Assessing Stealth and Sensed Base Editing in Human Hematopoietic Stem/Progenitor Cells

Genome editing represents a promising tool to manipulate human hematopoietic stem and progenitor cells (HSPCs) opening the possibility to correct hematopoietic diseases avoiding the risk of insertional mutagenesis and uncontrolled expression of the transgene, issues that emerged with retroviral and lentiviral gene therapy. Engineered nucleases such as CRISPR/Cas9 have enable targeted genetic manipulation in human HSPCs for therapeutic purposes. Still, nuclease-induced DNA double-strand breaks (DSBs) trigger p53-dependent DNA damage response affecting HSPC properties and may lead to unintended chromosomal rearrangements. Base editing (BE) holds the promise for precise editing by the introduction of specific single-nucleotide variants (SNVs) while bypassing the requirement for DSBs. In particular, base editors are composed by: i) a deamination domain that directly modifies nucleotides comprised within a defined editing window in one of the two genomic strands, and ii) a nickase Cas9 that introduces a single-strand break (SSB) on the other strand to promote more efficient base editing. Depending on the type of modification introduced editors are classified in Cytosine (C-) BE (C-G transition to T-A) and Adenine (A-) BE (A-T transition to G-C). However, a comprehensive characterization of efficiency, tolerability and genotoxicity of CBE and ABE in human HSPCs is lacking and is required to instruct the rationale towards safe and effective clinical translation. Here, we developed an optimized mRNA-based protocol for BE in human HSPCs and compared CBE4max, ABE8.20-m and Cas9 nuclease by targeting the same locus (B2M) using the same sgRNA. Common outcome for all editors is disruption of targeted gene expression, which is measured by flow cytometry and Next Generation Sequencing. ABE8.20-m showed higher efficiency than CBE4max and Cas9 nuclease at the target locus (up to 90, 40 and 50%), which was consistent across HSPC subpopulations comprising the most primitive compartment endowed with long term repopulation potential and cell sources (such as cord blood- and mobilized peripheral blood-derived HSPCs). Importantly, Cas9, but not CBE4max and ABE8.20m, treated HSPCs showed lower in-vitro clonogenic capacity than mock electroporated cells. Transcriptional analyses uncovered that CBE4max, but not ABE8.20-m, triggered p53 pathway activation, albeit at lower extent as compared to Cas9 and presumably consequent to a fraction of single-strand nicks turning into DSB upon DNA replication. Additionally, BE, and particularly CBE4max, upregulated the expression of interferon-stimulated genes, which was not ascribed to mRNA delivery. Remarkably, despite edited HSPCs showed long-term multilineage capacity in xenotransplanted mice, CBE4max edited cells tended to decrease over time in the graft pointing to some detrimental response to the treatment of the long-term engrafting HSC subset. Overall, our results prompt further investigation on BE sensing in human HSPCs. On-going studies are aimed to investigate clonal dynamics and genome integrity of base-edited HSPCs with the final goal of building confidence for their perspective clinical translation. Disclosures Naldini: Genenta Science: Consultancy, Current equity holder in publicly-traded company, Membership on an entity's Board of Directors or advisory committees, Other: Founder.

Rapid production of SaCas9 in plant-based cell-free lysate for activity testing

Cas9 nucleases have become the most versatile tool for genome editing projects in a broad range of organisms. The recombinant production of Cas9 nuclease is desirable for in vitro activity assays or the preparation of ribonucleoproteins (RNPs) for DNA-free genome editing approaches. For the rapid production of Cas9, we explored the use of a recently established cell-free lysate from tobacco (Nicotiana tabacum L.) BY-2 cells. Using this system, the 130-kDa Cas9 nuclease from Staphylococcus aureus (SaCas9) was produced and subsequently purified via affinity chromatography. The purified apoenzyme was supplemented with ten different sgRNAs, and the nuclease activity was confirmed by the linearization of plasmid DNA containing cloned DNA target sequences.

Alternative strategy to induce CRISPR-mediated genetic changes in hematopoietic cells

Acute Myeloid Leukaemia is a complex heterogenous disease caused by clonal expansion of undifferentiated myeloid precursors. Recently, several haematological models have been developed with CRISPR/Cas9, using viral vectors, because blood cells are hard to transfect. To avoid virus disadvantages, we have developed a strategy to generate CRISPR constructs, by means of PCR, which any lab equipped with basic technology can implement. These PCR-generated constructs enter easily into hard-to-transfect cells. After testing its functionality by editing MYBL2 gene in HEK293 cells, we successfully introduced the R172 mutation in IDH2 gene in NB4 cells that expresses constitutively the Cas9 nuclease. Comparing our methodology with ribonucleoprotein strategies, we found that mutation introduction efficiency was similar between both methodologies, and no off-target events were detected. Our strategy represents a valid alternative to introduce desired mutations in hard to transfect leukemic cells, avoiding using huge vectors or viral transduction.

