRISPRi implementation. These simple-to-use systems and protocols will allow the easy adoption of CRISPR technology by any laboratory.

Rationally optimized generation of integrated Escherichia coli with stable and high yield lycopene biosynthesis from heterologous mevalonate (MVA) and lycopene expression pathways

The stability and high productivity of heterogeneous terpenoid production in Escherichia coli expression system is one of the most key issues for its large scale industrialization. In the current study on taking lycopene biosynthesis as an example, an integrated Escherichia coli system has been generated successfully, which resulted into stable and high lycopene production. In this process, two modules of mevalonate (MVA) pathway and one module of lycopene expression pathway were completely integrated in the chromosome. Firstly, the copy number and integrated position of three modules of heterologous pathways were rationally optimized. Later, a strain DH416 equipped with heterogeneous expression pathways through chromosomal integration was efficiently derived from parental strain DH411. The evolving DH416 strain efficiently produced the lycopene level of 1.22 g/L (49.9 mg/g DCW) in a 5 L fermenter with mean productivity of 61.0 mg/L/h. Additionally, the integrated strain showed more genetic stability than the plasmid systems after successive 21st passage.

Application of CRISPR/Cas System in the Metabolic Engineering of Small Molecules

Clustered regularly interspaced short palindromic repeats (CRISPR) and their associated Cas protein technology area is rapidly growing technique for genome editing and modulation of transcription of several microbes. Successful engineering in microbes requires an emphasis on the aspect of efficiency and targeted aiming, which can be employed using CRISPR/Cas system. Hence, this type of system is used to modify the genome of several microbes such as yeast and bacteria. In recent years, CRISPR/Cas systems have been chosen for metabolic engineering in microbes due to their specificity, orthogonality, and efficacy. Therefore, we need to review the scheme which was acquired for the execution of the CRISPR/Cas system for the modification and metabolic engineering in yeast and bacteria. In this review, we highlighted the application of the CRISPR/Cas system which has been used for the production of small molecules in the microbial system that is chemically and biologically important.

Development and application of a highly efficient CRISPR-Cas9 system for genome engineering in Bacillus megaterium

Bacillus megaterium has become increasingly important for the biotechnological production of valuable compounds of industrial and pharmaceutical importance. Despite recent advances in rational strain design of B. megaterium, these studies have been largely impaired by the lack of molecular tools that are not state-of-the-art for comprehensive genome engineering approaches. In the current work, we describe the adaptation of the CRISPR-Cas9 vector pJOE8999 to enable efficient genome editing in B. megaterium. Crucial modifications comprise the exchange of promoter elements and associated ribosomal binding sites as well as the implementation of a 5-fluorouracil based counterselection system to facilitate proper plasmid curing. In addition, the functionality and performance of the new CRISPR-Cas9 vector pMOE was successfully evaluated by chromosomal disruption studies of the endogenous β-galactosidase gene (BMD_2126) and demonstrated an outstanding efficiency of 100 % based on combinatorial pheno- and genotype analyses. Furthermore, pMOE was applied for the genomic deletion of a steroid esterase gene (BMD_2256) that was identified among several other candidates as the gene encoding the esterase, which prevented accumulation of pharmaceutically important glucocorticoid esters. Recombinant expression of the bacterial chloramphenicol acetyltransferase 1 gene (cat1) in the resulting esterase deficient B. megaterium strain ultimately yielded C21-acetylated as well as novel C21-esterified derivates of cortisone.

Engineering an Optogenetic CRISPRi Platform for Improved Chemical Production

Microbial synthesis of chemicals typically requires the redistribution of metabolic flux toward the synthesis of targeted products. Dynamic control is emerging as an effective approach for solving the hurdles mentioned above. As light could control the cell behavior in a spatial and temporal manner, the optogenetic-CRISPR interference (opto-CRISPRi) technique that allocates the metabolic resources according to different optical signal frequencies will enable bacteria to be controlled between the growth phase and the production stage. In this study, we applied a blue light-sensitive protein EL222 to regulate the expression of the dCpf1-mediated CRISPRi system that turns off the competitive pathways and redirects the metabolic flux toward the heterologous muconic acid synthesis in Escherichia coli. We found that the opto-CRISPRi system dynamically regulating the suppression of the central metabolism and competitive pathways could increase the muconic acid production by 130%. These results demonstrated that the opto-CRISPRi platform is an effective method for enhancing chemical synthesis with broad utilities.

