Site-specific Double-strand Breaks Research Articles

Dose-dependent effects of histone methyltransferase NSD2 on site-specific double-strand break repair.

Histone modifications are catalyzed and recognized by specific proteins to regulate dynamic DNA metabolism processes. NSD2 is a histone H3 lysine 36 (H3K36)-specific methyltransferase that is associated with both various transcription regulators and DNA repair factors. Specifically, it has been implicated in the repair of DNA double-strand breaks (DSBs); however, the role of NSD2 during DSB repair remains enigmatic. Here, we show that NSD2 does not accumulate at DSB sites and that it is not further mobilized by DSB formation. Using three different DSB repair reporter systems, which contained the endonuclease site in the active thymidine kinase gene (TK) locus, we demonstrated separate dose-dependent effects of NSD2 on homologous recombination (HR), canonical-non-homologous end joining (c-NHEJ), and non-canonical-NHEJ (non-c-NHEJ). Endogenous NSD2 has a role in repressing non-c-NHEJ, without affecting DSB repair efficiency by HR or total NHEJ. Furthermore, overexpression of NSD2 promotes c-NHEJ repair and suppresses HR repair. Therefore, we propose that NSD2 has functions in chromatin integrity at the active regions during DSB repair.

The dCas9-based genome editing in Plasmodium yoelii.

Genetic editing is a powerful tool for functional characterization of genes in various organisms. With its simplicity and specificity, the CRISPR-Cas9 technology has become a popular editing tool, which introduces site-specific DNA double-strand breaks (DSBs), and then leverages the endogenous repair pathway for DSB repair via homology-directed repair (HDR) or the more error-prone non-homologous end joining (NHEJ) pathways. However, in the Plasmodium parasites, the lack of a typical NHEJ pathway selects for DSB repair through the HDR pathway when a homologous DNA template is available. The AT-rich nature of the Plasmodium genome exacerbates this drawback by making it difficult to clone longer homologous repair DNA templates. To circumvent these challenges, we adopted the hybrid catalytically inactive Cas9 (dCas9)-microbial single-stranded annealing proteins (SSAP) editor to the Plasmodium genome. In Plasmodium yoelii, we demonstrated the use of the dCas9-SSAP, as the cleavage-free gene editor, by targeted gene deletion and gene tagging, even using shorter homologous DNA templates. This dCas9-SSAP method with a shorter DNA template, which did not require DSBs, independent of HDR and NHEJ, would be a great addition to the existing genetic toolbox and could be deployed for the functional characterization of genes in Plasmodium, contributing to improving the ability of the malaria research community in characterizing more than half of genes with unknown functions.IMPORTANCEMalaria caused by Plasmodium parasites infection remains a serious threat to human health, with an estimated 249 million malaria cases and 608,000 deaths worldwide in 2022, according to the latest report from the World Health Organization (WHO). Here, we demonstrated the use of dCas9-single-stranded annealing protein, as the cleavage-free gene editor in Plasmodium yoelii, by targeted deletion and gene tagging, even using shorter homologous DNA templates. This method with a shorter DNA template, which did not require DSBs, independent of HDR and NHEJ, showing the potential significance in greatly improving our ability to elucidate gene functions, would contribute to assisting the malaria research community in deciphering more than half of genes with unknown functions to identify new drug and vaccine targets.

Nonhomologous tails direct heteroduplex rejection and mismatch correction during single-strand annealing in Saccharomyces cerevisiae.

Single-strand annealing (SSA) is initiated when a double strand break (DSB) occurs between two flanking repeated sequences, resulting in a deletion that leaves a single copy of the repeat. We studied budding yeast strains carrying two 200-bp URA3 sequences separated by 2.6 kb of spacer DNA (phage lambda) in which a site-specific DSB can be created by HO or Cas9 endonucleases. Repeat-mediated deletion requires removal of long 3'-ended single-stranded tails (flaps) by Rad1-Rad10 with the assistance of Msh2-Msh3, Saw1 and Slx4. A natural 3% divergence of unequally spaced heterologies between these repeats (designated F and A) causes a significant reduction in the frequency of SSA repair. This decrease is caused by heteroduplex rejection in which mismatches (MMs) in the annealed intermediate are recognized by the MutS (Msh2 and Msh6) components of the MM repair (MMR) pathway coupled to unwinding of the duplex by the Sgs1-Rmi1-Top3 helicase. MutL homologs, Mlh1-Pms1 (MutL), are not required for rejection but play their expected role in mismatch correction. Remarkably, heteroduplex rejection is very low in strains where the divergent repeats were immediately adjacent (Tailless strains) and the DSB was induced by Cas9. These results suggest that the presence of nonhomologous tails strongly stimulates heteroduplex rejection in SSA. DNA sequencing analysis of SSA products from the FA Tailed strain showed a gradient of correction favoring the sequence opposite each 3' end of the annealed strand. Mismatches located in the center of the repair intermediate were corrected by Msh2-Msh6 mediated mismatch correction, while correction of MMs at the extremity of the SSA intermediate often appears to use a different mechanism, possibly by 3' nonhomologous tail removal that includes part of the homologous sequence. In contrast, in FA Tailless strains there was a uniform repair of the MMs across the repeat. A distinctive pattern of correction was found in the absence of MSH2, in both Tailed and Tailless strains, different from the spectrum seen in a msh3Δ msh6Δ double mutant. Previous work has shown that SSA is Rad51-independent but dependent on the strand annealing activity of Rad52. However Rad52 becomes dispensable in a Tailless construct where the DSB is induced by Cas9 or in transformation of a plasmid where SSA occurs in the absence of nonhomologous tails.

