Editing the Genome Without Double-Stranded DNA Breaks

Current progress of genome editing in livestock

Historically, genetic engineering in livestock proved to be challenging. Without stable embryonic stem cell lines to utilize, somatic cell nuclear transfer (SCNT) had to be employed to produce many of the genetically engineered (GE) livestock models. Through the genetic engineering of somatic cells followed by SCNT, GE livestock models could be generated carrying site-specific modifications. Although successful, only a few GE livestock models were generated because of low efficiency and associated birth defects. Recently, there have been major strides in the development of genome editing tools: Zinc-Finger Nucleases (ZFNs), Transcription activator-like effector nucleases (TALENS), and Clustered regularly interspaced short palindromic repeats (CRISPR) and CRISPR-associated 9 (Cas9) system. These tools rely on the generation of a double strand DNA break, followed by one of two repair pathways: non-homologous end joining (NHEJ) or homology directed repair (HDR). Compared to the traditional approaches, these tools dramatically reduce time and effort needed to establish a GE animal. Another benefit of utilizing genome editing tools is the application of direct injection into developing embryos to induce targeted mutations, therefore, eliminating side effects associated with SCNT. Emerging technological advancements of genome editing systems have dramatically improved efficiency to generate GE livestock models for both biomedical and agricultural purposes. Although the efficiency of genome editing tools has revolutionized GE livestock production, improvements for safe and consistent application are desired. This review will provide an overview of genome editing techniques, as well as examples of GE livestock models for agricultural and biomedical purposes.

Cytosine base editor 4 but not adenine base editor generates off-target mutations in mouse embryos

Deaminase base editing has emerged as a tool to install or correct point mutations in the genomes of living cells in a wide range of organisms. However, the genome-wide off-target effects introduced by base editors in the mammalian genome have been examined in only one study. Here, we have investigated the fidelity of cytosine base editor 4 (BE4) and adenine base editors (ABE) in mouse embryos using unbiased whole-genome sequencing of a family-based trio cohort. The same sgRNA was used for BE4 and ABE. We demonstrate that BE4-edited mice carry an excess of single-nucleotide variants and deletions compared to ABE-edited mice and controls. Therefore, an optimization of cytosine base editors is required to improve its fidelity. While the remarkable fidelity of ABE has implications for a wide range of applications, the occurrence of rare aberrant C-to-T conversions at specific target sites needs to be addressed.

THE GORDON WILSON LECTURE: THE ETHICS OF HUMAN GENOME EDITING

Human genome editing has undergone major technological advances, raising the possibility of treating or preventing many illnesses. Somatic (nonheritable) genome editing, both in vitro and in vivo, is already being employed under a robust regulatory and ethical framework developed for human gene therapy. In contrast, the prospect of germline (heritable) genome editing is much more contentious, and there is currently no consensus on the proper path forward. The 2017 National Academy of Sciences (NAS) and National Academy of Medicine (NAM) report proposed a series of requirements designed to minimize ethical objections while allowing couples to accept the risks of genome editing in order to have a biologically related child without passing on a known genetic disorder. It is vital to prevent gene editing from resulting in unintended negative consequences for individuals with genetic variants. The utilization of genome editing to enhance human function is highly contentious; it may be better to focus on whether an edit creates an "unfair advantage" rather than an enhancement.

Correction of the aprt Gene Using Repair-Polypurine Reverse Hoogsteen Hairpins in Mammalian Cells

In this study, we describe the correction of single-point mutations in mammalian cells by repair-polypurine reverse Hoogsteen hairpins (repair-PPRHs). These molecules consist of (1) a PPRH hairpin core that binds to a polypyrimidine target sequence in the double-stranded DNA (dsDNA), producing a triplex structure, and (2) an extension sequence homologous to the DNA sequence to be repaired but containing the wild-type nucleotide instead of the mutation and acting as a donor DNA to correct the mutation. We repaired different point mutations in the adenosyl phosphoribosyl transferase (aprt) gene contained in different aprt-deficient Chinese hamster ovary (CHO) cell lines. Because we had previously corrected mutations in the dihydrofolate reductase (dhfr) gene, in this study, we demonstrate the generality of action of the repair-PPRHs. Repaired cells were analyzed by DNA sequencing, mRNA expression, and enzymatic activity to confirm the correction of the mutation. Moreover, whole-genome sequencing analyses did not detect any off-target effect in the repaired genome. We also performed gel-shift assays to show the binding of the repair-PPRH to the target sequence and the formation of a displacement-loop (D-loop) structure that can trigger a homologous recombination event. Overall, we demonstrate that repair-PPRHs achieve the permanent correction of point mutations in the dsDNA at the endogenous level in mammalian cells without off-target activity.

