DNA Repair Template Research Articles

Non-viral DNA delivery and TALEN editing correct the sickle cell mutation in hematopoietic stem cells

Sickle cell disease is a devastating blood disorder that originates from a single point mutation in the HBB gene coding for hemoglobin. Here, we develop a GMP-compatible TALEN-mediated gene editing process enabling efficient HBB correction via a DNA repair template while minimizing risks associated with HBB inactivation. Comparing viral versus non-viral DNA repair template delivery in hematopoietic stem and progenitor cells in vitro, both strategies achieve comparable HBB correction and result in over 50% expression of normal adult hemoglobin in red blood cells without inducing β-thalassemic phenotype. In an immunodeficient female mouse model, transplanted cells edited with the non-viral strategy exhibit higher engraftment and gene correction levels compared to those edited with the viral strategy. Transcriptomic analysis reveals that non-viral DNA repair template delivery mitigates P53-mediated toxicity and preserves high levels of long-term hematopoietic stem cells. This work paves the way for TALEN-based autologous gene therapy for sickle cell disease.

CRISPR-Cas9 Genome Editing of Rat Embryos using Adeno-Associated Virus (AAV) and 2-Cell Embryo Electroporation.

Genome editing technology is widely used to produce genetically modified animals, including rats. Cytoplasmic or pronuclear injection of DNA repair templates and CRISPR-Cas reagents is the most common delivery method into embryos. However, this type of micromanipulation necessitates access to specialized equipment, is laborious, and requires a certain level of technical skill. Moreover, microinjection techniques often result in lower embryo survival due to the mechanical stress on the embryo. In this protocol, we developed an optimized method to deliver large DNA repair templates to work in conjunction with CRISPR-Cas9 genome editing without the need for microinjection. This protocol combines AAV-mediated DNA delivery of single-stranded DNA donor templates along with the delivery of CRISPR-Cas9 ribonucleoprotein (RNP) by electroporation to modify 2-cell embryos. Using this novel strategy, we have successfully produced targeted knock-in rat models carrying insertion of DNA sequences from 1.2 to 3.0 kb in size with efficiencies between 42% and 90%.

The strategy of knock-in with homology-directed genome editing in the model ornamental plant Petunia using CRISPR/Cas9 ribonucleoprotein complex

The clustered regularly interspaced short palindromic repeats (CRISPR)/CRISPR-associated system 9 (Cas9) ribonucleoprotein (RNP) complex offers numerous advantages, including increased genome editing efficiency, reduced off-target effects, minimized risk of genomic integration, expanded applicability, and the ability to address specific ethical concerns. These benefits establish the CRISPR/Cas9 RNP complex as a valuable technique for precise genome editing in plant research and applications. Nevertheless, achieving precise knock-in using CRISPR/Cas9-mediated precision knock-in of genes by homology-directed repair (HDR) remains challenging due to the infrequent occurrence of HDR and the limited availability of DNA repair templates near double-strand breaks (DSBs). In this study, we presented the efficient utilization strategy of HDR-mediated gene knock-in through co-infection of the CRISPR/Cas9 RNP complex and donor plasmid in Petunia protoplast. We accomplish this by designing a donor plasmid targeting the chalcone synthase (CHS) in Petunia, consisting of a CRISPR RNA (crRNA)-PAM sequences flanked by a donor DNA cassette with homology arms. These findings offer insights into the application of these techniques in ornamental crops to enhance CRISPR/Cas9 RNA complex-mediated gene knock-in strategies.

Using Inducible Signaling Receptors to Increase Erythropoietic Output from Genome-Edited Hematopoietic Stem Cells

