Non-edited Cells Research Articles

Genetic and functional correction of argininosuccinate lyase deficiency using CRISPR adenine base editors

Argininosuccinate lyase deficiency (ASLD) is a recessive metabolic disorder caused by variants in ASL. In an essential step in urea synthesis, ASL breaks down argininosuccinate (ASA), a pathognomonic ASLD biomarker. The severe disease forms lead to hyperammonemia, neurological injury, and even early death. The current treatments are unsatisfactory, involving a strict low-protein diet, arginine supplementation, nitrogen scavenging, and in some cases, liver transplantation. An unmet need exists for improved, efficient therapies. Here, we show the potential of a lipid nanoparticle-mediated CRISPR approach using adenine base editors (ABEs) for ASLD treatment. To model ASLD, we first generated human-induced pluripotent stem cells (hiPSCs) from biopsies of individuals homozygous for the Finnish founder variant (c.1153C>T [p.Arg385Cys]) and edited this variant using the ABE. We then differentiated the hiPSCs into hepatocyte-like cells that showed a 1,000-fold decrease in ASA levels compared to those of isogenic non-edited cells. Lastly, we tested three different FDA-approved lipid nanoparticle formulations to deliver the ABE-encoding RNA and the sgRNA targeting the ASL variant. This approach efficiently edited the ASL variant in fibroblasts with no apparent cell toxicity and minimal off-target effects. Further, the treatment resulted in a significant decrease in ASA, to levels of healthy donors, indicating restoration of the urea cycle. Our work describes a highly efficient approach to editing the disease-causing ASL variant and restoring the function of the urea cycle. This method relies on RNA delivered by lipid nanoparticles, which is compatible with clinical applications, improves its safety profile, and allows for scalable production.

Abstract 1785: Multi-step engineering of gene-edited CAR T cells using RNA lipid nanoparticles

Abstract Autologous chimeric antigen receptor (CAR) T therapies utilize patient cells and can be limited by cell quality, and the high manufacturing burden of viral vectors. As such, there is a need for allogeneic, “off-the-shelf” CAR T cells to make these transformative treatments widely available. However, allogeneic therapies require multiple genetic engineering steps to express CAR and to delete proteins responsible for graft-versus-host disease. Messenger RNA (mRNA) is a promising approach for expression of therapeutic proteins and gene editing nucleases. In this work, we demonstrate a new method for multi-step engineering of gene-edited CAR T cells using RNA lipid nanoparticles (LNPs). LNPs encapsulating Spy-Cas9 mRNA, TCR and CD52 guide RNA (sgRNA), and CAR mRNA were produced using microfluidics. The CAR construct contained an anti-CD19 scFv binding domain and CD3ζ/4-1BB co-stimulatory domains. Microgram quantities of RNA LNPs were produced to optimize LNP packaging, cargo ratios, and sgRNA combinations. Lead candidates were scaled to milligrams. Purified human primary T cells were cultured, activated, and expanded in serum-free media in plates, flasks and bioreactors. CAR+, TCR− or CD52− cells were generated by addition of the corresponding LNP to activated cells. Cytotoxic killing was determined by co-culture assays with leukemia cells. Gene knockout, CAR expression, viability and cell killing were measured using flow-cytometry. CD19 CAR was selected as a relevant protein for expression, with TCR and CD52 proteins as gene knockout targets. Single-step addition of CAR LNPs to T cells resulted in transfection efficiencies of 95.0 ± 2.1% and high protein expression. Upon TCR or CD52 LNP addition to T cells, the onset of gene editing was within 48 hours, reaching single target knockout efficiencies of 92.3 ± 3.0% (TCR−), and double knockouts (TCR−/CD52−) of 74.5 ± 6.1%. Similar results were obtained when comparing different LNP batch sizes (microgram to milligram RNA) and cell culture vessels (125,000 to 45 million cells), demonstrating scalability of both the LNP production and cell treatment. Cell viabilities above 90% were maintained at all steps and for all RNA LNPs. Finally, as proof-of-concept for multi-step engineering, sequential addition of TCR LNPs and CAR LNPs resulted in simultaneous CAR expression and TCR gene knockout. These “off-the-shelf” gene-edited CAR T cells were functionally equivalent to non-edited cells in a B cell killing assay, efficiently clearing over 80% of leukemia target cells at a 1:1 ratio. Our findings demonstrate the advantages of LNPs for RNA delivery to T cells. The simple and gentle nature of LNP cell treatment allows for multiple genetic engineering steps for simultaneous expression and deletion of proteins. Furthermore, LNPs can be easily manufactured using microfluidics, enabling small-scale screening of RNA libraries and rapid scale-up of lead candidates for clinical translation. Citation Format: Samuel Clarke, R Geczy, A Balgi, S Park, R Zhao, M Swaminathan, R Tieu, N Hoang, C Webb, E Watt, M Wong, M Fujisawa, N Jain, Angela Zhang, Anitha Thomas. Multi-step engineering of gene-edited CAR T cells using RNA lipid nanoparticles [abstract]. In: Proceedings of the American Association for Cancer Research Annual Meeting 2023; Part 1 (Regular and Invited Abstracts); 2023 Apr 14-19; Orlando, FL. Philadelphia (PA): AACR; Cancer Res 2023;83(7_Suppl):Abstract nr 1785.

