CRISPR/dCas9-Mediated Multiplex Gene Repression in Streptomyces

Direct reprogramming of fibroblasts into functional hepatocytes via CRISPRa activation of endogenous Gata4 and Foxa3

BACKGROUND

The ability to generate functional hepatocytes without relying on donor liver organs holds significant therapeutic promise in the fields of regenerative medicine and potential liver disease treatments. Clustered regularly interspaced short palindromic repeats (CRISPR) activator (CRISPRa) is a powerful tool that can conveniently and efficiently activate the expression of multiple endogenous genes simultaneously, providing a new strategy for cell fate determination. The main purpose of this study is to explore the feasibility of applying CRISPRa for hepatocyte reprogramming and its application in the treatment of mouse liver fibrosis.

METHOD

The differentiation of mouse embryonic fibroblasts (MEFs) into functional induced hepatocyte-like cells (iHeps) was achieved by utilizing the CRISPRa synergistic activation mediator (SAM) system, which drove the combined expression of three endogenous transcription factors- Gata4, Foxa3 , and Hnf1a -or alternatively, the expression of two transcription factors, Gata4 and Foxa3 . In vivo , we injected adeno-associated virus serotype 6 (AAV6) carrying the CRISPRa SAM system into liver fibrotic Col1a1-CreER ; Cas9fl/fl mice, effectively activating the expression of endogenous Gata4 and Foxa3 in fibroblasts. The endogenous transcriptional activation of genes was confirmed using real-time quantitative polymerase chain reaction (RT-qPCR) and RNA-seq, and the morphology and characteristics of the induced hepatocytes were observed through microscopy. The level of hepatocyte reprogramming in vivo is detected by immunofluorescence staining, while the improvement of liver fibrosis is evaluated through Sirius red staining, alpha-smooth muscle actin (α-SMA) immunofluorescence staining, and blood alanine aminotransferase (ALT) examination.

RESULTS

Activation of only two factors, Gata4 and Foxa3 , via CRISPRa was sufficient to successfully induce the transformation of MEFs into iHeps. These iHeps could be expanded in vitro and displayed functional characteristics similar to those of mature hepatocytes, such as drug metabolism and glycogen storage. Additionally, AAV6-based delivery of the CRISPRa SAM system effectively induced the hepatic reprogramming from fibroblasts in mice with live fibrosis. After 8 weeks of induction, the reprogrammed hepatocytes comprised 0.87% of the total hepatocyte population in the mice, significantly reducing liver fibrosis.

CONCLUSION

CRISPRa-induced hepatocyte reprogramming may be a promising strategy for generating functional hepatocytes and treating liver fibrosis caused by hepatic diseases.

Enhancement of acyl-CoA precursor supply for increased avermectin B1a production by engineering meilingmycin polyketide synthase and key primary metabolic pathway genes

Avermectins (AVEs), a family of macrocyclic polyketides produced by Streptomyces avermitilis, have eight components, among which B1a is noted for its strong insecticidal activity. Biosynthesis of AVE "a" components requires 2-methylbutyryl-CoA (MBCoA) as starter unit, and malonyl-CoA (MalCoA) and methylmalonyl-CoA (MMCoA) as extender units. We describe here a novel strategy for increasing B1a production by enhancing acyl-CoA precursor supply. First, we engineered meilingmycin (MEI) polyketide synthase (PKS) for increasing MBCoA precursor supply. The loading module (using acetyl-CoA as substrate), extension module 7 (using MMCoA as substrate) and TE domain of MEI PKS were assembled to produce 2-methylbutyrate, providing the starter unit for B1a production. Heterologous expression of the newly designed PKS (termed Mei-PKS) in S. avermitilis wild-type (WT) strain increased MBCoA level, leading to B1a titer 262.2 μg/mL - 4.36-fold higher than WT value (48.9 μg/mL). Next, we separately inhibited three key nodes in essential pathways using CRISPRi to increase MalCoA and MMCoA levels in WT. The resulting strains all showed increased B1a titer. Combined inhibition of these key nodes in Mei-PKS expression strain increased B1a titer to 341.9 μg/mL. Overexpression of fatty acid β-oxidation pathway genes in the strain further increased B1a titer to 452.8 μg/mL - 8.25-fold higher than WT value. Finally, we applied our precursor supply strategies to high-yield industrial strain A229. The strategies, in combination, led to B1a titer 8836.4 μg/mL - 37.8% higher than parental A229 value. These findings provide an effective combination strategy for increasing AVE B1a production in WT and industrial S. avermitilis strains, and our precursor supply strategies can be readily adapted for overproduction of other polyketides.

An inducible CRISPRi circuit for tunable dynamic regulation of gene expression in Saccharopolyspora erythraea

Actinomyces are gram-positive bacteria known for their valuable secondary metabolites. Redirecting metabolic flux towards desired products in actinomycetes requires precise and dynamic regulation of gene expression. In this study, we integrated the CRISPR interference (CRISPRi) system with a cumate-inducible promoter to develop an inducible gene downregulation method in Saccharopolyspora erythraea, a prominent erythromycin-producing actinobacterium. The functionality of the cumate-inducible promoter was validated using the gusA gene as a reporter, and the successful inducible expression of the dCas9 gene was confirmed. The developed inducible CRISPRi strategy was then employed to downregulate the expression of target genes rppA in the wild-type strain NRRL2338 and sucC in the high erythromycin-producing strain E3. Through dynamic control of sucC expression, a significant enhancement in erythromycin production was achieved in strain E3. This study demonstrated the effectiveness of an inducible gene downregulation approach using CRISPRi and a cumate-inducible promoter, providing valuable insights for optimizing natural product production in actinomyces.

Genus Streptomyces: Recent advances for biotechnological purposes

Actinomycetes are a distinct group of filamentous bacteria. The Streptomyces genus within this group has been extensively studied over the years, with substantial contributions to society and science. This genus is known for its antimicrobial production, as well as antitumor, biopesticide, and immunomodulatory properties. Therefore, the extraordinary plasticity of the Streptomyces genus has inspired new research techniques. The newest way of exploring Streptomyces has comprised the discovery of new natural metabolites and the application of emerging tools such as CRISPR technology in drug discovery. In this narrative review, we explore relevant published literature concerning the ongoing novelties of the Streptomyces genus.

Rationally Improving Doramectin Production in Industrial Streptomyces avermitilis Strains

Avermectins (AVMs), a family of 16-membered macrocyclic macrolides produced by Streptomyces avermitilis, have been the most successful microbial natural antiparasitic agents in recent decades. Doramectin, an AVM derivative produced by S. avermitilis bkd^- mutants through cyclohexanecarboxylic acid (CHC) feeding, was commercialized as a veterinary antiparasitic drug by Pfizer Inc. Our previous results show that the production of avermectin and actinorhodin was affected by several other polyketide biosynthetic gene clusters in S. avermitilis and Streptomyces coelicolor, respectively. Thus, here, we propose a rational strategy to improve doramectin production via the termination of competing polyketide biosynthetic pathways combined with the overexpression of CoA ligase, providing precursors for polyketide biosynthesis. fadD17, an annotated putative cyclohex-1-ene-1-carboxylate:CoA ligase-encoding gene, was proven to be involved in the biosynthesis of doramectin. By sequentially removing three PKS (polyketide synthase) gene clusters and overexpressing FadD17 in the strain DM203, the resulting strain DM223 produced approximately 723 mg/L of doramectin in flasks, which was approximately 260% that of the original strain DM203 (approximately 280 mg/L). To summarize, our work demonstrates a novel viable approach to engineer doramectin overproducers, which might contribute to the reduction in the cost of this valuable compound in the future.

