Application of TALEs, CRISPR/Cas and sRNAs as trans-acting regulators in prokaryotes

Guide RNA structure design enables combinatorial CRISPRa programs for biosynthetic profiling

Engineering metabolism to efficiently produce chemicals from multi-step pathways requires optimizing multi-gene expression programs to achieve enzyme balance. CRISPR-Cas transcriptional control systems are emerging as important tools for programming multi-gene expression, but poor predictability of guide RNA folding can disrupt expression control. Here, we correlate efficacy of modified guide RNAs (scRNAs) for CRISPR activation (CRISPRa) in E. coli with a computational kinetic parameter describing scRNA folding rate into the active structure (rS = 0.8). This parameter also enables forward design of scRNAs, allowing us to design a system of three synthetic CRISPRa promoters that can orthogonally activate (>35-fold) expression of chosen outputs. Through combinatorial activation tuning, we profile a three-dimensional design space expressing two different biosynthetic pathways, demonstrating variable production of pteridine and human milk oligosaccharide products. This RNA design approach aids combinatorial optimization of metabolic pathways and may accelerate routine design of effective multi-gene regulation programs in bacterial hosts.

CRISPR/Cas9-Based Genome Editing and Its Application in Aspergillus Species

Aspergillus, a genus of filamentous fungi, is extensively distributed in nature and plays crucial roles in the decomposition of organic materials as an important environmental microorganism as well as in the traditional fermentation and food processing industries. Furthermore, due to their strong potential to secrete a large variety of hydrolytic enzymes and other natural products by manipulating gene expression and/or introducing new biosynthetic pathways, several Aspergillus species have been widely exploited as microbial cell factories. In recent years, with the development of next-generation genome sequencing technology and genetic engineering methods, the production and utilization of various homo-/heterologous-proteins and natural products in Aspergillus species have been well studied. As a newly developed genome editing technology, the clustered regularly interspaced short palindromic repeats/CRISPR-associated protein 9 (CRISPR/Cas9) system has been used to edit and modify genes in Aspergilli. So far, the CRISPR/Cas9-based approach has been widely employed to improve the efficiency of gene modification in the strain type Aspergillus nidulans and other industrially important and pathogenic Aspergillus species, including Aspergillus oryzae, Aspergillus niger, and Aspergillus fumigatus. This review highlights the current development of CRISPR/Cas9-based genome editing technology and its application in basic research and the production of recombination proteins and natural products in the Aspergillus species.

Carrier strategies boost the application of CRISPR/Cas system in gene therapy

Emerging clustered regularly interspaced short palindromic repeat/associated protein (CRISPR/Cas) genome editing technology shows great potential in gene therapy. However, proteins and nucleic acids suffer from enzymatic degradation in the physiological environment and low permeability into cells. Exploiting carriers to protect the CRISPR system from degradation, enhance its targeting of specific tissues and cells, and reduce its immunogenicity is essential to stimulate its clinical applications. Here, the authors review the state-of-the-art CRISPR delivery systems and their applications, and describe strategies to improve the safety and efficacy of CRISPR mediated genome editing, categorized by three types of cargo formats, that is, Cas: single-guide RNA ribonucleoprotein, Cas mRNA and single-guide RNA, and Cas plasmid expressing CRISPR/Cas systems. The authors hope this review will help develop safe and efficient nanomaterial-based carriers for CRISPR tools.

Efficient genome editing in filamentous fungi via an improved CRISPR-Cas9 ribonucleoprotein method facilitated by chemical reagents