Toward Developing a Yeast Cell Factory for the Production of Prenylated Flavonoids

Prenylated flavonoids possess a wide variety of biological activities, including estrogenic, antioxidant, antimicrobial, and anticancer activities. Hence, they have potential applications in food products, medicines, or supplements with health-promoting activities. However, the low abundance of prenylated flavonoids in nature is limiting their exploitation. Therefore, we investigated the prospect of producing prenylated flavonoids in the yeast Saccharomyces cerevisiae. As a proof of concept, we focused on the production of the potent phytoestrogen 8-prenylnaringenin. Introduction of the flavonoid prenyltransferase SfFPT from Sophora flavescens in naringenin-producing yeast strains resulted in de novo production of 8-prenylnaringenin. We generated several strains with increased production of the intermediate precursor naringenin, which finally resulted in a production of 0.12 mg L^-1 (0.35 μM) 8-prenylnaringenin under shake flask conditions. A number of bottlenecks in prenylated flavonoid production were identified and are discussed.

A transient reporter for editing enrichment (TREE) in human cells

Current approaches to identify cell populations that have been modified with deaminase base editing technologies are inefficient and rely on downstream sequencing techniques. In this study, we utilized a blue fluorescent protein (BFP) that converts to green fluorescent protein (GFP) upon a C-to-T substitution as an assay to report directly on base editing activity within a cell. Using this assay, we optimize various base editing transfection parameters and delivery strategies. Moreover, we utilize this assay in conjunction with flow cytometry to develop a transient reporter for editing enrichment (TREE) to efficiently purify base-edited cell populations. Compared to conventional cell enrichment strategies that employ reporters of transfection (RoT), TREE significantly improved the editing efficiency at multiple independent loci, with efficiencies approaching 80%. We also employed the BFP-to-GFP conversion assay to optimize base editor vector design in human pluripotent stem cells (hPSCs), a cell type that is resistant to genome editing and in which modification via base editors has not been previously reported. At last, using these optimized vectors in the context of TREE allowed for the highly efficient editing of hPSCs. We envision TREE as a readily adoptable method to facilitate base editing applications in synthetic biology, disease modeling, and regenerative medicine.

Engineered amphiphilic peptides enable delivery of proteins and CRISPR-associated nucleases to airway epithelia

The delivery of biologic cargoes to airway epithelial cells is challenging due to the formidable barriers imposed by its specialized and differentiated cells. Among cargoes, recombinant proteins offer therapeutic promise but the lack of effective delivery methods limits their development. Here, we achieve protein and SpCas9 or AsCas12a ribonucleoprotein (RNP) delivery to cultured human well-differentiated airway epithelial cells and mouse lungs with engineered amphiphilic peptides. These shuttle peptides, non-covalently combined with GFP protein or CRISPR-associated nuclease (Cas) RNP, allow rapid entry into cultured human ciliated and non-ciliated epithelial cells and mouse airway epithelia. Instillation of shuttle peptides combined with SpCas9 or AsCas12a RNP achieves editing of loxP sites in airway epithelia of ROSA^mT/mG mice. We observe no evidence of short-term toxicity with a widespread distribution restricted to the respiratory tract. This peptide-based technology advances potential therapeutic avenues for protein and Cas RNP delivery to refractory airway epithelial cells.

Delivering biological cargo to airway epithelial cells is very challenging. Here, the authors use engineered amphiphilic peptides to shuttle proteins and CRISPR RNPs into airway cells in vivo.