Establishment of a visual gene knockout system based on CRISPR/Cas9 for the rare actinomycete Nonomuraea gerenzanensis

OBJECTIVES

To develop a modified CRISPR/Cas9 system with the β-glucuronidase (GusA) reporter and a dual sgRNA cassette for Nonomuraea gerenzanensis (N. gerenzanensis).

RESULTS

With the aid of a visual GusA reporter, the complicated and tedious process of cloning and gene identification could be abandoned entirely in the genetic editing of N. gerenzanensis. Moreover, introducing a dual sgRNA cassette into the CRISPR/Cas9 system significantly improved gene deletion efficiency compared to the single sgRNA element. Furthermore, the length of the homologous flanking sequences set to the lowest value of 500 bp in this system could still reach the relatively higher conjugation transfer frequency.

CONCLUSIONS

The enhanced CRISPR/Cas9 system could efficiently perform genetic manipulation on the rare actinomycete N. gerenzanensis.

Characterization of the Biosynthetic Gene Cluster and Shunt Products Yields Insights into the Biosynthesis of Balmoralmycin

Angucyclines are a family of structurally diverse, aromatic polyketides with some members that exhibit potent bioactivity. Angucyclines have also attracted considerable attention due to the intriguing biosynthetic origins that underlie their structural complexity and diversity. Balmoralmycin (compound 1) represents a unique group of angucyclines that contain an angular benz[α]anthracene tetracyclic system, a characteristic C-glycosidic bond-linked deoxy-sugar (d-olivose), and an unsaturated fatty acid chain. In this study, we identified a Streptomyces strain that produces balmoralmycin and seven biosynthetically related coproducts (compounds 2-8). Four of the coproducts (compounds 5-8) are novel compounds that feature a highly oxygenated or fragmented lactone ring, and three of them (compounds 3-5) exhibited cytotoxicity against the human pancreatic cancer cell line MIA PaCa-2 with IC₅₀ values ranging from 0.9 to 1.2 μg/mL. Genome sequencing and CRISPR/dCas9-assisted gene knockdown led to the identification of the ~43 kb balmoralmycin biosynthetic gene cluster (bal BGC). The bal BGC encodes a type II polyketide synthase (PKS) system for assembling the angucycline aglycone, six enzymes for generating the deoxysugar d-olivose, and a hybrid type II/III PKS system for synthesizing the 2,4-decadienoic acid chain. Based on the genetic and chemical information, we propose a mechanism for the biosynthesis of balmoralmycin and the shunt products. The chemical and genetic studies yielded insights into the biosynthetic origin of the structural diversity of angucyclines. IMPORTANCE Angucyclines are structurally diverse aromatic polyketides that have attracted considerable attention due to their potent bioactivity and intriguing biosynthetic origin. Balmoralmycin is a representative of a small family of angucyclines with unique structural features and an unknown biosynthetic origin. We report a newly isolated Streptomyces strain that produces balmoralmycin in a high fermentation titer as well as several structurally related shunt products. Based on the chemical and genetic information, a biosynthetic pathway that involves a type II polyketide synthase (PKS) system, cyclases/aromatases, oxidoreductases, and other ancillary enzymes was established. The elucidation of the balmoralmycin pathway enriches our understanding of how structural diversity is generated in angucyclines and opens the door for the production of balmoralmycin derivatives via pathway engineering.

Two-Component Systems of Streptomyces coelicolor: An Intricate Network to Be Unraveled

Bacteria of the Streptomyces genus constitute an authentic biotech gold mine thanks to their ability to produce a myriad of compounds and enzymes of great interest at various clinical, agricultural, and industrial levels. Understanding the physiology of these organisms and revealing their regulatory mechanisms is essential for their manipulation and application. Two-component systems (TCSs) constitute the predominant signal transduction mechanism in prokaryotes, and can detect a multitude of external and internal stimuli and trigger the appropriate cellular responses for adapting to diverse environmental conditions. These global regulatory systems usually coordinate various biological processes for the maintenance of homeostasis and proper cell function. Here, we review the multiple TCSs described and characterized in Streptomyces coelicolor, one of the most studied and important model species within this bacterial group. TCSs are involved in all cellular processes; hence, unravelling the complex regulatory network they form is essential for their potential biotechnological application.

Enhancement of acarbose production by genetic engineering and fed-batch fermentation strategy in Actinoplanes sp. SIPI12-34

Background

Acarbose, as an alpha-glucosidase inhibitor, is widely used clinically to treat type II diabetes. In its industrial production, Actinoplanes sp. SE50/110 is used as the production strain. Lack of research on its regulatory mechanisms and unexplored gene targets are major obstacles to rational strain design. Here, transcriptome sequencing was applied to uncover more gene targets and rational genetic engineering was performed to increase acarbose production.

Results

In this study, with the help of transcriptome information, a TetR family regulator (TetR1) was identified and confirmed to have a positive effect on the synthesis of acarbose by promoting the expression of acbB and acbD. Some genes with low expression levels in the acarbose biosynthesis gene cluster were overexpressed and this resulted in a significant increase in acarbose yield. In addition, the regulation of metabolic pathways was performed to retain more glucose-1-phosphate for acarbose synthesis by weakening the glycogen synthesis pathway and strengthening the glycogen degradation pathway. Eventually, with a combination of multiple strategies and fed-batch fermentation, the yield of acarbose in the engineered strain increased 58% compared to the parent strain, reaching 8.04 g/L, which is the highest fermentation titer reported.

Conclusions

In our research, acarbose production had been effectively and steadily improved through genetic engineering based on transcriptome analysis and fed-batch culture strategy.

Graphical Abstract

Developing Genetic Engineering Techniques for Control of Seed Size and Yield

Many signaling pathways regulate seed size through the development of endosperm and maternal tissues, which ultimately results in a range of variations in seed size or weight. Seed size can be determined through the development of zygotic tissues (endosperm and embryo) and maternal ovules. In addition, in some species such as rice, seed size is largely determined by husk growth. Transcription regulator factors are responsible for enhancing cell growth in the maternal ovule, resulting in seed growth. Phytohormones induce significant effects on entire features of growth and development of plants and also regulate seed size. Moreover, the vegetative parts are the major source of nutrients, including the majority of carbon and nitrogen-containing molecules for the reproductive part to control seed size. There is a need to increase the size of seeds without affecting the number of seeds in plants through conventional breeding programs to improve grain yield. In the past decades, many important genetic factors affecting seed size and yield have been identified and studied. These important factors constitute dynamic regulatory networks governing the seed size in response to environmental stimuli. In this review, we summarized recent advances regarding the molecular factors regulating seed size in Arabidopsis and other crops, followed by discussions on strategies to comprehend crops' genetic and molecular aspects in balancing seed size and yield.