Reprogramming microorganisms for the biosynthesis of astaxanthin via metabolic engineering

There is an increasing demand for astaxanthin in food, feed, cosmetics and pharmaceutical applications because of its superior anti-oxidative and coloring properties. However, naturally produced astaxanthin is expensive, mainly due to low productivity and limited sources. Reprogramming of microorganisms for astaxanthin production via metabolic engineering is a promising strategy. We primarily focus on the application of synthetic biology, enzyme engineering and metabolic engineering in enhancing the synthesis and accumulation of astaxanthin in microorganisms in this review. We also discuss the biosynthetic pathways of astaxanthin within natural producers, and summarize the achievements and challenges in reprogramming microorganisms for enhancing astaxanthin production. This review illuminates recent biotechnological advances in microbial production of astaxanthin. Future perspectives on utilization of new technologies for boosting microbial astaxanthin production are also discussed.

CRISPR-based metabolic pathway engineering

A highly effective metabolic pathway is the key for an efficient cell factory. However, the engineered homologous or heterologous multi-gene pathway may be unbalanced, inefficient and causing the accumulation of potentially toxic intermediates. Therefore, pathways must be constructed optimally to minimize these negative effects and maximize catalytic efficiency. With the development of CRISPR technology, some of the problems of previous pathway engineering and genome editing techniques were resolved, providing higher efficiency, lower cost, and easily customizable targets. Moreover, CRISPR was demonstrated as robust and effective in various organisms including both prokaryotes and eukaryotes. In recent years, researchers in the field of metabolic engineering and synthetic biology have exploited various CRISPR-based pathway engineering approaches, which are both effective and convenient, as well as valuable from a theoretical standpoint. In this review, we systematically summarize novel pathway engineering techniques and strategies based on CRISPR nucleases system, CRISPR interference (CRISPRi), and CRISPR activation (CRISPRa), including figures and descriptions for easy understanding, with the aim to facilitate their broader application among fellow researchers.

Synthetic Biology on Acetogenic Bacteria for Highly Efficient Conversion of C1 Gases to Biochemicals

Synthesis gas, which is mainly produced from fossil fuels or biomass gasification, consists of C1 gases such as carbon monoxide, carbon dioxide, and methane as well as hydrogen. Acetogenic bacteria (acetogens) have emerged as an alternative solution to recycle C1 gases by converting them into value-added biochemicals using the Wood-Ljungdahl pathway. Despite the advantage of utilizing acetogens as biocatalysts, it is difficult to develop industrial-scale bioprocesses because of their slow growth rates and low productivities. To solve these problems, conventional approaches to metabolic engineering have been applied; however, there are several limitations owing to the lack of required genetic bioparts for regulating their metabolic pathways. Recently, synthetic biology based on genetic parts, modules, and circuit design has been actively exploited to overcome the limitations in acetogen engineering. This review covers synthetic biology applications to design and build industrial platform acetogens.

Synthetic biology, combinatorial biosynthesis, and chemo‑enzymatic synthesis of isoprenoids

Isoprenoids are a large class of natural products with myriad applications as bioactive and commercial compounds. Their diverse structures are derived from the biosynthetic assembly and tailoring of their scaffolds, ultimately constructed from two C5 hemiterpene building blocks. The modular logic of these platforms can be harnessed to improve titers of valuable isoprenoids in diverse hosts and to produce new-to-nature compounds. Often, this process is facilitated by the substrate or product promiscuity of the component enzymes, which can be leveraged to produce novel isoprenoids. To complement rational enhancements and even re-programming of isoprenoid biosynthesis, high-throughput approaches that rely on searching through large enzymatic libraries are being developed. This review summarizes recent advances and strategies related to isoprenoid synthetic biology, combinatorial biosynthesis, and chemo-enzymatic synthesis, focusing on the past 5 years. Emerging applications of cell-free biosynthesis and high-throughput tools are included that culminate in a discussion of the future outlook and perspective of isoprenoid biosynthetic engineering.