Gene Editing Profiles in 94 CRISPR-Cas9 Expressing T0 Transgenic Tobacco Lines Reveal High Frequencies of Chimeric Editing of the Target Gene.

Chimeric editing is often reported in gene editing. To assess how the general chimeric editing is, we created a transgenic tobacco line carrying a marker, beta-glucuronidase gene (gusA), introduced a CRISPR-Cas9 editing vector into the transgenic tobacco line for knocking out gusA, and then investigated the gusA editing efficiencies in T0 and subsequent generations. The editing vector carried a Cas9 gene, which was driven by the cauliflower mosaic virus 35S promoter, and two guide RNAs, gRNA1 and gRNA2, which were driven by Arabidopsis U6 (AtU6) and U3 (AtU3) promoter, respectively. The two gRNAs were designed to knock out a 42-nucleotide fragment of the coding region of gusA. The editing vector was transformed into gusA-containing tobacco leaves using Agrobacterium tumefaciens-mediated transformation and hygromycin selection. Hygromycin-resistant, independent T0 transgenic lines were used to evaluate gusA-editing efficiencies through histochemical GUS assays, polymerase chain reactions (PCR), and next-generation sequencing of PCR amplicons. Profiles of targeted sequences of 94 T0 transgenic lines revealed that these lines were regenerated from non-edited cells where subsequent editing occurred and created chimeric-edited cells in these lines during or after regeneration. Two of them had the target fragment of 42 bp pairs of nucleotides removed. Detail analysis showed that on-target mutations at the AtU6-gRNA1 site and the AtU3-gRNA2 site were found in 4.3% and 77.7% of T0 transgenic lines, respectively. To overcome the issue of extremely low editing efficiencies in T0 lines, we conducted a second round of shoot induction from the chimeric line(s) to enhance the success of obtaining lines with all or most cells edited. The mutation profiles in T0 transgenic lines provide valuable information to understand gene editing in plant cells with constitutively expressed CRISPR-Cas9 and gRNAs.