Base Editor Correction of COL7A1 in Recessive Dystrophic Epidermolysis Bullosa Patient-Derived Fibroblasts and iPSCs

Genome editing represents a promising strategy for the therapeutic correction of COL7A1 mutations that cause recessive dystrophic epidermolysis bullosa (RDEB). DNA cleavage followed by homology-directed repair (HDR) using an exogenous template has previously been used to correct COL7A1 mutations. HDR rates can be modest, and the double-strand DNA breaks that initiate HDR commonly result in accompanying undesired insertions and deletions (indels). To overcome these limitations, we applied an A•T→G•C adenine base editor (ABE) to correct two different COL7A1 mutations in primary fibroblasts derived from RDEB patients. ABE enabled higher COL7A1 correction efficiencies than previously reported HDR efforts. Moreover, ABE obviated the need for a repair template, and minimal indels or editing at off-target sites was detected. Base editing restored the endogenous type VII collagen expression and function in vitro. We also treated induced pluripotent stem cells (iPSCs) derived from RDEB fibroblasts with ABE. The edited iPSCs were differentiated into mesenchymal stromal cells, a cell population with therapeutic potential for RDEB. In a mouse teratoma model, the skin derived from ABE-treated iPSCs showed the proper deposition of C7 at the dermal-epidermal junction in vivo. These demonstrate that base editing provides an efficient and precise genome editing method for autologous cell engineering for RDEB.

Base Editor Correction of COL7A1 in Recessive Dystrophic Epidermolysis Bullosa Patient-Derived Fibroblasts and iPSCs

Genome editing represents a promising strategy for the therapeutic correction of COL7A1 mutations that cause recessive dystrophic epidermolysis bullosa (RDEB). DNA cleavage followed by homology-directed repair (HDR) using an exogenous template has previously been used to correct COL7A1 mutations. HDR rates can be modest, and the double-strand DNA breaks that initiate HDR commonly result in accompanying undesired insertions and deletions (indels). To overcome these limitations, we applied an A•T→G•C adenine base editor (ABE) to correct two different COL7A1 mutations in primary fibroblasts derived from RDEB patients. ABE enabled higher COL7A1 correction efficiencies than previously reported HDR efforts. Moreover, ABE obviated the need for a repair template, and minimal indels or editing at off-target sites was detected. Base editing restored the endogenous type VII collagen expression and function in vitro. We also treated induced pluripotent stem cells (iPSCs) derived from RDEB fibroblasts with ABE. The edited iPSCs were differentiated into mesenchymal stromal cells, a cell population with therapeutic potential for RDEB. In a mouse teratoma model, the skin derived from ABE-treated iPSCs showed the proper deposition of C7 at the dermal-epidermal junction in vivo. These demonstrate that base editing provides an efficient and precise genome editing method for autologous cell engineering for RDEB.

Barriers to genome editing with CRISPR in bacteria

Genome editing is essential for probing genotype-phenotype relationships and for enhancing chemical production and phenotypic robustness in industrial bacteria. Currently, the most popular tools for genome editing couple recombineering with DNA cleavage by the CRISPR nuclease Cas9 from Streptococcus pyogenes. Although successful in some model strains, CRISPR-based genome editing has been slow to extend to the multitude of industrially relevant bacteria. In this review, we analyze existing barriers to implementing CRISPR-based editing across diverse bacterial species. We first compare the efficacy of current CRISPR-based editing strategies. Next, we discuss alternatives when the S. pyogenes Cas9 does not yield colonies. Finally, we describe different ways bacteria can evade editing and how elucidating these failure modes can improve CRISPR-based genome editing across strains. Together, this review highlights existing obstacles to CRISPR-based editing in bacteria and offers guidelines to help achieve and enhance editing in a wider range of bacterial species, including non-model strains.

CRISPR-Pass: Gene Rescue of Nonsense Mutations Using Adenine Base Editors

A nonsense mutation is a substitutive mutation in a DNA sequence that causes a premature termination during translation and produces stalled proteins, resulting in dysfunction of a gene. Although it usually induces severe genetic disorders, there are no definite methods for inducing read through of premature termination codons (PTCs). Here, we present a targeted tool for bypassing PTCs, named CRISPR-pass, that uses CRISPR-mediated adenine base editors. CRISPR-pass, which should be applicable to 95.5% of clinically significant nonsense mutations in the ClinVar database, rescues protein synthesis in patient-derived fibroblasts, suggesting potential clinical utility.

Genome editing in large animals: current status and future prospects

Large animals (non-human primates, livestock and dogs) are playing important roles in biomedical research, and large livestock animals serve as important sources of meat and milk. The recently developed programmable DNA nucleases have revolutionized the generation of gene-modified large animals that are used for biological and biomedical research. In this review, we briefly introduce the recent advances in nuclease-meditated gene editing tools, and we outline these editing tools' applications in human disease modeling, regenerative medicine and agriculture. Additionally, we provide perspectives regarding the challenges and prospects of the new genome editing technology.

