Perspective pubs.acs.org/acschemicalbiology
Genome Editing: Insights from Chemical Biology to Support Safe and Transformative Therapeutic Applications Renee D. Wegrzyn,*,† Andrew H. Lee,‡ Amy L. Jenkins,§ Colby D. Stoddard,∥ and Anne E. Cheever‡ †
Defense Advanced Research Projects Agency (DARPA), 675 N. Randolph St., Arlington, Virginia 22203, United States Booz Allen Hamilton, 3811 Fairfax Dr. Suite 600, Arlington, Virginia 22203, United States § Schafer: A Belcan Company, 3811 Fairfax Dr., Arlington, Virginia 22203, United States ∥ Quantitative Scientific Solutions, 4601 N. Fairfax Dr. Suite 1200, Arlington, Virginia 22203, United States ‡
ABSTRACT: Programmable nuclease-based genome editing technologies, including the clustered, regularly interspaced, short palindromic repeats (CRISPR)/Cas9 system, are becoming an essential component of many applications ranging from agriculture to medicine. However, fundamental limitations currently prevent the widespread, safe, and practical use of genome editors, especially for human disease interventions. These limitations include off-target effects, a lack of control over editing activity, suboptimal DNA repair outcomes, insufficient target conversion, and inadequate delivery performance. This perspective focuses on the potential for biological chemistry to address these limitations such that newly developed genome editing technologies can enable the broadest range of potential future applications. Equally important will be the development of these powerful technologies within a relevant ethical framework that emphasizes safety and responsible innovation.
including heritable genetic disorders, that will ultimately require the use of genome editors in vivo. Beyond heritable genetic disorders, new opportunities are emerging to apply genome editors to combat a variety of infectious disease challenges, including viral infections8 and antimicrobial resistance.9 While these approaches may one day enable treatment or prophylactic options for routine infections, their use in otherwise healthy patients will be contingent on more stringent biosafety requirements and significant improvements in performance. Therefore, it is paramount that these technical advances be addressed early in the development timeline of CRISPR technologies to fully utilize genome editing tools for therapeutic benefit. Recent demonstrations of small molecule and novel macromolecular design strategies to modulate the activity of genome editors provide the foundation to deliver disruptive capabilities for safe and effective genome editing technologies. These insights support the development of design rules to engineer new editing enzymes and functionalities, first-in-class molecular inhibitors of gene editing activity, improved controls to refine outcomes of DNA repair events, and enhanced formulations for effective in vivo delivery. Importantly, when
Programmable nuclease-based genome editors, including the CRISPR/Cas9 system, enable researchers to modify an organism’s genomic material in a manner that is increasingly targeted, rapid, and cost-effective. CRISPR/Cas9 gene editing tools have not only enabled significant advancements in genetic research, including manipulation of previously inaccessible genomes, but have also set the groundwork for transformative applications in the fields of disease vector control, agriculture, and biomedicine. Among the most compelling of genome editing applications are those that seek to enable novel therapeutic intervention strategies to promote human health. Initial preclinical and clinical work on therapeutic applications of gene editors, including T-cell immunotherapies to treat cancers,1 sickle cell disease,2 and reduction of HIV burden in patients,3 have underscored the potential for these tools to provide a novel and disruptive means to address otherwise intractable human diseases. Despite promising early results, these studies have also revealed significant challenges that are associated with the current suite of genome editing technologies, including the presence of off-target effects (both predicted and unanticipated4), a lack of precise control over genome editing activity and repair outcomes,5 low efficiency of target conversion,6 and in vivo delivery limitations.7 These challenges highlight the performance and biosafety concerns that have so far limited clinical successes for genome editors to the ex vivo manipulation of cells for therapeutic benefit3 and hinder progress toward new solutions to treat a range of conditions, © 2017 American Chemical Society
Special Issue: Chemical Biology of CRISPR Received: August 9, 2017 Accepted: October 9, 2017 Published: October 9, 2017 333
DOI: 10.1021/acschembio.7b00689 ACS Chem. Biol. 2018, 13, 333−342
Perspective
ACS Chemical Biology
that do not induce DSBs, have also been developed to create DNA nicks (Cas9-D10A12,13 and Cas9-FokI14), which can lead to higher HDR frequencies15 and reduced off-target effects.14 The DNA repair processes that are triggered by DSBs increase the probability of unintended effects both at target and nontarget sites, from disruption of nontarget gene function through single nucleotide polymorphisms and frameshifts to chromosomal rearrangements and genome instability.16 The field lacks the ability to easily remediate or correct these types of off-target mutations, particularly in therapeutic applications. Therefore, new approaches are necessary to mitigate these potential risks. One way to address the challenges associated with DNA repair and reduce the risk of unintended effects has been through the development of noncanonical genome editing systems, or Cas9 variants, that leverage programmable DNA target binding without DNA cleavage activity. Catalytically inactive or “dead” Cas9 (dCas9)17 serves as a central platform or scaffold for building novel noncanonical functionality. Novel effector domains, such as cytosine deamination for base editing;18 epigenetic modifiers such as p300,19 Tet1,20 DNMT3a,21 LSD1,22 and MQ123 to alter gene expression activity; or transcriptional activator/repressor domains such as VP64,24 VPR,25 and KRAB26,27 can be fused directly to dCas9 to activate or repress target gene expression. These noncanonical editors have the potential to generate first-in-class therapeutics that can be single dose, yet elicit long-term cellular responses that are ultimately reversible. Other approaches may modulate the transcriptome directly through RNA cleavage, thus avoiding DNA targets altogether (e.g., C2c2/Cas13a28,29). These tools, while already demonstrated in vitro,28,29 will require the development and refinement of methods to carefully modulate editor activity and the discovery of genetic targets for tunable gene expression therapies. It is conceivable that noncanonical genome editingbased therapeutics could be used for a diverse range of applications, from temporarily conferring protection against infectious diseases to mitigating acute or chronic pain in a nonaddictive manner to treating cancers or other complex ailments. Together, the development of transcriptome and epigenome editors would enable safer targeted, transient, and reversible therapies. Modulating Cellular Processes. To achieve fundamental improvements in the design of safe and effective gene editors, it is also important to consider the cellular repair processes that can influence the outcome of a given genome editing event. The current toolbox of “precision genome editors” is relatively efficient at inducing a targeted DNA lesion (a site-specific DSB); however, the subsequent repair processes that must proceed to achieve the actual edit (i.e., replacement, deletion, or revision of the target sequence) are subject to the host cell context and can be inefficient and error-prone. In somatic cells, resolution of DSBs is biased toward NHEJ, with reported frequencies of 20−60% in targeted cells30,31 compared to 0.5− 20% for HDR,30−32 where lower efficiency is attributable to the need for codelivery of a donor template. NHEJ repair is active at all stages of the cell cycle,33 whereas HDR is typically active in S and G2 phases when DNA is replicating. Therefore, while gene editing strategies can be designed to favor certain repair pathways, they cannot yet be fully controlled. To enter the next phase of true “precision” genome editing, new advances are required to control cellular repair outcomes of CRISPR-mediated editing in a deterministic manner. One
considering current challenges to the future translation of genome editing technologies to the clinic, the most comprehensive solutions will also address ethical and societal concerns that are associated with these powerful genome editing technologies.
■
FUNDAMENTAL IMPROVEMENTS IN GENOME EDITING DESIGNS FOR SAFETY The development of genome editors for enhanced efficacy and biosafety requires a detailed biophysical and mechanistic understanding of editor activity and necessitates design efforts in the context of the natural cellular repair processes that will help determine the outcomes of editing events. Fundamental technical improvements such as the development of multiple genome editing systems with varying functionalities to match applications, deterministic control of cellular repair outcomes of CRISPR-mediated editing, and advancements in measuring offtarget effects are all required to move the field toward safe and effective clinical applications. Engineering CRISPR Systems. The canonical CRISPR/ Cas9 system consists of two components: Cas9 nuclease protein and a single guide RNA (sgRNA or gRNA; Figure 1).