DNA double-strand break (DSB) repair induced by the RNA-programmed nuclease Cas9 has become a popular method for genome editing. Direct genome editing via Cas9-CRISPR gRNA (guide RNA) ribonucleoprotein (RNP) complexes assembled in vitro has also been successful in some fungi. However, the efficiency of direct RNP transformation into fungal protoplasts is currently too low. Here, we report an optimized genome editing approach for filamentous fungi based on RNPs facilitated by adding chemical reagents. We increased the transformation efficiency of RNPs significantly by adding Triton X-100 and prolonging the incubation time, and the editing efficiency reached 100% in Trichoderma reesei and Cordyceps militaris. The optimized RNP-based method also achieved efficient (56.52%) homologous recombination integration with short homology arms (20 bp) and gene disruption (7.37%) that excludes any foreign DNA (selection marker) in T. reesei. In particular, after adding reagents related to mitosis and cell division, the further optimized protocol showed an increased ratio of edited homokaryotic transformants (from 0% to 40.0% for inositol and 71.43% for benomyl) from Aspergillus oryzae, which contains multinucleate spores and protoplasts. Furthermore, the multi-target engineering efficiency of the optimized RNP transformation method was similar to those of methods based on in vivo expression of Cas9. This newly established genome editing system based on RNPs may be widely applicable to construction of genome-edited fungi for the food and medical industries, and has good prospects for commercialization.

Post-Transcriptional Control in the Regulation of Polyhydroxyalkanoates Synthesis

The large production of non-degradable petrol-based plastics has become a major global issue due to its environmental pollution. Biopolymers produced by microorganisms such as polyhydroxyalkanoates (PHAs) are gaining potential as a sustainable alternative, but the high cost associated with their industrial production has been a limiting factor. Post-transcriptional regulation is a key step to control gene expression in changing environments and has been reported to play a major role in numerous cellular processes. However, limited reports are available concerning the regulation of PHA accumulation in bacteria, and many essential regulatory factors still need to be identified. Here, we review studies where the synthesis of PHA has been reported to be regulated at the post-transcriptional level, and we analyze the RNA-mediated networks involved. Finally, we discuss the forthcoming research on riboregulation, synthetic, and metabolic engineering which could lead to improved strategies for PHAs synthesis in industrial production, thereby reducing the costs currently associated with this procedure.

Tailor-made sRNAs: a plasmid tool to control the expression of target mRNAs in Pseudomonas putida

Pseudomonas putida is a highly attractive production system for industrial needs. However, for its improvement as a biocatalyst at the industrial level, modulation of its gene expression is urgently needed. We report the construction of a plasmid expressing a small RNA-based system with the potential to be used for different purposes. Due to the small RNAs modular composition, the design facilities and ability to tune gene expression, they constitute a powerful tool in genetic and metabolic engineering. In the tool presented here, customized sRNAs are expressed from a plasmid and specifically directed to any region of a chosen target. Expression of these customized sRNAs is shown to differentially modulate the level of endogenous and heterologous reporter genes. The antisense interaction of the sRNA with the mRNA produces different outcomes. Depending on the particularity of each sRNA-target mRNA pair, we demonstrate the duality of this system, which is able either to decrease or increase the expression of the same given gene. This system combines high specificity with the potential to be widely applied, due to its predicted ability to modulate the expression of virtually any given gene. This plasmid can be used to redesign P. putida metabolism, fulfilling an important industrial gap.

Converting a Periplasmic Binding Protein into a Synthetic Biosensing Switch through Domain Insertion

All biosensing platforms rest on two pillars: specific biochemical recognition of a particular analyte and transduction of that recognition into a readily detectable signal. Most existing biosensing technologies utilize proteins that passively bind to their analytes and therefore require wasteful washing steps, specialized reagents, and expensive instruments for detection. To overcome these limitations, protein engineering strategies have been applied to develop new classes of protein-based sensor/actuators, known as protein switches, responding to small molecules. Protein switches change their active state (output) in response to a binding event or physical signal (input) and therefore show a tremendous potential to work as a biosensor. Synthetic protein switches can be created by the fusion between two genes, one coding for a sensor protein (input domain) and the other coding for an actuator protein (output domain) by domain insertion. The binding of a signal molecule to the engineered protein will switch the protein function from an “off” to an “on” state (or vice versa) as desired. The molecular switch could, for example, sense the presence of a metabolite, pollutant, or a biomarker and trigger a cellular response. The potential sensing and response capabilities are enormous; however, the recognition repertoire of natural switches is limited. Thereby, bioengineers have been struggling to expand the toolkit of molecular switches recognition repertoire utilizing periplasmic binding proteins (PBPs) as protein-sensing components. PBPs are a superfamily of bacterial proteins that provide interesting features to engineer biosensors, for instance, immense ligand-binding diversity and high affinity, and undergo large conformational changes in response to ligand binding. The development of these protein switches has yielded insights into the design of protein-based biosensors, particularly in the area of allosteric domain fusions. Here, recent protein engineering approaches for expanding the versatility of protein switches are reviewed, with an emphasis on studies that used PBPs to generate novel switches through protein domain insertion.