NT-CRISPR, combining natural transformation and CRISPR-Cas9 counterselection for markerless and scarless genome editing in Vibrio natriegens

The fast-growing bacterium Vibrio natriegens has recently gained increasing attention as a novel chassis organism for fundamental research and biotechnology. To fully harness the potential of this bacterium, highly efficient genome editing methods are indispensable to create strains tailored for specific applications. V. natriegens is able to take up free DNA and incorporate it into its genome by homologous recombination. This highly efficient natural transformation is able to mediate uptake of multiple DNA fragments, thereby allowing for multiple simultaneous edits. Here, we describe NT-CRISPR, a combination of natural transformation with CRISPR-Cas9 counterselection. In two temporally distinct steps, we first performed a genome edit by natural transformation and second, induced CRISPR-Cas9 targeting the wild type sequence, and thus leading to death of non-edited cells. Through cell killing with efficiencies of up to 99.999%, integration of antibiotic resistance markers became dispensable, enabling scarless and markerless edits with single-base precision. We used NT-CRISPR for deletions, integrations and single-base modifications with editing efficiencies of up to 100%. Further, we confirmed its applicability for simultaneous deletion of multiple chromosomal regions. Lastly, we showed that the near PAM-less Cas9 variant SpG Cas9 is compatible with NT-CRISPR and thereby broadens the target spectrum.

HEK293T Cells with TFAM Disruption by CRISPR-Cas9 as a Model for Mitochondrial Regulation.

The mitochondrial transcription factor A (TFAM) is considered a key factor in mitochondrial DNA (mtDNA) copy number. Given that the regulation of active copies of mtDNA is still not fully understood, we investigated the effects of CRISPR-Cas9 gene editing of TFAM in human embryonic kidney (HEK) 293T cells on mtDNA copy number. The aim of this study was to generate a new in vitro model by CRISPR-Cas9 system by editing the TFAM locus in HEK293T cells. Among the resulting single-cell clones, seven had high mutation rates (67–96%) and showed a decrease in mtDNA copy number compared to control. Cell staining with Mitotracker Red showed a reduction in fluorescence in the edited cells compared to the non-edited cells. Our findings suggest that the mtDNA copy number is directly related to TFAM control and its disruption results in interference with mitochondrial stability and maintenance.

CRISPRroots: on- and off-target assessment of RNA-seq data in CRISPR-Cas9 edited cells.

The CRISPR-Cas9 genome editing tool is used to study genomic variants and gene knockouts, and can be combined with transcriptomic analyses to measure the effects of such alterations on gene expression. But how can one be sure that differential gene expression is due to a successful intended edit and not to an off-target event, without performing an often resource-demanding genome-wide sequencing of the edited cell or strain? To address this question we developed CRISPRroots: CRISPR–Cas9-mediated edits with accompanying RNA-seq data assessed for on-target and off-target sites. Our method combines Cas9 and guide RNA binding properties, gene expression changes, and sequence variants between edited and non-edited cells to discover potential off-targets. Applied on seven public datasets, CRISPRroots identified critical off-target candidates that were overlooked in all of the corresponding previous studies. CRISPRroots is available via https://rth.dk/resources/crispr.

Development of a Cas12a-Based Genome Editing Tool for Moderate Thermophiles.

The ability of CRISPR-Cas12a nucleases to function reliably in a wide range of species has been key to their rapid adoption as genome engineering tools. However, so far, Cas12a nucleases have been limited for use in organisms with growth temperatures up to 37 °C. Here, we biochemically characterize three Cas12a orthologs for their temperature stability and activity. We demonstrate that Francisella novicida Cas12a (FnCas12a) has great biochemical potential for applications that require enhanced stability, including use at temperatures >37°C. Furthermore, by employing the moderate thermophilic bacterium Bacillus smithii as our experimental platform, we demonstrate that FnCas12a is active in vivo at temperatures up to 43°C. Subsequently, we develop a single-plasmid FnCas12a-based genome editing tool for B. smithii, combining the FnCas12a targeting system with plasmid-borne homologous recombination (HR) templates that carry the desired modifications. Culturing of B. smithii cells at 45°C allows for the uninhibited realization of the HR-based editing step, while a subsequent culturing step at reduced temperatures induces the efficient counterselection of the non-edited cells by FnCas12a. The developed gene-editing tool yields gene-knockout mutants within 3 days, and does not require tightly controllable expression of FnCas12a to achieve high editing efficiencies, indicating its potential for other (thermophilic) bacteria and archaea, including those with minimal genetic toolboxes. Altogether, our findings provide new biochemical insights into three widely used Cas12a nucleases, and establish the first Cas12a-based bacterial genome editing tools for moderate thermophilic microorganisms.