Lineage tracing using a Cas9-deaminase barcoding system targeting endogenous L1 elements

Determining cell lineage and function is critical to understanding human physiology and pathology. Although advances in lineage tracing methods provide new insight into cell fate, defining cellular diversity at the mammalian level remains a challenge. Here, we develop a genome editing strategy using a cytidine deaminase fused with nickase Cas9 (nCas9) to specifically target endogenous interspersed repeat regions in mammalian cells. The resulting mutation patterns serve as a genetic barcode, which is induced by targeted mutagenesis with single-guide RNA (sgRNA), leveraging substitution events, and subsequent read out by a single primer pair. By analyzing interspersed mutation signatures, we show the accurate reconstruction of cell lineage using both bulk cell and single-cell data. We envision that our genetic barcode system will enable fine-resolution mapping of organismal development in healthy and diseased mammalian states.

A universal method for sensitive and cell-free detection of CRISPR-associated nucleases

A multitude of biological applications for CRISPR-associated (Cas) nucleases have propelled the development of robust cell-based methods for quantitation of on- and off-target activities of these nucleases. However, emerging applications of these nucleases require cell-free methods that are simple, sensitive, cost effective, high throughput, multiplexable, and generalizable to all classes of Cas nucleases. Current methods for cell-free detection are cumbersome, expensive, or require sophisticated sequencing technologies, hindering their widespread application beyond the field of life sciences. Developing such cell-free assays is challenging for multiple reasons, including that Cas nucleases are single-turnover enzymes that must be present in large excess over their substrate and that different classes of Cas nucleases exhibit wildly different operating mechanisms. Here, we report the development of a cell-free method wherein Cas nuclease activity is amplified via an in vitro transcription reaction that produces a fluorescent RNA:small-molecule adduct. We demonstrate that our method is sensitive, detecting activity from low nanomolar concentrations of several families of Cas nucleases, and can be conducted in a high-throughput microplate fashion with a simple fluorescent-based readout. We provide a mathematical framework for quantifying the activities of these nucleases and demonstrate two applications of our method, namely the development of a logic circuit and the characterization of an anti-CRISPR protein. We anticipate our method will be valuable to those studying Cas nucleases and will allow the application of Cas nuclease beyond the field of life sciences.

Engineered CRISPR-Cas12a variants with increased activities and improved targeting ranges for gene, epigenetic and base editing

Broad use of CRISPR-Cas12a (formerly Cpf1) nucleases¹ has been hindered by the requirement for an extended TTTV protospacer adjacent motif (PAM)². To address this limitation, we engineered an enhanced Acidaminococcus sp. Cas12a variant (enAsCas12a) that has a substantially expanded targeting range, enabling targeting of many previously inaccessible PAMs. On average, enAsCas12a exhibits two-fold higher genome editing activity on sites with canonical TTTV PAMs compared to wild-type AsCas12a, and we successfully grafted a subset of mutations from enAsCas12a onto other previously described AsCas12a variants³ to enhance their activities. enAsCas12a improves the efficiency of multiplex gene editing, endogenous gene activation, and C-to-T base editing, and we engineered a high-fidelity version of enAsCas12a (enAsCas12a-HF1) to reduce off-target effects. Both enAsCas12a and enAsCas12a-HF1 function in HEK293T and primary human T cells when delivered as ribonucleoprotein (RNP) complexes. Collectively, enAsCas12a provides an optimized version of Cas12a that should enable wider application of Cas12a enzymes for gene and epigenetic editing. [AU: Revised abstract OK?]

Simultaneous targeting of linked loci in mouse embryos using base editing

A particular challenge in genome engineering has been the simultaneous introduction of mutations into linked (located on the same chromosome) loci. Although CRISPR/Cas9 has been widely used to mutate individual sites, its application in simultaneously targeting of linked loci is limited as multiple nearby double-stranded DNA breaks created by Cas9 routinely result in the deletion of sequences between the cleavage sites. Base editing is a newer form of genome editing that directly converts C∙G-to-T∙A, or A∙T-to-G∙C, base pairs without introducing double-stranded breaks, thus opening the possibility to generate linked mutations without disrupting the entire locus. Through the co-injection of two base editors and two sgRNAs into mouse zygotes, we introduced C∙G-to-T∙A transitions into two cytokine-sensing transcription factor binding sites separated by 9 kb. We determined that one enhancer activates the two flanking genes in mammary tissue during pregnancy and lactation. The ability to introduce linked mutations simultaneously in one step into the mammalian germline has implications for a wide range of applications, including the functional analysis of linked cis-elements creating disease models and correcting pathogenic mutations.