Figure 1. Schematic drawing of canonical CRISPR/Cas9 genome editing mechanism and outcomes. The Cas9:guide RNA (dark gray and yellow, respectively) ribonucleoprotein binds a target DNA sequence at the protospacer adjacent motif (PAM) sequence, opens the DNA double helix, and hybridizes the guide RNA to the target sequence (protospacer) before generating a DNA double-strand break (DSB). DSBs are typically resolved by either the error-prone nonhomologous end joining (NHEJ) or homology-directed repair (HDR) DNA repair pathways.
The Cas9:gRNA ribonucleoprotein (RNP) first detects and binds to protospacer-adjacent-motif (PAM) sites, then interrogates adjacent protospacer sequence complementarity through RNA hybridization and generates a DNA doublestrand break (DSB) through its endonuclease activity.10 DSBs are resolved primarily through the error-prone nonhomologous end joining (NHEJ) pathway or the homology-directed repair (HDR) pathway. NHEJ can generate small insertions or deletions (indels) and disrupt target gene function. In contrast, HDR uses a donor DNA sequence as a template for repair, which can be leveraged to introduce a user-defined sequence into a locus.11 Cas9 nuclease variants, such as Cas9 nickases 334
DOI: 10.1021/acschembio.7b00689 ACS Chem. Biol. 2018, 13, 333−342
Perspective
ACS Chemical Biology
Figure 2. Strategies to control Cas9 activity. CRISPR/Cas9 activity can be controlled using multiple strategies including to activate Cas9 through domain tags and gRNA controllers or inhibit Cas9 function. (A) Split Cas9: Cas9 is divided into two monomers fused to interacting domains (domains 1 and 2), which dimerize upon introduction of a stimulus such as a ligand or light. Intein Cas9: intein protein fusion into Cas9 blocks Cas9 activity in the absence of a ligand. Ligand binding triggers post-translational splicing of the intein domain, resulting in functional Cas9. Destabilizing domains (DD): DD-Cas9 fusion leads to Cas9 instability and proteosomal degradation. Addition of ligand stabilizes DD-Cas9 to its functional form. Domain replacement: Cas9 REC2 domain replacement with ligand-dependent protein−protein interaction domains prevents Cas9 activity until ligand is introduced. (B) Chemical modification or gRNA truncation help enable stability and reduced off-targets, respectively. (C) Aptazyme control: Ligand-dependent aptazyme-gRNA constructs enables aptazyme self-cleavage and release of a functional Cas9 upon ligand introduction. Effector control: appending binding sequences (Casilio) or hairpin structures (Scaffold) to gRNAs enables effector protein binding and target gene modulation. (D) Inhibition of Cas9 can occur during gRNA binding, DNA binding, or DNA cleavage. Small molecules, competing nucleic acids, or anti-CRISPR proteins could be employed at any stage to prevent or halt gene editing.
strategy with particular utility for ex vivo applications involves the use of small molecules that function to synchronize the cell cycle34,35 or otherwise bias repair processes by disrupting DNA cleavage activity or targeting endogenous cellular activities to increase desired editing outcomes.5,36,37 For example, inhibiting NHEJ by blocking the DNA binding capability of DNA Ligase
IV with the small molecule inhibitor Scr7 has been shown in some cases to decrease NHEJ and increase HDR efficiencies.5,37 Use of the β3-adrenergic receptor partial agonist L755507 has also been shown to improve HDR efficiencies, although the mechanism by which it does so remains unclear.38,39 Small molecule-mediated cell-cycle arrest followed 335
DOI: 10.1021/acschembio.7b00689 ACS Chem. Biol. 2018, 13, 333−342
Perspective
ACS Chemical Biology
including complex or indirect off-target outcomes such as alterations of the epigenome after conversion of pathogenic genetic target with a DNA editor, remains a challenge. Many approaches are expensive, require a reference genome, or are not scalable to multiple gRNAs or sites. A recent provocative study reported the ultimately unsubstantiated finding of very high frequency Cas9-mediated off-target effects in organisms,50,51 emphasizing the urgent need for better tools and standards for prediction, detection, and mitigation of off-target effects induced by gene editors before clinical use of these tools becomes the norm.
by the timed delivery of CRISPR/Cas9 RNPs has demonstrated HDR-mediated editing levels of up to 38% with no detectable off-target editing.35 These initial results highlight the potential utility of leveraging chemical compounds to obtain desired DNA repair outcomes, particularly for ex vivo applications where protein engineering and discovery of novel genome editors may fall short. Small molecule mediated solutions to synchronize the cell cycle and optimize repair outcomes are useful for ex vivo applications, but may be toxic in vivo. Therefore, alternative strategies are required to modulate DNA repair pathway choice. One approach could use covalent fusion of protein domains or small molecule moieties to editing machinery to modulate pathway choice at the site of editing. An initial demonstration of this capability involved the tethering of a cytidine deaminase enzyme to a Cas9 nickase to convert cytidine to uridine at the target site, eliminating the need for a DSB or donor template to achieve “base edits” with efficiencies ranging from 15 to 75%.18 Additionally, post-translational regulation of CRISPR/Cas9 through fusion with the replication licensing factor Geminin that limits activity of the fusion in a cell-cycle dependent manner increased the rate of HDR to 87%.40 Finally, careful design and choice of CRISPR components can bias repair to desired outcomes. For instance, Cas9-nickase or Cpf1generated DSBs with 5′ or 3′ overhang DNA ends can affect repair pathway choice to favor HDR over NHEJ.15,41 The length of donor homology arms can help dictate the preference for less common pathways, such as microhomology-mediated end-joining (MMEJ), 42−44 while linear dsDNA donor templates can promote synthesis-dependent strand annealing HDR over double Holliday junction-mediated HDR.45 Reducing Off-Targets. Finally, advancements in measuring off-target effects will be critical for improving the safety of both ex vivo and in vivo therapeutic applications. We consider offtarget effects broadly, inclusive of unintended genetic, transcriptomic, or epigenetic changes at nontarget sites across the genome, unintended outcomes at the target site itself, and genome editing that may occur in nontarget cells or tissues. Off-target screening methods with a scalable capability and with orders of magnitude greater sensitivity than existing methods would dramatically improve potential therapeutic models. More comprehensive functional assays will elucidate the relevance and impact of a given off-target effect, which can vary significantly depending on genome site, cell or tissue context, and even stage of development. Sensitive and accurate measurement of off-target activity, and ultimately function, will enable improvements to genome editing systems and help define the need for upstream mitigation measures such as the careful selection of gRNAs and an appropriate gene editor variant for the task at hand. Current approaches for unbiased, genome-wide measurement of DSBs include Genome-wide Unbiased Identifications of DSBs Evaluated by Sequencing (GUIDE-seq),4 High-Throughput Genome-wide Translocation Sequencing (HTGTS),16 Digested genome sequencing (Digenome-seq),46 and Breaks Labeling Enrichments on Streptavidin and next generation Sequencing (BLESS).47 More recent tools such as the circularization for in vitro reporting of cleavage effects by sequencing (CIRCLE-seq)48 and selective enrichment and identification of tagged genomic DNA ends by sequencing (SITE-seq)49 have further improved off-target detection and analysis. Despite these varied approaches to measuring offtarget effects, accurately predicting the off-target cleavage sites,
■
CONTROLLABLE GENOME EDITING The ability to control the activity of genome editors in a predictable and stringent manner will facilitate advances toward in vivo therapeutic applications. In contrast to ex vivo and in vitro uses of genome editors, where undesirable outcomes can largely be screened and discarded, in vivo uses will require the highest standards of performance and lowest tolerance for error. The development of controllers of genome editing activity has been approached through various strategies that provide some ability to define the temporal and spatial parameters conducive to genome editor activity. To date, these strategies have focused primarily on transcriptional controls for inducible expression of the editors, direct engineering of the enzymes or gRNAs to be responsive to stimuli, and discovery of molecules that can inhibit function of the editing complex. Post-Translational Control. The state of the art for genome editing control has focused on engineering posttranslational control strategies in which Cas9 is modified to enable control through the introduction of a stimulus. Unlike regulation of transcriptional control, post-translational control offers a higher resolution of user-defined control and less reliance on inherent cellular processes such as transcription, translation, and protein degradation. Control of Cas9 has been demonstrated using light,52 temperature,53 intein ligand dependency,54,55 dimerization dependency,55−58 destabilization domains,59,60 and domain replacement61 (Figure 2a). In addition to engineering Cas9, these strategies (excluding light and temperature) rely on small molecule induction. These chemical controllers (e.g., rapamycin, trimethoprim, and tamoxifen) have provided the foundation for chemically mediated post-translational control in vivo. However, implementation of these control systems in vivo will require medicinal chemistry optimizations or discovery of novel compounds to improve binding affinities, pharmacokinetic properties, and toxicological profiles. These challenges represent opportunities for modern chemistry to make advances in genome editing capabilities. Control through gRNA Engineering. In addition to modulating Cas9 protein, gRNA engineering has been actively pursued to improve genome editing control (Figure 2b and c). Beginning with the original fusion of Cas9 RNA components to form the sgRNA,12 gRNA development has expanded to include gRNA truncations to decrease off-target effects62 and create synthetic circuits.63 RNA hairpin engineering has been shown to modulate transcription by recruiting RNA-binding transcriptional modifiers such as VP64,64−66 KRAB,65 or PUFeffector fusion proteins.67 5′ and 3′ chemically modified gRNAs can increase gRNA stability and improve genome editing.68,69 Small molecule control has been engineered into gRNA sequences appended with ligand-activated self-cleaving apta336
