Ligation-assisted homologous recombination enables precise genome editing by deploying both MMEJ and HDR

CRISPR/Cas12a is a single effector nuclease that, like CRISPR/Cas9, has been harnessed for genome editing based on its ability to generate targeted DNA double strand breaks (DSBs). Unlike the blunt-ended DSB generated by Cas9, Cas12a generates sticky-ended DSB that could potentially aid precise genome editing, but this unique feature has thus far been underutilized. In the current study, we found that a short double-stranded DNA (dsDNA) repair template containing a sticky end that matched one of the Cas12a-generated DSB ends and a homologous arm sharing homology with the genomic region adjacent to the other end of the DSB enabled precise repair of the DSB and introduced a desired nucleotide substitution. We termed this strategy ‘Ligation-Assisted Homologous Recombination’ (LAHR). Compared to the single-stranded oligo deoxyribonucleotide (ssODN)-mediated homology directed repair (HDR), LAHR yields relatively high editing efficiency as demonstrated for both a reporter gene and endogenous genes. We found that both HDR and microhomology-mediated end joining (MMEJ) mechanisms are involved in the LAHR process. Our LAHR genome editing strategy, extends the repertoire of genome editing technologies and provides a broader understanding of the type and role of DNA repair mechanisms involved in genome editing.     

Targeting DNA polymerase to DNA double-strand breaks reduces DNA deletion size and increases templated insertions generated by CRISPR/Cas9

Most insertions or deletions generated by CRISPR/Cas9 (clustered regularly interspaced short palindromic repeats/CRISPR-associated protein 9) endonucleases are short (<25 bp), but unpredictable on-target long DNA deletions (>500 bp) can be observed. The possibility of generating long on-target DNA deletions poses safety risks to somatic genome editing and makes the outcomes of genome editing less predictable. Methods for generating refined mutations are desirable but currently unavailable. Here, we show that fusing Escherichia coli DNA polymerase I or the Klenow fragment to Cas9 greatly increases the frequencies of 1-bp deletions and decreases >1-bp deletions or insertions. Importantly, doing so also greatly decreases the generation of long deletions, including those >2 kb. In addition, templated insertions (the insertion of the nucleotide 4 nt upstream of the protospacer adjacent motif) were increased relative to other insertions. Counteracting DNA resection was one of the mechanisms perturbing deletion sizes. Targeting DNA polymerase to double-strand breaks did not increase off-targets or base substitution rates around the cleavage sites, yet increased editing efficiency in primary cells. Our strategy makes it possible to generate refined DNA mutations for improved safety without sacrificing efficiency of genome editing.     

Kinetics of the CRISPR-Cas9 effector complex assembly and the role of 3′-terminal segment of guide RNA

CRISPR-Cas9 is widely applied for genome engineering in various organisms. The assembly of single guide RNA (sgRNA) with the Cas9 protein may limit the Cas9/sgRNA effector complex function. We developed a FRET-based assay for detection of CRISPR–Cas9 complex binding to its targets and used this assay to investigate the kinetics of Cas9 assembly with a set of structurally distinct sgRNAs. We find that Cas9 and isolated sgRNAs form the effector complex efficiently and rapidly. Yet, the assembly process is sensitive to the presence of moderate concentrations of non-specific RNA competitors, which considerably delay the Cas9/sgRNA complex formation, while not significantly affecting already formed complexes. This observation suggests that the rate of sgRNA loading into Cas9 in cells can be determined by competition between sgRNA and intracellular RNA molecules for the binding to Cas9. Non-specific RNAs exerted particularly large inhibitory effects on formation of Cas9 complexes with sgRNAs bearing shortened 3′-terminal segments. This result implies that the 3′-terminal segment confers sgRNA the ability to withstand competition from non-specific RNA and at least in part may explain the fact that use of sgRNAs truncated for the 3′-terminal stem loops leads to reduced activity during genomic editing.     