Plant-based biosensors for detecting CRISPR-mediated genome engineering

CRISPR/Cas has recently emerged as the most reliable system for genome engineering in various species. However, concerns about risks associated with CRISPR/Cas9 technology are increasing on potential unintended DNA changes that might accidentally arise from CRISPR gene editing. Developing a system that can detect and report the presence of active CRIPSR/Cas tools in biological systems is therefore very necessary. Here, we developed the real-time detection systems that can spontaneously indicate CRISPR-Cas tools for genome editing and gene regulation including CRISPR/Cas9 nuclease, base editing, prime editing and CRISPRa in plants. Using the fluorescence-based molecular biosensors, we demonstrated that the activities of CRISPR/Cas9 nuclease, base editing, prime editing and CRIPSRa can be effectively detected in transient expression via protoplast transformation and leaf infiltration (in Arabidopsis, poplar, and tobacco) and stable transformation in Arabidopsis.

MULTIPLEXED SIV-SPECIFIC PAIRED RNA-GUIDED CAS9 NICKASES INACTIVATE PROVIRAL DNA

Human and simian immunodeficiency virus infections establish lifelong reservoir of cells harboring an integrated proviral genome. Genome editing CRISPR-associated Cas9 nucleases, combined with SIV-specific guiding RNA (gRNA) molecules, inactive integrated provirus DNA in vitro and in animal models. We generated RNA-guided Cas9 nucleases (RGNu) and nickases (RGNi) targeting conserved SIV regions with no homology in the human or rhesus macaque genome. Assays in cells co-transfected with SIV provirus and plasmids coding for RGNus identified SIV LTR, TAR, and RSS regions as the most effective at virus suppression; RGNi targeting these same regions inhibited virus production significantly. Multiplex plasmids that co-expressed these three RGNu (Nu3), or six (three pairs) RGNi (Ni6), were more efficient at virus suppression than any combination of individual RGNu and RGNi plasmids. Both Nu3 and Ni6 plasmids were tested in lymphoid cells chronically infected with SIVmac239, and whole genome sequencing was used to determine on- and off-target mutations. Treatment with these all-in-one plasmids resulted in similar levels of mutations of viral sequences from the cellular genome; Nu3 induced indels at the 3 SIV-specific sites, whereas for Ni6 indels were present at the LTR and TAR sites. Levels of off-target effects detected by two different algorithms were indistinguishable from background mutations. In summary, we demonstrate that Cas9 nickase in association with gRNA pairs can specifically eliminate parts of the integrated provirus DNA; also, we show that careful design of an all-in-one plasmid coding for 3 gRNAs and Cas9 nuclease inhibits SIV production with undetectable off-target mutations making these tools a desirable prospect for moving into animal studies. Importance: Our approach to HIV cure, utilizing the translatable SIV/rhesus macaque model system, aims at provirus inactivation and its removal with the least possible off-target side effects. We developed single molecules that delivered either three truncated SIV-specific gRNAs along with Cas9 nuclease, or three pairs of SIV-specific gRNAs (six individual gRNAs) along with Cas9 nickase to enhance efficacy of on-target mutagenesis. Whole genome sequencing demonstrated effective SIV sequence mutation and inactivation, and absence of demonstrable off-target mutations. These results open the possibility to employ Cas9 variants that introduce single-strand DNA breaks to eliminate integrated proviral DNA.

Small molecule inhibition of ATM kinase increases CRISPR-Cas9 1-bp insertion frequency

Mutational outcomes following CRISPR-Cas9-nuclease cutting in mammalian cells have recently been shown to be predictable and, in certain cases, skewed toward single genotypes. However, the ability to control these outcomes remains limited, especially for 1-bp insertions, a common and therapeutically relevant class of repair outcomes. Here, through a small molecule screen, we identify the ATM kinase inhibitor KU-60019 as a compound capable of reproducibly increasing the fraction of 1-bp insertions relative to other Cas9 repair outcomes. Small molecule or genetic ATM inhibition increases 1-bp insertion outcome fraction across three human and mouse cell lines, two Cas9 species, and dozens of target sites, although concomitantly reducing the fraction of edited alleles. Notably, KU-60019 increases the relative frequency of 1-bp insertions to over 80% of edited alleles at several native human genomic loci and improves the efficiency of correction for pathogenic 1-bp deletion variants. The ability to increase 1-bp insertion frequency adds another dimension to precise template-free Cas9-nuclease genome editing.