DOI: 10.1021/acschembio.7b00689 ACS Chem. Biol. 2018, 13, 333−342
Perspective
ACS Chemical Biology zymes such that RNP activity is ligand-dependent.70 RNA aptamers can bind a variety of ligands, either engineered or natural.71 Using these systems, it is conceivable that complex and fine-tuned control of genome editing activity can be combinatorially arranged to develop tools ranging from more precise therapeutics to complex logic gates.72 Small Molecule-Mediated Control. Small molecules and macromolecular chemistry can be leveraged at multiple stages of genome editor assembly and DNA targeting to impart control over activity (Figure 2c). These molecules would be categorized based on their mechanism of action, including inhibition of DNA or RNA binding, DNA cleavage, or conformational changes in the enzyme itself. For example, inhibition of Cas9 has been demonstrated with bacteriophageencoded anti-CRISPR proteins that act by blocking dsDNA binding and cleavage.73−75 These findings demonstrate that RNA-guided nucleases can be targets for small protein inhibitors; however, a proof of concept that small molecules can be applied to specifically inhibit (or alternatively enhance) Cas9 enzyme activity has so far not been demonstrated. Practical or therapeutic use of anti-CRISPR proteins is subject to the limitations of Cas9 use itself; namely, expression, delivery, and immunogenicity. Small molecules can largely surmount these hurdles given their ability to transverse cell membranes and avoid immune detection. Furthermore, there is a unique opportunity for small molecules not only to prevent or fully suppress gene editing activity but also to fine-tune the window of gene editor activity to enable high on-target editing while limiting or entirely preventing off-target effects. Similar to the state of the art in drug discovery, high-throughput screening of small molecule libraries in conjunction with structure optimization may yield molecules capable of high-resolution temporal and dosedependent inhibition of genome editors. Iterative screening would enable the identification of a range of molecules that display unique activities including broad specificity against classes of enzymes (e.g., RNA-guided nucleases) or more specific molecules that target a single nuclease (e.g., SaCas9). A large portfolio of inhibitory or modulatory compounds would enable customizable control and expand genome editing capabilities for basic research and practical applications. This highlights that chemical biology will serve a significant role in pursuing the diversity and scale of opportunities in controlling genome editors.
Figure 3. Genome editor delivery strategies for in vivo applications using RNA, DNA, or RNP. Delivery of gene editors ranges across four primary methods: (A) direct nucleic acid delivery using mechanical methods (e.g., electroporation, sonoporation, hydrodynamic injection); (B) viral vector-based delivery methods (e.g., adeno-associated virus); (C) lipid-based delivery and uptake via endocytosis or micropinocytosis (depicted); and (D) chemically mediated delivery based on polymer carriers depicting update via endocytosis (depicted) or micropinocytosis. *Cargo delivered by viral vector, lipid-based systems, and polymer-based systems can be RNAs, DNAs, or RNPs.
short-lived, transient delivery formulations; other applications, such as CRISPR activators or repressors that may need to occupy genome positions for longer durations of time, will require longer-lived formulations. Given the limited efficacy of current gene editing tools, it is also likely that repeated administration will be required for therapeutic benefit, introducing significant immune challenges. Unique Delivery Cargo. Biologics, such as Cas9, therapeutic proteins, and nucleic acids, often suffer from poor pharmacokinetic and pharmacodynamics (PK/PD) properties due to their rapid degradation by serum proteases and nucleases. To date, advances in macromolecule delivery are largely attributed to the increased use of nucleic acid-based therapeutics such as siRNA and DNA/RNA vaccines.77−79 However, CRISPR/Cas9 technology possesses several fundamental differences as compared to currently utilized nucleic acid and protein technology that will require a new set of delivery tools for its widespread adaptation for medical applications. For example, the bacterial origin of Cas9 and other genome editors may lead to rapid immune clearance, thereby impairing desired PK/PD properties. Through subcellular localization, proteins or nucleic acids, such as Cas9, can be blocked from cellular uptake and are often trapped inside the endosome following endocytosis.80 An additional consideration for the delivery of CRISPR/Cas9 constructs is that applications of genome editors in therapy often include only a minimal number of targets inside a cell where transient expression is highly desirable to enable enough time to convert a target, while minimizing the window of time within which offtarget effects may occur. For instance, lentiviral-delivered self-
■
TRANSLATION TO CLINICAL APPLICATIONS Genome editing technologies for ex vivo applications3 or xenotransplantation76 are already starting to, or have the potential to, address unmet clinical needs. However, achieving the goal of applying genome editors in patients for medical applications that do not lend themselves to correction through ex vivo therapies will rely largely on the ability to safely and effectively deliver the desired construct in a targeted, and likely transient, manner. To fully realize the potential of genome editors for therapeutic applications, new delivery mechanisms and formulations must be developed that allow for their safe and effective use in vivo. The development of delivery modalities for gene editors presents unique challenges and opportunities, including the possibility of delivering the editor as a nucleic acid or RNP (see Figure 3), and applications for both short- and long-lived formulations. For example, some applications, such as targeting gene sequences for correction, introduction, or removal of an epigenetic mark, will require 337
DOI: 10.1021/acschembio.7b00689 ACS Chem. Biol. 2018, 13, 333−342
Perspective
ACS Chemical Biology targeting “hit and go” Cas9 constructs, which include gRNAs that target Cas9 directly, have been shown to decrease Cas9 protein half-life and reduce off-target effects.81 This strategy may in fact be useful where initial high editing efficiency is not necessary for a desired outcome due to growth advantages conferred by an introduced allele.82 The need for transient delivery of genome editors for some applications presents an advantage in that many of the currently employed delivery methods have a short half-life. This has hindered their use in other nucleic acid applications, such as vaccines or siRNA but may prove beneficial for genome editors where short-duration delivery formulations may be a design feature. Viral Vectors. Approaches for the delivery of genome editors to date have relied on the standard delivery toolbox, including viral vectors, nucleic acids, and packaged ribonucleoproteins (RNPs; Figure 3a,b). A large majority of current approaches focus on the use of viral vectors, including lentiviral and adeno-associated virus (AAV;83−85 Figure 3b). While viral vectors can readily facilitate targeted cellular entry through viral tropism and can produce quantities of editors sufficient for therapeutic applications, they often face many challenges. In particular, viral packaging size limits represent a significant barrier and often necessitate the cotransfection of CRISPR components (e.g., AAV-split-Cas986) to reconstitute a fulllength Cas9 RNP in vivo, thereby reducing the probability of a desired gene editing outcome. While alternative smaller genome editors and modified systems may alleviate concerns surrounding packaging size, viral vectors are still hindered by the presence of pre-existing immunity to the vector or by persistent expression of cargo. While increased longevity of expression is advantageous for other therapeutic modalities, the need for transient expression to limit off-target effects by genome editors largely restricts the delivery of such constructs via viral vectors and limits their utility for widespread in vivo applications in healthy subjects.83,85,87,88 Chemical Clinical Formulations. Chemical approaches to delivery offer advantages over viral vectors in that they do not possess restrictions on packaging size, and there is no preexisting immunity in patients that would render the delivery modality less effective. Additionally, manufacturing of chemical formulations has the potential for scale-up and rapid response that is not currently afforded by viral vectors. Genome editors represent a particularly interesting challenge and opportunity for chemists focused on facilitating macromolecular delivery; namely, the fact that editors can be delivered as RNPs, as encoded DNA, or as mRNA constructs. This flexibility allows for novel approaches to delivery that are not afforded by the limited size and functionality constraints inherent to current RNA and DNA delivery technologies. The delivery of CRISPR/Cas9 or other genome editors via plasmid or mini-circle DNA has been demonstrated in proof-ofconcept in vivo animal experiments.82 DNA delivery has largely depended upon mechanical techniques, such as electroporation, sonoporation, or hydrodynamic injection, to ensure delivery into the nucleus of target cells89,90 (Figure 3a). While this technology possesses several advantages over viral vectors, including essentially no limitation on packaging size and a simplified manufacturing pathway, there are fundamental barriers to the use of plasmid DNA in therapeutic settings. For example, plasmid DNA, while not integrating, does provide extended expression of the encoded constructs, which may lead to off-target effects. Additionally, the use of an electroporation device in vivo often results in cellular damage. Finally, delivery