Programmable site-specific DNA double-strand breaks via PNA-assisted prokaryotic Argonautes.

Programmable site-specific nucleases promise to unlock myriad applications in basic biology research, biotechnology and gene therapy. Gene-editing systems have revolutionized our ability to engineer genomes across diverse eukaryotic species. However, key challenges, including delivery, specificity and targeting organellar genomes, pose barriers to translational applications. Here, we use peptide nucleic acids (PNAs) to facilitate precise DNA strand invasion and unwinding, enabling prokaryotic Argonaute (pAgo) proteins to specifically bind displaced single-stranded DNA and introduce site-specific double-strand breaks (DSBs) independent of the target sequence. We named this technology PNA-assisted pAgo editing (PNP editing) and determined key parameters for designing PNP editors to efficiently generate programable site-specific DSBs. Our design allows the simultaneous use of multiple PNP editors to generate multiple site-specific DSBs, thereby informing design considerations for potential in vitro and in vivo applications, including genome editing.

Improving Precise Genome Editing Using Donor DNA/gRNA Hybrid Duplex Generated by Complementary Bases.

In precise genome editing, site-specific DNA double-strand breaks (DSBs) induced by the CRISPR/Cas9 system are repaired via homology-directed repair (HDR) using exogenous donor DNA templates. However, the low efficiency of HDR-mediated genome editing is a barrier to widespread use. In this study, we created a donor DNA/guide RNA (gRNA) hybrid duplex (DGybrid) that was composed of sequence-extended gRNA and single-stranded oligodeoxynucleotide (ssODN) combined with complementary bases without chemical modifications to increase the concentration of donor DNA at the cleavage site. The efficiency of genome editing using DGybrid was evaluated in Saccharomyces cerevisiae. The results show a 1.8-fold (from 35% to 62%) improvement in HDR-mediated editing efficiency compared to genome editing in which gRNA and donor DNA were introduced separately. In addition, analysis of the nucleic acid introduction efficiency using flow cytometry indicated that both RNA and ssODNs are efficiently incorporated into cells together by using the DNA/RNA hybrid. Our technique would be preferred as a universal and concise tool for improving the efficiency of HDR-mediated genome editing.

Highly efficient generation of isogenic pluripotent stem cell models using prime editing.

The recent development of prime editing (PE) genome engineering technologies has the potential to significantly simplify the generation of human pluripotent stem cell (hPSC)-based disease models. PE is a multicomponent editing system that uses a Cas9-nickase fused to a reverse transcriptase (nCas9-RT) and an extended PE guide RNA (pegRNA). Once reverse transcribed, the pegRNA extension functions as a repair template to introduce precise designer mutations at the target site. Here, we systematically compared the editing efficiencies of PE to conventional gene editing methods in hPSCs. This analysis revealed that PE is overall more efficient and precise than homology-directed repair of site-specific nuclease-induced double-strand breaks. Specifically, PE is more effective in generating heterozygous editing events to create autosomal dominant disease-associated mutations. By stably integrating the nCas9-RT into hPSCs we achieved editing efficiencies equal to those reported for cancer cells, suggesting that the expression of the PE components, rather than cell-intrinsic features, limit PE in hPSCs. To improve the efficiency of PE in hPSCs, we optimized the delivery modalities for the PE components. Delivery of the nCas9-RT as mRNA combined with synthetically generated, chemically-modified pegRNAs and nicking guide RNAs improved editing efficiencies up to 13-fold compared with transfecting the PE components as plasmids or ribonucleoprotein particles. Finally, we demonstrated that this mRNA-based delivery approach can be used repeatedly to yield editing efficiencies exceeding 60% and to correct or introduce familial mutations causing Parkinson's disease in hPSCs.