Introduction: Oxygen is delivered to all tissues in the body through the hemoglobin molecule uniquely present in red blood cells (RBCs). Proper production and function of RBCs is therefore critical to human health. However, genetic defects of the hematologic system such as anemias and hemoglobin disorders (broadly termed “hemoglobinopathies”) are among the most common genetic disorders in the world. Therefore, developing strategies to regulate RBC development may allow researchers and clinicians to increase therapeutic potential of hematologic cell and gene therapies. Erythropoietin (EPO) signaling is indispensable to RBC development. In its native form, two EPO receptor (EPOR) monomers dimerize in the presence of EPO cytokine to initiate a downstream signaling cascade. Prior work has shown that conformational requirements of EPOR dimerization are rather “loose” and signaling may also be initiated by agonistic diabodies or in the context of chimeric receptors. While these chimeric receptors have limited clinical applicability, inducible dimerization domains such as engineered FKBPs are currently being used in clinical trials in the form of inducible safety switches. Due to the modular nature of cytokine receptors, we hypothesized that inducible dimerization domains (such as the FKBP domain) could be paired with endogenous receptors and signaling pathways to generate chimeric receptors that place RBC development under the control of a bioavailable small molecule - and therefore in the control researchers and clinicians. This study presents a set of chimeric, synthetic receptors that support HSPC output towards the erythroid lineage via a non-immunogenic, non-toxic small molecule (AP30187, hereafter termed “BB”). Methods: In-vitro editing was performed in primary human CD34+ HSPCs by electroporating Cas9 protein complexed with sgRNA along with an AAV6 DNA repair template to facilitate gene insertion via homology-directed repair (HDR). Bulk edited CD34+ HSPCs were then differentiated into RBCs using established differentiation protocols in the presence and absence of both erythropoietin and BB. Differentiation was determined by staining for RBC surface markers (CD71 and GPA) and subsequent flow cytometry analysis. Erythropoietic output was determined by cell density counts. Allele frequency of the edited cell population was determined through droplet digital PCR. Results: We generated seven FKBP-EPOR chimeric receptors - with the basic structure of SFFV-[FKBP-EPOR]-2A-YFP-bGH - that placed the FKBP binding domain at different regions of the endogenous receptor (Figure 1) and analyzed their ability to support RBC differentiation in the absence of EPO but with BB. Out of the seven constructs we found a set of receptors - 1.4 and 1.5 - that demonstrated erythropoietin-independent, BB-mediated erythroid differentiation at 108% and 94% mock RBC differentiation respectively. Construct 1.5 - dubbed inducible EPOR (iEPOR) - was chosen over 1.4 for further optimization because it was also no longer EPO responsive. We further optimized the design of iEPOR by 1) introducing a secretory signal peptide to improve trafficking of the receptor to the cell surface; 2) inclusion of a naturally occurring EPOR truncation that results in supraphysiological RBC differentiation; and 3) modification of the genomic insertion site and promoter of the iEPOR cassette to generate a toolbox of expression profiles. Our results suggest that iEPOR mimics the native EPOR signaling cascade with high fidelity and sensitivity according to bulk RNA-Seq data. We also observe a significant increase in erythropoietic output for iEPOR version 1.4 (6.7-fold by day 11 of RBC differentiation in +BB over -BB conditions, P=0.003), which was further enhanced by the above modifications for vector iEPOR version 3.5.2 (12.9-fold by day 11 of RBC differentiation in +BB over -BB conditions, P<0.0001). Additionally, iEPOR is demonstrated to be erythropoietin-independent, dose-responsive, and tunable via expression from different loci and promoters. Conclusion:iEPORs represent a powerful new tool to alter the cellular dynamics of red blood cell development. The work demonstrated in this study has the potential to dramatically improve the efficacies of existing gene therapies, offer a route to EPO-free ex-vivo RBC production, as well as serve as a new tool by which to better understand stem cell biology and differentiation.

The enhancement of CRISPR/Cas9 gene editing using metformin

The CRISPR/Cas9 technology is a revolutionary tool that can be used to edit the genome. Specifically, the genome of hematopoietic stem cells (HSCs) could be edited to correct monogenic blood disorders as well as produce immunotherapies. However, the efficiency of editing HSCs remains low. To overcome this hurdle, we set out to investigate the use of metformin, an FDA-approved drug, to enhance gene modification. We assessed the effect of metformin on the growth of two hematopoietic cell lines: a myeloid-erythroid leukemic cell line (K562 cells) representative of the myeloid population and an immortalized T lymphocyte cell line (Jurkat cells) representative of the lymphoid population. No significant difference in growth patterns was observed in concentrations up to 10 mM metformin in both cell lines. We then assessed the ability of two different concentrations of metformin (0.001 mM or 1 mM), based on our observations, to enhance both (1) the cutting efficiency of Cas9 and (2) the targeting efficiency with the use of a donor DNA repair template. The cutting efficiency of Cas9 was significantly enhanced in a total of five guide RNAs (four specific to a platelet locus and one specific to an erythroid locus) following treatment. In addition, an enhancement in targeting was observed with the use of a GFP-containing donor DNA repair template with both concentrations. Overall, a greater than two-fold increase in GFP expression was noted in cells treated with metformin. This suggests that metformin, an FDA-approved drug, could be added to existing protocols to enhance CRISPR/Cas9 gene editing.