Engineering of succinyl-CoA metabolism in view of succinylation regulation to improve the erythromycin production

As a novel protein post-translational modification (PTM), lysine succinylation is widely involved in metabolism regulation by altering the activity of catalytic enzymes. Inactivating succinyl-CoA synthetase in Saccharopolyspora erythraea HL3168 E3 was proved significantly inducing the global protein hypersuccinylation. To investigate the effects, succinylome of the mutant strain E3ΔsucC was identified by using a high-resolution mass spectrometry-based proteomics approach. PTMomics analyses suggested the important roles of succinylation on protein biosynthesis, carbon metabolism, and antibiotics biosynthesis in S. erythraea. Enzymatic experiments in vivo and in vitro were further conducted to determine the succinylation regulation in the TCA cycle. We found out that the activity of aconitase (SACE_3811) was significantly inhibited by succinylation in E3ΔsucC, which probably led to the extracellular accumulation of pyruvate and citrate during the fermentation. Enzyme structural analyses indicated that the succinylation of K278 and K373, conservative lysine residues locating around the protein binding pocket, possibly affects the activity of aconitase. To alleviate the metabolism changes caused by succinyl-CoA synthetase inactivation and protein hypersuccinylation, CRISPR interference (CRISPRi) was applied to mildly downregulate the transcription level of gene sucC in E3. The erythromycin titer of the CRISPRi mutant E3-sucC-sg1 was increased by 54.7% compared with E3, which was 1200.5 mg/L. Taken together, this work not only expands our knowledge of succinylation regulation in the TCA cycle, but also validates that CRISPRi is an efficient strategy on the metabolic engineering of S. erythraea. KEY POINTS: • We reported the first systematic profiling of the S. erythraea succinylome. • We found that the succinylation regulation on the activity of aconitase. • We enhanced the production of erythromycin by using CRISPRi to regulate the transcription of gene sucC.

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.

Overview of advances in CRISPR/deadCas9 technology and its applications in human diseases

Prokaryotes possess an adaptive immune system using various CRISPR associated (Cas) genes to make an archive of records from invading phages and eliminate them upon re-exposure when specialized Cas proteins cut foreign DNA into small pieces. On the basis of the different types of Cas proteins, CRISPR systems seen in some prokaryotic genomes, are different to each other. It has been proved that CRISPR has a great potential for genome engineering. Studies have also demonstrated that in comparison to the preceding genome engineering tools CRISPR/Cas systems can be harnessed as a flexible tool with easy multiplexing and scaling ability. Recent studies suggest that CRISPR/Cas systems can also be used for non-genome engineering roles. Isolation and identification of new Cas proteins or modification of existing ones are effectively increasing the number of CRISPR applications and helps its development. D10A and H840A mutations at RuvC and HNH endonuclease domains of wild type Streptococcus pyogenes Cas9 (SpCas9) respectively creates a nuclease, dead Cas9 (dCas9) molecule, that does not cut target DNA but still retains its capability for binding to target DNA based on the gRNA targeting sequence. In this article we review the potentials of this enzyme, dCas9, toward development of the applications of CRISPR/dCas9 technology in fields such as; visualization of genomic loci, disease diagnosis and transcriptional repression and activation.

CRISPR-Based Approaches for Gene Regulation in Non-Model Bacteria

CRISPR interference (CRISPRi) and CRISPR activation (CRISPRa) have become ubiquitous approaches to control gene expression in bacteria due to their simple design and effectiveness. By regulating transcription of a target gene(s), CRISPRi/a can dynamically engineer cellular metabolism, implement transcriptional regulation circuitry, or elucidate genotype-phenotype relationships from smaller targeted libraries up to whole genome-wide libraries. While CRISPRi/a has been primarily established in the model bacteria Escherichia coli and Bacillus subtilis, a growing numbering of studies have demonstrated the extension of these tools to other species of bacteria (here broadly referred to as non-model bacteria). In this mini-review, we discuss the challenges that contribute to the slower creation of CRISPRi/a tools in diverse, non-model bacteria and summarize the current state of these approaches across bacterial phyla. We find that despite the potential difficulties in establishing novel CRISPRi/a in non-model microbes, over 190 recent examples across eight bacterial phyla have been reported in the literature. Most studies have focused on tool development or used these CRISPRi/a approaches to interrogate gene function, with fewer examples applying CRISPRi/a gene regulation for metabolic engineering or high-throughput screens and selections. To date, most CRISPRi/a reports have been developed for common strains of non-model bacterial species, suggesting barriers remain to establish these genetic tools in undomesticated bacteria. More efficient and generalizable methods will help realize the immense potential of programmable CRISPR-based transcriptional control in diverse bacteria.

A Study of Type II ɛ-PL Degrading Enzyme (pldII) in Streptomyces albulus through the CRISPRi System

ε-Poly-L-lysine (ε-PL) is a widely used antibacterial peptide polymerized of 25–35 L-lysine residues. The antibacterial effect of ε-PL is closely related to the polymerization degree. However, the mechanism of ε-PL degradation in S. albulus remains unclear. This study utilized the integrative plasmid pSET152-based CRISPRi system to transcriptionally repress the ε-PL degrading enzyme (pldII). The expression of pldII is regulated by changing the recognition site of dCas9. Through the ε-PL bacteriostatic experiments of repression strains, it was found that the repression of pldII improves the antibacterial effect of the ε-PL product. The consecutive MALDI-TOF-MS results confirmed that the molecular weight distribution of the ε-PL was changed after repression. The repression strain S1 showed a particular peak with a polymerization degree of 44, and other repression strains also generated ε-PL with a polymerization degree of over 40. Furthermore, the homology modeling and substrate docking of pldII, a typical endo-type metallopeptidase, were performed to resolve the degradation mechanism of ε-PL in S. albulus. The hydrolysis of ε-PL within pldII, initiated from the N-terminus by two amino acid-binding residues, Thr194 and Glu281, led to varying levels of polymerization of ε-PL.

New approaches to moderate CRISPR-Cas9 activity: Addressing issues of cellular uptake and endosomal escape

CRISPR-Cas9 is rapidly entering molecular biology and biomedicine as a promising gene-editing tool. A unique feature of CRISPR-Cas9 is a single-guide RNA directing a Cas9 nuclease toward its genomic target. Herein, we highlight new approaches for improving cellular uptake and endosomal escape of CRISPR-Cas9. As opposed to other recently published works, this review is focused on non-viral carriers as a means to facilitate the cellular uptake of CRISPR-Cas9 through endocytosis. The majority of non-viral carriers, such as gold nanoparticles, polymer nanoparticles, lipid nanoparticles, and nanoscale zeolitic imidazole frameworks, is developed with a focus toward optimizing the endosomal escape of CRISPR-Cas9 by taking advantage of the acidic environment in the late endosomes. Among the most broadly used methods for in vitro and ex vivo ribonucleotide protein transfection are electroporation and microinjection. Thus, other delivery formats are warranted for in vivo delivery of CRISPR-Cas9. Herein, we specifically revise the use of peptide and nanoparticle-based systems as platforms for CRISPR-Cas9 delivery in vivo. Finally, we highlight future perspectives of the CRISPR-Cas9 gene-editing tool and the prospects of using non-viral vectors to improve its bioavailability and therapeutic potential.