of plasmid DNA via electroporation or other mechanical techniques in a targeted manner to specific cells and tissues is difficult, again increasing the likelihood of off-target effects. The chemical delivery of CRISPR/Cas9 as RNA or RNP constructs may help ensure transient, targeted expression of genome editors in vivo. Therapeutic applications of siRNA and nucleic acid vaccines spurred the development of modern chemical delivery formulations, such as lipid nanoparticle (LNP) and pluronics polymers, for the transport of siRNA, mRNA, and replicating RNA into the cytoplasm (Figure 3c, d). LNP formulations are often composed of lipids containing ionizable amines, which not only result in cellular uptake through endocytosis or micropinocytosis, but also facilitate endosomal escape through a variety of poorly understood mechanisms.91−94 While advances in macromolecular delivery have facilitated the adaptation of nucleic acid-based technologies, there are still several barriers to their widespread use for the delivery of CRISPR/Cas9 constructs.88 Currently available chemical formulations often result in induced toxicity and immunogenicity, including the development of antidrug antibodies against the expressed protein, resulting in suboptimal PK/PD qualities. An additional consideration is the use of genome editors in healthy subjects, where much lower levels of immunogenicity and toxicity can be tolerated. Cas9 has been shown to evoke cellular and humoral immune responses in wildtype mice.86 Pre-existing immunity to nucleases found in the microbiome (S. pyogenes and S. aureus) may exist. Therefore, immune responses may be a significant limiting factor for therapeutic uses where (a) the rapid clearance of the Cas9 may lower the the half-life threshold needed for transient activity in vivo and (b) adaptive immunity against Cas9 would reduce the efficacy of repeated treatments. Three distinct methods could be employed to alleviate the immune response: immune-silent delivery modalities enabling editor entry into cells without triggering immune clearance, “humanization” of the editor proteins to prevent immune response, or induction of immune tolerance prior to administration of the therapy.95 Lipid and polymer toxicity is often alleviated or minimized by the use of biodegradable formulations with readily degraded ester or amide bonds.96,97 While these formulations have been successful at limiting some of the associated toxicity, there is much work to be done to ensure safe and effective delivery in healthy subjects, including the synthesis and screening of biodegradable polymers and lipids with an even safer toxicity profile. The advent of new targeting modalities, such as the small peptide targeting groups discovered using in vivo phage display98 or the use of lipids with tropisms for specific tissues and organs, coupled with biodegradable polymers and lipids, can increase specificity while decreasing toxicity. The identification, synthesis, and testing of such modalities will require advances in the coming years to ensure the greater adaptation of genome editing tools for in vivo use.
■
ETHICS AND THE FUTURE OF THERAPEUTIC GENOME EDITING To ensure a bright future for new interventions that utilize genome editing technologies, the development of strategies that sufficiently address the relevant legal, ethical, environmental, dual-use, and responsible innovation (LEEDR) concerns of genome editing are as important as technological advances. Despite the logarithmic growth of the CRISPR-mediated genome editing field, clinical therapeutic interventions using these tools are still in the early phases of development and still 338
DOI: 10.1021/acschembio.7b00689 ACS Chem. Biol. 2018, 13, 333−342
Perspective
ACS Chemical Biology
Figure 4. End-to-end gene editing platform. A comprehensive gene editing platform for clinical applications requires advances in delivery (left), inhibitors of activity and design rules for editors (middle), and controls for DNA repair (right). Improving these technologies will enable more effective and safer in vivo gene editing therapeutics.
gies can be used to explore more ambitious goals of treatment of complex multigenic diseases such as neurodegenerative disorders, scalable production of delivery methods that may require significant chemical and process engineering improvements, and development of novel in silico systems to help predict next-generation genome editor functionality and process large bioinformatics data sets. Given that the rate of progress in genome editing is rapid and current technical hurdles will likely be quickly surmounted, there are and will be many opportunities for chemical biology to contribute to advancing safe, ethical, and responsible use of current and future tools.
involve laboratory manipulations that are difficult, timeconsuming, and expensive. Overcoming these technical challenges will take time, and during this nascent stage of development, it is imperative to lay a foundation that also comprehensively addresses these unique LEEDR challenges and establishes best practices for the safe use of genome editing tools, rather than apply patches and fixes ex post facto. Early LEEDR solutions will not only delineate the standards for the responsible use of genome editing therapies but also drive the field toward positive, beneficial human applications. Efforts to address concerns associated with the clinical use (or misuse) of these tools, including the recent Human Genome Editing study conducted by the National Academy of Sciences and Medicine99 and recommendations published by professional societies such as the American College of Medical Genetics and Genomics,100 are helpful starting points to open the dialogue on LEEDR topics and establish best practices. For example, there is general agreement that given the premature nature of clinical demonstration of the tools and significant ethical concerns, human germline editing should not be attempted in the near term, or possibly ever. Today, engagement with stakeholders representing patient advocacy groups, medical professionals, government regulators, and industry is essential not only to educate them on the opportunities and risks of genome editing tools but also, importantly, to guide technology development with thoughtful consideration of the spectrum of those who might be impacted. Through development of technologies such as anti-CRISPR small molecules or tissue-specific delivery systems, researchers and innovators help define the risks and opportunities of genome editing at the earliest inception of these tools. The pursuit of safe and effective therapeutic genome editing tools is built upon a deep understanding of the fundamental cellular and molecular processes underlying genome editing, the ability to navigate or control those processes to avoid detrimental outcomes and ensure highly specific editing, and accurate delivery of genome editing tools for full therapeutic benefit (Figure 4). Ultimately, these technological breakthroughs, amplified by effective LEEDR development, will bridge the gap to future therapeutic applications. Advances in these areas will also provide the foundation from which genome editing technolo-
■
AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. ORCID
Andrew H. Lee: 0000-0003-0963-423X Notes
The authors declare the following competing financial interest(s): A.H.L. and A.E.C. are employees of Booz Allen Hamilton. A.L.J. is an employee of Schafer, a Belcan Company. C.D.S. is an employee of Quantitative Scientific Solutions.
■
ACKNOWLEDGMENTS The authors thank T. Kilbride (Spire Communications) for critical reading of the manuscript. The opinions expressed herein belong solely to the authors. They do not represent and should not be interpreted as being those of or endorsed by the Defense Advanced Research Projects Agency, the Department of Defense, or any other branch of the federal government.
■
REFERENCES
(1) Ren, J., Liu, X., Fang, C., Jiang, S., June, C. H., and Zhao, Y. (2017) Multiplex genome editing to generate universal CAR T cells resistant to PD1 inhibition. Clin. Cancer Res. 23, 2255−2266. (2) DeWitt, M. A., Magis, W., Bray, N. L., Wang, T., Berman, J. R., Urbinati, F., Heo, S.-J., Mitros, T., Muñoz, D. P., Boffelli, D., Kohn, D. B., Walters, M. C., Carroll, D., Martin, D. I. K., and Corn, J. E. (2016) Selection-free genome editing of the sickle mutation in human adult hematopoietic stem/progenitor cells. Sci. Transl. Med. 8, 360ra134− 360ra134.