Repair of mismatched templates during Rad51-dependent Break-Induced Replication.

Using budding yeast, we have studied Rad51-dependent break-induced replication (BIR), where the invading 3' end of a site-specific double-strand break (DSB) and a donor template share 108 bp of homology that can be easily altered. BIR still occurs about 10% as often when every 6th base is mismatched as with a perfectly matched donor. Here we explore the tolerance of mismatches in more detail, by examining donor templates that each carry 10 mismatches, each with different spatial arrangements. Although 2 of the 6 arrangements we tested were nearly as efficient as the evenly-spaced reference, 4 were significantly less efficient. A donor with all 10 mismatches clustered at the 3' invading end of the DSB was not impaired compared to arrangements where mismatches were clustered at the 5' end. Our data suggest that the efficiency of strand invasion is principally dictated by thermodynamic considerations, i.e., by the total number of base pairs that can be formed; but mismatch position-specific effects are also important. We also addressed an apparent difference between in vitro and in vivo strand exchange assays, where in vitro studies had suggested that at a single contiguous stretch of 8 consecutive bases was needed to be paired for stable strand pairing, while in vivo assays using 108-bp substrates found significant recombination even when every 6th base was mismatched. Now, using substrates of either 90 or 108 nt-the latter being the size of the in vivo templates-we find that in vitro D-loop results are very similar to the in vivo results. However, there are still notable differences between in vivo and in vitro assays that are especially evident with unevenly-distributed mismatches. Mismatches in the donor template are incorporated into the BIR product in a strongly polar fashion up to ~40 nucleotides from the 3' end. Mismatch incorporation depends on the 3'→ 5' proofreading exonuclease activity of DNA polymerase δ, with little contribution from Msh2/Mlh1 mismatch repair proteins, or from Rad1-Rad10 flap nuclease or the Mph1 helicase. Surprisingly, the probability of a mismatch 27 nt from the 3' end being replaced by donor sequence was the same whether the preceding 26 nucleotides were mismatched every 6th base or fully homologous. These data suggest that DNA polymerase δ "chews back" the 3' end of the invading strand without any mismatch-dependent cues from the strand invasion structure. However, there appears to be an alternative way to incorporate a mismatch at the first base at the 3' end of the donor.

Correction of Fanconi Anemia Mutations Using Digital Genome Engineering.

Fanconi anemia (FA) is a rare genetic disease in which genes essential for DNA repair are mutated. Both the interstrand crosslink (ICL) and double-strand break (DSB) repair pathways are disrupted in FA, leading to patient bone marrow failure (BMF) and cancer predisposition. The only curative therapy for the hematological manifestations of FA is an allogeneic hematopoietic cell transplant (HCT); however, many (>70%) patients lack a suitable human leukocyte antigen (HLA)-matched donor, often resulting in increased rates of graft-versus-host disease (GvHD) and, potentially, the exacerbation of cancer risk. Successful engraftment of gene-corrected autologous hematopoietic stem cells (HSC) circumvents the need for an allogeneic HCT and has been achieved in other genetic diseases using targeted nucleases to induce site specific DSBs and the correction of mutated genes through homology-directed repair (HDR). However, this process is extremely inefficient in FA cells, as they are inherently deficient in DNA repair. Here, we demonstrate the correction of FANCA mutations in primary patient cells using ‘digital’ genome editing with the cytosine and adenine base editors (BEs). These Cas9-based tools allow for C:G > T:A or A:T > C:G base transitions without the induction of a toxic DSB or the need for a DNA donor molecule. These genetic corrections or conservative codon substitution strategies lead to phenotypic rescue as illustrated by a resistance to the alkylating crosslinking agent Mitomycin C (MMC). Further, FANCA protein expression was restored, and an intact FA pathway was demonstrated by downstream FANCD2 monoubiquitination induction. This BE digital correction strategy will enable the use of gene-corrected FA patient hematopoietic stem and progenitor cells (HSPCs) for autologous HCT, obviating the risks associated with allogeneic HCT and DSB induction during autologous HSC gene therapy.