Recombinase-independent chromosomal rearrangements between dispersed inverted repeats in Saccharomyces cerevisiae meiosis.

DNA double-strand break (DSB) repair by homologous recombination (HR) uses a DNA template with similar sequence to restore genetic identity. Allelic DNA repair templates can be found on the sister chromatid or homologous chromosome. During meiotic recombination, DSBs preferentially repair from the homologous chromosome, with a proportion of HR events generating crossovers. Nevertheless, regions of similar DNA sequence exist throughout the genome, providing potential DNA repair templates. When DSB repair occurs at these non-allelic loci (termed ectopic recombination), chromosomal duplications, deletions and rearrangements can arise. Here, we characterize in detail ectopic recombination arising between a dispersed pair of inverted repeats in wild-type Saccharomyces cerevisiae at both a local and a chromosomal scale-the latter identified via gross chromosomal acentric and dicentric chromosome rearrangements. Mutation of the DNA damage checkpoint clamp loader Rad24 and the RecQ helicase Sgs1 causes an increase in ectopic recombination. Unexpectedly, additional mutation of the RecA orthologues Rad51 and Dmc1 alters-but does not abolish-the type of ectopic recombinants generated, revealing a novel class of inverted chromosomal rearrangement driven by the single-strand annealing pathway. These data provide important insights into the role of key DNA repair proteins in regulating DNA repair pathway and template choice during meiosis.

Interrogation of human microglial phagocytosis by CRISPR genome editing.

Microglia are an integral part of central nervous system, but our understanding of microglial biology is limited due to the challenges in obtaining and culturing primary human microglia. HMC3 is an important cell line for studying human microglia because it is readily accessible and straightforward to maintain in standard laboratories. Although HMC3 is widely used for microglial research, a robust genetic method has not been described. Here, we report a CRISPR genome editing platform, by the electroporation of Cas9 ribonucleoproteins (Cas9 RNP) and synthetic DNA repair templates, to enable rapid and precise genetic modifications of HMC3. For proof-of-concept demonstrations, we targeted the genes implicated in the regulation of amyloid beta (Aβ) and glioblastoma phagocytosis in microglia. We showed that CRISPR genome editing could enhance the phagocytic activities of HMC3. We performed CRISPR gene knockout (KO) in HMC3 by the electroporation of pre-assembled Cas9 RNP. Co-introduction of DNA repair templates allowed site-specific knock-in (KI) of an epitope tag, a synthetic promoter and a fluorescent reporter gene. The editing efficiencies were determined genotypically by DNA sequencing and phenotypically by immunofluorescent staining and flow cytometry. The gene-edited HMC3 cells were examined in vitro by fluorescent Aβ and glioblastoma phagocytosis assays. Our platform enabled robust single (>90%) and double (>70%) KO without detectable off-target editing by high throughput DNA sequencing. We also inserted a synthetic SFFV promoter to efficiently upregulate the expression of endogenous CD14 and TREM2 genes associated with microglial phagocytosis. The CRISPR-edited HMC3 showed stable phenotypes and enhanced phagocytosis of fluorescence-labeled Aβ1-42 peptides. Confocal microscopy further confirmed the localization of Aβ1-42 aggregates in the acidified lysosomes. HMC3 mutants also changed the phagocytic characteristic toward apoptotic glioblastoma cells. CRISPR genome editing by Cas9 RNP electroporation is a robust approach to genetically modify HMC3 for functional studies such as the interrogation of Aβ and tumor phagocytosis, and is readily adoptable to investigate other aspects of microglial biology.

PTPN22 R620W gene editing in T cells enhances low-avidity TCR responses.

