CRISPR base editing of cis-regulatory elements enables the perturbation of neurodegeneration-linked genes.

CRISPR technology has demonstrated broad utility for controlling target gene expression; however, there remains a need for strategies capable of modulating expression via the precise editing of non-coding regulatory elements. Here, we demonstrate that CRISPR base editors, a class of gene-modifying proteins capable of creating single-base substitutions in DNA, can be used to perturb gene expression via their targeted mutagenesis of cis-acting sequences. Using the promoter region of the human huntingtin (HTT) gene as an initial target, we show that editing of the binding site for the transcription factor NF-κB led to a marked reduction in HTT gene expression in base-edited cell populations. We found that these gene perturbations were persistent and specific, as a transcriptome-wide RNA analysis revealed minimal off-target effects resulting from the action of the base editor protein. We further demonstrate that this base-editing platform could influence gene expression in vivo as its delivery to a mouse model of Huntington's disease led to a potent decrease in HTT mRNA in striatal neurons. Finally, to illustrate the applicability of this concept, we target the amyloid precursor protein, showing that multiplex editing of its promoter region significantly perturbed its expression. These findings demonstrate the potential for base editors to regulate target gene expression.

Treatment of a Mouse Model of ALS by In Vivo Base Editing.

Amyotrophic lateral sclerosis (ALS) is a debilitating and fatal disorder that can be caused by mutations in the superoxide dismutase 1 (SOD1) gene. Although ALS is currently incurable, CRISPR base editors hold the potential to treat the disease through their ability to create nonsense mutations that can permanently disable the expression of the mutant SOD1 gene. However, the restrictive carrying capacity of adeno-associated virus (AAV) vectors has limited their therapeutic application. In this study, we establish an intein-mediated trans-splicing system that enables in vivo delivery of cytidine base editors (CBEs) consisting of the widely used Cas9 protein from Streptococcus pyogenes. We show that intrathecal injection of dual AAV particles encoding a split-intein CBE engineered to trans-splice and introduce a nonsense-coding substitution into a mutant SOD1 gene prolonged survival and markedly slowed the progression of disease in the G93A-SOD1 mouse model of ALS. Adult animals treated by this split-intein CRISPR base editor had a reduced rate of muscle atrophy, decreased muscle denervation, improved neuromuscular function, and up to 40% fewer SOD1 immunoreactive inclusions at end-stage mice compared to control mice. This work expands the capabilities of single-base editors and demonstrates their potential for gene therapy.

Targeted gene knock-in by homology-directed genome editing using Cas9 ribonucleoprotein and AAV donor delivery.

Realizing the full potential of genome editing requires the development of efficient and broadly applicable methods for delivering programmable nucleases and donor templates for homology-directed repair (HDR). The RNA-guided Cas9 endonuclease can be introduced into cells as a purified protein in complex with a single guide RNA (sgRNA). Such ribonucleoproteins (RNPs) can facilitate the high-fidelity introduction of single-base substitutions via HDR following co-delivery with a single-stranded DNA oligonucleotide. However, combining RNPs with transgene-containing donor templates for targeted gene addition has proven challenging, which in turn has limited the capabilities of the RNP-mediated genome editing toolbox. Here, we demonstrate that combining RNP delivery with naturally recombinogenic adeno-associated virus (AAV) donor vectors enables site-specific gene insertion by homology-directed genome editing. Compared to conventional plasmid-based expression vectors and donor templates, we show that combining RNP and AAV donor delivery increases the efficiency of gene addition by up to 12-fold, enabling the creation of lineage reporters that can be used to track the conversion of striatal neurons from human fibroblasts in real time. These results thus illustrate the potential for unifying nuclease protein delivery with AAV donor vectors for homology-directed genome editing.

CRISPR-mediated Activation of Latent HIV-1 Expression.

Complete eradication of HIV-1 infection is impeded by the existence of cells that harbor chromosomally integrated but transcriptionally inactive provirus. These cells can persist for years without producing viral progeny, rendering them refractory to immune surveillance and antiretroviral therapy and providing a permanent reservoir for the stochastic reactivation and reseeding of HIV-1. Strategies for purging this latent reservoir are thus needed to eradicate infection. Here, we show that engineered transcriptional activation systems based on CRISPR/Cas9 can be harnessed to activate viral gene expression in cell line models of HIV-1 latency. We further demonstrate that complementing Cas9 activators with latency-reversing compounds can enhance latent HIV-1 transcription and that epigenome modulation using CRISPR-based acetyltransferases can also promote viral gene activation. Collectively, these results demonstrate that CRISPR systems are potentially effective tools for inducing latent HIV-1 expression and that their use, in combination with antiretroviral therapy, could lead to improved therapies for HIV-1 infection.