Advance trends in targeting homology-directed repair for accurate gene editing: An inclusive review of small molecules and modified CRISPR-Cas9 systems

Introduction: Clustered regularly interspaced short palindromic repeat and its associated protein (CRISPR-Cas)-based technologies generate targeted modifications in host genome by inducing site-specific double-strand breaks (DSBs) that can serve as a substrate for homology-directed repair (HDR) in both in vitro and in vivo models. HDR pathway could enhance incorporation of exogenous DNA templates into the CRISPR-Cas9-mediated DSB site. Owing to low rate of HDR pathway, the efficiency of accurate genome editing is diminished. Enhancing the efficiency of HDR can provide fast, easy, and accurate technologies based on CRISPR-Cas9 technologies. Methods: The current study presents an overview of attempts conducted on the precise genome editing strategies based on small molecules and modified CRISPR-Cas9 systems. Results: In order to increase HDR rate in targeted cells, several logical strategies have been introduced such as generating CRISPR effector chimeric proteins, anti-CRISPR proteins, modified Cas9 with donor template, and using validated synthetic or natural small molecules for either inhibiting non-homologous end joining (NHEJ), stimulating HDR, or synchronizing cell cycle. Recently, high-throughput screening methods have been applied for identification of small molecules which along with the CRISPR system can regulate precise genome editing through HDR. Conclusion: The stimulation of HDR components or inhibiting NHEJ can increase the accuracy of CRISPR-Cas-mediated engineering systems. Generating chimeric programmable endonucleases provide this opportunity to direct DNA template close proximity of CRISPR-Cas-mediated DSB. Small molecules and their derivatives can also proficiently block or activate certain DNA repair pathways and bring up novel perspectives for increasing HDR efficiency, especially in human cells. Further, high throughput screening of small molecule libraries could result in more discoveries of promising chemicals that improve HDR efficiency and CRISPR-Cas9 systems.

Abstract 5688: RNA-mediated DNA repair: A novel repair pathway in homologous recombination-deficient cancers

Abstract Genome instability has long been considered the primary driver of most cancer types. A double strand break (DSB) in DNA can have deleterious consequences for a cell, which if not repaired faithfully, can lead to mutations and chromosomal rearrangements, or even cell death. DSBs can be processed by several DNA repair pathways, of which homologous recombination (HR) is the preferred method due to its error-free nature. HR uses an intact homologous DNA sequence as a template for recovering the information lost at the break site. A significant proportion of all cancers, especially triple-negative breast, ovarian pancreatic and prostate cancers, have loss of function alterations affecting genes involved in HR-mediated DNA repair. Alternate repair pathways operate when HR is defective in tumors, but the pathways operative in this context remain a matter of contention. Previous work in vivo in yeast and in vitro systems has established a new role of RNA in DNA repair. Owing to its abundance in the cell and its sequence similarity to parental DNA, we sought to define whether RNA can act as a template for the repair of DSBs in human cells. We developed a novel high throughput assay to test if DNA breaks can be repaired using RNA as an alternative template in mammalian cells. Human cells were transfected with a guide RNA cloned in a Cas9 expression vector to generate a site-specific DSB at the AAVS1 locus, a safe harbour, in the human genome. Furthermore, a donor template in the form of DNA or RNA (homologous to the DSB locus) containing a unique mutational signature was provided at the time of transfection. The unique mutational signature enables us to determine if the donor has been utilized as a template for DNA repair. Using this assay, we demonstrate that cells can use a spliced RNA transcript as a functional template to repair a DSB. We have identified that Rev3L, a key component of the translesion synthesis polymerase Pol Zeta (ζ), has a novel reverse-transcriptase activity in human cells and can help repair the DSB using RNA as a template. Further characterization of this repair pathway and its associated mutational scar will provide new insights into the mutational signatures seen in HR-defective cancers, enabling a better understanding of the DNA repair pathways upregulated in these tumours. The proposed studies could help prioritize novel therapeutic approaches by exploiting synthetic lethality in HR-deficient cancers as well as HR-proficient cancers when used in combinatorial cancer therapy. Citation Format: Manisha Jalan, Juber Patel, Kyrie S Olsen, Sana Ahmed-Seghir, Daniel S Higginson, Jorge S Reis-Filho, Nadeem Riaz, Simon N Powell. RNA-mediated DNA repair: A novel repair pathway in homologous recombination-deficient cancers [abstract]. In: Proceedings of the American Association for Cancer Research Annual Meeting 2022; 2022 Apr 8-13. Philadelphia (PA): AACR; Cancer Res 2022;82(12_Suppl):Abstract nr 5688.