A genetic variant in the gene PTPN22 (R620W, rs2476601) is strongly associated with increased risk for multiple autoimmune diseases and linked to altered TCR regulation and T cell activation. Here, we utilize Crispr/Cas9 gene editing with donor DNA repair templates in human cord blood-derived, naive T cells to generate PTPN22 risk edited (620W), non-risk edited (620R), or knockout T cells from the same donor. PTPN22 risk edited cells exhibited increased activation marker expression following non-specific TCR engagement, findings that mimicked PTPN22 KO cells. Next, using lentiviral delivery of T1D patient-derived TCRs against the pancreatic autoantigen, islet-specific glucose-6 phosphatase catalytic subunit-related protein (IGRP), we demonstrate that loss of PTPN22 function led to enhanced signaling in T cells expressing a lower avidity self-reactive TCR, but not a high-avidity TCR. In this setting, loss of PTPN22 mediated enhanced proliferation and Th1 skewing. Importantly, expression of the risk variant in association with a lower avidity TCR also increased proliferation relative to PTPN22 non-risk T cells. Together, these findings suggest that, in primary human T cells, PTPN22 rs2476601 contributes to autoimmunity risk by permitting increased TCR signaling and activation in mildly self-reactive T cells, thereby potentially expanding the self-reactive T cell pool and skewing this population toward an inflammatory phenotype.

MDM2 antagonists promote CRISPR/Cas9-mediated precise genome editing in sheep primary cells

CRISPR-Cas9-mediated genome editing in sheep is of great use in both agricultural and biomedical applications. While targeted gene knockout by CRISPR-Cas9 through non-homologous end joining (NHEJ) has worked efficiently, the knockin efficiency via homology-directed repair (HDR) remains lower, which severely hampers the application of precise genome editing in sheep. Here, in sheep fetal fibroblasts (SFFs), we optimized several key parameters that affect HDR, including homology arm (HA) length and the amount of double-stranded DNA (dsDNA) repair template; we also observed synchronization of SFFs in G2/M phase could increase HDR efficiency. Besides, we identified three potent small molecules, RITA, Nutlin3, and CTX1, inhibitors of p53-MDM2 interaction, that caused activation of the p53 pathway, resulting in distinct G2/M cell-cycle arrest in response to DNA damage and improved CRISPR-Cas9-mediated HDR efficiency by 1.43- to 4.28-fold in SFFs. Furthermore, we demonstrated that genetic knockout of p53 could inhibit HDR in SFFs by suppressing the expression of several key factors involved in the HDR pathway, such as BRCA1 and RAD51. Overall, this study offersan optimized strategy for the usage of dsDNA repair template, more importantly, the application of MDM2 antagonists provides a simple and efficient strategy to promote CRISPR/Cas9-mediated precise genome editing in sheep primary cells.

PCR cloning Intermediated Gibson assembly (PIG) for Constructing DNA Repair Templates in CRISPR-Cas9 Based Gene Editing.

The CRISPR-Cas9 gene-editing system has revolutionized genome engineering, allowing precise modifications to be made in a wide range of organisms. One significant challenge associated with CRISPR-Cas9 mediated gene editing is the construction of DNA repair templates containing homology arms, a screenable marker and a tag sequence of interest. Here, we present an efficient, two-step strategy to generate DNA repair templates in approximately one week, facilitating rapid and precise genome engineering applications.

A Simple, Improved Method for Scarless Genome Editing of Budding Yeast Using CRISPR-Cas9.

Until recently, the favored method for making directed modifications to the budding yeast genome involved the introduction of a DNA template carrying the desired genetic changes along with a selectable marker, flanked by homology arms. This approach both limited the ability to make changes within genes due to disruption by the introduced selectable marker and prevented the use of that selectable marker for subsequent genomic manipulations. Following the discovery of CRISPR-Cas9-mediated genome editing, protocols were developed for modifying any DNA region of interest in a similar single transformation step without the need for a permanent selectable marker. This approach involves the generation of a DNA double-strand break (DSB) at the desired genomic location by the Cas9 nuclease, expressed on a plasmid which also expresses the guide RNA (gRNA) sequence directing the location of the DSB. The DSB is subsequently repaired via homologous recombination using a PCR-derived DNA repair template. Here, we describe in detail an improved method for incorporation of the gRNA-encoding DNA sequences into the Cas9 expression plasmid. Using Golden Gate cloning, annealed oligonucleotides bearing unique single-strand DNA overhangs are ligated into directional restriction enzyme sites. We describe the use of this CRISPR-Cas9 genome editing protocol to introduce multiple types of directed genetic changes into the yeast genome.