Precision Control of CRISPR-Cas9 Using Small Molecules and Light

The CRISPR (clustered regularly interspaced short palindromic repeat)-Cas system is an adaptive immune system of bacteria that has furnished several RNA-guided DNA endonucleases (e.g., Cas9) that are revolutionizing the field of genome engineering. Cas9 is being used to effect genomic alterations as well as in gene drives, where a particular trait may be propagated through a targeted species population over several generations. The ease of targeting catalytically impaired Cas9 to any genomic loci has led to development of technologies for base editing, chromatin imaging and modeling, epigenetic editing, and gene regulation. Unsurprisingly, Cas9 is being developed for numerous applications in biotechnology and biomedical research and as a gene therapy agent for multiple pathologies. There is a need for precise control of Cas9 activity over several dimensions, including those of dose, time, and space in these applications. Such precision controls, which are required of therapeutic agents, are particularly important for Cas9 as off-target effects, chromosomal translocations, immunogenic response, genotoxicity, and embryonic mosaicism are observed at elevated levels and with prolonged activity of Cas9. Here, we provide a perspective on advances in the precision control of Cas9 over aforementioned dimensions using external stimuli (e.g., small molecules or light) for controlled activation, inhibition, or degradation of Cas9.

Clinical applications of CRISPR-based genome editing and diagnostics

Clustered regularly interspaced short palindromic repeats (CRISPR)-driven genome editing has rapidly transformed preclinical biomedical research by eliminating the underlying genetic basis of many diseases in model systems and facilitating the study of disease etiology. Translation to the clinic is under way, with announced or impending clinical trials utilizing ex vivo strategies for anticancer immunotherapy or correction of hemoglobinopathies. These exciting applications represent just a fraction of what is theoretically possible for this emerging technology, but many technical hurdles must be overcome before CRISPR-based genome editing technology can reach its full potential. One exciting recent development is the use of CRISPR systems for diagnostic detection of genetic sequences associated with pathogens or cancer. We review the biologic origins and functional mechanism of CRISPR systems and highlight several current and future clinical applications of genome editing.

CRISPR Base Editing in Induced Pluripotent Stem Cells

Induced pluripotent stem cells (iPSCs) have demonstrated tremendous potential in numerous disease modeling and regenerative medicine-based therapies. The development of innovative gene transduction and editing technologies has further augmented the potential of iPSCs. Cas9-cytidine deaminases, for example, have developed as an alternative strategy to integrate single-base mutations (C → T or G → A transitions) at specific genomic loci. In this chapter, we specifically describe CRISPR (clustered regularly interspaced short palindromic repeats) base editing in iPSCs for editing precise locations in the genome. This state-of-the-art approach enables highly efficient and accurate modifications in genes. Thus, this technique not only has the potential to have biotechnology and therapeutic applications but also the ability to reveal underlying mechanisms regarding pathologies caused by specific mutations.

Delivering on the promise of gene editing for cystic fibrosis

In this review, we describe a path for translation of gene editing into therapy for cystic fibrosis (CF). Cystic fibrosis results from mutations in the CFTR gene, with one allele predominant in patient populations. This simple, genetic etiology makes gene editing appealing for treatment of this disease. There already have been success in applying this approach to cystic fibrosis in cell and animal models, although these advances have been modest in comparison to advances for other disease.

Less than six years after its first demonstration in animals, CRISPR/Cas gene editing is in early clinical trials for several disorders. Most clinical trials, thus far, attempt to edit genes in cells of the blood lineages. The advantage of the blood is that the stem cells are known, can be isolated, edited, selected, expanded, and returned to the body. The likely next trials will be in the liver, which is accessible to many delivery methods. For cystic fibrosis, the biggest hurdle is to deliver editors to other, less accessible organs. We outline a path by which delivery can be improved.

The translation of new therapies doesn't occur in isolation, and the development of gene editors is occurring as advances in gene therapy and small molecule therapeutics are being made. The advances made in gene therapy may help develop delivery vehicles for gene editing, although major improvements are needed. Conversely, the approval of effective small molecule therapies for many patients with cystic fibrosis will raise the bar for translation of gene editing.