Genome Engineering Using Adeno-associated Virus: Basic and Clinical Research Applications.

In addition to their broad potential for therapeutic gene delivery, adeno-associated virus (AAV) vectors possess the innate ability to stimulate homologous recombination in mammalian cells at high efficiencies. This process--referred to as AAV-mediated gene targeting--has enabled the introduction of a diverse array of genomic modifications both in vitro and in vivo. With the recent emergence of targeted nucleases, AAV-mediated genome engineering is poised for clinical translation. Here, we review key properties of AAV vectors that underscore its unique utility in genome editing. We highlight the broad range of genome engineering applications facilitated by this technology and discuss the strong potential for unifying AAV with targeted nucleases for next-generation gene therapy.

Expanding the zinc-finger recombinase repertoire: directed evolution and mutational analysis of serine recombinase specificity determinants.

The serine recombinases are a diverse family of modular enzymes that promote high-fidelity DNA rearrangements between specific target sites. Replacement of their native DNA-binding domains with custom-designed Cys2-His2 zinc-finger proteins results in the creation of engineered zinc-finger recombinases (ZFRs) capable of achieving targeted genetic modifications. The flexibility afforded by zinc-finger domains enables the design of hybrid recombinases that recognize a wide variety of potential target sites; however, this technology remains constrained by the strict recognition specificities imposed by the ZFR catalytic domains. In particular, the ability to fully reprogram serine recombinase catalytic specificity has been impeded by conserved base requirements within each recombinase target site and an incomplete understanding of the factors governing DNA recognition. Here we describe an approach to complement the targeting capacity of ZFRs. Using directed evolution, we isolated mutants of the ß and Sin recombinases that specifically recognize target sites previously outside the scope of ZFRs. Additionally, we developed a genetic screen to determine the specific base requirements for site-specific recombination and showed that specificity profiling enables the discovery of unique genomic ZFR substrates. Finally, we conducted an extensive and family-wide mutational analysis of the serine recombinase DNA-binding arm region and uncovered a diverse network of residues that confer target specificity. These results demonstrate that the ZFR repertoire is extensible and highlights the potential of ZFRs as a class of flexible tools for targeted genome engineering.

Targeted gene knockout by direct delivery of zinc-finger nuclease proteins.

Zinc-finger nucleases (ZFNs) are versatile reagents that have redefined genome engineering. Realizing the full potential of this technology requires the development of safe and effective methods for delivering ZFNs into cells. We demonstrate the intrinsic cell-penetrating capabilities of the standard ZFN architecture and show that direct delivery of ZFNs as proteins leads to efficient endogenous gene disruption in various mammalian cell types with minimal off-target effects.

Synthetic zinc finger proteins: the advent of targeted gene regulation and genome modification technologies.

The understanding of gene regulation and the structure and function of the human genome increased dramatically at the end of the 20th century. Yet the technologies for manipulating the genome have been slower to develop. For instance, the field of gene therapy has been focused on correcting genetic diseases and augmenting tissue repair for more than 40 years. However, with the exception of a few very low efficiency approaches, conventional genetic engineering methods have only been able to add auxiliary genes to cells. This has been a substantial obstacle to the clinical success of gene therapies and has also led to severe unintended consequences in several cases. Therefore, technologies that facilitate the precise modification of cellular genomes have diverse and significant implications in many facets of research and are essential for translating the products of the Genomic Revolution into tangible benefits for medicine and biotechnology. To address this need, in the 1990s, we embarked on a mission to develop technologies for engineering protein-DNA interactions with the aim of creating custom tools capable of targeting any DNA sequence. Our goal has been to allow researchers to reach into genomes to specifically regulate, knock out, or replace any gene. To realize these goals, we initially focused on understanding and manipulating zinc finger proteins. In particular, we sought to create a simple and straightforward method that enables unspecialized laboratories to engineer custom DNA-modifying proteins using only defined modular components, a web-based utility, and standard recombinant DNA technology. Two significant challenges we faced were (i) the development of zinc finger domains that target sequences not recognized by naturally occurring zinc finger proteins and (ii) determining how individual zinc finger domains could be tethered together as polydactyl proteins to recognize unique locations within complex genomes. We and others have since used this modular assembly method to engineer artificial proteins and enzymes that activate, repress, or create defined changes to user-specified genes in human cells, plants, and other organisms. We have also engineered novel methods for externally controlling protein activity and delivery, as well as developed new strategies for the directed evolution of protein and enzyme function. This Account summarizes our work in these areas and highlights independent studies that have successfully used the modular assembly approach to create proteins with novel function. We also discuss emerging alternative methods for genomic targeting, including transcription activator-like effectors (TALEs) and CRISPR/Cas systems, and how they complement the synthetic zinc finger protein technology.