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.

Targeted Gene Mutations in the Forest Pathogen Dothistroma septosporum Using CRISPR/Cas9.

Dothistroma needle blight, caused by Dothistroma septosporum, has increased in incidence and severity over the last few decades and is now one of the most important global diseases of pines. Disease resistance breeding could be accelerated by knowledge of pathogen virulence factors and their host targets. However, this is hindered due to inefficient targeted gene disruption in D. septosporum, which is required for virulence gene characterisation. Here we report the first successful application of CRISPR/Cas9 gene editing to a Dothideomycete forest pathogen, D. septosporum. Disruption of the dothistromin pathway regulator gene AflR, with a known phenotype, was performed using nonhomologous end-joining repair with an efficiency of >90%. Transformants with a range of disruption mutations in AflR were produced. Disruption of Ds74283, a D. septosporum gene encoding a secreted cell death elicitor, was also achieved using CRISPR/Cas9, by using a specific donor DNA repair template to aid selection where the phenotype was unknown. In this case, 100% of screened transformants were identified as disruptants. In establishing CRISPR/Cas9 as a tool for gene editing in D. septosporum, our research could fast track the functional characterisation of candidate virulence factors in D. septosporum and helps set the foundation for development of this technology in other forest pathogens.

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.

An update on precision genome editing by homology-directed repair in plants.

Beneficial alleles derived from local landraces or related species, or even orthologs from other plant species, are often caused by differences of one or several single-nucleotide polymorphisms or indels in either the promoter region or the encoding region of a gene and often account for major differences in agriculturally important traits. Clustered regularly interspaced short palindromic repeats-associated endonuclease Cas9 system (CRISPR/Cas9)-mediated precision genome editing enables targeted allele replacement or insertion of flag or foreign genes at specific loci via homology-directed repair (HDR); however, HDR efficiency is low due to the intrinsic rare occurrence of HDR and insufficient DNA repair template in the proximity of a double-stranded break (DSB). Precise replacement of the targeted gene with elite alleles from landraces or relatives into a commercial variety through genome editing has been a holy grail in the crop genome editing field. In this update, we briefly summarize CRISPR/Cas-mediated HDR in plants. We describe diverse strategies to improve HDR efficiency by manipulating the DNA repair pathway, timing DSB induction, and donor delivery, and so on. Lastly, we outline open questions and challenges in HDR-mediated precision genome editing in both plant biological research and crop improvement.

Prime Editing for Inherited Retinal Diseases.

Inherited retinal diseases (IRDs) are chronic, hereditary disorders that lead to progressive degeneration of the retina. Disease etiology originates from a genetic mutation—inherited or de novo—with a majority of IRDs resulting from point mutations. Given the plethora of IRDs, to date, mutations that cause these dystrophies have been found in approximately 280 genes. However, there is currently only one FDA-approved gene augmentation therapy, Luxturna (voretigene neparvovec-rzyl), available to patients with RPE65-mediated retinitis pigmentosa (RP). Although clinical trials for other genes are underway, these techniques typically involve gene augmentation rather than genome surgery. While gene augmentation therapy delivers a healthy copy of DNA to the cells of the retina, genome surgery uses clustered regularly interspaced short palindromic repeats (CRISPR)-based technology to correct a specific genetic mutation within the endogenous genome sequence. A new technique known as prime editing (PE) applies a CRISPR-based technology that possesses the potential to correct all twelve possible transition and transversion mutations as well as small insertions and deletions. EDIT-101, a CRISPR-based therapy that is currently in clinical trials, uses double-strand breaks and nonhomologous end joining to remove the IVS26 mutation in the CEP290 gene. Preferably, PE does not cause double-strand breaks nor does it require any donor DNA repair template, highlighting its unparalleled efficiency. Instead, PE uses reverse transcriptase and Cas9 nickase to repair mutations in the genome. While this technique is still developing, with several challenges yet to be addressed, it offers promising implications for the future of IRD treatment.

