Genome Editing: Insights from Chemical Biology to Support Safe and

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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 ACS Chem. Biol., Just Accepted Manuscript • DOI: 10.1021/acschembio.7b00689 • Publication Date (Web): 09 Oct 2017 Downloaded from http://pubs.acs.org on October 11, 2017

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Title: Genome Editing: Insights from Chemical Biology to Support Safe and Transformative Therapeutic Applications Author List: Renee D. Wegrzyn*1, Andrew H. Lee2, Amy L. Jenkins3, Colby D. Stoddard4, and Anne E. Cheever2 Author Information: 1. Defense Advanced Research Projects Agency (DARPA); 675 N. Randolph St., Arlington, VA 22203 2. Booz Allen Hamilton; 3811 Fairfax Dr. Suite 600, Arlington, VA 22203 3. Schafer: A Belcan Company; 3811 Fairfax Dr., Arlington, VA 22203 4. Quantitative Scientific Solutions; 4601 N. Fairfax Dr. Suite 1200, Arlington, VA 22203 *Corresponding author: [email protected] 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 including the potential for off-target effects, limited control of editing activity and subsequent DNA repair outcomes, and insufficient target conversion and delivery performance prevent the widespread, safe, and practical use of genome editors, especially for human disease interventions. 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. Key words: CRISPR/Cas9, genome editing, biosafety, delivery, therapeutics, DNA repair, LEEDR, bioethics Introduction 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 costeffective. 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 pre-clinical and clinical work on therapeutic applications of gene editors, including T cell immunotherapies to treat cancers1, sickle cell disease2, and reduction of HIV burden in patients3, 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), lack of precise control over genome editing activity and repair outcomes5, low efficiency of target conversion6, and in vivo delivery limitations7. 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 towards new solutions to treat a range of conditions, including heritable genetic disorders, that will ultimately require the use of genome editors in vivo. Beyond heritable genetic disorders, new opportunities are emerging 1 ACS Paragon Plus Environment


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to apply genome editors to combat a variety of infectious disease challenges, including viral infections8 and antimicrobial resistance9. 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 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 CRISPRmediated editing, and advancements in measuring off-target 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). 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 double-strand break (DSB) through its endonuclease activity10. DSBs are resolved primarily through the error-prone non-homologous 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 donor DNA sequence as a template for repair, which can be leveraged to introduce user-defined sequence into a locus11. Cas9 nuclease variants, such as Cas9 nickases 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 effects14. The DNA repair processes that are triggered by DSBs increase the probability of unintended effects both at target and non-target sites, from disruption of non-target gene function through single nucleotide polymorphisms and frameshifts to chromosomal rearrangements and genome instability16. 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. 2 ACS Paragon Plus Environment

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One way to address the challenges associated with DNA repair and reduce the risk of unintended effects has been through the development of non-canonical 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 non-canonical functionality. Novel effector domains, such as cytosine deamination for base editing18, epigenetic modifiers such as p30019, Tet120, DNMT3a21, LSD122, and MQ123 to alter gene expression activity, or transcriptional activator/repressor domains such as VP6424, VPR25, and KRAB26,27 can be fused directly to dCas9 to activate or repress target gene expression. These non-canonical epigenome 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 vitro28,29, 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 non-canonical genome editing-based 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 non-addictive 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 towards NHEJ, with reported frequencies of 20%-60% in targeted cells30,31 compared to 0.5% - 20% for HDR30–32, where lower efficiency is attributable to the need for co-delivery of a donor template. NHEJ repair is active at all stages of the cell cycle33, 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 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 outcomes5,36,37. 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 efficiencies5,37. 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 unclear38,39. Small molecule-mediated cell-cycle arrest followed by the timed delivery of CRISPR/Cas9 RNPs has demonstrated HDR-mediated editing levels of up to 38% with no detectable off-target editing35. These initial results highlight the potential utility of leveraging

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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 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 15-75%18. Additionally, post-translational regulation of CRISPR/Cas9 through fusion with the replication licensing factor Geminin that limit 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 Cpf1-generated DSBs with 5’ or 3’ overhang DNA ends can affect repair pathway choice to favor HDR over NHEJ15,41. 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 junctionmediated HDR45. 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 off-target effects broadly, inclusive of unintended genetic, transcriptomic, or epigenetic changes at non-target sites across the genome, unintended outcomes at the target site itself, and genome editing that may occur in non-target cells or tissues. Offtarget 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 off-target effects, accurately predicting the off-target cleavage sites, 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 organisms50,51,