339
DOI: 10.1021/acschembio.7b00689 ACS Chem. Biol. 2018, 13, 333−342
Perspective
ACS Chemical Biology (3) Tebas, P., Stein, D., Tang, W. W., Frank, I., Wang, S. Q., Lee, G., Spratt, S. K., Surosky, R. T., Giedlin, M. A., Nichol, G., Holmes, M. C., Gregory, P. D., Ando, D. G., Kalos, M., Collman, R. G., Binder-Scholl, G., Plesa, G., Hwang, W.-T., Levine, B. L., and June, C. H. (2014) Gene editing of CCR5 in autologous CD4 T cells of persons infected with HIV. N. Engl. J. Med. 370, 901−910. (4) Tsai, S. Q., Zheng, Z., Nguyen, N. T., Liebers, M., Topkar, V. V., Thapar, V., Wyvekens, N., Khayter, C., Iafrate, A. J., Le, L. P., Aryee, M. J., and Joung, J. K. (2014) GUIDE-seq enables genome-wide profiling of off-target cleavage by CRISPR-Cas nucleases. Nat. Biotechnol. 33, 187−197. (5) Chu, V. T., Weber, T., Wefers, B., Wurst, W., Sander, S., Rajewsky, K., and Kühn, R. (2015) Increasing the efficiency of homology-directed repair for CRISPR-Cas9-induced precise gene editing in mammalian cells. Nat. Biotechnol. 33, 543−548. (6) Miyaoka, Y., Berman, J. R., Cooper, S. B., Mayerl, S. J., Chan, A. H., Zhang, B., Karlin-Neumann, G. A., and Conklin, B. R. (2016) Systematic quantification of HDR and NHEJ reveals effects of locus, nuclease, and cell type on genome-editing. Sci. Rep. 6, 23549. (7) Ran, F. A., Cong, L., Yan, W. X., Scott, D. a., Gootenberg, J. S., Kriz, A. J., Zetsche, B., Shalem, O., Wu, X., Makarova, K. S., Koonin, E. V., Sharp, P. a., and Zhang, F. (2015) In vivo genome editing using Staphylococcus aureus Cas9. Nature 520, 186−190. (8) Ramanan, V., Shlomai, A., Cox, D. B. T., Schwartz, R. E., Michailidis, E., Bhatta, A., Scott, D. A., Zhang, F., Rice, C. M., and Bhatia, S. N. (2015) CRISPR/Cas9 cleavage of viral DNA efficiently suppresses hepatitis B virus. Sci. Rep. 5, 10833. (9) Bikard, D., Euler, C. W., Jiang, W., Nussenzweig, P. M., Goldberg, G. W., Duportet, X., Fischetti, V. A., and Marraffini, L. A. (2014) Exploiting CRISPR-Cas nucleases to produce sequence-specific antimicrobials. Nat. Biotechnol. 32, 1146−1150. (10) Sternberg, S. H., Redding, S., Jinek, M., Greene, E. C., and Doudna, J. A. (2014) DNA interrogation by the CRISPR RNA-guided endonuclease Cas9. Nature 507, 62−67. (11) Symington, L. S., and Gautier, J. (2011) Double-Strand Break End Resection and Repair Pathway Choice. Annu. Rev. Genet. 45, 247− 271. (12) Jinek, M., Chylinski, K., Fonfara, I., Hauer, M., Doudna, J. A., and Charpentier, E. (2012) A Programmable Dual-RNA − Guided DNA Endonuclease in Adaptice Bacterial Immunity. Science 337, 816− 822. (13) Ran, F. A., Hsu, P. D., Lin, C. Y., Gootenberg, J. S., Konermann, S., Trevino, A. E., Scott, D. A., Inoue, A., Matoba, S., Zhang, Y., and Zhang, F. (2013) Double nicking by RNA-guided CRISPR cas9 for enhanced genome editing specificity. Cell 154, 1380−1389. (14) Guilinger, J. P., Thompson, D. B., and Liu, D. R. (2014) Fusion of catalytically inactive Cas9 to FokI nuclease improves the specificity of genome modification. Nat. Biotechnol. 32, 577−582. (15) Bothmer, A., Phadke, T., Barrera, L. A., Margulies, C. M., Lee, C. S., Buquicchio, F., Moss, S., Abdulkerim, H. S., Selleck, W., Jayaram, H., Myer, V. E., and Cotta-Ramusino, C. (2017) Characterization of the interplay between DNA repair and CRISPR/Cas9-induced DNA lesions at an endogenous locus. Nat. Commun. 8, 13905. (16) Frock, R. L., Hu, J., Meyers, R. M., Ho, Y.-J., Kii, E., and Alt, F. W. (2014) Genome-wide detection of DNA double-stranded breaks induced by engineered nucleases. Nat. Biotechnol. 33, 179−186. (17) Qi, L. S., Larson, M. H., Gilbert, L. A., Doudna, J. A., Weissman, J. S., Arkin, A. P., and Lim, W. A. (2013) Repurposing CRISPR as an RNA-Guided Platform for Sequence-Specific Control of Gene Expression NIH Public Access. Cell 152, 1173−1183. (18) Komor, A. C., Kim, Y. B., Packer, M. S., Zuris, J. A., and Liu, D. R. (2016) Programmable editing of a target base in genomic DNA without double-stranded DNA cleavage. Nature 533, 420−424. (19) Hilton, I. B., D’Ippolito, A. M., Vockley, C. M., Thakore, P. I., Crawford, G. E., Reddy, T. E., and Gersbach, C. A. (2015) Epigenome editing by a CRISPR-Cas9-based acetyltransferase activates genes from promoters and enhancers. Nat. Biotechnol. 33, 510−517.