Strain engineering for high-level 5-aminolevulinic acid production in Escherichia coli

Herein, we report the development of a microbial bioprocess for high-level production of 5-aminolevulinic acid (5-ALA), a valuable non-proteinogenic amino acid with multiple applications in medical, agricultural, and food industries, using Escherichia coli as a cell factory. We first implemented the Shemin (i.e., C4) pathway for heterologous 5-ALA biosynthesis in E. coli. To reduce, but not to abolish, the carbon flux toward essential tetrapyrrole/porphyrin biosynthesis, we applied clustered regularly interspersed short palindromic repeats interference (CRISPRi) to repress hemB expression, leading to extracellular 5-ALA accumulation. We then applied metabolic engineering strategies to direct more dissimilated carbon flux toward the key precursor of succinyl-CoA for enhanced 5-ALA biosynthesis. Using these engineered E. coli strains for bioreactor cultivation, we successfully demonstrated high-level 5-ALA biosynthesis from glycerol (~30 g L^-1 ) under both microaerobic and aerobic conditions, achieving up to 5.95 g L^-1 (36.9% of the theoretical maximum yield) and 6.93 g L^-1 (50.9% of the theoretical maximum yield) 5-ALA, respectively. This study represents one of the most effective bio-based production of 5-ALA from a structurally unrelated carbon to date, highlighting the importance of integrated strain engineering and bioprocessing strategies to enhance bio-based production.

Application of different types of CRISPR/Cas-based systems in bacteria

As important genome editing tools, CRISPR/Cas systems, especially those based on type II Cas9 and type V Cas12a, are widely used in genetic and metabolic engineering of bacteria. However, the intrinsic toxicity of Cas9 and Cas12a-mediated CRISPR/Cas tools can lead to cell death in some strains, which led to the development of endogenous type I and III CRISPR/Cas systems. However, these systems are hindered by complicated development and limited applications. Thus, further development and optimization of CRISPR/Cas systems is needed. Here, we briefly summarize the mechanisms of different types of CRISPR/Cas systems as genetic manipulation tools and compare their features to provide a reference for selecting different CRISPR/Cas tools. Then, we show the use of CRISPR/Cas technology for bacterial strain evolution and metabolic engineering, including genome editing, gene expression regulation and the base editor tool. Finally, we offer a view of future directions for bacterial CRISPR/Cas technology.

Microbial Cell Factory for Efficiently Synthesizing Plant Natural Products via Optimizing the Location and Adaptation of Pathway on Genome Scale

Plant natural products (PNPs) possess important pharmacological activities and are widely used in cosmetics, health care products, and as food additives. Currently, most PNPs are mainly extracted from cultivated plants, and the yield is limited by the long growth cycle, climate change and complex processing steps, which makes the process unsustainable. However, the complex structure of PNPs significantly reduces the efficiency of chemical synthesis. With the development of metabolic engineering and synthetic biology, heterologous biosynthesis of PNPs in microbial cell factories offers an attractive alternative. Based on the in-depth mining and analysis of genome and transcriptome data, the biosynthetic pathways of a number of natural products have been successfully elucidated, which lays the crucial foundation for heterologous production. However, there are several problems in the microbial synthesis of PNPs, including toxicity of intermediates, low enzyme activity, multiple auxotrophic dependence, and uncontrollable metabolic network. Although various metabolic engineering strategies have been developed to solve these problems, optimizing the location and adaptation of pathways on the whole-genome scale is an important strategy in microorganisms. From this perspective, this review introduces the application of CRISPR/Cas9 in editing PNPs biosynthesis pathways in model microorganisms, the influences of pathway location, and the approaches for optimizing the adaptation between metabolic pathways and chassis hosts for facilitating PNPs biosynthesis.