Mechanistic and genetic basis of single-strand templated repair at Cas12a-induced DNA breaks in Chlamydomonas reinhardtii

Single-stranded oligodeoxynucleotides (ssODNs) are widely used as DNA repair templates in CRISPR/Cas precision genome editing. However, the underlying mechanisms of single-strand templated DNA repair (SSTR) are inadequately understood, constraining rational improvements to precision editing. Here we study SSTR at CRISPR/Cas12a-induced DNA double-strand breaks (DSBs) in the eukaryotic model green microalga Chlamydomonas reinhardtii. We demonstrate that ssODNs physically incorporate into the genome during SSTR at Cas12a-induced DSBs. This process is genetically independent of the Rad51-dependent homologous recombination and Fanconi anemia pathways, is strongly antagonized by non-homologous end-joining, and is mediated almost entirely by the alternative end-joining enzyme polymerase θ. These findings suggest differences in SSTR between C. reinhardtii and animals. Our work illustrates the promising potentially of C. reinhardtii as a model organism for studying nuclear DNA repair.

Introduction and Genetic Correction of Calreticulin Mutations in Human Hematopoietic Stem and Progenitor Cells Sheds Light on MPN Pathogenesis

Introduction:Recurrent mutations in calreticulin (CALR) are present in 70% to 80% of essential thrombocythemia (ET) and primary myelofibrosis (PMF) patients without a JAK2 or MPL mutation. Despite recent advances in understanding mutant CALR, the detailed mechanisms are not fully elucidated, and current knowledge is mainly based on transgenic mouse models or human cancer cell lines. Thus, to more faithfully model MPN pathogenesis, we first aimed to introduce heterozygous type-1 and type-2 CALR mutations into healthy human hematopoietic stem and progenitor cells (HSPCs) via targeted CRISPR/Cas9-mediated gene knock-in (KI) and investigate its impact on HSPC function in vitro and in vivo. Second, we aimed to correct CALR mutations in patient-derived HSPCs to study their dependence on the initial driver event to exert an MPN phenotype.Methods:We used CRISPR/Cas9 to introduce heterozygous CALR mutations into the endogenous gene locus of healthy cord blood-derived HSPCs. Our approach is based on homologous recombination using DNA repair templates delivered by adeno-associated virus serotype 6 (AAV6). Briefly, Cas9-sgRNA ribonucleoprotein (RNP) was used to cut the DNA. Simultaneously AAV6, carrying either a mutation-bearing or a wildtype control cDNA, was co-delivered to allow for targeted in-frame integration. This way, mutant CALR remains under the control of the endogenous promoter. Concurrent integration of a fluorescent reporter downstream of the mutated exon, enabled purification and tracking of modified cells via flow cytometry. Purified CRISPR-modified HSPCs were used for in vitro collagen-based colony-forming assays, proliferation and differentiation assays in liquid culture, and intrafemoral transplantation into immunodeficient NSG mice to assess their pathogenic potential.Results:Our CRISPR/Cas9 KI strategy enabled us to efficiently generate and enrich for heterozygous CALR mutant human HSPCs. Modified cells harbor the mutation at the endogenous CALR locus with intact gene regulatory regions. Correct integration and transcript expression were confirmed on DNA and RNA level by sanger sequencing. Additionally, CALR mutant protein expression was confirmed via immunohistochemistry using a diagnostically approved mutant-specific antibody.Type-1 and type-2 CALR mutations led to TPO-independent growth of CD34 + HSPC-derived cells and a two-fold (p<0.01) increase of megakaryocyte colonies in collagen-based media compared to wildtype control KI. These findings were corroborated by significantly enhanced CD41 + CD42b + megakaryocyte formation of CALR mutant HSPCs upon liquid culture differentiation.When transplanted into sublethally irradiated immunodeficient NSG mice, CALR mutant HSPCs showed robust engraftment in the bone marrow with a myeloid lineage skewing, outcompetition of wildtype cells and increased formation of CALR mutant CD41 + megakaryocyte progenitors.To investigate, if removal of type-1 and type-2 CALR mutations can ameliorate MPNs, we utilized our KI strategy to correct both CALR mutations in MPN patient-derived HSPCs by replacing them with wildtype sequences. A successful correction was confirmed on DNA and RNA level and by the absence of mutant CALR protein. Opposite to the results from introducing CALR mutations, correcting the mutations led to a two-fold decrease in megakaryocyte colony formation. Interestingly this was only seen in ET and post-ET MF samples, whereas primary MF samples were unaffected, underscoring the importance of other secondary genetic driver events in the pathogenesis of primary MF.Conclusion:Our system allows us to investigate human MPN pathogenesis prospectively and shed light on the transforming mechanisms of mutant CALR in primary HSPCs. We could show that CALR mutations prime HSPCs toward the formation of platelet-producing megakaryocytes. Genetic correction of CALR mutations in MPN patient-derived HSPCs revealed a dependence on the oncogenic mutant CALR driver event in ET and post-ET MF patients, opening the possibility of an ex vivo gene correction approach to remove mutant CALR in patient-derived HSPCs . Lastly, since MPN patient-derived cells have notoriously low engraftment potential in mice, our CRISPR/Cas9-engineered CALR mutant model also provides a powerful new strategy to generate MPN xenotransplants with defined genotypes for the evaluation of novel therapies. DisclosuresGreinix: Celgene: Consultancy; Therakos: Consultancy; Takeda: Consultancy; Sanofi: Consultancy; Novartis: Consultancy. Sill: Astellas: Consultancy, Membership on an entity's Board of Directors or advisory committees; Celgene: Consultancy, Membership on an entity's Board of Directors or advisory committees; AbbVie: Consultancy, Membership on an entity's Board of Directors or advisory committees. Zebisch: Novartis: Consultancy; AbbVie: Consultancy; Celgene: Consultancy, Honoraria. Reinisch: Celgene: Research Funding; Pfizer: Consultancy.