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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. Controllable Genome Editing The ability to control the activity of genome editors in a predictable and stringent manner will facilitate advances towards 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 post-translational control strategies in which Cas9 is modified to enable control through the introduction of a stimulus. Posttranslational control, unlike regulation of transcriptional control, offers a higher resolution of userdefined control and less reliance on inherent cellular processes such as transcription, translation, and protein degradation. Control of Cas9 has been demonstrated using light52, temperature53, intein liganddependency54,55, dimerization-dependency55–58, destabilization domains59,60, 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 2c). Beginning with the original fusion of Cas9 RNA components to form the sgRNA12, gRNA development has expanded to include gRNA truncations to decrease off-target effects62 and create synthetic circuits63. RNA hairpin engineering has been shown to modulate transcription by recruiting RNA-binding transcriptional modifiers such as VP6464–66, KRAB65, or PUFeffector fusion proteins67. 5’ and 3’ chemically-modified gRNAs can increase gRNA stability and improve genome editing68,69 . Small molecule control has been engineered into gRNA sequences appended with ligand-activated self-cleaving aptazymes such that RNP activity is ligand-dependent70 . RNA aptamers can bind a variety of ligands, either engineered or natural71. 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 gates72. Small molecule-mediated control



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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 bacteriophage-encoded anti-CRISPR proteins that act by blocking dsDNA binding and cleavage73–75. These findings demonstrate that RNA-guided nucleases can be targets for small protein inhibitors; however, 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 dose-dependent 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 (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. 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 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 6 ACS Paragon Plus Environment

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nucleic acid-based therapeutics such as siRNA and DNA/RNA vaccines77–79. 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 endocytosis80. 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 off-target effects may occur. For instance, lentiviral-delivered self-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 effects81. 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 allele82. 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 (DNA or RNA), 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 co-transfection of CRISPR components (e.g., AAV-split-Cas986) to reconstitute a full-length 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 subjects83,85,87,88. 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 pre-existing 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 messenger RNA 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. 7 ACS Paragon Plus Environment


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The delivery of CRISPR/Cas9 or other genome editors via plasmid or mini-circle DNA has been demonstrated in proof-of-concept in vivo animal experiments82. 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, 3d). 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 mechanisms91–94. 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 constructs88. Currently available chemical formulations often result in induced toxicity and immunogenicity, including the development of antidrug antibodies against the expressed protein, resulting in sub-optimal 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 mice86. Therefore, 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 than 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 therapy95. Lipid and polymer toxicity is often alleviated or minimized by the use of biodegradable formulations with readily degraded ester or amide bonds96,97. 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


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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 involve laboratory manipulations that are difficult, time-consuming, 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 towards 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 Genomics100, 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 technologies 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 datasets. 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.

Conflict of Interest 9 ACS Paragon Plus Environment


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AHL and AEC are employees of Booz Allen Hamilton. ALJ is an employee of Schafer: A Belcan Company. CDS is an employee of Quantitative Scientific Solutions.

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Figure Legends Graphical Table of Contents: Outline of the potential opportunities for chemical biology to support safe and responsible advancement of genome editing applications. Figure 1. Schematic drawing of canonical CRISPR/Cas9 genome editing mechanism and outcomes. The Cas9:guide RNA (dark grey 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 non-homologous end joining (NHEJ) or homology-directed repair (HDR) DNA repair pathways. 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: Liganddependent 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 enable 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. 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. 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.


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Graphical Table of Contents: Outline of the potential opportunities for chemical biology to support safe and responsible advancement of genome editing applications. 118x59mm (300 x 300 DPI)

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Figure 1. Schematic drawing of canonical CRISPR/Cas9 genome editing mechanism and outcomes. The Cas9:guide RNA (dark grey 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 non-homologous end joining (NHEJ) or homology-directed repair (HDR) DNA repair pathways. 113x91mm (300 x 300 DPI)


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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 enable 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.



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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. 137x142mm (300 x 300 DPI)



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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. 118x59mm (300 x 300 DPI)


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Genome Editing: Insights from Chemical Biology to Support Safe and

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