Development of CRISPR Interference (CRISPRi) Platform for Metabolic Engineering of Leuconostoc citreum and Its Application for Engineering Riboflavin Biosynthesis

Leuconostoc citreum, a hetero-fermentative type of lactic acid bacteria, is a crucial probiotic candidate because of its ability to promote human health. However, inefficient gene manipulation tools limit its utilization in bioindustries. We report, for the first time, the development of a CRISPR (Clustered Regularly Interspaced Short Palindromic Repeats) interference (CRISPRi) system for engineering L. citreum. For reliable expression, the expression system of synthetic single guide RNA (sgRNA) and the deactivated Cas9 of Streptococcus pyogenes (SpdCas9) were constructed in a bicistronic design (BCD) platform using a high-copy-number plasmid. The expression of SpdCas9 and sgRNA was optimized by examining the combination of two synthetic promoters and Shine-Dalgarno sequences; the strong expression of sgRNA and the weak expression of SpdCas9 exhibited the most significant downregulation (20-fold decrease) of the target gene (sfGFP), without cell growth retardation caused by SpdCas9 overexpression. The feasibility of the optimized CRISPRi system was demonstrated by modulating the biosynthesis of riboflavin. Using the CRISPRi system, the expression of ribF and folE genes was downregulated (3.3-fold and 5.6-fold decreases, respectively), thereby improving riboflavin production. In addition, the co-expression of the rib operon was introduced and the production of riboflavin was further increased up to 1.7 mg/L, which was 1.53 times higher than that of the wild-type strain.

CRISPR-Cas9/CRISPRi tools for cell factory construction in E. coli

The innovative CRISPR-Cas based genome editing technology provides some functionality and advantages such as the high efficiency and specificity as well as ease of handling. Both aspects of the CRISPR-Cas9 system including genetic engineering and gene regulation are advantageously applicable to the construction of microbial cell factories. As one of the most extensively used cell factories, E. coli has been engineered to produce various high value-added chemical compounds such as pharmaceuticals, biochemicals, and biofuels. Therefore, to improve the production of valuable metabolites, many investigations have been performed by focusing on CRISPR-Cas- based metabolic engineering of this host. In the current review, the biology underlying CRISPR-Cas9 system was briefly explained and then the applications of CRISPR-Cas9/CRISPRi tools were considered for cell factory construction in E. coli.

Conventional genetic manipulation of desulfurizing bacteria and prospects of using CRISPR-Cas systems for enhanced desulfurization activity

Highly active and stable biocatalysts are the prerequisite for industrial scale application of the biodesulfurization process. Scientists are making efforts for increasing the desulfurizing activity of native strains by employing various genetic engineering approaches. Nevertheless, the achieved desulfurization rate is lower than the industrial requirements. Thus, there is a dire need to use efficient genetic tools for precise genome editing of desulfurizing bacteria for enhanced efficiency. In comparison to the previously used genetic engineering tools the newly developed CRISPR-Cas is a more efficient and simple genetic tool that has been successfully applied for targeted genome modification of eukaryotes as well as prokaryotes. In this paper, we have reviewed the approaches, previously used to enhance the biodesulfurization rates of the sulfur metabolizing microorganisms and have discussed the potential of CRISPR-Cas systems in engineering desulfurizing biocatalysts. We have also proposed a model to construct competent desulfurizing recombinants involving use of CRISPR-Cas technology. The model can be used to over-express the dsz genes under a constitutive promoter in a suitable heterologous host, to get a steady expression of desulfurization pathway. This may serve as an inducement to develop better performing desulfurizing recombinant strains using CRISPR-Cas systems, which can be helpful in increasing the rate of biodesulfurization in future.

CRISPR Tools for Physiology and Cell State Changes: Potential of Transcriptional Engineering and Epigenome Editing

Given the large amount of genome-wide data that have been collected during the last decades, a good understanding of how and why cells change during development, homeostasis, and disease might be expected. Unfortunately, the opposite is true; triggers that cause cellular state changes remain elusive, and the underlying molecular mechanisms are poorly understood. Although genes with the potential to influence cell states are known, the historic dependency on methods that manipulate gene expression outside the endogenous chromatin context has prevented us from understanding how cells organize, interpret, and protect cellular programs. Fortunately, recent methodological innovations are now providing options to answer these outstanding questions, by allowing to target and manipulate individual genomic and epigenomic loci. In particular, three experimental approaches are now feasible due to DNA targeting tools, namely, activation and/or repression of master transcription factors in their endogenous chromatin context; targeting transcription factors to endogenous, alternative, or inaccessible sites; and finally, functional manipulation of the chromatin context. In this article, we discuss the molecular basis of DNA targeting tools and review the potential of these new technologies before we summarize how these have already been used for the manipulation of cellular states and hypothesize about future applications.