A comprehensive approach to zinc-finger recombinase customization enables genomic targeting in human cells.

Zinc-finger recombinases (ZFRs) represent a potentially powerful class of tools for targeted genetic engineering. These chimeric enzymes are composed of an activated catalytic domain derived from the resolvase/invertase family of serine recombinases and a custom-designed zinc-finger DNA-binding domain. The use of ZFRs, however, has been restricted by sequence requirements imposed by the recombinase catalytic domain. Here, we combine substrate specificity analysis and directed evolution to develop a diverse collection of Gin recombinase catalytic domains capable of recognizing an estimated 3.77 × 10(7) unique DNA sequences. We show that ZFRs assembled from these engineered catalytic domains recombine user-defined DNA targets with high specificity, and that designed ZFRs integrate DNA into targeted endogenous loci in human cells. This study demonstrates the feasibility of generating customized ZFRs and the potential of ZFR technology for a diverse range of applications, including genome engineering, synthetic biology and gene therapy.

Enhancing the specificity of recombinase-mediated genome engineering through dimer interface redesign.

Despite recent advances in genome engineering made possible by the emergence of site-specific endonucleases, there remains a need for tools capable of specifically delivering genetic payloads into the human genome. Hybrid recombinases based on activated catalytic domains derived from the resolvase/invertase family of serine recombinases fused to Cys2-His2 zinc-finger or TAL effector DNA-binding domains are a class of reagents capable of achieving this. The utility of these enzymes, however, has been constrained by their low overall targeting specificity, largely due to the formation of side-product homodimers capable of inducing off-target modifications. Here, we combine rational design and directed evolution to re-engineer the serine recombinase dimerization interface and generate a recombinase architecture that reduces formation of these undesirable homodimers by >500-fold. We show that these enhanced recombinases demonstrate substantially improved targeting specificity in mammalian cells and achieve rates of site-specific integration similar to those previously reported for site-specific nucleases. Additionally, we show that enhanced recombinases exhibit low toxicity and promote the delivery of the human coagulation factor IX and α-galactosidase genes into endogenous genomic loci with high specificity. These results provide a general means for improving hybrid recombinase specificity by protein engineering and illustrate the potential of these enzymes for basic research and therapeutic applications.

Expanding the scope of site-specific recombinases for genetic and metabolic engineering.

Site-specific recombinases are tremendously valuable tools for basic research and genetic engineering. By promoting high-fidelity DNA modifications, site-specific recombination systems have empowered researchers with unprecedented control over diverse biological functions, enabling countless insights into cellular structure and function. The rigid target specificities of many sites-specific recombinases, however, have limited their adoption in fields that require highly flexible recognition abilities. As a result, intense effort has been directed toward altering the properties of site-specific recombination systems by protein engineering. Here, we review key developments in the rational design and directed molecular evolution of site-specific recombinases, highlighting the numerous applications of these enzymes across diverse fields of study.

Chimeric TALE recombinases with programmable DNA sequence specificity.

Site-specific recombinases are powerful tools for genome engineering. Hyperactivated variants of the resolvase/invertase family of serine recombinases function without accessory factors, and thus can be re-targeted to sequences of interest by replacing native DNA-binding domains (DBDs) with engineered zinc-finger proteins (ZFPs). However, imperfect modularity with particular domains, lack of high-affinity binding to all DNA triplets, and difficulty in construction has hindered the widespread adoption of ZFPs in unspecialized laboratories. The discovery of a novel type of DBD in transcription activator-like effector (TALE) proteins from Xanthomonas provides an alternative to ZFPs. Here we describe chimeric TALE recombinases (TALERs): engineered fusions between a hyperactivated catalytic domain from the DNA invertase Gin and an optimized TALE architecture. We use a library of incrementally truncated TALE variants to identify TALER fusions that modify DNA with efficiency and specificity comparable to zinc-finger recombinases in bacterial cells. We also show that TALERs recombine DNA in mammalian cells. The TALER architecture described herein provides a platform for insertion of customized TALE domains, thus significantly expanding the targeting capacity of engineered recombinases and their potential applications in biotechnology and medicine.

Structure-guided reprogramming of serine recombinase DNA sequence specificity.