Modeling the Development of SRSF2 Mutated Myeloid Malignancies By CRISPR/Cas9 Mediated Genome Engineering of Primary Human Hematopoietic Stem and Progenitor Cells

Introduction: Acute Myeloid Leukemia (AML) is a malignant disease of the bone marrow that can arise from a premalignant condition called clonal hematopoiesis of indeterminate potential (CHIP). Mutations in Serine and Arginine-rich Splicing Factor 2 (SRSF2) are detected in CHIP and mediate a high risk for AML development. Here we used CRISPR/Cas9-mediated genome engineering to introduce a heterozygous SRSF2P95H mutation into primary human hematopoietic stem and progenitor cells (HSPCs) and investigated its functional consequences using both in vitro and in vivo assays.Methods: We used CRISPR/Cas9 technology to introduce a heterozygous mutant (mut) SRSF2P95H into the endogenous SRSF2 gene locus of healthy cord blood HSPCs. Our approach is based on homologous recombination using DNA repair templates delivered by adeno-associated virus serotype 6 (AAV6) (Figure A). This allows for targeted in-frame integration of mut and/or wildtype (WT) SRSF2 cDNA under the control of the endogenous SRSF2 promoter. Notably, an integrated fluorescent reporter enables the isolation and tracking of heterozygously mutated HSPCs (Figure B). Methylcellulose colony and long-term competition assays of SRSF2 mut and WT HSPCs were performed in vitro. Cells were analyzed by flow cytometry and characterized cytomorphologically. In addition, bulk RNA-seq analyses were performed to characterize differential gene expression and abnormal splicing events. Xenotransplantation into NSG-SGM3 mice was performed in order to assess stem cell characteristics and the in vivo leukemogenic potential of SRSF2 mut HSPCs. Finally, we investigated the mutation-specific effect of the splicing inhibitor Indisulam to determine if SRSF2 mut cells are particularly vulnerable to splicing inhibition.Results: Colony assays (n=9) revealed impaired erythroid and increased monocytic differentiation of SRSF2 mut HSPCs. Quantification of colonies showed a lower frequency of erythroid BFU-E in SRSF2 mut compared to SRSF2 WT HSPCs (mean ± SD; 33.3 ± 12.5% vs. 17.4 ± 10.8%, p=0.00002). In contrast, the frequency of myeloid CFU-M colonies was higher in SRSF2 mut HSPCs compared to SRSF2 WT HSPCs (38.3 ± 7.3% vs. 22.6 ± 6.8%, p = 0.0003) (Figure C). Long-term in vitro competition assays revealed an outgrowth of SRSF2 mut over WT cells in 2 out of 7 donors. Strikingly, after three months of in vitro culture, in one donor, the SRSF2 mut cells developed a blast-like morphology with strong CD34 expression (Figure D). To assess stem cell characteristics and the leukemogenic potential in vivo, we transplanted SRSF2 mut HSPCs from 4 different donors into immunodeficient NSG-SGM3 mice (n=11). SRSF2 mut cells showed a myeloid-skewed engraftment. Cytomorphologic analysis of long-term engrafted SRSF2 mut myeloid cells revealed dysplastic changes such as nuclear abnormalities and extensive cytoplasmic vacuolization. In 4 out of 11 xenografts, human engraftment substantially increased over time with a parallel outgrowth of the SRSF2 mut clone and the appearance of blast-like cells resembling transformation into myeloid leukemia (Figure E). Comparative RNA-seq analysis identified 138 differentially spliced genes, with exon skipping being the dominant altered splicing type. Gene ontology (GO) analysis on differentially expressed