A robust CRISPR–Cas9-based fluorescent reporter assay for the detection and quantification of DNA double-strand break repair

DNA double-strand breaks (DSBs) are highly cytotoxic lesions that can lead to chromosome rearrangements, genomic instability and cell death. Consequently, cells have evolved multiple mechanisms to efficiently repair DSBs to preserve genomic integrity. We have developed a DSB repair assay system, designated CDDR (CRISPR–Cas9-based Dual-fluorescent DSB Repair), that enables the detection and quantification of DSB repair outcomes in mammalian cells with high precision. CDDR is based on the introduction and subsequent resolution of one or two DSB(s) in an intrachromosomal fluorescent reporter following the expression of Cas9 and sgRNAs targeting the reporter. CDDR can discriminate between high-fidelity (HF) and error-prone non-homologous end-joining (NHEJ), as well as between proximal and distal NHEJ repair. Furthermore, CDDR can detect homology-directed repair (HDR) with high sensitivity. Using CDDR, we found HF-NHEJ to be strictly dependent on DNA Ligase IV, XRCC4 and XLF, members of the canonical branch of NHEJ pathway (c-NHEJ). Loss of these genes also stimulated HDR, and promoted error-prone distal end-joining. Deletion of the DNA repair kinase ATM, on the other hand, stimulated HF-NHEJ and suppressed HDR. These findings demonstrate the utility of CDDR in characterizing the effect of repair factors and in elucidating the balance between competing DSB repair pathways.     

利用CRISPR/Cas9技术对人多能干细胞进行高效基因组编辑

CRISPR/Cas9技术提供了一个全新的基因组编辑体系。本文利用CRISPR/Cas9平台,在人胚胎干细胞株中对选取的一段特定基因组区域进行了多种基因组编辑：通过在基因编码框中引入移码突变进行基因敲除;通过单链DNA提供外源模板经由同源重组定点敲入FLAG序列;通过同时靶向多个位点诱导基因组大片段删除。研究结果表明CRISPR/Cas9可以对多能干细胞进行高效基因编辑,获得的突变干细胞株有助于对基因和基因组区域的功能进行分析和干细胞疾病模型的建立。     

Single nucleotide substitutions effectively block Cas9 and allow for scarless genome editing in Caenorhabditis elegans

In Caenorhabditis elegans, germline injection of Cas9 complexes is reliably used to achieve genome editing through homology-directed repair of Cas9-generated DNA breaks. To prevent Cas9 from targeting repaired DNA, additional blocking mutations are often incorporated into homologous repair templates. Cas9 can be blocked either by mutating the PAM sequence that is essential for Cas9 activity or by mutating the guide sequence that targets Cas9 to a specific genomic location. However, it is unclear how many nucleotides within the guide sequence should be mutated, since Cas9 can recognize “off-target” sequences that are imperfectly paired to its guide. In this study, we examined whether single-nucleotide substitutions within the guide sequence are sufficient to block Cas9 and allow for efficient genome editing. We show that a single mismatch within the guide sequence effectively blocks Cas9 and allows for recovery of edited animals. Surprisingly, we found that a low rate of edited animals can be recovered without introducing any blocking mutations, suggesting a temporal block to Cas9 activity in C. elegans. Furthermore, we show that the maternal genome of hermaphrodite animals is preferentially edited over the paternal genome. We demonstrate that maternally provided haplotypes can be selected using balancer chromosomes and propose a method of mutant isolation that greatly reduces screening efforts postinjection. Collectively, our findings expand the repertoire of genome editing strategies in C. elegans and demonstrate that extraneous blocking mutations are not required to recover edited animals when the desired mutation is located within the guide sequence.     

CRISPR/Cas9 systems targeting β-globin and CCR5 genes have substantial off-target activity

The ability to precisely modify endogenous genes can significantly facilitate biological studies and disease treatment, and the clustered regularly interspaced short palindromic repeats (CRISPR) systems have the potential to be powerful tools for genome engineering. However, the target specificity of CRISPR systems is largely unknown. Here we demonstrate that CRISPR/Cas9 systems targeting the human hemoglobin β and C-C chemokine receptor type 5 genes have substantial off-target cleavage, especially within the hemoglobin δ and C-C chemokine receptor type 2 genes, respectively, causing gross chromosomal deletions. The guide strands of the CRISPR/Cas9 systems were designed to have a range of mismatches with the sequences of potential off-target sites. Off-target analysis was performed using the T7 endonuclease I mutation detection assay and Sanger sequencing. We found that the repair of the on-and off-target cleavage resulted in a wide variety of insertions, deletions and point mutations. Therefore, CRISPR/Cas9 systems need to be carefully designed to avoid potential off-target cleavage sites, including those with mismatches to the 12-bases proximal to the guide strand protospacer-adjacent motif.     