Routine manipulation of cellular genomes is contingent upon the development of proteins and enzymes with programmable DNA sequence specificity. Here we describe the structure-guided reprogramming of the DNA sequence specificity of the invertase Gin from bacteriophage Mu and Tn3 resolvase from Escherichia coli. Structure-guided and comparative sequence analyses were used to predict a network of amino acid residues that mediate resolvase and invertase DNA sequence specificity. Using saturation mutagenesis and iterative rounds of positive antibiotic selection, we identified extensively redesigned and highly convergent resolvase and invertase populations in the context of engineered zinc-finger recombinase (ZFR) fusion proteins. Reprogrammed variants selectively catalyzed recombination of nonnative DNA sequences > 10,000-fold more effectively than their parental enzymes. Alanine-scanning mutagenesis revealed the molecular basis of resolvase and invertase DNA sequence specificity. When used as rationally designed ZFR heterodimers, the reprogrammed enzyme variants site-specifically modified unnatural and asymmetric DNA sequences. Early studies on the directed evolution of serine recombinase DNA sequence specificity produced enzymes with relaxed substrate specificity as a result of randomly incorporated mutations. In the current study, we focused our mutagenesis exclusively on DNA determinants, leading to redesigned enzymes that remained highly specific and directed transgene integration into the human genome with > 80% accuracy. These results demonstrate that unique resolvase and invertase derivatives can be developed to site-specifically modify the human genome in the context of zinc-finger recombinase fusion proteins.

Site-selective labeling of a lysine residue in human serum albumin.

Conjugation to human serum albumin (HSA) has emerged as a powerful approach for extending the in vivo half-life of many small molecule and peptide/protein drugs. Current HSA conjugation strategies, however, can often yield heterogeneous mixtures with inadequate pharmacokinetics, low efficacies, and variable safety profiles. Here, we designed and synthesized analogues of TAK-242, a small molecule inhibitor of Toll-like receptor 4, that primarily reacted with a single lysine residue of HSA. These TAK-242-based cyclohexene compounds demonstrated robust reactivity, and Lys64 was identified as the primary conjugation site. A bivalent HSA conjugate was also prepared in a site-specific manner. Additionally, HSA-cyclohexene conjugates maintained higher levels of stability both in human plasma and in mice than the corresponding maleimide conjugates. This new conjugation strategy promises to broadly enhance the performance of HSA conjugates for numerous applications.

SPLICER: A Highly Efficient Base Editing Toolbox That Enables In Vivo Therapeutic Exon Skipping.

Exon skipping technologies enable exclusion of targeted exons from mature mRNA transcripts, which has broad applications in molecular biology, medicine, and biotechnology. Existing exon skipping techniques include antisense oligonucleotides, targetable nucleases, and base editors, which, while effective for specific applications at some target exons, remain hindered by shortcomings, including transient effects for oligonucleotides, genotoxicity for nucleases and inconsistent exon skipping for base editors. To overcome these limitations, we created SPLICER, a toolbox of next-generation base editors consisting of near-PAMless Cas9 nickase variants fused to adenosine or cytosine deaminases for the simultaneous editing of splice acceptor (SA) and splice donor (SD) sequences. Synchronized SA and SD editing with SPLICER improves exon skipping, reduces aberrant outcomes, including cryptic splicing and intron retention, and enables skipping of exons refractory to single splice-site editing. To demonstrate the therapeutic potential of SPLICER, we targeted APP exon 17, which encodes the amino acid residues that are cleaved to form the Aß plaques in Alzheimer's disease. SPLICER reduced the formation of Aß42 peptides in vitro and enabled efficient exon skipping in a mouse model of Alzheimer's disease. Overall, SPLICER is a widely applicable and efficient toolbox for exon skipping with broad therapeutic applications.

In vivo CRISPR base editing for treatment of Huntington's disease.

Huntington's disease (HD) is an inherited and ultimately fatal neurodegenerative disorder caused by an expanded polyglutamine-encoding CAG repeat within exon 1 of the huntingtin (HTT) gene, which produces a mutant protein that destroys striatal and cortical neurons. Importantly, a critical event in the pathogenesis of HD is the proteolytic cleavage of the mutant HTT protein by caspase-6, which generates fragments of the N-terminal domain of the protein that form highly toxic aggregates. Given the role that proteolysis of the mutant HTT protein plays in HD, strategies for preventing this process hold potential for treating the disorder. By screening 141 CRISPR base editor variants targeting splice elements in the HTT gene, we identified platforms capable of producing HTT protein isoforms resistant to caspase-6-mediated proteolysis via editing of the splice acceptor sequence for exon 13. When delivered to the striatum of a rodent HD model, these base editors induced efficient exon skipping and decreased the formation of the N-terminal fragments, which in turn reduced HTT protein aggregation and attenuated striatal and cortical atrophy. Collectively, these results illustrate the potential for CRISPR base editing to decrease the toxicity of the mutant HTT protein for HD.