Protein Engineering Strategies to Expand CRISPR-Cas9 Applications

The development of precise and modulated methods for customized manipulation of DNA is an important objective for the study and engineering of biological processes and is essential for the optimization of gene therapy, metabolic flux, and synthetic gene networks. The clustered regularly interspaced short palindromic repeat- (CRISPR-) associated protein 9 is an RNA-guided site-specific DNA-binding complex that can be reprogrammed to specifically interact with a desired DNA sequence target. CRISPR-Cas9 has been used in a wide variety of applications ranging from basic science to the clinic, such as gene therapy, gene regulation, modifying epigenomes, and imaging chromosomes. Although Cas9 has been successfully used as a precise tool in all these applications, some limitations have also been reported, for instance (i) a strict dependence on a protospacer-adjacent motif (PAM) sequence, (ii) aberrant off-target activity, (iii) the large size of Cas9 is problematic for CRISPR delivery, and (iv) lack of modulation of protein binding and endonuclease activity, which is crucial for precise spatiotemporal control of gene expression or genome editing. These obstacles hinder the use of CRISPR for disease treatment and in wider biotechnological applications. Protein-engineering approaches offer solutions to overcome the limitations of Cas9 and generate robust and efficient tools for customized DNA manipulation. Here, recent protein-engineering approaches for expanding the versatility of the Streptococcus pyogenes Cas9 (SpCas9) is reviewed, with an emphasis on studies that improve or develop novel protein functions through domain fusion or splitting, rational design, and directed evolution.

Recent advances in synthetic biology of cyanobacteria

Cyanobacteria are attractive hosts that can be engineered for the photosynthetic production of fuels, fine chemicals, and proteins from CO₂. Moreover, the responsiveness of these photoautotrophs towards different environmental signals, such as light, CO₂, diurnal cycle, and metals make them potential hosts for the development of biosensors. However, engineering these hosts proves to be a challenging and lengthy process. Synthetic biology can make the process of biological engineering more predictable through the use of standardized biological parts that are well characterized and tools to assemble them. While significant progress has been made with model heterotrophic organisms, many of the parts and tools are not portable in cyanobacteria. Therefore, efforts are underway to develop and characterize parts derived from cyanobacteria. In this review, we discuss the reported parts and tools with the objective to develop cyanobacteria as cell factories or biosensors. We also discuss the issues related to characterization, tunability, portability, and the need to develop enabling technologies to engineer this "green" chassis.

Transcription control engineering and applications in synthetic biology

In synthetic biology, researchers assemble biological components in new ways to produce systems with practical applications. One of these practical applications is control of the flow of genetic information (from nucleic acid to protein), a.k.a. gene regulation. Regulation is critical for optimizing protein (and therefore activity) levels and the subsequent levels of metabolites and other cellular properties. The central dogma of molecular biology posits that information flow commences with transcription, and accordingly, regulatory tools targeting transcription have received the most attention in synthetic biology. In this mini-review, we highlight many past successes and summarize the lessons learned in developing tools for controlling transcription. In particular, we focus on engineering studies where promoters and transcription terminators (cis-factors) were directly engineered and/or isolated from DNA libraries. We also review several well-characterized transcription regulators (trans-factors), giving examples of how cis- and trans-acting factors have been combined to create digital and analogue switches for regulating transcription in response to various signals. Last, we provide examples of how engineered transcription control systems have been used in metabolic engineering and more complicated genetic circuits. While most of our mini-review focuses on the well-characterized bacterium Escherichia coli, we also provide several examples of the use of transcription control engineering in non-model organisms. Similar approaches have been applied outside the bacterial kingdom indicating that the lessons learned from bacterial studies may be generalized for other organisms.