Multiplex gene precise editing and large DNA fragment deletion by the CRISPR‐Cas9‐TRAMA system in edible mushroom Cordyceps militaris

The medicinal mushroom Cordyceps militaris contains abundant valuable bioactive ingredients that have attracted a great deal of attention in the pharmaceutical and cosmetic industries. However, the development of this valuable mushroom faces the obstacle of lacking powerful genomic engineering tools. Here, by excavating the endogenous tRNA‐processed element, introducing the extrachromosomal plasmid and alongside with homologous template, we develop a marker‐free CRISPR‐Cas9‐TRAMA genomic editing system to achieve the multiplex gene precise editing and large synthetic cluster deletion in C. militaris. We further operated editing in the synthetases of cordycepin and ergothioneine to demonstrate the application of Cas9‐TRAMA system in protein modification, promoter strength evaluation and 10 kb metabolic synthetic cluster deletion. The Cas9‐TRAMA system provides a scalable method for excavating the valuable metabolic resource of medicinal mushrooms and constructing a mystical cellular pathway to elucidate the complex cell behaviours of the edible mushroom.

This study identified the endogenic tRNA processed element of mushroom crop Cordyceps militaris, and proved the in vivo generation of gRNA by promoter PtrpC could highly improve the editing efficiency of genomic editing. By introducing the extra‐chromosomal plasmid and alongside with homologous template, a marker‐free CRISPR‐Cas9‐TRAMA genomic editing system was built to achieve the multiplex gene precise editing and large synthetic cluster deletion in C. militaris.     

The role of Cas8?in type?I CRISPR interference

CRISPR (clustered regularly interspaced short palindromic repeat) systems provide bacteria and archaea with adaptive immunity to repel invasive genetic elements. Type I systems use ‘cascade’ [CRISPR-associated (Cas) complex for antiviral defence] ribonucleoprotein complexes to target invader DNA, by base pairing CRISPR RNA (crRNA) to protospacers. Cascade identifies PAMs (protospacer adjacent motifs) on invader DNA, triggering R-loop formation and subsequent DNA degradation by Cas3. Cas8 is a candidate PAM recognition factor in some cascades. We analysed Cas8 homologues from type IB CRISPR systems in archaea Haloferax volcanii (Hvo) and Methanothermobacter thermautotrophicus (Mth). Cas8 was essential for CRISPR interference in Hvo and purified Mth Cas8 protein responded to PAM sequence when binding to nucleic acids. Cas8 interacted physically with Cas5–Cas7–crRNA complex, stimulating binding to PAM containing substrates. Mutation of conserved Cas8 amino acid residues abolished interference in vivo and altered catalytic activity of Cas8 protein in vitro. This is experimental evidence that Cas8 is important for targeting Cascade to invader DNA.     

Rational guide RNA engineering for small-molecule control of CRISPR/Cas9 and gene editing

It is important to control CRISPR/Cas9 when sufficient editing is obtained. In the current study, rational engineering of guide RNAs (gRNAs) is performed to develop small-molecule-responsive CRISPR/Cas9. For our purpose, the sequence of gRNAs are modified to introduce ligand binding sites based on the rational design of ligand–RNA pairs. Using short target sequences, we demonstrate that the engineered RNA provides an excellent scaffold for binding small molecule ligands. Although the ‘stem–loop 1’ variants of gRNA induced variable cleavage activity for different target sequences, all ‘stem–loop 3’ variants are well tolerated for CRISPR/Cas9. We further demonstrate that this specific ligand–RNA interaction can be utilized for functional control of CRISPR/Cas9 in vitro and in human cells. Moreover, chemogenetic control of gene editing in human cells transfected with all-in-one plasmids encoding Cas9 and designer gRNAs is demonstrated. The strategy may become a general approach for generating switchable RNA or DNA for controlling other biological processes.     