Targeted plasmid integration into the human genome by an engineered zinc-finger recombinase.

The development of new methods for gene addition to mammalian genomes is necessary to overcome the limitations of conventional genetic engineering strategies. Although a variety of DNA-modifying enzymes have been used to directly catalyze the integration of plasmid DNA into mammalian genomes, there is still an unmet need for enzymes that target a single specific chromosomal site. We recently engineered zinc-finger recombinase (ZFR) fusion proteins that integrate plasmid DNA into a synthetic target site in the human genome with exceptional specificity. In this study, we present a two-step method for utilizing these enzymes in any cell type at randomly-distributed target site locations. The piggyBac transposase was used to insert recombinase target sites throughout the genomes of human and mouse cell lines. The ZFR efficiently and specifically integrated a transfected plasmid into these genomic target sites and into multiple transposons within a single cell. Plasmid integration was dependent on recombinase activity and the presence of recombinase target sites. This work demonstrates the potential for broad applicability of the ZFR technology in genome engineering, synthetic biology and gene therapy.

Delivering Base Editors In Vivo by Adeno-Associated Virus Vectors.

CRISPR base editors are genome-modifying proteins capable of creating single-base substitutions in DNA but without the requirement for a DNA double-strand break. Given their ability to precisely edit DNA, they hold tremendous therapeutic potential. Here, we describe procedures for delivering base editors in vivo via adeno-associated virus (AAV) vectors, a promising engineered gene delivery vehicle capable of transducing a range of cell types and tissues. We provide step by step protocols for (i) designing and validating base editing systems, (ii) packaging base editors into recombinant AAV vector particles, (iii) delivering AAV to the central nervous system via intrathecal injection, and (iv) quantifying base editing frequencies by next-generation sequencing.

Mitigating a TDP-43 proteinopathy by targeting ataxin-2 using RNA-targeting CRISPR effector proteins.

The TDP-43 proteinopathies, which include amyotrophic lateral sclerosis and frontotemporal dementia, are a devastating group of neurodegenerative disorders that are characterized by the mislocalization and aggregation of TDP-43. Here we demonstrate that RNA-targeting CRISPR effector proteins, a programmable class of gene silencing agents that includes the Cas13 family of enzymes and Cas7-11, can be used to mitigate TDP-43 pathology when programmed to target ataxin-2, a modifier of TDP-43-associated toxicity. In addition to inhibiting the aggregation and transit of TDP-43 to stress granules, we find that the in vivo delivery of an ataxin-2-targeting Cas13 system to a mouse model of TDP-43 proteinopathy improved functional deficits, extended survival, and reduced the severity of neuropathological hallmarks. Further, we benchmark RNA-targeting CRISPR platforms against ataxin-2 and find that high-fidelity forms of Cas13 possess improved transcriptome-wide specificity compared to Cas7-11 and a first-generation effector. Our results demonstrate the potential of CRISPR technology for TDP-43 proteinopathies.

Mitigating a TDP-43 proteinopathy by targeting ataxin-2 using RNA-targeting CRISPR effector proteins.

The TDP-43 proteinopathies, which include amyotrophic lateral sclerosis and frontotemporal dementia, are a devastating group of neurodegenerative disorders that are characterized by the mislocalization and aggregation of TDP-43. Here we demonstrate that RNA-targeting CRISPR effector proteins, a programmable class of gene silencing agents that includes the Cas13 family of enzymes and Cas7-11, can be used to mitigate TDP-43 pathology when programmed to target ataxin-2, a modifier of TDP-43-associated toxicity. In addition to inhibiting the aggregation and transit of TDP-43 to stress granules, we find that the in vivo delivery of an ataxin-2-targeting Cas13 system to a mouse model of TDP-43 proteinopathy improved functional deficits, extended survival, and reduced the severity of neuropathological hallmarks. Further, we benchmark RNA-targeting CRISPR platforms against ataxin-2 and find that high-fidelity forms of Cas13 possess improved transcriptome-wide specificity compared to Cas7-11 and a first-generation effector. Our results demonstrate the potential of CRISPR technology for TDP-43 proteinopathies.