CRISPR-Cas9 globin editing can induce megabase-scale copy-neutral losses of heterozygosity in hematopoietic cells

CRISPR-Cas9 is a promising technology for gene therapy. However, the ON-target genotoxicity of CRISPR-Cas9 nuclease due to DNA double-strand breaks has received little attention and is probably underestimated. Here we report that genome editing targeting globin genes induces megabase-scale losses of heterozygosity (LOH) from the globin CRISPR-Cas9 cut-site to the telomere (5.2 Mb). In established lines, CRISPR-Cas9 nuclease induces frequent terminal chromosome 11p truncations and rare copy-neutral LOH. In primary hematopoietic progenitor/stem cells, we detect 1.1% of clones (7/648) with acquired megabase LOH induced by CRISPR-Cas9. In-depth analysis by SNP-array reveals the presence of copy-neutral LOH. This leads to 11p15.5 partial uniparental disomy, comprising two Chr11p15.5 imprinting centers (H19/IGF2:IG-DMR/IC1 and KCNQ1OT1:TSS-DMR/IC2) and impacting H19 and IGF2 expression. While this genotoxicity is a safety concern for CRISPR clinical trials, it is also an opportunity to model copy-neutral-LOH for genetic diseases and cancers.

Efficient CRISPR/Cas9 Genome Editing in Alfalfa Using a Public Germplasm

Because its ability to acquire large amounts of nitrogen by symbiosis, tetraploid alfalfa is the main source of vegetable proteins in meat and milk production systems in temperate regions. Alfalfa cultivation also adds fixed nitrogen to the soil, improving the production of non-legumes in crop rotation and reducing the use of nitrogen fertilizers derived from fossil fuel. Despite its economic and ecological relevance, alfalfa genetics remains poorly understood, limiting the development of public elite germplasm. In this brief article, we reported the high-efficiency of alfalfa mutagenesis by using the public clone C23 and the CRISPR/Cas9 system. Around half of the GUS overexpressing plants (35S-GUS under C23 genomic background) transformed with an editing plasmid containing two sgRNAs against the GUS gene and the Cas9 nuclease exhibited absence of GUS activity. Nucleotide analysis showed that the inactivation of GUS in CRISPR/Cas9-editing events were produced via different modifications in the GUS gene, including frameshift and non-sense mutations. Using the CRISPR/Cas9 system and two sgRNAs, we have also edited the alfalfa gene NOD26, generating plants with different doses of alleles at this locus, including complete gene knockout at high efficiency (11%). Finally, we discuss the potential applications of genome-editing technologies to polyploid research and to alfalfa improvement public programs.

Mismatch Intolerance of 5′-Truncated sgRNAs in CRISPR/Cas9 Enables Efficient Microbial Single-Base Genome Editing

The CRISPR/Cas9 system has recently emerged as a useful gene-specific editing tool. However, this approach occasionally results in the digestion of both the DNA target and similar DNA sequences due to mismatch tolerance, which remains a significant drawback of current genome editing technologies. However, our study determined that even single-base mismatches between the target DNA and 5′-truncated sgRNAs inhibited target recognition. These results suggest that a 5′-truncated sgRNA/Cas9 complex could be used to negatively select single-base-edited targets in microbial genomes. Moreover, we demonstrated that the 5′-truncated sgRNA method can be used for simple and effective single-base editing, as it enables the modification of individual bases in the DNA target, near and far from the 5′ end of truncated sgRNAs. Further, 5′-truncated sgRNAs also allowed for efficient single-base editing when using an engineered Cas9 nuclease with an expanded protospacer adjacent motif (PAM; 5′-NG), which may enable whole-genome single-base editing.

An Efficient and Cost-effective Purification Methodology for Cas9 Nuclease

With an ever-increasing demand for laboratory-grade Cas9 proteins by many groups advancing the use of CRISPR technology, a more efficient and scalable process for generating the proteins, coupled with rapid purification methods is in urgent demand. Here, we introduce a modified methodology for rapid purification of active SaCas9 protein within 24 hours. The product has over 90% protein purity. The simplicity and cost-effectiveness of such methodology will enable general labs to produce a sizable amount of Cas9 proteins, further accelerating the advancement of CRISPR/Cas9-based research.