Engineering Modular Polyketide Biosynthesis in Streptomyces Using CRISPR/Cas: A Practical Guide

The CRISPR/Cas system, which has been widely applied to organisms ranging from microbes to animals, is currently being adapted for use in Streptomyces bacteria. In this case, it is notably applied to rationally modify the biosynthetic pathways giving rise to the polyketide natural products, which are heavily exploited in the medical and agricultural arenas. Our aim here is to provide the potential user with a practical guide to exploit this approach for manipulating polyketide biosynthesis, by treating key experimental aspects including vector choice, design of the basic engineering components, and trouble-shooting.

Repurposing CRISPR-Cas Systems as Genetic Tools for the Enterobacteriales

Over the last decade, the study of CRISPR-Cas systems has progressed from a newly discovered bacterial defense mechanism to a diverse suite of genetic tools that have been applied across all domains of life. While the initial applications of CRISPR-Cas technology fulfilled a need to more precisely edit eukaryotic genomes, creative "repurposing" of this adaptive immune system has led to new approaches for genetic analysis of microorganisms, including improved gene editing, conditional gene regulation, plasmid curing and manipulation, and other novel uses. The main objective of this review is to describe the development and current state-of-the-art use of CRISPR-Cas techniques specifically as it is applied to members of the Enterobacteriales. While many of the applications covered have been initially developed in Escherichia coli, we also highlight the potential, along with the limitations, of this technology for expanding the availability of genetic tools in less-well-characterized non-model species, including bacterial pathogens.

Harnessing intercellular signals to engineer the soil microbiome

Covering: Focus on 2015 to 2020Plant and soil microbiomes consist of diverse communities of organisms from across kingdoms and can profoundly affect plant growth and health. Natural product-based intercellular signals govern important interactions between microbiome members that ultimately regulate their beneficial or harmful impacts on the plant. Exploiting these evolved signalling circuits to engineer microbiomes towards beneficial interactions with crops is an attractive goal. There are few reports thus far of engineering the intercellular signalling of microbiomes, but this article argues that it represents a tremendous opportunity for advancing the field of microbiome engineering. This could be achieved through the selection of synergistic consortia in combination with genetic engineering of signal pathways to realise an optimised microbiome.

Improved dsDNA recombineering enables versatile multiplex genome engineering of kilobase-scale sequences in diverse bacteria

Recombineering assisted multiplex genome editing generally uses single-stranded oligonucleotides for site directed mutational changes. It has proven highly efficient for functional screens and to optimize microbial cell factories. However, this approach is limited to relatively small mutational changes. Here, we addressed the challenges involved in the use of double-stranded DNA substrates for multiplex genome engineering. Recombineering is mediated by phage single-strand annealing proteins annealing ssDNAs into the replication fork. We apply this insight to facilitate the generation of ssDNA from the dsDNA substrate and to alter the speed of replication by elevating the available deoxynucleoside triphosphate (dNTP) levels. Intracellular dNTP concentration was elevated by ribonucleotide reductase overexpression or dNTP addition to establish double-stranded DNA Recombineering-assisted Multiplex Genome Engineering (dReaMGE), which enables rapid and flexible insertional and deletional mutagenesis at multiple sites on kilobase scales in diverse bacteria without the generation of double-strand breaks or disturbance of the mismatch repair system. dReaMGE can achieve combinatorial genome engineering works, for example, alterations to multiple biosynthetic pathways, multiple promoter or gene insertions, variations of transcriptional regulator combinations, within a few days. dReaMGE adds to the repertoire of bacterial genome engineering to facilitate discovery, functional genomics, strain optimization and directed evolution of microbial cell factories.

Flaviolin-Like Gene Cluster Deletion Optimized the Butenyl-Spinosyn Biosynthesis Route in Saccharopolyspora pogona

Reduction and optimization of the microbial genome is an important strategy for constructing synthetic biological chassis cells and overcoming obstacles in natural product discovery and production. However, it is of great challenge to discover target genes that can be deleted and optimized due to the complicated genome of actinomycetes. Saccharopolyspora pogona can produce butenyl-spinosyn during aerobic fermentation, and its genome contains 32 different gene clusters. This suggests that there is a large amount of potential competitive metabolism in S. pogona, which affects the biosynthesis of butenyl-spinosyn. By analyzing the genome of S. pogona, six polyketide gene clusters were identified. From those, the complete deletion of clu13, a flaviolin-like gene cluster, generated a high butenyl-spinosyn-producing strain. Production of this strain was 4.06-fold higher than that of the wildtype strain. Transcriptome profiling revealed that butenyl-spinosyn biosynthesis was not primarily induced by the polyketide synthase RppA-like but was related to hypothetical protein Sp1764. However, the repression of sp1764 was not enough to explain the enormous enhancement of butenyl-spinosyn yields in S. pogona-Δclu13. After the comparative proteomic analysis of S. pogona-Δclu13 and S. pogona, two proteins, biotin carboxyl carrier protein (BccA) and response regulator (Reg), were investigated, whose overexpression led to great advantages of butenyl-spinosyn biosynthesis. In this way, we successfully discovered three key genes that obviously optimize the biosynthesis of butenyl-spinosyn. Gene cluster simplification performed in conjunction with multiomics analysis is of great practical significance for screening dominant chassis strains and optimizing secondary metabolism. This work provided an idea about screening key factors and efficient construction of production strains.

Uncovering and Engineering a Mini-Regulatory Network of the TetR-Family Regulator SACE_0303 for Yield Improvement of Erythromycin in Saccharopolyspora erythraea

Erythromycins produced by Saccharopolyspora erythraea have broad-spectrum antibacterial activities. Recently, several TetR-family transcriptional regulators (TFRs) were identified to control erythromycin production by multiplex control modes; however, their regulatory network remains poorly understood. In this study, we report a novel TFR, SACE_0303, positively correlated with erythromycin production in Sac. erythraea. It directly represses its adjacent gene SACE_0304 encoding a MarR-family regulator and indirectly stimulates the erythromycin biosynthetic gene eryAI and resistance gene ermE. SACE_0304 negatively regulates erythromycin biosynthesis by directly inhibiting SACE_0303 as well as eryAI and indirectly repressing ermE. Then, the SACE_0303 binding site within the SACE_0303-SACE_0304 intergenic region was defined. Through genome scanning combined with in vivo and in vitro experiments, three additional SACE_0303 target genes (SACE_2467 encoding cation-transporting ATPase, SACE_3156 encoding a large transcriptional regulator, SACE_5222 encoding α-ketoglutarate permease) were identified and proved to negatively affect erythromycin production. Finally, by coupling CRISPRi-based repression of those three targets with SACE_0304 deletion and SACE_0303 overexpression, we performed stepwise engineering of the SACE_0303-mediated mini-regulatory network in a high-yield strain, resulting in enhanced erythromycin production by 67%. In conclusion, the present study uncovered the regulatory network of a novel TFR for control of erythromycin production and provides a multiplex tactic to facilitate the engineering of industrial actinomycetes for yield improvement of antibiotics.