Bacterial Genetic Engineering by Means of Recombineering for Reverse Genetics.

Serving a robust platform for reverse genetics enabling the in vivo study of gene functions primarily in enterobacteriaceae, recombineering -or recombination-mediated genetic engineering-represents a powerful and relative straightforward genetic engineering tool. Catalyzed by components of bacteriophage-encoded homologous recombination systems and only requiring short ∼40–50 base homologies, the targeted and precise introduction of modifications (e.g., deletions, knockouts, insertions and point mutations) into the chromosome and other episomal replicons is empowered. Furthermore, by its ability to make use of both double- and single-stranded linear DNA editing substrates (e.g., PCR products or oligonucleotides, respectively), lengthy subcloning of specific DNA sequences is circumvented. Further, the more recent implementation of CRISPR-associated endonucleases has allowed for more efficient screening of successful recombinants by the selective purging of non-edited cells, as well as the creation of markerless and scarless mutants. In this review we discuss various recombineering strategies to promote different types of gene modifications, how they are best applied, and their possible pitfalls.

GOTI, a method to identify genome-wide off-target effects of genome editing in mouse embryos.

Genome editing holds great potential for correcting pathogenic mutations. We developed a method called GOTI (genome-wide off-target analysis by two-cell embryo injection) to detect off-target mutations by editing one blastomere of two-cell mouse embryos using either CRISPR-Cas9 or base editors. GOTI directly compares edited and non-edited cells without the interference of genetic background and thus could detect potential off-target variants with high sensitivity. Notably, the GOTI method was designed to detect potential off-target variants of any genome editing tools by the combination of experimental and computational approaches, which is critical for accurate evaluation of the safety of genome editing tools. Here we provide a detailed protocol for GOTI, including mice mating, two-cell embryo injection, embryonic day 14.5 embryo digestion, fluorescence-activated cell sorting, whole-genome sequencing and data analysis. To enhance the utility of GOTI, we also include a computational workflow called GOTI-seq (https://github.com/sydaileen/GOTI-seq) for the sequencing data analysis, which can generate the final genome-wide off-target variants from raw sequencing data directly. The protocol typically takes 20 d from the mice mating to sequencing and 7 d for sequencing data analysis.

3117 – GPRASP2 REGULATES CXCR4 STABILITY AND SIGNAL TRANSDUCTION IN HUMAN HEMATOPOIETIC CELLS WITH IMPLICATIONS FOR HSPC TRANSPLANTATION

Transplantation of hematopoietic stem and progenitor cells (HSPCs) is a life-saving therapy that is limited by donor availability and low cell numbers in sources such as cord blood. Advances in our understanding of the biochemical pathways that determine HSPC engraftment will allow for the optimization of transplantation strategies. We previously found that knockdown of GPRASP (G-protein Coupled Receptor Associated Sorting Protein) protein family members increases CXCR4 abundance and transplantation efficiency in murine HSPCs. Here, we demonstrate that GPRASP2 facilitates the acute agonist-induced degradation of CXCR4 in human hematopoietic cells and limits signaling downstream of CXCR4. Initial data also indicate that loss of GPRASP2 may increase the transplantation efficiency of human CD34+ cells. An antibody protection assay performed in HL60 cells demonstrated that knockdown of GPRASP2 significantly increases stability of CXCR4 following SDF-1 treatment. Phosphoflow experiments were performed to quantify the impact of GPRASP2 on signaling pathways downstream of CXCR4 including AKT, ERK1/2, p38, and JAK/STAT. SDF-1 treatment primarily resulted in phosphorylation of ERK1/2 and this was significantly increased by GPRASP2 knockdown. To test the role of GPRASP2 in human HSPC transplantation, CD34+ cells were edited by CRISPR/Cas9 with sgRNAs targeting GPRASP2 or the AAVS1 safe harbor site and then xenotransplanted into NBSGW mice. Initial results indicate that GPRASP2 KO results in higher human chimerism than is seen for AAVS1-targeted cells. In addition to assessing chimerism between transplants, changes in INDEL frequency within each transplant will be tracked for 16 weeks to determine the fitness of GPRASP2 knockout cells as compared to non-edited cells. This work reveals novel aspects of CXCR4 regulation by GPRASP2 and illuminates a potential point of intervention for the optimization of HSPC transplantation.