Fast cleavage action unveils 3D dynamics and kinetics of DNA repair in telomeres

Understanding the three-dimensional (3D) landscape of the DNA damage response (DDR) is crucial to unveiling genome maintenance mechanisms. Furthermore, accurate kinetic analysis of the DDR mechanisms grants better comprehension of the cellular response from lethal DNA lesions. To achieve the necessary temporal resolution and induce precise, controllable DSBs, we recently developed a novel technology, termed very fast CRISPR (vfCRISPR). This technique enables controllable, site-specific double-strand DNA break (DSB) generation in living cells within only a few seconds of photoactivation of bound Cas9.

Priming Human Hematopoietic Stem and Progenitor Cells for Cas9/Sgrna and rAAV6-Mediated Homologous Recombination

Highly efficient engineered nuclease-mediated gene editing through homologous recombination (HR) in hematopoietic stem and progenitor cells (HSPCs) gives rise to the opportunity of developing gene corrective therapies for hematological disorders. The β-hemoglobinopathies, such as sickle cell disease (SCD) and β-thalassemia, are inherited blood disorders that manifest because of mutations in the beta-globin gene (HBB) and afflict millions of people worldwide. The only curative treatment for these blood disorders is allogeneic-hematopoietic stem cell transplant (allo-HSCT) from a healthy donor, which comes with severe limitations, such as a shortage of immunologically matched donors, graft-versus-host disease, and graft rejection. With the lack of effective curative therapies, it is postulated that autologous HSCT of HBB gene corrected HSPCs would be a potential one-time curative treatment for patients afflicted with β-hemoglobinopathies without the risk of graft versus host diseases and graft rejection, as well as avoiding the need to find an immunologically matched donor. Our previous work demonstrated a methodology to efficiently target the beta-globin gene (HBB) in HSPCs by inducing a site-specific double strand break (DSB) with the RNA-guided (sgRNA) clustered regularly interspaced short palindromic repeats (CRISPR)-Cas9 nuclease (Cas9/sgRNA) system while supplying an homologous HBB gene corrective donor template via recombinant adeno-associated virus serotype 6 (rAAV6). Here, we further build upon that work to identify the critical parameters to reproducibly achieve high levels of HR in HSPCs. By optimizing Cas9 ribonucleoprotein (RNP) concentration and electroporation conditions, we were able to induce DSBs in long-term repopulating hematopoietic stem cells (LT-HSCs), notably in serial transplants engrafting into immunodeficient non-obese diabetic (NOD)- severe combined immunodeficiency (SCID) Il2rg−/− (NSG) mice where we found ~49% of human cells had INDELS in the HBB gene, matching our input, implying the CRISPR/Cas9 system is effective at inducing double strand breaks at the HBB locus in long term repopulating stem cells. We identified that by transducing HSPCs with rAAV6 post-electroporation, there was a two-fold electroporation-aided transduction (EAT) of rAAV6 endocytosis and subsequently higher HR levels. Perhaps the most interesting finding is that when HSPCs are cultured at low densities (100,000 cells/mL) prior to HBB targeting, HSPC expansion rates are significantly positively correlated with HR frequencies. Building upon that work, we show that culturing HBB -targeted HSPCs at low cell densities in the presence of the small molecules, UM171 and SR1, stimulates the expansion of gene-edited HSPCs as measured by higher engraftment levels in NSG mice. This work serves not only as an optimized protocol for genome editing HSPCs at the HBB locus for the treatment of β-hemoglobinopathies, but also as a foundation for editing HSPCs at other loci for both basic and translational research. DisclosuresPorteus:CRISPRTx: Equity Ownership.

A Precise Tool to Edit Human Hematopoietic Stem Cells at High Efficiency and Decipher and Optimize the Different Double Strand Break Repair Outcomes