Data-driven network models for genetic circuits from time-series data with incomplete measurements

Synthetic gene networks are frequently conceptualized and visualized as static graphs. This view of biological programming stands in stark contrast to the transient nature of biomolecular interaction, which is frequently enacted by labile molecules that are often unmeasured. Thus, the network topology and dynamics of synthetic gene networks can be difficult to verify in vivo or in vitro, due to the presence of unmeasured biological states. Here we introduce the dynamical structure function as a new mesoscopic, data-driven class of models to describe gene networks with incomplete measurements of state dynamics. We develop a network reconstruction algorithm and a code base for reconstructing the dynamical structure function from data, to enable discovery and visualization of graphical relationships in a genetic circuit diagram as time-dependent functions rather than static, unknown weights. We prove a theorem, showing that dynamical structure functions can provide a data-driven estimate of the size of crosstalk fluctuations from an idealized model. We illustrate this idea with numerical examples. Finally, we show how data-driven estimation of dynamical structure functions can explain failure modes in two experimentally implemented genetic circuits, a previously reported in vitro genetic circuit and a new E. coli-based transcriptional event detector.

Synthetic Cellobiose-Inducible Regulatory Systems Allow Tight and Dynamic Controls of Gene Expression in Streptomyces

Precise control of microbial gene expression is crucial for synthetic biotechnological applications. This is particularly true for the bacterial genus Streptomyces, major producers of diverse natural products including many antibiotics. Although a plethora of genetic tools have been developed for Streptomyces, there is still an urgent need for effective gene induction systems. We herein created two novel cellobiose-inducible regulatory systems referred to as Cel-RS1 and Cel-RS2. The regulatory systems are based upon the well-characterized repressor/operator pair CebR/cebO from Streptomyces scabies and the well-defined constitutive kasO* promoter. Both Cel-RS1 and Cel-RS2 exhibit a high level of induced reporter activity and virtually no leaky expression in three model Streptomyces species, which are commonly used as surrogate hosts for expression of natural product biosynthetic gene clusters. Cel-RS2 has been proven successful for programmable control of gene expression and controllable production of specialized metabolites in multiple Streptomyces species. The strategy can be used to expand the toolkit of inducible regulatory systems that will be broadly applicable to various Streptomyces.

Development of a pyrF-based counterselectable system for targeted gene deletion in Streptomyces rimosus

Streptomyces produces many valuable and important biomolecules with clinical and pharmaceutical applications. The development of simple and highly efficient gene editing tools for genetic modification of Streptomyces is highly desirable. In this study, we developed a screening system for targeted gene knockout using a uracil auxotrophic host (ΔpyrF) resistant to the highly toxic uracil analog of 5-fluoroorotic acid (5-FOA) converted by PyrF, and a non-replicative vector pKC1132-pyrF carrying the complemented pyrF gene coding for orotidine-5'-phosphate decarboxylase. The pyrF gene acts as a positive selection and counterselection marker for recombinants during genetic modifications. Single-crossover homologous integration mutants were selected on minimal medium without uracil by reintroducing pyrF along with pKC1132-pyrF into the genome of the mutant ΔpyrF at the targeted locus. Double-crossover recombinants were generated, from which the pyrF gene, plasmid backbone, and targeted gene were excised through homologous recombination exchange. These recombinants were rapidly screened by the counterselection agent, 5-FOA. We demonstrated the feasibility and advantage of using this pyrF-based screening system through deleting the otcR gene, which encodes the cluster-situated regulator that directly activates oxytetracycline biosynthesis in Streptomyces rimosus M4018. This system provides a new genetic tool for investigating the genetic characteristics of Streptomyces species.

Genetic manipulation of pathogenic Leptospira: CRISPR interference (CRISPRi)-mediated gene silencing and rapid mutant recovery at 37 °C

Leptospirosis is a neglected, widespread zoonosis caused by pathogenic species of the genus Leptospira, and is responsible for 60,000 deaths per year. Pathogenic mechanisms of leptospirosis remain poorly understood mainly because targeted mutations or gene silencing in pathogenic Leptospira continues to be inherently inefficient, laborious, costly and difficult to implement. In addition, pathogenic leptospires are highly fastidious and the selection of mutants on solid agar media can take up to 6 weeks. The catalytically inactive Cas9 (dCas9) is an RNA-guided DNA-binding protein from the Streptococcus pyogenes CRISPR/Cas system and can be used for gene silencing, in a strategy termed CRISPR interference (CRISPRi). Here, this technique was employed to silence genes encoding major outer membrane proteins of pathogenic L. interrogans. Conjugation protocols were optimized using the newly described HAN media modified for rapid mutant recovery at 37 °C in 3% CO2 within 8 days. Complete silencing of LipL32 and concomitant and complete silencing of both LigA and LigB outer membrane proteins were achieved, revealing for the first time that Lig proteins are involved in pathogenic Leptospira serum resistance. Gene silencing in pathogenic leptospires and rapid mutant recovery will facilitate novel studies to further evaluate and understand pathogenic mechanisms of leptospirosis.

Systems and synthetic biology to elucidate secondary metabolite biosynthetic gene clusters encoded in Streptomyces genomes

Covering: 2010 to 2020 Over the last few decades, Streptomyces have been extensively investigated for their ability to produce diverse bioactive secondary metabolites. Recent advances in Streptomyces research have been largely supported by improvements in high-throughput technology 'omics'. From genomics, numerous secondary metabolite biosynthetic gene clusters were predicted, increasing their genomic potential for novel bioactive compound discovery. Additional omics, including transcriptomics, translatomics, interactomics, proteomics and metabolomics, have been applied to obtain a system-level understanding spanning entire bioprocesses of Streptomyces, revealing highly interconnected and multi-layered regulatory networks for secondary metabolism. The comprehensive understanding derived from this systematic information accelerates the rational engineering of Streptomyces to enhance secondary metabolite production, integrated with the exploitation of the highly efficient 'Design-Build-Test-Learn' cycle in synthetic biology. In this review, we describe the current status of omics applications in Streptomyces research to better understand the organism and exploit its genetic potential for higher production of valuable secondary metabolites and novel secondary metabolite discovery.

Developing an endogenous quorum-sensing based CRISPRi circuit for autonomous and tunable dynamic regulation of multiple targets in Streptomyces

Quorum-sensing (QS) mediated dynamic regulation has emerged as an effective strategy for optimizing product titers in microbes. However, these QS-based circuits are often created on heterologous systems and require careful tuning via a tedious testing/optimization process. This hampers their application in industrial microbes. Here, we design a novel QS circuit by directly integrating an endogenous QS system with CRISPRi (named EQCi) in the industrial rapamycin-producing strain Streptomyces rapamycinicus. EQCi combines the advantages of both the QS system and CRISPRi to enable tunable, autonomous, and dynamic regulation of multiple targets simultaneously. Using EQCi, we separately downregulate three key nodes in essential pathways to divert metabolic flux towards rapamycin biosynthesis and significantly increase its titers. Further application of EQCi to simultaneously regulate these three key nodes with fine-tuned repression strength boosts the rapamycin titer by ∼660%, achieving the highest reported titer (1836 ± 191 mg/l). Notably, compared to static engineering strategies, which result in growth arrest and suboptimal rapamycin titers, EQCi-based regulation substantially promotes rapamycin titers without affecting cell growth, indicating that it can achieve a trade-off between essential pathways and product synthesis. Collectively, this study provides a convenient and effective strategy for strain improvement and shows potential for application in other industrial microorganisms.