Engineering Modular Biosensors to Confer Metabolite-Responsive Regulation of Transcription

Efforts to engineer microbial factories have benefitted from mining biological diversity and high throughput synthesis of novel enzymatic pathways, yet screening and optimizing metabolic pathways remain rate-limiting steps. Metabolite-responsive biosensors may help to address these persistent challenges by enabling the monitoring of metabolite levels in individual cells and metabolite-responsive feedback control. We are currently limited to naturally evolved biosensors, which are insufficient for monitoring many metabolites of interest. Thus, a method for engineering novel biosensors would be powerful, yet we lack a generalizable approach that enables the construction of a wide range of biosensors. As a step toward this goal, we here explore several strategies for converting a metabolite-binding protein into a metabolite-responsive transcriptional regulator. By pairing a modular protein design approach with a library of synthetic promoters and applying robust statistical analyses, we identified strategies for engineering biosensor-regulated bacterial promoters and for achieving design-driven improvements of biosensor performance. We demonstrated the feasibility of this strategy by fusing a programmable DNA binding motif (zinc finger module) with a model ligand binding protein (maltose binding protein), to generate a novel biosensor conferring maltose-regulated gene expression. This systematic investigation provides insights that may guide the development of additional novel biosensors for diverse synthetic biology applications.

CRISPR/Cas9-Based Efficient Genome Editing in Clostridium ljungdahlii, an Autotrophic Gas-Fermenting Bacterium

Acetogenic bacteria have the potential to convert single carbon gases (CO and CO2) into a range of bulk chemicals and fuels. Realization of their full potential is being impeded by the absence of effective genetic tools for high throughput genome modification. Here we report the development of a highly efficient CRISPR/Cas9 system for rapid genome editing of Clostridium ljungdahlii, a paradigm for the commercial production of ethanol from synthesis gas. Following the experimental selection of two promoters (Pthl and ParaE) for expression of cas9 and the requisite single guide RNA (sgRNA), the efficiency of system was tested by making precise deletions of four genes, pta, adhE1, ctf and pyrE. Deletion efficiencies were 100%, >75%, 100% and >50%, respectively. The system overcomes the deficiencies of currently available tools (more rapid, no added antibiotic resistance gene, scarless and minimal polar effects) and will find utility in other acetogens, including the pathogen Clostridium difficile.

Development of Design Rules for Reliable Antisense RNA Behavior in E. coli

A key driver of synthetic biology is the development of designable genetic parts with predictable behaviors that can be quickly implemented in complex genetic systems. However, the intrinsic complexity of gene regulation can make the rational design of genetic parts challenging. This challenge is apparent in the design of antisense RNA (asRNA) regulators. Though asRNAs are well-known regulators, the literature governing their design is conflicting and leaves the synthetic biology community without clear asRNA design rules. The goal of this study is to perform a comprehensive experimental characterization and statistical analysis of 121 unique asRNA regulators in order to resolve the conflicts that currently exist in the literature. asRNAs usually consist of two regions, the Hfq binding site and the target binding region (TBR). First, the behaviors of several high-performing Hfq binding sites were compared, in terms of their ability to improve repression efficiencies and their orthogonality. Next, a large-scale analysis of TBR design parameters identified asRNA length, the thermodynamics of asRNA-mRNA complex formation, and the percent of target mismatch as key parameters for TBR design. These parameters were used to develop simple asRNA design rules. Finally, these design rules were applied to construct both a simple and a complex genetic circuit containing different asRNAs, and predictable behavior was observed in both circuits. The results presented in this study will drive synthetic biology forward by providing useful design guidelines for the construction of asRNA regulators with predictable behaviors.