The CRISPR/Cas9 discovery has revolutionized precise editing of the genome. However, Cas9-induced site-specific double-strand breaks (DSB) are predominantly repaired by the ‘error-prone’ non-homologous end-joining (NHEJ)/alternate NHEJ (aNHEJ) mechanisms, with relatively low precise correction using homology-directed repair (HDR). The HDR-corrected cells can be selected with drugs/surface reporter in cell lines, induced pluripotent cells, mouse embryonic cells. However, selection of gene edited/corrected hematopoietic stem cells (HSC) is not feasible in vitro. HSC remain the most promising targets because of their unique self-renewing, expansion and differentiation potential. Assessment and optimization of editing/correction of the rare human HSC comprising 1-2% of the CD34+ hematopoietic stem and progenitor cell (HSPC) population is particularly challenging, as their selection before/after often results in insufficient cell numbers for detailed molecular analysis; and selection post-editing for clinical translation can reduce recovery/engraftment potential from increased processing. Currently, the readout of the edited HSC is by molecular analysis for NHEJ- and aNHEJ-induced insertions/deletions and HDR detection of total CD34+ cells and assessment of long term repopulating potential (LTRP) in in vivo models. Herein, we report a unique platform that can precisely quantify NHEJ, aNHEJ, HDR and random integration of the HDR donor template in the human HSC using a flow cytometry. We targeted the CD45 surface reporter (PTPRC), present on all leukocytes and HSPC, to induce a DSB in exon 2 of the PTPRC gene using CRISPR/Cas9, and an in-frame promoter-less GFP containing homology template and a downstream tNGFR expression cassette (GFP/tNGFR HDR donor). HSC and HSPC that repaired Cas9-induced DSB via NHEJ on a single allele in diploid cells lost CD45 expression by flow cytometry. Furthermore, the gRNA/Cas9 induced DSB site was designed to have a 3bp micro-homology so that DSB repaired via aNHEJ resulted in cells express CD45 in frame, but a 6bp deletion detectable by TIDE assay. Provision of the GFP/tNGFR HDR donor repair template resulted in GFP and CD45 expressing, but not tNGFR expressing cells, but if the GFP/tNGFR HDR donor integrated randomly, cells were tNGFR+. Using this system, we optimized editing of hematopoietic progenitor cell lines (K562, Kasumi cells) and EBV transformed B-lymphoblasts to >95% efficiency, and molecular analysis of total and CD45+, and CD45- and CD45+GFP+ cell populations confirmed the precision of the flow cytometry based quantification of NHEJ, aNHEJ. In fact, molecular analysis of the CD45+ cells led to the discovery that K562 cells had a high rate of aNHEJ, while this editing via aNHEJ was absent in LCL- and Kasumi- B lymphoid cells. Inhibition of the aNHEJ pathway with small molecule inhibitors/siRNA resulted in loss of the 6bp aNHEJ-mediated deletion in K562 cells. We also found that random integration in LCL, Kasumi and CD34+ cells was a rare event, but occurred with high frequency in K562 cells. Due to the rapid detection of editing events by flow cytometry, we were able to optimize different editing and cell parameters in both cord blood and mobilized peripheral blood derived CD34+ HSPC and CD34+38-90+RA- HSC populations to achieve high viability and 80-87% editing efficiency (CD45-) in HSPC and HSC, and the FACS-based results were similar to molecular analysis. The edited CD34+ HSPC, when injected into NSG mice resulted in robust human cell engraftment, with 45±10% total engraftment, and 39±16% edited cell engraftment (n=12 mice). Using this platform, we improved HDR frequency to 30% and reduced NHEJ events to 60% in hematopoietic cell lines and similar optimization of HDR in HSPC/HSC is underway. In summary, we have developed a novel platform that allows rapid quantification of gene editing by flow cytometry of human HSC at a single cell and a single allele level, in a single step; with concurrent molecular analysis that identifies different DNA repair pathways utilized by different cell types. This platform allows rapid identification of edited events in the rare HSC population, rather than bulk CD34+ HSPC. Furthermore, ability to sort the edited HSC allows precise determination of the genotoxicity of gene editing, and optimization of DSB repair pathway needed, and optimize editing parameters that result in engraftable edited HSC with LTRP. DisclosuresNo relevant conflicts of interest to declare.

CRISPR-Cas-mediated chromosome engineering for crop improvement and synthetic biology.

Plant breeding relies on the presence of genetic variation, as well as on the ability to break or stabilize genetic linkages between traits. The development of the genome-editing tool clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR-associated protein (Cas) has allowed breeders to induce genetic variability in a controlled and site-specific manner, and to improve traits with high efficiency. However, the presence of genetic linkages is a major obstacle to the transfer of desirable traits from wild species to their cultivated relatives. One way to address this issue is to create mutants with deficiencies in the meiotic recombination machinery, thereby enhancing global crossover frequencies between homologous parental chromosomes. Although this seemed to be a promising approach at first, thus far, no crossover frequencies could be enhanced in recombination-cold regions of the genome. Additionally, this approach can lead to unintended genomic instabilities due to DNA repair defects. Therefore, efforts have been undertaken to obtain predefined crossovers between homologues by inducing site-specific double-strand breaks (DSBs) in meiotic, as well as in somatic plant cells using CRISPR-Cas tools. However, this strategy has not been able to produce a substantial number of heritable homologous recombination-based crossovers. Most recently, heritable chromosomal rearrangements, such as inversions and translocations, have been obtained in a controlled way using CRISPR-Cas in plants. This approach unlocks a completely new way of manipulating genetic linkages, one in which the DSBs are induced in somatic cells, enabling the formation of chromosomal rearrangements in the megabase range, by DSB repair via non-homologous end-joining. This technology might also enable the restructuring of genomes more globally, resulting in not only the obtainment of synthetic plant chromosome, but also of novel plant species.