(20) Xu, X., Tao, Y., Gao, X., Zhang, L., Li, X., Zou, W., Ruan, K., Wang, F., Xu, G.-L., and Hu, R. (2016) A CRISPR-based approach for targeted DNA demethylation. Cell Discovery 2, 16009. (21) McDonald, J. I., Celik, H., Rois, L. E., Fishberger, G., Fowler, T., Rees, R., Kramer, A., Martens, A., Edwards, J. R., and Challen, G. A. (2016) Reprogrammable CRISPR/Cas9-based system for inducing site-specific DNA methylation. Biol. Open 5, 866−874. (22) Kearns, N. a, Pham, H., Tabak, B., Genga, R. M., Silverstein, N. J., Garber, M., and Maehr, R. (2015) Functional annotation of native enhancers with a Cas9-histone demethylase fusion. Nat. Methods 12, 401−403. (23) Lei, Y., Zhang, X., Su, J., Jeong, M., Gundry, M. C., Huang, Y.H., Zhou, Y., Li, W., and Goodell, M. A. (2017) Targeted DNA methylation in vivo using an engineered dCas9-MQ1 fusion protein. Nat. Commun. 8, 16026. (24) Mali, P., Aach, J., Stranges, P. B., Esvelt, K. M., Moosburner, M., Kosuri, S., Yang, L., and Church, G. M. (2013) CAS9 transcriptional activators for target specificity screening and paired nickases for cooperative genome engineering. Nat. Biotechnol. 31, 833−838. (25) Chavez, A., Scheiman, J., Vora, S., Pruitt, B. W., Tuttle, M., P R Iyer, E., Lin, S., Kiani, S., Guzman, C. D., Wiegand, D. J., TerOvanesyan, D., Braff, J. L., Davidsohn, N., Housden, B. E., Perrimon, N., Weiss, R., Aach, J., Collins, J. J., and Church, G. M. (2015) Highly efficient Cas9-mediated transcriptional programming. Nat. Methods 12, 326−328. (26) Thakore, P. I., Song, L., Safi, A., Shivakumar, K., Kabadi, A. M., Reddy, T. E., Crawford, G. E., Gersbach, C. A., and D'Ippolito, A. M. (2015) Highly Specific Epigenome Editing by CRISPR/Cas9 Repressors for Silencing of Distal Regulatory Elements. Nat. Methods 12, 1143−1149. (27) Gilbert, L. A., Horlbeck, M. A., Adamson, B., Villalta, J. E., Chen, Y., Whitehead, E. H., Guimaraes, C., Panning, B., Ploegh, H. L., Bassik, M. C., Qi, L. S., Kampmann, M., and Weissman, J. S. (2014) GenomeScale CRISPR-Mediated Control of Gene Repression and Activation. Cell 159, 647−661. (28) Abudayyeh, O. O., Gootenberg, J. S., Konermann, S., Joung, J., Slaymaker, I. M., Cox, D. B. T., Shmakov, S., Makarova, K. S., Semenova, E., Minakhin, L., Severinov, K., Regev, A., Lander, E. S., Koonin, E. V., and Zhang, F. (2016) C2c2 is a single-component programmable RNA-guided RNA-targeting CRISPR effector. Science 353, aaf5573. (29) East-Seletsky, A., O’Connell, M. R., Knight, S. C., Burstein, D., Cate, J. H. D., Tjian, R., and Doudna, J. A. (2016) Two distinct RNase activities of CRISPR-C2c2 enable guide-RNA processing and RNA detection. Nature 538, 270−273. (30) Yang, H., Wang, H., Shivalila, C. S., Cheng, A. W., Shi, L., and Jaenisch, R. (2013) One-step generation of mice carrying reporter and conditional alleles by CRISPR/cas-mediated genome engineering. Cell 154, 1370−1379. (31) Wang, H., Yang, H., Shivalila, C. S., Dawlaty, M. M., Cheng, A. W., Zhang, F., and Jaenisch, R. (2013) One-step generation of mice carrying mutations in multiple genes by CRISPR/cas-mediated genome engineering. Cell 153, 910−918. (32) Mali, P., Yang, L., Esvelt, K. M., Aach, J., Guell, M., DiCarlo, J. E., Norville, J. E., and Church, G. M. (2013) RNA-guided human genome engineering via Cas9. Science 339, 823−826. (33) Mao, Z., Jiang, Y., Liu, X., Seluanov, A., and Gorbunova, V. (2009) DNA Repair by Homologous Recombination, But Not by Nonhomologous End Joining, Is Elevated in Breast Cancer Cells. Neoplasia 11, 683−691. (34) Yang, D., Scavuzzo, M. A., Chmielowiec, J., Sharp, R., Bajic, A., and Borowiak, M. (2016) Enrichment of G2/M cell cycle phase in human pluripotent stem cells enhances HDR-mediated gene repair with customizable endonucleases. Sci. Rep. 6, 21264. (35) Lin, S., Staahl, B. T., Alla, R. K., and Doudna, J. A. (2014) Enhanced homology-directed human genome engineering by controlled timing of CRISPR/Cas9 delivery. eLife 3, e04766. 340
DOI: 10.1021/acschembio.7b00689 ACS Chem. Biol. 2018, 13, 333−342
Perspective
ACS Chemical Biology (36) Song, J., Yang, D., Xu, J., Zhu, T., Chen, Y. E., and Zhang, J. (2016) RS-1 enhances CRISPR/Cas9- and TALEN-mediated knockin efficiency. Nat. Commun. 7, 10548. (37) Maruyama, T., Dougan, S. K., Truttmann, M. C., Bilate, A. M., Ingram, J. R., and Ploegh, H. L. (2015) Increasing the efficiency of precise genome editing with CRISPR-Cas9 by inhibition of nonhomologous end joining. Nat. Biotechnol. 33, 538−542. (38) Pinder, J., Salsman, J., and Dellaire, G. (2015) Nuclear domain “knock-in” screen for the evaluation and identification of small molecule enhancers of CRISPR-based genome editing. Nucleic Acids Res. 43, 9379−9392. (39) Yu, C., Liu, Y., Ma, T., Liu, K., Xu, S., Zhang, Y., Liu, H., La Russa, M., Xie, M., Ding, S., and Qi, L. S. (2015) Small molecules enhance crispr genome editing in pluripotent stem cells. Cell Stem Cell 16, 142−147. (40) Gutschner, T., Haemmerle, M., Genovese, G., Draetta, G. F., and Chin, L. (2016) Post-translational Regulation of Cas9 during G1 Enhances Homology-Directed Repair. Cell Rep. 14, 1555−1566. (41) Zetsche, B., Gootenberg, J. S., Abudayyeh, O. O., Slaymaker, I. M., Makarova, K. S., Essletzbichler, P., Volz, S. E., Joung, J., Van Der Oost, J., Regev, A., Koonin, E. V., and Zhang, F. (2015) Cpf1 Is a Single RNA-Guided Endonuclease of a Class 2 CRISPR-Cas System. Cell 163, 759−771. (42) Sakuma, T., Nakade, S., Sakane, Y., Suzuki, K.-I. T., and Yamamoto, T. (2015) MMEJ-assisted gene knock-in using TALENs and CRISPR-Cas9 with the PITCh systems. Nat. Protoc. 11, 118−133. (43) Yao, X., Wang, X., Liu, J., Hu, X., Shi, L., Shen, X., Ying, W., Sun, X., Wang, X., Huang, P., and Yang, H. (2017) CRISPR/Cas9 − Mediated Precise Targeted Integration In Vivo Using a Double Cut Donor with Short Homology Arms. EBioMedicine 20, 19−26. (44) Yao, X., Wang, X., Hu, X., Liu, Z., Liu, J., Zhou, H., Shen, X., Wei, Y., Huang, Z., Ying, W., Wang, Y., Nie, Y.-H., Zhang, C.-C., Li, S., Cheng, L., Wang, Q., Wu, Y., Huang, P., Sun, Q., Shi, L., and Yang, H. (2017) Homology-mediated end joining-based targeted integration using CRISPR/Cas9. Cell Res. 27, 801. (45) Paix, A., Folkmann, A., Goldman, D. H., Kulaga, H., Rasoloson, D., Paidemarry, S., Green, R., and Seydoux, G. (2017) Precision genome editing using synthesis-dependent repair of Cas9-induced DNA breaks. bioRxiv, DOI: 10.1101/161109. (46) Kim, D., Bae, S., Park, J., Kim, E., Kim, S., Yu, H. R., Hwang, J., Kim, J.-I., and Kim, J.-S. (2015) Digenome-seq: genome-wide profiling of CRISPR-Cas9 off-target effects in human cells. Nat. Methods 12, 237−243. (47) Crosetto, N., Mitra, A., Silva, M. J., Bienko, M., Dojer, N., Wang, Q., Karaca, E., Chiarle, R., Skrzypczak, M., Ginalski, K., Pasero, P., Rowicka, M., and Dikic, I. (2013) Nucleotide-resolution DNA doublestrand break mapping by next-generation sequencing. Nat. Methods 10, 361−365. (48) Tsai, S. Q., Nguyen, N. T., Malagon-Lopez, J., Topkar, V. V., Aryee, M. J., and Joung, J. K. (2017) CIRCLE-seq: a highly sensitive in vitro screen for genome-wide CRISPR−Cas9 nuclease off-targets. Nat. Methods 14, 607. (49) Cameron, P., Fuller, C. K., Donohoue, P. D., Jones, B. N., Thompson, M. S., Carter, M. M., Gradia, S., Vidal, B., Garner, E., Slorach, E. M., Lau, E., Banh, L. M., Lied, A. M., Edwards, L. S., Settle, A. H., Capurso, D., Llaca, V., Deschamps, S., Cigan, M., Young, J. K., and May, A. P. (2017) Mapping the genomic landscape of CRISPR− Cas9 cleavage. Nat. Methods 14, 600−606. (50) Schaefer, K. A., Wu, W.-H., Colgan, D. F., Tsang, S. H., Bassuk, A. G., and Mahajan, V. B. (2017) Unexpected mutations after CRISPR−Cas9 editing in vivo. Nat. Methods 14, 547−548. (51) Lareau, C. A., Clement, K., Hsu, J. Y., Pattanayak, V., and Keith, J. (2017) Unexpected mutations after CRISPR-Cas9 editing in vivo” are most likely pre- existing sequence variants and not nucleaseinduced mutations. bioRxiv, DOI: 10.1101/159707. (52) Nihongaki, Y., Kawano, F., Nakajima, T., and Sato, M. (2015) Photoactivatable CRISPR-Cas9 for optogenetic genome editing. Nat. Biotechnol. 33, 755−760.