Extending CRISPR-Cas9 Technology from Genome Editing to Transcriptional Engineering in the Genus Clostridium

The discovery and exploitation of the prokaryotic adaptive immunity system based on clustered regularly interspaced short palindromic repeats (CRISPRs) and CRISPR-associated (Cas) proteins have revolutionized genetic engineering. CRISPR-Cas tools have enabled extensive genome editing as well as efficient modulation of the transcriptional program in a multitude of organisms. Progress in the development of genetic engineering tools for the genus Clostridium has lagged behind that of many other prokaryotes, presenting the CRISPR-Cas technology an opportunity to resolve a long-existing issue. Here, we applied the Streptococcus pyogenes type II CRISPR-Cas9 (SpCRISPR-Cas9) system for genome editing in Clostridium acetobutylicum DSM792. We further explored the utility of the SpCRISPR-Cas9 machinery for gene-specific transcriptional repression. For proof-of-concept demonstration, a plasmid-encoded fluorescent protein gene was used for transcriptional repression in C. acetobutylicum Subsequently, we targeted the carbon catabolite repression (CCR) system of C. acetobutylicum through transcriptional repression of the hprK gene encoding HPr kinase/phosphorylase, leading to the coutilization of glucose and xylose, which are two abundant carbon sources from lignocellulosic feedstocks. Similar approaches based on SpCRISPR-Cas9 for genome editing and transcriptional repression were also demonstrated in Clostridium pasteurianum ATCC 6013. As such, this work lays a foundation for the derivation of clostridial strains for industrial purposes.

IMPORTANCE

After recognizing the industrial potential of Clostridium for decades, methods for the genetic manipulation of these anaerobic bacteria are still underdeveloped. This study reports the implementation of CRISPR-Cas technology for genome editing and transcriptional regulation in Clostridium acetobutylicum, which is arguably the most common industrial clostridial strain. The developed genetic tools enable simpler, more reliable, and more extensive derivation of C. acetobutylicum mutant strains for industrial purposes. Similar approaches were also demonstrated in Clostridium pasteurianum, another clostridial strain that is capable of utilizing glycerol as the carbon source for butanol fermentation, and therefore can be arguably applied in other clostridial strains.

CRISPR interference as a titratable, trans-acting regulatory tool for metabolic engineering in the cyanobacterium Synechococcus sp. strain PCC 7002

Trans-acting regulators provide novel opportunities to study essential genes and regulate metabolic pathways. We have adapted the clustered regularly interspersed palindromic repeats (CRISPR) system from Streptococcus pyogenes to repress genes in trans in the cyanobacterium Synechococcus sp. strain PCC 7002 (hereafter PCC 7002). With this approach, termed CRISPR interference (CRISPRi), transcription of a specific target sequence is repressed by a catalytically inactive Cas9 protein recruited to the target DNA by base-pair interactions with a single guide RNA that is complementary to the target sequence. We adapted this system for PCC 7002 and achieved conditional and titratable repression of a heterologous reporter gene, yellow fluorescent protein. Next, we demonstrated the utility of finely tuning native gene expression by downregulating the abundance of phycobillisomes. In addition, we created a conditional auxotroph by repressing synthesis of the carboxysome, an essential component of the carbon concentrating mechanism cyanobacteria use to fix atmospheric CO₂. Lastly, we demonstrated a novel strategy for increasing central carbon flux by conditionally downregulating a key node in nitrogen assimilation. The resulting cells produced 2-fold more lactate than a baseline engineered cell line, representing the highest photosynthetically generated productivity to date. This work is the first example of titratable repression in cyanobacteria using CRISPRi, enabling dynamic regulation of essential processes and manipulation of flux through central carbon metabolism. This tool facilitates the study of essential genes of unknown function and enables groundbreaking metabolic engineering capability, by providing a straightforward approach to redirect metabolism and carbon flux in the production of high-value chemicals.

Tobacco Etch Virus protease: A shortcut across biotechnologies

About thirty years ago, studies on the RNA genome of Tobacco Etch Virus revealed the presence of an efficient and specific protease, called Tobacco Etch Virus protease (TEVp), that was part of the Nuclear Inclusion a (NIa) enzyme. TEVp is an efficient and specific protease of 27kDa that has become a valuable biotechnological tool. Nowadays TEVp is a unique endopeptidase largely exploited in biotechnology from industrial applications to in vitro and in vivo cellular studies. A number of TEVp mutants with different rate of cleavage, stability and specificity have been reported. Similarly, a panel of different target cleavage sites, derived from the canonical ENLYFQ-G/S site, has been established. In this review we describe these aspects of TEVp and some of its multiple applications. A particular focus is on the use and molecular biology of TEVp in living cells and organisms.