Elucidation of the Activation Pathways of ScyA1/ScyR1, an Aco/ArpA-Like System That Regulates the Expression of Nemadectin and Other Secondary Metabolic Biosynthetic Genes

The quorum-sensing system, consisting of an autoregulator synthase (AfsA or Aco homolog) and an autoregulator receptor (ArpA homolog), has been reported to be universally involved in regulating secondary metabolism in streptomycetes. Although the autoregulator synthase is thought to activate antibiotic production, the activation pathway remains poorly understood. Streptomyces cyaneogriseus ssp. noncyanogenus NMWT1 produces nemadectin, which is widely used as a biopesticide and veterinary drug due to its potent nematocidal activity. Here, we identified the Aco/ArpA-like system ScyA1/ScyR1, the ArpA homolog ScyR2 and the AfsA/ArpA-like system ScyA3/ScyR3 as important regulators of nemadectin production in NMWT1. Genetic experiments revealed that these five genes positively regulate nemadectin production, with scyA1 and scyR1 having the most potent effects. Importantly, ScyA1 is an upstream regulator of scyR1 and promotes nemadectin production and sporulation by activating scyR1 transcription. Intriguingly, scyR1 silencing in NMWT1 up-regulated 12 of the 17 secondary metabolite biosynthetic core genes present in the NMWT1 genome, suggesting that ScyR1 mainly to be a repressor of secondary metabolism. In conclusion, our findings unveiled the regulatory pathways adopted by the quorum-sensing system, and provided the basis for a method to enhance antibiotic production and to activate the expression of cryptic biosynthetic gene clusters.

System-level understanding of gene expression and regulation for engineering secondary metabolite production in Streptomyces

The gram-positive bacterium, Streptomyces, is noticed for its ability to produce a wide array of pharmaceutically active compounds through secondary metabolism. To discover novel bioactive secondary metabolites and increase the production, Streptomyces species have been extensively studied for the past decades. Among the cellular components, RNA molecules play important roles as the messengers for gene expression and diverse regulations taking place at the RNA level. Thus, the analysis of RNA-level regulation is critical to understanding the regulation of Streptomyces’ metabolism and secondary metabolite production. A dramatic advance in Streptomyces research was made recently, by exploiting high-throughput technology to systematically understand RNA levels. In this review, we describe the current status of the system-wide investigation of Streptomyces in terms of RNA, toward expansion of its genetic potential for secondary metabolite synthesis.

An Update on Molecular Tools for Genetic Engineering of Actinomycetes-The Source of Important Antibiotics and Other Valuable Compounds

The first antibiotic-producing actinomycete (Streptomyces antibioticus) was described by Waksman and Woodruff in 1940. This discovery initiated the "actinomycetes era", in which several species were identified and demonstrated to be a great source of bioactive compounds. However, the remarkable group of microorganisms and their potential for the production of bioactive agents were only partially exploited. This is caused by the fact that the growth of many actinomycetes cannot be reproduced on artificial media at laboratory conditions. In addition, sequencing, genome mining and bioactivity screening disclosed that numerous biosynthetic gene clusters (BGCs), encoded in actinomycetes genomes are not expressed and thus, the respective potential products remain uncharacterized. Therefore, a lot of effort was put into the development of technologies that facilitate the access to actinomycetes genomes and activation of their biosynthetic pathways. In this review, we mainly focus on molecular tools and methods for genetic engineering of actinomycetes that have emerged in the field in the past five years (2015-2020). In addition, we highlight examples of successful application of the recently developed technologies in genetic engineering of actinomycetes for activation and/or improvement of the biosynthesis of secondary metabolites.

High efficiency CRISPR/Cas9 genome editing system with an eliminable episomal sgRNA plasmid in Pichia pastoris

Pichia pastoris is a methylotrophic yeast in which host heterologous expression of proteins has been developed owing to the strong inducible alcohol oxidase promoter (PAOX1). However, it is difficult to manipulate the genome in P. pastoris. Based on previous attempts to apply the CRISPR/Cas9 system in P. pastoris, a CRISPR/Cas9 system with episomal sgRNA plasmid was developed and 100 % genome editing efficiency, high multicopy gene editing and stable multigene editing were obtained without a sharp decline caused by multi-sgRNA. And 28/34 (∼82 %) sgRNAs tested were effective. The CGG may have a slightly higher and more stable cleavage efficiency than the other three NGG motifs, and a low GC content may be preferable for higher cleavage efficiency. This provides researchers with a stable genome editing tool that shows a high editing efficiency, shortening the experimentation period. Furthermore, we introduced dCas9 into P. pastoris and achieved target gene interference, expanding the CRISPR/Cas9 toolbox in P. pastoris.

Multiplex genome editing using a dCas9-cytidine deaminase fusion in Streptomyces

CRISPR/Cas-mediated genome editing has greatly facilitated the study of gene function in Streptomyces. However, it could not be efficiently employed in streptomycetes with low homologous recombination (HR) ability. Here, a deaminase-assisted base editor dCas9-CDA-UL_str was developed in Streptomyces, which comprises the nuclease-deficient Cas9 (dCas9), the cytidine deaminase from Petromyzon marinus (PmCDA1), the uracil DNA glycosylase inhibitor (UGI) and the protein degradation tag (LVA tag). Using dCas9-CDA-UL_str, we achieved single-, double- and triple-point mutations (cytosine-to-thymine substitutions) at target sites in Streptomyces coelicolor with efficiency up to 100%, 60% and 20%, respectively. This base editor was also demonstrated to be highly efficient for base editing in the industrial strain, Streptomyces rapamycinicus, which produces the immunosuppressive agent rapamycin. Compared with base editors derived from the cytidine deaminase rAPOBEC1, the PmCDA1-assisted base editor dCas9-CDA-UL_str could edit cytosines preceded by guanosines with high efficiency, which is a great advantage for editing Streptomyces genomes (with high GC content). Collectively, the base editor dCas9-CDA-UL_str could be employed for efficient multiplex genome editing in Streptomyces. Since the dCas9-CDA-UL_str-based genome editing is independent of HR-mediated DNA repair, we believe this technology will greatly facilitate functional genome research and metabolic engineering in Streptomyces strains with weak HR ability.

Multiplex genome editing of microorganisms using CRISPR-Cas

Microbial production of chemical compounds often requires highly engineered microbial cell factories. During the last years, CRISPR-Cas nucleases have been repurposed as powerful tools for genome editing. Here, we briefly review the most frequently used CRISPR-Cas tools and describe some of their applications. We describe the progress made with respect to CRISPR-based multiplex genome editing of industrial bacteria and eukaryotic microorganisms. We also review the state of the art in terms of gene expression regulation using CRISPRi and CRISPRa. Finally, we summarize the pillars for efficient multiplexed genome editing and present our view on future developments and applications of CRISPR-Cas tools for multiplex genome editing.