(53) Richter, F., Fonfara, I., Bouazza, B., Schumacher, C. H., Bratovič, M., Charpentier, E., and Mö glich, A. (2016) Engineering of temperature- and light-switchable Cas9 variants. Nucleic Acids Res. 44, 10003−10014. (54) Davis, K. M., Pattanayak, V., Thompson, D. B., Zuris, J. A., and Liu, D. R. (2015) Small molecule−triggered Cas9 protein with improved genome-editing specificity. Nat. Chem. Biol. 11, 316−318. (55) Truong, D. J. J., Kühner, K., Kühn, R., Werfel, S., Engelhardt, S., Wurst, W., and Ortiz, O. (2015) Development of an intein-mediated split-Cas9 system for gene therapy. Nucleic Acids Res. 43, 6450−6458. (56) Zetsche, B., Volz, S. E., and Zhang, F. (2015) A split-Cas9 architecture for inducible genome editing and transcription modulation. Nat. Biotechnol. 33, 139−142. (57) Nguyen, D. P., Miyaoka, Y., Gilbert, L. A., Mayerl, S. J., Lee, B. H., Weissman, J. S., Conklin, B. R., and Wells, J. A. (2016) Ligandbinding domains of nuclear receptors facilitate tight control of split CRISPR activity. Nat. Commun. 7, 12009. (58) Oakes, B. L., Nadler, D. C., Flamholz, A., Fellmann, C., Staahl, B. T., Doudna, J. A., and Savage, D. F. (2016) Profiling of engineering hotspots identifies an allosteric CRISPR-Cas9 switch. Nat. Biotechnol. 34, 646−651. (59) Senturk, S., Shirole, N. H., Nowak, D. G., Corbo, V., Pal, D., Vaughan, A., Tuveson, D. A., Trotman, L. C., Kinney, J. B., and Sordella, R. (2017) Rapid and tunable method to temporally control gene editing based on conditional Cas9 stabilization. Nat. Commun. 8, 14370. (60) Maji, B., Moore, C. L., Zetsche, B., Volz, S. E., Zhang, F., Shoulders, M. D., and Choudhary, A. (2016) Multidimensional chemical control of CRISPR−Cas9. Nat. Chem. Biol. 13, 9−11. (61) Rose, J. C., Stephany, J. J., Valente, W. J., Trevillian, B. M., Dang, H. V., Bielas, J. H., Maly, D. J., and Fowler, D. M. (2017) Rapidly inducible Cas9 and DSB-ddPCR to probe editing kinetics. Nat. Methods 14, 891. (62) Fu, Y., Sander, J. D., Reyon, D., Cascio, V. M., and Joung, J. K. (2014) Improving CRISPR-Cas nuclease specificity using truncated guide RNAs. Nat. Biotechnol. 32, 279−284. (63) Kiani, S., Chavez, A., Tuttle, M., Hall, R. N., Chari, R., TerOvanesyan, D., Qian, J., Pruitt, B. W., Beal, J., Vora, S., Buchthal, J., Kowal, E. J. K., Ebrahimkhani, M. R., Collins, J. J., Weiss, R., and Church, G. (2015) Cas9 gRNA engineering for genome editing, activation and repression. Nat. Methods 12, 1051−1054. (64) Konermann, S., Brigham, M. D., Trevino, A. E., Joung, J., Abudayyeh, O. O., Barcena, C., Hsu, P. D., Habib, N., Gootenberg, J. S., Nishimasu, H., Nureki, O., and Zhang, F. (2014) Genome-scale transcriptional activation by an engineered CRISPR-Cas9 complex. Nature 517, 583−588. (65) Zalatan, J. G., Lee, M. E., Almeida, R., Gilbert, L. A., Whitehead, E. H., La Russa, M., Tsai, J. C., Weissman, J. S., Dueber, J. E., Qi, L. S., and Lim, W. A. (2015) Engineering complex synthetic transcriptional programs with CRISPR RNA scaffolds. Cell 160, 339−350. (66) Shechner, D. M., Hacisuleyman, E., Younger, S. T., and Rinn, J. L. (2015) Multiplexable, locus-specific targeting of long RNAs with CRISPR-Display. Nat. Methods 12, 664−670. (67) Cheng, A. W., Jillette, N., Lee, P., Plaskon, D., Fujiwara, Y., Wang, W., Taghbalout, A., and Wang, H. (2016) Casilio: a versatile CRISPR-Cas9-Pumilio hybrid for gene regulation and genomic labeling. Cell Res. 26, 254−257. (68) Hendel, A., Bak, R. O., Clark, J. T., Kennedy, A. B., Ryan, D. E., Roy, S., Steinfeld, I., Lunstad, B. D., Kaiser, R. J., Wilkens, A. B., Bacchetta, R., Tsalenko, A., Dellinger, D., Bruhn, L., and Porteus, M. H. (2015) Chemically modified guide RNAs enhance CRISPR-Cas genome editing in human primary cells. Nat. Biotechnol. 33, 985−989. (69) Dever, D. P., Bak, R. O., Reinisch, A., Camarena, J., Washington, G., Nicolas, C. E., Pavel-Dinu, M., Saxena, N., Wilkens, A. B., Mantri, S., Uchida, N., Hendel, A., Narla, A., Majeti, R., Weinberg, K. I., and Porteus, M. H. (2016) CRISPR/Cas9 β-globin gene targeting in human haematopoietic stem cells. Nature 539, 384−389. 341
DOI: 10.1021/acschembio.7b00689 ACS Chem. Biol. 2018, 13, 333−342
Perspective
ACS Chemical Biology (70) Tang, W., Hu, J. H., and Liu, D. R. (2017) Aptazyme-embedded guide RNAs enable ligand-responsive genome editing and transcriptional activation. Nat. Commun. 8, 15939. (71) Link, K. H., and Breaker, R. R. (2009) Engineering ligandresponsive gene-control elements: lessons learned from natural riboswitches. Gene Ther. 16, 1189−1201. (72) Gander, M. W., Vrana, J. D., Voje, W. E., Carothers, J. M., and Klavins, E. (2017) Digital logic circuits in yeast with CRISPR-dCas9 NOR gates. Nat. Commun. 8, 15459. (73) Bondy-Denomy, J., Garcia, B., Strum, S., Du, M., Rollins, M. F., Hidalgo-Reyes, Y., Wiedenheft, B., Maxwell, K. L., and Davidson, A. R. (2015) Multiple mechanisms for CRISPR−Cas inhibition by antiCRISPR proteins. Nature 526, 136−139. (74) Dong, D., Guo, M., Wang, S., Zhu, Y., Wang, S., Xiong, Z., Yang, J., Xu, Z., and Huang, Z. (2017) Structural basis of CRISPR−SpyCas9 inhibition by an anti-CRISPR protein. Nature 546, 436−439. (75) Chowdhury, S., Carter, J., Rollins, M. C. F., Golden, S. M., Jackson, R. N., Hoffmann, C., Nosaka, L., Bondy-Denomy, J., Maxwell, K. L., Davidson, A. R., Fischer, E. R., Lander, G. C., and Wiedenheft, B. (2017) Structure Reveals Mechanisms of Viral Suppressors that Intercept a CRISPR RNA-Guided Surveillance Complex. Cell 169, 47−57. (76) Niu, D., Wei, H.-J., Lin, L., George, H., Wang, T., Lee, I.-H., Zhao, H.