Local nucleosome dynamics and eviction following a double-strand break are reversible by NHEJ-mediated repair in the absence of DNA replication

We interrogated at nucleotide resolution the spatiotemporal order of chromatin changes that occur immediately following a site-specific double-strand break (DSB) upstream of the PHO5 locus and its subsequent repair by nonhomologous end joining (NHEJ). We observed the immediate eviction of a nucleosome flanking the break and the repositioning of adjacent nucleosomes away from the break. These early chromatin events were independent of the end-processing Mre11-Rad50-Xrs2 (MRX) complex and preceded the MRX-dependent broad eviction of histones and DNA end-resectioning that extends up to ∼8 kb away from the break. We also examined the temporal dynamics of NHEJ-mediated repair in a G1-arrested population. Concomitant with DSB repair by NHEJ, we observed the redeposition and precise repositioning of nucleosomes at their originally occupied positions. This re-establishment of the prelesion chromatin landscape suggests that a DNA replication-independent mechanism exists to preserve epigenome organization following DSB repair.

Development of a CRISPR/Cpf1 system for targeted gene disruption in Aspergillus aculeatus TBRC 277

BackgroundCRISPR-Cas genome editing technologies have revolutionized biotechnological research particularly in functional genomics and synthetic biology. As an alternative to the most studied and well-developed CRISPR/Cas9, a new class 2 (type V) CRISPR-Cas system called Cpf1 has emerged as another versatile platform for precision genome modification in a wide range of organisms including filamentous fungi.ResultsIn this study, we developed AMA1-based single CRISPR/Cpf1 expression vector that targets pyrG gene in Aspergillus aculeatus TBRC 277, a wild type filamentous fungus and potential enzyme-producing cell factory. The results showed that the Cpf1 codon optimized from Francisella tularensis subsp. novicida U112, FnCpf1, works efficiently to facilitate RNA-guided site-specific DNA cleavage. Specifically, we set up three different guide crRNAs targeting pyrG gene and demonstrated that FnCpf1 was able to induce site-specific double-strand breaks (DSBs) followed by an endogenous non-homologous end-joining (NHEJ) DNA repair pathway which caused insertions or deletions (indels) at these site-specific loci.ConclusionsThe use of FnCpf1 as an alternative class II (type V) nuclease was reported for the first time in A. aculeatus TBRC 277 species. The CRISPR/Cpf1 system developed in this study highlights the feasibility of CRISPR/Cpf1 technology and could be envisioned to further increase the utility of the CRISPR/Cpf1 in facilitating strain improvements as well as functional genomics of filamentous fungi.

DNA-free genome editing

<p indent=0mm>CRISPR (clustered regularly interspaced short palindromic repeats)-Cas systems have been extensively used for gene editing of many plant species, including model plants such as <italic>Arabidopsis</italic> and rice, and major crops such as maize and wheat, as well as woody plants like poplar, bamboo, apple and citrus, etc. This technique has not only significantly promoted basic research, but has also facilitated the agronomic traits improvement of economically important crops or trees. Typically, the CRISPR/Cas system required its integration and expression of exogenous DNA in the genome of host plants, which lead to the cleavage of site-specific DNA double-strand breaks, and introduced the genome modifications during the DNA repair process. However, the integration of CRISPR/Cas elements brings several concerns for commercialization and basic research. First, the inserted CRISPR/Cas cassette is constitutively expressed in plant cells, and may increase the chance of off-target changes, as well as make it difficult to evaluate the stability and penetrance of the initially observed phenotypes. Second, plants with a foreign gene integrated into its genome were regarded as classic genetically modified organisms, which may potentially lead to the escape of the gene into the environment, leading to legislation concerns and strict government policy regulation about such organisms. Therefore, one of the main challenges in using the CRISPR/Cas system in commercial agriculture is to improve the plant’s agronomic traits by genome editing, meanwhile obtaining transgene-free mutant plants with stable transmissible properties. DNA-free genome editing technology in plants is new and began in 2015, and since then great advances have been made on producing transgenic-free genome-edited plants. The exogenous CRISPR/Cas cassette and other transgenes could be eliminated by multiple approaches after the target gene has been edited. For example, the CRISPR/Cas expression cassette can be removed through genetic segregation for plants that are reproduced sexually, and the end products are CRISPR-edited but transgene-free. Transgene-free gene-edited plants may also be generated by non-transgenic approaches, such as transient expression of mRNA encoding Cas nuclease and guide RNA without DNA integration, preassembled ribonucleoprotein and sgRNA complex transfection, fluorescence marker-assisted transgene cassette selection and elimination, virus-mediated and nano-material mediated genome editing technologies. By using these methods, people can obtain the genome-edited end plant product with increased agronomic traits, while no additional foreign DNA fragments have been introduced into the plant genome. Until now, at least 14 transgene-free and genome-edited plant species have been generated using this technology. In this review, we have highlighted these achievements, including methods, applications and the possibilities of their commercialization in plant breeding. We also discuss the current problems in using this technology, such as very low efficiency, limited gene modification types and labor-consuming aspects. With the development of new plant cell transformation methods and the applications of new typical CRISPR/Cas systems, the potential strategies for improving this revolutionary technology in the future are also discussed.