Targeting fidelity of adenine and cytosine base editors in mouse embryos

Base editing directly converts a target base pair into a different base pair in the genome of living cells without introducing double-stranded DNA breaks. While cytosine base editors (CBE) and adenine base editors (ABE) are used to install and correct point mutations in a wide range of organisms, the extent and distribution of off-target edits in mammalian embryos have not been studied in detail. We analyze on-target and proximal off-target editing at 13 loci by a variety of CBEs and ABE in more than 430 alleles generated from mouse zygotic injections using newly generated and published sequencing data. ABE predominantly generates anticipated A•T-to-G•C edits. Among CBEs, SaBE3 and BE4, result in the highest frequencies of anticipated C•G-to-T•A products relative to editing byproducts. Together, these findings highlight the remarkable fidelity of ABE in mouse embryos and identify preferred CBE variants when fidelity in vivo is critical.

CRISPR base editors: genome editing without double-stranded breaks

The CRISPR (clustered regularly interspaced short palindromic repeat)/Cas9 adaptive immunity system has been harnessed for genome editing applications across eukaryotic species, but major drawbacks, such as the inefficiency of precise base editing and off-target activities, remain. A catalytically inactive Cas9 variant (dead Cas9, dCas9) has been fused to diverse functional domains for targeting genetic and epigenetic modifications, including base editing, to specific DNA sequences. As base editing does not require the generation of double-strand breaks, dCas9 and Cas9 nickase have been used to target deaminase domains to edit specific loci. Adenine and cytidine deaminases convert their respective nucleotides into other DNA bases, thereby offering many possibilities for DNA editing. Such base-editing enzymes hold great promise for applications in basic biology, trait development in crops, and treatment of genetic diseases. Here, we discuss recent advances in precise gene editing using different platforms as well as their potential applications in basic biology and biotechnology.

CRISPR base editors: genome editing without double-stranded breaks

The CRISPR (clustered regularly interspaced short palindromic repeat)/Cas9 adaptive immunity system has been harnessed for genome editing applications across eukaryotic species, but major drawbacks, such as the inefficiency of precise base editing and off-target activities, remain. A catalytically inactive Cas9 variant (dead Cas9, dCas9) has been fused to diverse functional domains for targeting genetic and epigenetic modifications, including base editing, to specific DNA sequences. As base editing does not require the generation of double-strand breaks, dCas9 and Cas9 nickase have been used to target deaminase domains to edit specific loci. Adenine and cytidine deaminases convert their respective nucleotides into other DNA bases, thereby offering many possibilities for DNA editing. Such base-editing enzymes hold great promise for applications in basic biology, trait development in crops, and treatment of genetic diseases. Here, we discuss recent advances in precise gene editing using different platforms as well as their potential applications in basic biology and biotechnology.

BE-PLUS: a new base editing tool with broadened editing window and enhanced fidelity

Base editor (BE), containing a cytidine deaminase and catalytically defective Cas9, has been widely used to perform base editing. However, the narrow editing window of BE limits its utility. Here, we developed a new editing technology named as base editor for programming larger C to U (T) scope (BE-PLUS) by fusing 10 copies of GCN4 peptide to nCas9(D10A) for recruiting scFv-APOBEC-UGI-GB1 to the target sites. The new system achieves base editing with a broadened window, resulting in an increased genome-targeting scope. Interestingly, the new system yielded much fewer unwanted indels and non-C-to-T conversions. We also demonstrated its potential use in gene disruption across the whole genome through induction of stop codons (iSTOP). Taken together, the BE-PLUS system offers a new editing tool with increased editing window and enhanced fidelity.

Advancing Metabolic Engineering of Saccharomyces cerevisiae Using the CRISPR/Cas System

Thanks to its ease of use, modularity, and scalability, the clustered regularly interspaced short palindromic repeats (CRISPR) system has been increasingly used in the design and engineering of Saccharomyces cerevisiae, one of the most popular hosts for industrial biotechnology. This review summarizes the recent development of this disruptive technology for metabolic engineering applications, including CRISPR-mediated gene knock-out and knock-in as well as transcriptional activation and interference. More importantly, multi-functional CRISPR systems that combine both gain- and loss-of-function modulations for combinatorial metabolic engineering are highlighted.