Deploying MMEJ using MENdel in precision gene editing applications for gene therapy and functional genomics

Gene-editing experiments commonly elicit the error-prone non-homologous end joining for DNA double-strand break (DSB) repair. Microhomology-mediated end joining (MMEJ) can generate more predictable outcomes for functional genomic and somatic therapeutic applications. We compared three DSB repair prediction algorithms – MENTHU, inDelphi, and Lindel – in identifying MMEJ-repaired, homogeneous genotypes (PreMAs) in an independent dataset of 5,885 distinct Cas9-mediated mouse embryonic stem cell DSB repair events. MENTHU correctly identified 46% of all PreMAs available, a ∼2- and ∼60-fold sensitivity increase compared to inDelphi and Lindel, respectively. In contrast, only Lindel correctly predicted predominant single-base insertions. We report the new algorithm MENdel, a combination of MENTHU and Lindel, that achieves the most predictive coverage of homogeneous out-of-frame mutations in this large dataset. We then estimated the frequency of Cas9-targetable homogeneous frameshift-inducing DSBs in vertebrate coding regions for gene discovery using MENdel. 47 out of 54 genes (87%) contained at least one early frameshift-inducing DSB and 49 out of 54 (91%) did so when also considering Cas12a-mediated deletions. We suggest that the use of MENdel helps researchers use MMEJ at scale for reverse genetics screenings and with sufficient intra-gene density rates to be viable for nearly all loss-of-function based gene editing therapeutic applications.     

High-efficiency and multilocus targeted integration in CHO cells using CRISPR-mediated donor nicking and DNA repair inhibitors

Efforts to leverage clustered regularly interspaced short palindromic repeats/CRISPR-associated protein 9 (CRISPR/Cas9) for targeted genomic modifications in mammalian cells are limited by low efficiencies and heterogeneous outcomes. To aid method optimization, we developed an all-in-one reporter system, including a novel superfolder orange fluorescent protein (sfOrange), to simultaneously quantify gene disruption, site-specific integration (SSI), and random integration (RI). SSI strategies that utilize different donor plasmid formats and Cas9 nuclease variants were evaluated for targeting accuracy and efficiency in Chinese hamster ovary cells. Double-cut and double-nick donor formats significantly improved targeting accuracy by 2.3–8.3-fold and 19–22-fold, respectively, compared to standard circular donors. Notably, Cas9-mediated donor linearization was associated with increased RI events, whereas donor nicking minimized RI without sacrificing SSI efficiency and avoided low-fidelity outcomes. A screen of 10 molecules that modulate the major mammalian DNA repair pathways identified two inhibitors that further enhance targeting accuracy and efficiency to achieve SSI in 25% of transfected cells without selection. The optimized methods integrated transgene expression cassettes with 96% efficiency at a single locus and with 53%–55% efficiency at two loci simultaneously in selected clones. The CRISPR-based tools and methods developed here could inform the use of CRISPR/Cas9 in mammalian cell lines, accelerate mammalian cell line engineering, and support advanced recombinant protein production applications.     

Epigenetic features drastically impact CRISPR–Cas9 efficacy in plants

CRISPR–Cas9-mediated genome editing has been widely adopted for basic and applied biological research in eukaryotic systems. While many studies consider DNA sequences of CRISPR target sites as the primary determinant for CRISPR mutagenesis efficiency and mutation profiles, increasing evidence reveals the substantial role of chromatin context. Nonetheless, most prior studies are limited by the lack of sufficient epigenetic resources and/or by only transiently expressing CRISPR–Cas9 in a short time window. In this study, we leveraged the wealth of high-resolution epigenomic resources in Arabidopsis (Arabidopsis thaliana) to address the impact of chromatin features on CRISPR–Cas9 mutagenesis using stable transgenic plants. Our results indicated that DNA methylation and chromatin features could lead to substantial variations in mutagenesis efficiency by up to 250-fold. Low mutagenesis efficiencies were mostly associated with repressive heterochromatic features. This repressive effect appeared to persist through cell divisions but could be alleviated through substantial reduction of DNA methylation at CRISPR target sites. Moreover, specific chromatin features, such as H3K4me1, H3.3, and H3.1, appear to be associated with significant variation in CRISPR–Cas9 mutation profiles mediated by the non-homologous end joining repair pathway. Our findings provide strong evidence that specific chromatin features could have substantial and lasting impacts on both CRISPR–Cas9 mutagenesis efficiency and DNA double-strand break repair outcomes.

Epigenetic features substantially influence genome editing efficiency and DNA repair outcomes.     