CRISPR Tools To Control Gene Expression in Bacteria

CRISPR-Cas systems have been engineered as powerful tools to control gene expression in bacteria. The most common strategy relies on the use of Cas effectors modified to bind target DNA without introducing DNA breaks. These effectors can either block the RNA polymerase or recruit it through activation domains. Here, we discuss the mechanistic details of how Cas effectors can modulate gene expression by blocking transcription initiation or acting as transcription roadblocks. CRISPR-Cas tools can be further engineered to obtain fine-tuned control of gene expression or target multiple genes simultaneously. Several caveats in using these tools have also been revealed, including off-target effects and toxicity, making it important to understand the design rules of engineered CRISPR-Cas effectors in bacteria. Alternatively, some types of CRISPR-Cas systems target RNA and could be used to block gene expression at the posttranscriptional level. Finally, we review applications of these tools in high-throughput screens and the progress and challenges in introducing CRISPR knockdown to other species, including nonmodel bacteria with industrial or clinical relevance. A deep understanding of how CRISPR-Cas systems can be harnessed to control gene expression in bacteria and build powerful tools will certainly open novel research directions.

Sensing and responding to diverse extracellular signals: an updated analysis of the sensor kinases and response regulators of Streptomyces species

Streptomyces venezuelae is a Gram-positive, filamentous actinomycete with a complex developmental life cycle. Genomic analysis revealed that S. venezuelae encodes a large number of two-component systems (TCSs): these consist of a membrane-bound sensor kinase (SK) and a cognate response regulator (RR). These proteins act together to detect and respond to diverse extracellular signals. Some of these systems have been shown to regulate antimicrobial biosynthesis in Streptomyces species, making them very attractive to researchers. The ability of S. venezuelae to sporulate in both liquid and solid cultures has made it an increasingly popular model organism in which to study these industrially and medically important bacteria. Bioinformatic analysis identified 58 TCS operons in S. venezuelae with an additional 27 orphan SK and 18 orphan RR genes. A broader approach identified 15 of the 58 encoded TCSs to be highly conserved in 93 Streptomyces species for which high-quality and complete genome sequences are available. This review attempts to unify the current work on TCS in the streptomycetes, with an emphasis on S. venezuelae.

Challenges and Advances in Genome Editing Technologies in Streptomyces

The genome of Streptomyces encodes a high number of natural product (NP) biosynthetic gene clusters (BGCs). Most of these BGCs are not expressed or are poorly expressed (commonly called silent BGCs) under traditional laboratory experimental conditions. These NP BGCs represent an unexplored rich reservoir of natural compounds, which can be used to discover novel chemical compounds. To activate silent BGCs for NP discovery, two main strategies, including the induction of BGCs expression in native hosts and heterologous expression of BGCs in surrogate Streptomyces hosts, have been adopted, which normally requires genetic manipulation. So far, various genome editing technologies have been developed, which has markedly facilitated the activation of BGCs and NP overproduction in their native hosts, as well as in heterologous Streptomyces hosts. In this review, we summarize the challenges and recent advances in genome editing tools for Streptomyces genetic manipulation with a focus on editing tools based on clustered regularly interspaced short palindrome repeat (CRISPR)/CRISPR-associated protein (Cas) systems. Additionally, we discuss the future research focus, especially the development of endogenous CRISPR/Cas-based genome editing technologies in Streptomyces.

Modulating cancer stemness provides luminal a breast cancer cells with HER2 positive-like features

Breast cancers can be classified into luminal A, luminal B, HER2 positive and triple-negative subtypes, each with a distinct therapeutic response. Tumor stemness drives cancer malignancy that challenges cancer control. Understanding the revolutionary relationships driven by tumor stemness among breast cancer subtypes is fundamental to identifying feasible therapeutic modalities for each breast cancer subtype. Utilizing the endogenous tRNA-processing system, we established a multiplexing CRISPR/dCas9 system in breast cancer cells, and applied it to a four-gene panel controlling cell potency, i.e., OCT4, KLF, MYC, SOX2. The stable cell strain, OKMS#1 was obtained through concomitantly over-expressing these genes in luminal A breast cancer cells. OKMS#1 cells showed increased invasion, proliferation and cancer stemness, shared similar drug response pattern with HER2 positive cells, and exhibited altered MAPK and enhanced NFkB signaling. This study contributes in providing an efficient multiplexing CRISPR/dCas9 system that enriches our genetic modulation tool box, and suggests that HER2 positive cells are potential progenitors of luminal A cells and that these two breast cancer subtypes may share similar treatment strategies once rewired between the two states. Our results also implicate that triple negative breast cancer cells, though sharing similar cancer stemness with HER2 positive cells, represent a distinct type of disease and require unique treatment solutions.

Establishment and application of multiplexed CRISPR interference system in Bacillus licheniformis

Bacillus licheniformis has been regarded as an outstanding microbial cell factory for the production of biochemicals and enzymes. Due to lack of genetic tools to repress gene expression, metabolic engineering and gene function elucidation are limited in this microbe. In this study, an integrated CRISPR interference (CRISPRi) system was constructed in B. licheniformis. Several endogenous genes, including yvmC, cypX, alsD, pta, ldh, and essential gene rpsC, were severed as the targets to test this CRISPRi system, and the repression efficiencies were ranged from 45.02 to 94.00%. Moreover, the multiple genes were simultaneously repressed with high efficiency using this CRISPRi system. As a case study, the genes involved in by-product synthetic and L-valine degradation pathways were selected as the silence targets to redivert metabolic flux toward L-valine synthesis. Repression of acetolactate decarboxylase (alsD) and leucine dehydrogenase (bcd) led to 90.48% and 80.09 % increases in L-valine titer, respectively. Compared with the control strain DW9i△leuA (1.47 g/L and 1.79 g/L), the L-valine titers of combinatorial strain DW9i△leuA/pHYi-alsD-bcd were increased by 1.27-fold and 2.89-fold, respectively, in flask and bioreactor. Collectively, this work provides a feasible approach for multiplex metabolic engineering and functional genome studies of B. licheniformis.

Enhancement of FK520 production in Streptomyces hygroscopicus by combining traditional mutagenesis with metabolic engineering

FK520 (ascomycin), a 23-membered macrolide with immunosuppressive activity, is produced by Streptomyces hygroscopicus. The problem of low yield and high impurities (mainly FK523) limits the industrialized production of FK520. In this study, the FK520 yield was significantly improved by strain mutagenesis and genetic engineering. First, a FK520 high-producing strain SFK-6-33 (2432.2 mg/L) was obtained from SFK-36 (1588.4 mg/L) through ultraviolet radiation mutation coupled with streptomycin resistance screening. The endogenous crotonyl-CoA carboxylase/reductase (FkbS) was found to play an important role in FK520 biosynthesis, identified with CRISPR/dCas9 inhibition system. FkbS was overexpressed in SFK-6-33 to obtain the engineered strain SFK-OfkbS, which produced 2817.0 mg/L of FK520 resulting from an increase in intracellular ethylmalonyl-CoA levels. In addition, the FK520 levels could be further increased with supplementation of crotonic acid in SFK-OfkbS. Overexpression of acetyl-CoA carboxylase (ACCase), used for the synthesis of malonyl-CoA, was also investigated in SFK-6-33, which improved the FK520 yield to 3320.1 mg/L but showed no significant inhibition in FK523 production. To further enhance FK520 production, FkbS and ACCase combinatorial overexpression strain SFK-OASN was constructed; the FK520 production increased by 44.4% to 3511.4 mg/L, and the FK523/FK520 ratio was reduced from 9.6 to 5.6% compared with that in SFK-6-33. Finally, a fed-batch culture was carried out in a 5-L fermenter, and the FK520 yield reached 3913.9 mg/L at 168 h by feeding glycerol, representing the highest FK520 yield reported thus far. These results demonstrated that traditional mutagenesis combined with metabolic engineering was an effective strategy to improve FK520 production.