Harnessing "A Billion Years of Experimentation": The Ongoing Exploration and Exploitation of CRISPR-Cas Immune Systems

The famed physicist-turned-biologist, Max Delbrück, once remarked that, for physicists, "the field of bacterial viruses is a fine playground for serious children who ask ambitious questions." Early discoveries in that playground helped establish molecular genetics, and half a century later, biologists delving into the same field have ushered in the era of precision genome engineering. The focus has of course shifted-from bacterial viruses and their mechanisms of infection to the bacterial hosts and their mechanisms of immunity-but it is the very same evolutionary arms race that continues to awe and inspire researchers worldwide. In this review, we explore the remarkable diversity of CRISPR-Cas adaptive immune systems, describe the molecular components that mediate nucleic acid targeting, and outline the use of these RNA-guided machines for biotechnology applications. CRISPR-Cas research has yielded far more than just Cas9-based genome-editing tools, and the wide-reaching, innovative impacts of this fascinating biological playground are sure to be felt for years to come.

Programmable base editing of A•T to G•C in genomic DNA without DNA cleavage

The spontaneous deamination of cytosine is a major source of transitions from C•G to T•A base pairs, which account for half of known pathogenic point mutations in humans. The ability to efficiently convert targeted A•T base pairs to G•C could therefore advance the study and treatment of genetic diseases. The deamination of adenine yields inosine, which is treated as guanine by polymerases, but no enzymes are known to deaminate adenine in DNA. Here we describe adenine base editors (ABEs) that mediate the conversion of A•T to G•C in genomic DNA. We evolved a transfer RNA adenosine deaminase to operate on DNA when fused to a catalytically impaired CRISPR-Cas9 mutant. Extensive directed evolution and protein engineering resulted in seventh-generation ABEs that convert targeted A•T base pairs efficiently to G•C (approximately 50% efficiency in human cells) with high product purity (typically at least 99.9%) and low rates of indels (typically no more than 0.1%). ABEs introduce point mutations more efficiently and cleanly, and with less off-target genome modification, than a current Cas9 nuclease-based method, and can install disease-correcting or disease-suppressing mutations in human cells. Together with previous base editors, ABEs enable the direct, programmable introduction of all four transition mutations without double-stranded DNA cleavage.

Programmable base editing of A•T to G•C in genomic DNA without DNA cleavage

The spontaneous deamination of cytosine is a major source of transitions from C•G to T•A base pairs, which account for half of known pathogenic point mutations in humans. The ability to efficiently convert targeted A•T base pairs to G•C could therefore advance the study and treatment of genetic diseases. The deamination of adenine yields inosine, which is treated as guanine by polymerases, but no enzymes are known to deaminate adenine in DNA. Here we describe adenine base editors (ABEs) that mediate the conversion of A•T to G•C in genomic DNA. We evolved a transfer RNA adenosine deaminase to operate on DNA when fused to a catalytically impaired CRISPR-Cas9 mutant. Extensive directed evolution and protein engineering resulted in seventh-generation ABEs that convert targeted A•T base pairs efficiently to G•C (approximately 50% efficiency in human cells) with high product purity (typically at least 99.9%) and low rates of indels (typically no more than 0.1%). ABEs introduce point mutations more efficiently and cleanly, and with less off-target genome modification, than a current Cas9 nuclease-based method, and can install disease-correcting or disease-suppressing mutations in human cells. Together with previous base editors, ABEs enable the direct, programmable introduction of all four transition mutations without double-stranded DNA cleavage.

Programmable base editing of A•T to G•C in genomic DNA without DNA cleavage

The spontaneous deamination of cytosine is a major source of transitions from C•G to T•A base pairs, which account for half of known pathogenic point mutations in humans. The ability to efficiently convert targeted A•T base pairs to G•C could therefore advance the study and treatment of genetic diseases. The deamination of adenine yields inosine, which is treated as guanine by polymerases, but no enzymes are known to deaminate adenine in DNA. Here we describe adenine base editors (ABEs) that mediate the conversion of A•T to G•C in genomic DNA. We evolved a transfer RNA adenosine deaminase to operate on DNA when fused to a catalytically impaired CRISPR-Cas9 mutant. Extensive directed evolution and protein engineering resulted in seventh-generation ABEs that convert targeted A•T base pairs efficiently to G•C (approximately 50% efficiency in human cells) with high product purity (typically at least 99.9%) and low rates of indels (typically no more than 0.1%). ABEs introduce point mutations more efficiently and cleanly, and with less off-target genome modification, than a current Cas9 nuclease-based method, and can install disease-correcting or disease-suppressing mutations in human cells. Together with previous base editors, ABEs enable the direct, programmable introduction of all four transition mutations without double-stranded DNA cleavage.