Double-strand break toxicity is chromatin context independent

Cells respond to double-strand breaks (DSBs) by activating DNA damage response pathways, including cell cycle arrest. We have previously shown that a single double-strand break generated via CRISPR/Cas9 is sufficient to delay cell cycle progression and compromise cell viability. However, we also found that the cellular response to DSBs can vary, independent of the number of lesions. This implies that not all DSBs are equally toxic, and raises the question if the location of a single double-strand break could influence its toxicity. To systematically investigate if DSB-location is a determinant of toxicity we performed a CRISPR/Cas9 screen targeting 6237 single sites in the human genome. Next, we developed a data-driven framework to design CRISPR/Cas9 sgRNA (crRNA) pools targeting specific chromatin features. The chromatin context was defined using ChromHMM states, Lamin-B1 DAM-iD, DNAseI hypersensitivity, and RNA-sequencing data. We computationally designed 6 distinct crRNA pools, each containing 10 crRNAs targeting the same chromatin state. We show that the toxicity of a DSB is highly similar across the different ChromHMM states. Rather, we find that the major determinants of toxicity of a sgRNA are cutting efficiency and off-target effects. Thus, chromatin features have little to no effect on the toxicity of a single CRISPR/Cas9-induced DSB.     

Efficient Mutagenesis by Cas9 Protein-Mediated Oligonucleotide Insertion and Large-Scale Assessment of Single-Guide RNAs

The CRISPR/Cas9 system has been implemented in a variety of model organisms to mediate site-directed mutagenesis. A wide range of mutation rates has been reported, but at a limited number of genomic target sites. To uncover the rules that govern effective Cas9-mediated mutagenesis in zebrafish, we targeted over a hundred genomic loci for mutagenesis using a streamlined and cloning-free method. We generated mutations in 85% of target genes with mutation rates varying across several orders of magnitude, and identified sequence composition rules that influence mutagenesis. We increased rates of mutagenesis by implementing several novel approaches. The activities of poor or unsuccessful single-guide RNAs (sgRNAs) initiating with a 5′ adenine were improved by rescuing 5′ end homogeneity of the sgRNA. In some cases, direct injection of Cas9 protein/sgRNA complex further increased mutagenic activity. We also observed that low diversity of mutant alleles led to repeated failure to obtain frame-shift mutations. This limitation was overcome by knock-in of a stop codon cassette that ensured coding frame truncation. Our improved methods and detailed protocols make Cas9-mediated mutagenesis an attractive approach for labs of all sizes.     

Marker-free quantification of repair pathway utilization at Cas9-induced double-strand breaks

Genome integrity and genome engineering require efficient repair of DNA double-strand breaks (DSBs) by non-homologous end joining (NHEJ), homologous recombination (HR), or alternative end-joining pathways. Here we describe two complementary methods for marker-free quantification of DSB repair pathway utilization at Cas9-targeted chromosomal DSBs in mammalian cells. The first assay features the analysis of amplicon next-generation sequencing data using ScarMapper, an iterative break-associated alignment algorithm to classify individual repair products based on deletion size, microhomology usage, and insertions. The second assay uses repair pathway-specific droplet digital PCR assays (‘PathSig-dPCR’) for absolute quantification of signature DSB repair outcomes. We show that ScarMapper and PathSig-dPCR enable comprehensive assessment of repair pathway utilization in different cell models, after a variety of experimental perturbations. We use these assays to measure the differential impact of DNA end resection on NHEJ, HR and polymerase theta-mediated end joining (TMEJ) repair. These approaches are adaptable to any cellular model system and genomic locus where Cas9-mediated targeting is feasible. Thus, ScarMapper and PathSig-dPCR allow for systematic fate mapping of a targeted DSB with facile and accurate quantification of DSB repair pathway choice at endogenous chromosomal loci.     

Genome editing in mammalian cells using the CRISPR type I-D nuclease

Adoption of CRISPR–Cas systems, such as CRISPR–Cas9 and CRISPR–Cas12a, has revolutionized genome engineering in recent years; however, application of genome editing with CRISPR type I—the most abundant CRISPR system in bacteria—remains less developed. Type I systems, such as type I-E, and I-F, comprise the CRISPR-associated complex for antiviral defense (‘Cascade’: Cas5, Cas6, Cas7, Cas8 and the small subunit) and Cas3, which degrades the target DNA; in contrast, for the sub-type CRISPR–Cas type I-D, which lacks a typical Cas3 nuclease in its CRISPR locus, the mechanism of target DNA degradation remains unknown. Here, we found that Cas10d is a functional nuclease in the type I-D system, performing the role played by Cas3 in other CRISPR–Cas type I systems. The type I-D system can be used for targeted mutagenesis of genomic DNA in human cells, directing both bi-directional long-range deletions and short insertions/deletions. Our findings suggest the CRISPR–Cas type I-D system as a unique effector pathway in CRISPR that can be repurposed for genome engineering in eukaryotic cells.     