A single Ho-induced double-strand break at the MAT locus is lethal in Candida glabrata

Mating-type switching is a complex mechanism that promotes sexual reproduction in Saccharomycotina. In the model species Saccharomyces cerevisiae, mating-type switching is initiated by the Ho endonuclease that performs a site-specific double-strand break (DSB) at MAT, repaired by homologous recombination (HR) using one of the two silent mating-type loci, HMLalpha and HMRa. The reasons why all the elements of the mating-type switching system have been conserved in some Saccharomycotina, that do not show a sexual cycle nor mating-type switching, remain unknown. To gain insight on this phenomenon, we used the yeast Candida glabrata, phylogenetically close to S. cerevisiae, and for which no spontaneous and efficient mating-type switching has been observed. We have previously shown that expression of S. cerevisiae's Ho (ScHo) gene triggers mating-type switching in C. glabrata, but this leads to massive cell death. In addition, we unexpectedly found, that not only MAT but also HML was cut in this species, suggesting the formation of multiple chromosomal DSBs upon HO induction. We now report that HMR is also cut by ScHo in wild-type strains of C. glabrata. To understand the link between mating-type switching and cell death in C. glabrata, we constructed strains mutated precisely at the Ho recognition sites. We find that even when HML and HMR are protected from the Ho-cut, introducing a DSB at MAT is sufficient to induce cell death, whereas one DSB at HML or HMR is not. We demonstrate that mating-type switching in C. glabrata can be triggered using CRISPR-Cas9, without high lethality. We also show that switching is Rad51-dependent, as in S. cerevisiae, but that donor preference is not conserved in C. glabrata. Altogether, these results suggest that a DSB at MAT can be repaired by HR in C. glabrata, but that repair is prevented by ScHo.

Hedgehog signaling enables repair of ribosomal DNA double-strand breaks.

Ribosomal DNA (rDNA) consists of highly repeated sequences that are prone to incurring damage. Delays or failure of rDNA double-strand break (DSB) repair are deleterious, and can lead to rDNA transcriptional arrest, chromosomal translocations, genomic losses, and cell death. Here, we show that the zinc-finger transcription factor GLI1, a terminal effector of the Hedgehog (Hh) pathway, is required for the repair of rDNA DSBs. We found that GLI1 is activated in triple-negative breast cancer cells in response to ionizing radiation (IR) and localizes to rDNA sequences in response to both global DSBs generated by IR and site-specific DSBs in rDNA. Inhibiting GLI1 interferes with rDNA DSB repair and impacts RNA polymerase I activity and cell viability. Our findings tie Hh signaling to rDNA repair and this heretofore unknown function may be critically important in proliferating cancer cells.

Monitoring 5'-End Resection at Site-Specific Double-Strand Breaks by Southern Blot Analysis.

DNA double-strand break (DSB) is one of the most deleterious types of DNA lesions threatening genome integrity. Cells have evolved several exquisite pathways to repair these breaks. Homologous recombination (HR) is an essential DSB repair mechanism that utilizes an intact homologous sequence as a template to repair DSBs with high fidelity. To initiate the HR repair, the 5'-ends of DSBs have to be nucleolytically cleaved by nucleases to generate 3'-single-strand DNA (ssDNA). Exposed 3'-ssDNA recruits the ssDNA binding protein complex RPA to activate the DNA damage checkpoint. RPA is subsequently replaced by Rad51 recombinase to form Rad51 nucleoprotein filament that catalyzes strand invasion and formation of the D-loop. Processing of 5'-ends (called resection) is a crucial step that determines the choice of repair pathways. Here we introduce an assay for monitoring the dynamics of resection at different locations from a site-specific DSB in yeast.