Simultaneous repression of multiple bacterial genes using nonrepetitive extra-long sgRNA arrays

Engineering cellular phenotypes often requires the regulation of many genes. When using CRISPR interference, coexpressing many single-guide RNAs (sgRNAs) triggers genetic instability and phenotype loss, due to the presence of repetitive DNA sequences. We stably coexpressed 22 sgRNAs within nonrepetitive extra-long sgRNA arrays (ELSAs) to simultaneously repress up to 13 genes by up to 3,500-fold. We applied biophysical modeling, biochemical characterization and machine learning to develop toolboxes of nonrepetitive genetic parts, including 28 sgRNA handles that bind Cas9. We designed ELSAs by combining nonrepetitive genetic parts according to algorithmic rules quantifying DNA synthesis complexity, sgRNA expression, sgRNA targeting and genetic stability. Using ELSAs, we created three highly selective phenotypes in Escherichia coli, including redirecting metabolism to increase succinic acid production by 150-fold, knocking down amino acid biosynthesis to create a multi-auxotrophic strain and repressing stress responses to reduce persister cell formation by 21-fold. ELSAs enable simultaneous and stable regulation of many genes for metabolic engineering and synthetic biology applications.

Characterization of Cas proteins for CRISPR-Cas editing in streptomycetes

Application of the well-characterized Streptococcus pyogenes CRISPR-Cas9 system in actinomycetes streptomycetes has enabled high-efficiency multiplex genome editing and CRISPRi-mediated transcriptional regulation in these prolific bioactive metabolite producers. Nonetheless, SpCas9 has its limitations and can be ineffective depending on the strains and target sites. Here, we built and tested alternative CRISPR-Cas constructs based on the standalone pCRISPomyces-2 editing plasmid. We showed that Streptococcus thermophilus CRISPR1 Cas9 (sth1Cas9), Staphylococcus aureus Cas9 (saCas9), and Francisella tularensis subsp. novicida U112 Cpf1 (fnCpf1) are functional in multiple streptomycetes, enabling efficient homology-directed repair-mediated knock-in and deletion. In strains where spCas9 was nonfunctional, these alternative Cas systems enabled precise genomic modifications within biosynthetic gene clusters for the discovery, production, and diversification of natural products. These additional Cas proteins provide us with the versatility to overcome the limitations of individual CRISPR-Cas systems for genome editing and transcriptional regulation of these industrially important bacteria.

A CRISPR-Cas9 Strategy for Activating the Saccharopolyspora erythraea Erythromycin Biosynthetic Gene Cluster with Knock-in Bidirectional Promoters

The regulation of biosynthetic pathways is a universal strategy for industrial strains that overproduce metabolites. Erythromycin produced by Saccharopolyspora erythraea has extensive clinical applications. In this study, promoters of the erythromycin biosynthesis gene cluster were tested by reporter mCherry. The SACE_0720 ( eryBIV)-SACE_0721 ( eryAI) spacer was selected as a target regulatory region, and bidirectional promoters with dual single guide RNAs (sgRNAs) were knocked-in using the clustered regularly interspaced short palindromic repeats (CRISPR)-Cas9 method. qPCR results indicated that knock-in of Pj23119-PkasO, which replaced the native promoter, enabled biosynthetic gene cluster activation, with eryBIV and eryAI expression increased 32 and 79 times, respectively. High performance liquid chromatography results showed that, compared with the wild-type strain, the yield of erythromycin was increased (58.3%) in bidirectional promoter knock-in recombinant strains. On the basis of the activated strain Ab::Pj23119-PkasO, further investigation showed that CRISPR-based interference of sdhA gene affected erythromycin biosynthesis and cell growth. Finally, regulating the culture temperature to optimize the inhibition intensity of sdhA further increased the yield by 15.1%. In summary, this study showed that bidirectional promoter knock-in and CRISPR interference could regulate gene expression in S. erythraea. This strategy has potential application for biosynthetic gene cluster activation and gene regulation in Actinobacteria.

Recent advances in natural products exploitation in Streptomyces via synthetic biology

Natural products of microbial origin have proven to be the wellspring of clinically useful compounds for human therapeutics. Streptomyces species are predominant sources of bioactive compounds, most of which serve as potential drug candidates. While the exploitation of natural products has been severely reduced over the past two decades, the growing crisis of evolution and dissemination of drug resistant pathogens have again attracted great interest in this field. The emerging synthetic biology has been heralded as a new bioengineering platform to discover novel bioactive compounds and expand bioactive natural products diversity and production. Herein, we review recent advances in the natural products exploitation of Streptomyces with the applications of synthetic biology from three major aspects, including recently developed synthetic biology tools, natural products biosynthetic pathway engineering strategies as well as chassis host modifications.

Gene silencing based on RNA-guided catalytically inactive Cas9 (dCas9): a new tool for genetic engineering in Leptospira

Leptospirosis is a worldwide zoonosis caused by pathogenic bacteria of the genus Leptospira, which also includes free-living saprophyte strains. Many aspects of leptospiral basic biology and virulence mechanisms remain unexplored mainly due to the lack of effective genetic tools available for these bacteria. Recently, the type II CRISPR/Cas system from Streptococcus pyogenes has been widely used as an efficient genome engineering tool in bacteria by inducing double-strand breaks (DSBs) in the desired genomic targets caused by an RNA-guided DNA endonuclease called Cas9, and the DSB repair associated machinery. In the present work, plasmids expressing heterologous S. pyogenes Cas9 in L. biflexa cells were generated, and the enzyme could be expressed with no apparent toxicity to leptospiral cells. However, L. biflexa cells were unable to repair RNA-guided Cas9-induced DSBs. Thus, we used a catalytically dead Cas9 (dCas9) to obtain gene silencing rather than disruption, in a strategy called CRISPR interference (CRISPRi). We demonstrated complete gene silencing in L. biflexa cells when both dCas9 and single-guide RNA (sgRNA) targeting the coding strand of the β-galactosidase gene were expressed simultaneously. Furthermore, when the system was applied for silencing the dnaK gene, no colonies were recovered, indicating that DnaK protein is essential in Leptospira. In addition, flagellar motor switch FliG gene silencing resulted in reduced bacterial motility. To the best of our knowledge, this is the first work applying the CRISPRi system in Leptospira and spirochetes in general, expanding the tools available for understanding leptospiral biology.