CRISPR-on system for the activation of the endogenous human INS gene

Advances in the field of epigenetics have allowed the design of new therapeutic strategies to address complex diseases such as type 1 diabetes (T1D). Clustered regularly interspaced short palindromic repeats (CRISPR)-on is a novel and powerful RNA-guided transcriptional activator system that can turn on specific gene expression; however, it remains unclear whether this system can be widely used or whether its use will be restricted depending on cell types, methylation promoter statuses or the capacity to modulate chromatin state. Our results revealed that the CRISPR-on system fused with transcriptional activators (dCas9-VP160) activated endogenous human INS, which is a silenced gene with a fully methylated promoter. Similarly, we observed a synergistic effect on gene activation when multiple single guide RNAs were used, and the transcriptional activation was maintained until day 21. Regarding the epigenetic profile, the targeted promoter gene did not exhibit alteration in its methylation status but rather exhibited altered levels of H3K9ac following treatment. Importantly, we showed that dCas9-VP160 acts on patients' cells in vitro, particularly the fibroblasts of patients with T1D.

Programmable control of bacterial gene expression with the combined CRISPR and antisense RNA system

A central goal of synthetic biology is to implement diverse cellular functions by predictably controlling gene expression. Though research has focused more on protein regulators than RNA regulators, recent advances in our understanding of RNA folding and functions have motivated the use of RNA regulators. RNA regulators provide an advantage because they are easier to design and engineer than protein regulators, potentially have a lower burden on the cell and are highly orthogonal. Here, we combine the CRISPR system from Streptococcus pyogenes and synthetic antisense RNAs (asRNAs) in Escherichia coli strains to repress or derepress a target gene in a programmable manner. Specifically, we demonstrate for the first time that the gene target repressed by the CRISPR system can be derepressed by expressing an asRNA that sequesters a small guide RNA (sgRNA). Furthermore, we demonstrate that tunable levels of derepression can be achieved (up to 95%) by designing asRNAs that target different regions of a sgRNA and by altering the hybridization free energy of the sgRNA-asRNA complex. This new system, which we call the combined CRISPR and asRNA system, can be used to reversibly repress or derepress multiple target genes simultaneously, allowing for rational reprogramming of cellular functions.

A transcription activator-like effector (TALE) induction system mediated by proteolysis

Simple and predictable trans-acting regulatory tools are needed in the fields of synthetic biology and metabolic engineering to build complex genetic circuits and optimize the levels of native and heterologous gene products. Transcription activator-like effectors (TALEs) are bacterial virulence factors that have recently gained traction in biotechnology applications due to their customizable DNA binding specificity. In this work we expand the versatility of these transcription factors to create an inducible TALE system by inserting tobacco-etch virus (TEV) protease recognition sites into the TALE backbone. The resulting engineered TALEs maintain transcriptional repression of their target genes in Escherichia coli, but are degraded following the induction of the TEV protease, thereby promoting expression of the previously repressed target gene of interest. This TALE-TEV technology enables both repression and induction of plasmid or chromosomal target genes in a manner analogous to traditional repressor proteins but with the added flexibility of being operator agnostic.

On the road to synthetic life: the minimal cell and genome-scale engineering

Synthetic biology employs rational engineering principles to build biological systems from the libraries of standard, well characterized biological parts. Biological systems designed and built by synthetic biologists fulfill a plethora of useful purposes, ranging from better healthcare and energy production to biomanufacturing. Recent advancements in the synthesis, assembly and "booting-up" of synthetic genomes and in low and high-throughput genome engineering have paved the way for engineering on the genome-wide scale. One of the key goals of genome engineering is the construction of minimal genomes consisting solely of essential genes (genes indispensable for survival of living organisms). Besides serving as a toolbox to understand the universal principles of life, the cell encoded by minimal genome could be used to build a stringently controlled "cell factory" with a desired phenotype. This review provides an update on recent advances in the genome-scale engineering with particular emphasis on the engineering of minimal genomes. Furthermore, it presents an ongoing discussion to the scientific community for better suitability of minimal or robust cells for industrial applications.