genes revealed “Acute Myeloid Leukemia” among the most enriched terms (p-val = 8.2E-07, min FDR = 1.486E-04). When testing the SRSF2-mutation specific effect of the splicing inhibitor Indisulam, SRSF2 mut HSPCs show a significantly lower IC-50 than WT cells (977nM vs. 3574 nM). Strikingly, in competition- and CFU-assays, Indisulam preferentially eradicates SRSF2 mut hematopoietic cells, while sparing WT cells.Conclusion: Using our CRISPR/Cas9 approach, we can successfully introduce heterozygous SRSF2P95H mutants in primary human HSPCs. Mutant SRSF2P95H leads to increased monocytic differentiation, impaired erythroid differentiation, and phenocopy SRSF2P95H driven diseases in patients. Importantly, we show for the first time that the SRSF2 mutation alone is sufficient to induce dysplastic features and even transform healthy human HSPCs into AML-like blasts. Our model allows the identification and therapeutic investigation of specific cellular vulnerabilities caused by SRSF2 mutations and highlights Indisulam as a potential compound to specifically treat individuals carrying a SRSF2 mutation. [Display omitted] DisclosuresEdiriwickrema: Nanosive SAS: Patents & Royalties. Greinix: Novartis: Consultancy; Celgene: Consultancy; Takeda: Consultancy; Sanofi: Consultancy; Therakos: Consultancy. Sill: Astellas: Consultancy, Membership on an entity's Board of Directors or advisory committees; Celgene: Consultancy, Membership on an entity's Board of Directors or advisory committees; AbbVie: Consultancy, Membership on an entity's Board of Directors or advisory committees. Zebisch: Celgene: Consultancy, Honoraria; AbbVie: Consultancy; Novartis: Consultancy. Majeti: BeyondSpring Inc.: Membership on an entity's Board of Directors or advisory committees; CircBio Inc.: Membership on an entity's Board of Directors or advisory committees; Kodikaz Therapeutic Solutions Inc.: Membership on an entity's Board of Directors or advisory committees; Coherus Biosciences: Membership on an entity's Board of Directors or advisory committees; Acuta Capital Partners: Consultancy; Gilead: Patents & Royalties: inventor on a number of patents related to CD47 cancer immunotherapy licensed to Gilead Sciences, Inc.. Reinisch: Pfizer: Consultancy; Celgene: Research Funding.

Genome editing of Corynebacterium glutamicum mediated with Cpf1 plus Ku/LigD.

Corynebacterium glutamicum (C. glutamicum) has been harnessed for multi-million-ton scale production of glutamate and lysine. To further increase its amino acid production for fermentation industry, there is an acute need to develop next-generation genome manipulation tool for its metabolic engineering. All reported methods for genome editing triggered with CRISPR-Cas are based on the homologous recombination. While, it requires the generation of DNA repair template, which is a bottle-neck for its extensive application. In this study, we developed a method for gene knockout in C. glutamicum via CRISPR-Cpf1-coupled non-homologous end-joining (CC-NHEJ). Specifically, CRISPR-Cpf1 introduced double-strandbreaks in the genome of C. glutamicum, which was further repaired by ectopically expressed two NHEJ key proteins (Mycobacterium tuberculosis Ku and ligase D). We provide the proof of concept, for CC-NHEJ, by the successful knockout of the crtYf/e gene in C. glutamicum with the efficiency of 22.00 ± 5.56%, or something like that. The present study reported a novel genome manipulation method for C. glutamicum.