-Y., Wang, Y., Kan, Y., Shrock, E., Lesha, E., Wang, G., Luo, Y., Qing, Y., Jiao, D., Zhao, H., Zhou, X., Wang, S., Wei, H., Güell, M., Church, G. M., and Yang, L. (2017) Inactivation of porcine endogenous retrovirus in pigs using CRISPR-Cas9. Science (Washington, DC, U. S.) 357, 1303. (77) Elliott, S. T., Kallewaard, N. L., Benjamin, E., Wachter-Rosati, L., McAuliffe, J. M., Patel, A., Smith, T. R., Schultheis, K., Park, D. H., Flingai, S., Wise, M. C., et al. (2017) DMAb inoculation of synthetic cross reactive antibodies protects against lethal influenza A and B infections. npj Vaccines 2, 18. (78) Flingai, S., Plummer, E. M., Patel, A., Shresta, S., Mendoza, J. M., Broderick, K. E., Sardesai, N. Y., Muthumani, K., and Weiner, D. B. (2015) Protection against dengue disease by synthetic nucleic acid antibody prophylaxis/immunotherapy. Sci. Rep. 5, 1−9. (79) Muthumani, K., Block, P., Flingai, S., Muruganantham, N., Chaaithanya, I. K., Tingey, C., Wise, M., Reuschel, E. L., Chung, C., Muthumani, A., Sarangan, G., Srikanth, P., Khan, A. S., Vijayachari, P., Sardesai, N. Y., Kim, J. J., Ugen, K. E., and Weiner, D. B. (2016) Rapid and Long-Term Immunity Elicited by DNA-Encoded Antibody Prophylaxis and DNA Vaccination Against Chikungunya Virus. J. Infect. Dis. 214, 369−378. (80) Kay, M. A. (2011) State-of-the-art gene-based therapies: the road ahead. Nat. Rev. Genet. 12, 316−328. (81) Petris, G., Casini, A., Montagna, C., Lorenzin, F., Prandi, D., Romanel, A., Zasso, J., Conti, L., Demichelis, F., and Cereseto, A. (2017) Hit and go CAS9 delivered through a lentiviral based selflimiting circuit. Nat. Commun. 8, 15334. (82) Yin, H., Xue, W., Chen, S., Bogorad, R. L., Benedetti, E., Grompe, M., Koteliansky, V., Sharp, P. A., Jacks, T., and Anderson, D. G. (2014) Genome editing with Cas9 in adult mice corrects a disease mutation and phenotype. Nat. Biotechnol. 32, 551−553. (83) Wang, M., Glass, Z. A., and Xu, Q. (2017) Non-viral delivery of genome-editing nucleases for gene therapy. Gene Ther. 24, 144. (84) Wang, L., Li, F., Dang, L., Liang, C., Wang, C., He, B., Liu, J., Li, D., Wu, X., Xu, X., Lu, A., and Zhang, G. (2016) In vivo delivery systems for therapeutic genome editing. Int. J. Mol. Sci. 17, 626. (85) Mout, R., Ray, M., Lee, Y. W., Scaletti, F., and Rotello, V. M. (2017) In Vivo Delivery of CRISPR/Cas9 for Therapeutic Gene Editing: Progress and Challenges. Bioconjugate Chem. 28, 880−884. (86) Chew, W. L., Tabebordbar, M., Cheng, J. K. W., Mali, P., Wu, E. Y., Ng, A. H. M., Zhu, K., Wagers, A. J., and Church, G. M. (2016) A multifunctional AAV−CRISPR−Cas9 and its host response. Nat. Methods 13, 868−874. (87) Yin, H., Song, C.-Q., Dorkin, J. R., Zhu, L. J., Li, Y., Wu, Q., Park, A., Yang, J., Suresh, S., Bizhanova, A., Gupta, A., Bolukbasi, M. F., Walsh, S., Bogorad, R. L., Gao, G., Weng, Z., Dong, Y., Koteliansky, V.,
Wolfe, S. A., Langer, R., Xue, W., and Anderson, D. G. (2016) Therapeutic genome editing by combined viral and non-viral delivery of CRISPR system components in vivo. Nat. Biotechnol. 34, 328−333. (88) Yin, H., Kauffman, K. J., and Anderson, D. G. (2017) Delivery technologies for genome editing. Nat. Rev. Drug Discovery 16, 387. (89) Sardesai, N. Y., and Weiner, D. B. (2011) Electroporation delivery of DNA vaccines: Prospects for success. Curr. Opin. Immunol. 23, 421−429. (90) Ding, X., Stewart, M. P., Sharei, A., Weaver, J. C., Langer, R. S., and Jensen, K. F. (2017) High-throughput nuclear delivery and rapid expression of DNA via mechanical and electrical cell-membrane disruption. Nat. Biomed. Eng. 1, 39. (91) Zhang, S., Zhi, D., and Huang, L. (2012) Lipid-based vectors for siRNA delivery. J. Drug Target. 20, 724−735. (92) Akinc, A., Zumbuehl, A., Goldberg, M., Leshchiner, E. S., Busini, V., Hossain, N., Bacallado, S. a, Nguyen, D. N., Fuller, J., Alvarez, R., Borodovsky, A., Borland, T., Constien, R., de Fougerolles, A., Dorkin, J. R., Narayanannair Jayaprakash, K., Jayaraman, M., John, M., Koteliansky, V., Manoharan, M., Nechev, L., Qin, J., Racie, T., Raitcheva, D., Rajeev, K. G., Sah, D. W. Y., Soutschek, J., Toudjarska, I., Vornlocher, H.-P., Zimmermann, T. S., Langer, R., and Anderson, D. G. (2008) A combinatorial library of lipid-like materials for delivery of RNAi therapeutics. Nat. Biotechnol. 26, 561−569. (93) Gilleron, J., Querbes, W., Zeigerer, A., Borodovsky, A., Marsico, G., Schubert, U., Manygoats, K., Seifert, S., Andree, C., Stöter, M., Epstein-Barash, H., Zhang, L., Koteliansky, V., Fitzgerald, K., Fava, E., Bickle, M., Kalaidzidis, Y., Akinc, A., Maier, M., and Zerial, M. (2013) Image-based analysis of lipid nanoparticle-mediated siRNA delivery, intracellular trafficking and endosomal escape. Nat. Biotechnol. 31, 638−646. (94) Sahay, G., Querbes, W., Alabi, C., Eltoukhy, A., Sarkar, S., Zurenko, C., Karagiannis, E., Love, K., Chen, D., Zoncu, R., Buganim, Y., Schroeder, A., Langer, R., and Anderson, D. G. (2013) Efficiency of siRNA delivery by lipid nanoparticles is limited by endocytic recycling. Nat. Biotechnol. 31, 653−658. (95) Kishimoto, T. K., Ferrari, J. D., Lamothe, R. A., Kolte, P. N., Griset, A. P., O ’neil, C., Chan, V., Browning, E., Chalishazar, A., Kuhlman, W., Fu, F.-N., Viseux, N., Altreuter, D. H., Johnston, L., and Maldonado, R. A. (2016) Improving the efficacy and safety of biologic drugs with tolerogenic nanoparticles. Nat. Nanotechnol. 11, 890−899. (96) Maier, M. a, Jayaraman, M., Matsuda, S., Liu, J., Barros, S., Querbes, W., Tam, Y. K., Ansell, S. M., Kumar, V., Qin, J., Zhang, X., Wang, Q., Panesar, S., Hutabarat, R., Carioto, M., Hettinger, J., Kandasamy, P., Butler, D., Rajeev, K. G., Pang, B., Charisse, K., Fitzgerald, K., Mui, B. L., Du, X., Cullis, P., Madden, T. D., Hope, M. J., Manoharan, M., and Akinc, A. (2013) Biodegradable lipids enabling rapidly eliminated lipid nanoparticles for systemic delivery of RNAi therapeutics. Mol. Ther. 21, 1570−1578. (97) Zhu, L., and Mahato, R. I. (2010) Lipid and polymeric carriermediated nucleic acid delivery. Expert Opin. Drug Delivery 7, 1209− 1226. (98) Pasqualini, R., and Ruoslahti, E. (1996) Organ targeting in vivo using phage display peptide libraries. Nature 380, 364−366. (99) National Academies of Sciences Engineering and Medicine. (2017) Human Genome Editing: Science, Ethics, and Governance, The National Academies Press, Washington, DC. (100) ACMG Board of Directors. (2016) Direct-to-consumer genetic testing: a revised position statement of the American College of Medical Genetics and Genomics. Genet. Med. 18, 207−208.
342
DOI: 10.1021/acschembio.7b00689 ACS Chem. Biol. 2018, 13, 333−342