Improvement of baculovirus as protein expression vector and as biopesticide by CRISPR/Cas9 editing

The clustered regularly interspaced short palindromic repeats (CRISPR) system–associated Cas9 endonuclease is a molecular tool that enables specific sequence editing with high efficiency. In this study, we have explored the use of CRISPR/Cas9 system for the engineering of baculovirus. We have shown that the delivering of Cas9-single guide RNA ribonucleoprotein (RNP) complex with or without DNA repair template into Sf21 insect cells through lipofection might be efficient to produce knockouts as well as knock-ins into the baculovirus. To evaluate potential application of our CRISPR/Cas9 method to improve baculovirus as protein expression vector and as biopesticide, we attempted to knockout several genes from a recombinant AcMNPV form used in the baculovirus expression system as well as in a natural occurring viral isolate from the same virus. We have additionally confirmed the adaptation of this methodology for the generation of viral knock-ins in specific regions of the viral genome. Analysis of the generated mutants revealed that the editing efficiency and the type of changes was variable but relatively high. Depending on the targeted gene, the editing rate ranged from 10% to 40%. This study established the first report revealing the potential of CRISPR/Cas9 for genome editing in baculovirus, contributing to the engineering of baculovirus as a protein expression vector as well as a biological control agent.     

Systematic in vitro specificity profiling reveals nicking defects in natural and engineered CRISPR–Cas9 variants

Cas9 is an RNA-guided endonuclease in the bacterial CRISPR–Cas immune system and a popular tool for genome editing. The commonly used Streptococcus pyogenes Cas9 (SpCas9) is relatively non-specific and prone to off-target genome editing. Other Cas9 orthologs and engineered variants of SpCas9 have been reported to be more specific. However, previous studies have focused on specificity of double-strand break (DSB) or indel formation, potentially overlooking alternative cleavage activities of these Cas9 variants. In this study, we employed in vitro cleavage assays of target libraries coupled with high-throughput sequencing to systematically compare cleavage activities and specificities of two natural Cas9 variants (SpCas9 and Staphylococcus aureus Cas9) and three engineered SpCas9 variants (SpCas9 HF1, HypaCas9 and HiFi Cas9). We observed that all Cas9s tested could cleave target sequences with up to five mismatches. However, the rate of cleavage of both on-target and off-target sequences varied based on target sequence and Cas9 variant. In addition, SaCas9 and engineered SpCas9 variants nick targets with multiple mismatches but have a defect in generating a DSB, while SpCas9 creates DSBs at these targets. Overall, these differences in cleavage rates and DSB formation may contribute to varied specificities observed in genome editing studies.     

Characterization of Genomic Deletion Efficiency Mediated by Clustered Regularly Interspaced Palindromic Repeats (CRISPR)/Cas9 Nuclease System in Mammalian Cells

The clustered regularly interspaced palindromic repeats (CRISPR)/CRISPR-associated (Cas) 9 nuclease system has provided a powerful tool for genome engineering. Double strand breaks may trigger nonhomologous end joining repair, leading to frameshift mutations, or homology-directed repair using an extrachromosomal template. Alternatively, genomic deletions may be produced by a pair of double strand breaks. The efficiency of CRISPR/Cas9-mediated genomic deletions has not been systematically explored. Here, we present a methodology for the production of deletions in mammalian cells, ranging from 1.3 kb to greater than 1 Mb. We observed a high frequency of intended genomic deletions. Nondeleted alleles are nonetheless often edited with inversions or small insertion/deletions produced at CRISPR recognition sites. Deleted alleles also typically include small insertion/deletions at predicted deletion junctions. We retrieved cells with biallelic deletion at a frequency exceeding that of probabilistic expectation. We demonstrate an inverse relationship between deletion frequency and deletion size. This work suggests that CRISPR/Cas9 is a robust system to produce a spectrum of genomic deletions to allow investigation of genes and genetic elements.