Characterization of post-edited cells modified in the TFAM gene by CRISPR/Cas9 technology in the bovine model.

Gene editing in large animal models for future applications in translational medicine and food production must be deeply investigated for an increase of knowledge. The mitochondrial transcription factor A (TFAM) is a member of the HMGB subfamily that binds to mtDNA promoters. This gene maintains mtDNA, and it is essential for the initiation of mtDNA transcription. Lately, we generated a new cell line through the disruption of the TFAM gene in bovine fibroblast cells by CRISPR/Cas 9 technology. We showed that the CRISPR/Cas9 design was efficient through the generation of heterozygous mutant clones. In this context, once this gene regulates the mtDNA replication specificity, the study aimed to determine if the post-edited cells are capable of in vitro maintenance and assess if they present changes in mtDNA copies and mitochondrial membrane potential after successive passages in culture. The post-edited cells were expanded in culture, and we performed a growth curve, doubling time, cell viability, mitochondrial DNA copy number, and mitochondrial membrane potential assays. The editing process did not make cell culture unfeasible, even though cell growth rate and viability were decreased compared to control since we observed the cells grow well when cultured in a medium supplemented with uridine and pyruvate. They also exhibited a classical fibroblastoid appearance. The RT-qPCR to determine the mtDNA copy number showed a decrease in the edited clones compared to the non-edited ones (control) in different cell passages. Cell staining with Mitotracker Green and red suggests a reduction in red fluorescence in the edited cells compared to the non-edited cells. Thus, through characterization, we demonstrated that the TFAM gene is critical to mitochondrial maintenance due to its interference in the stability of the mitochondrial DNA copy number in different cell passages and membrane potential confirming the decrease in mitochondrial activity in cells edited in heterozygosis.

Mycoplasma pneumoniae Genome Editing Based on Oligo Recombineering and Cas9-Mediated Counterselection.

Mycoplasma species share a set of features, such as lack of a cell wall, streamlined genomes, simplified metabolism, and the use of a deviant genetic code, that make them attractive approximations of what a chassis strain should ideally be. Among them, Mycoplasma pneumoniae arises as a candidate for synthetic biology projects, as it is one of the most deeply characterized bacteria. However, the historical paucity of tools for editing Mycoplasma genomes has precluded the establishment of M. pneumoniae as a suitable chassis strain. Here, we developed an oligonucleotide recombineering method for this strain based on GP35, a ssDNA recombinase originally encoded by a Bacillus subtilis-associated phage. GP35-mediated oligo recombineering is able to carry out point mutations in the M. pneumoniae genome with an efficiency as high as 2.7 × 10–2, outperforming oligo recombineering protocols developed for other bacteria. Gene deletions of different sizes showed a decreasing power trend between efficiency and the scale of the attempted edition. However, the editing rates for all modifications increased when CRISPR/Cas9 was used to counterselect nonedited cells. This allowed edited clones carrying chromosomal deletions of up to 1.8 kb to be recovered with little to no screening of survivor cells. We envision this technology as a major step toward the use of M. pneumoniae, and possibly other Mycoplasmas, as synthetic biology chassis strains.

Characterisation of endogenous Claudin-1 expression, motility and susceptibility to hepatitis C virus in CRISPR knock-in cells.

Claudin-1 (CLDN1) is a four-span transmembrane protein localised at cell-cell tight junctions (TJs), playing an important role in epithelial impermeability and tissue homoeostasis under physiological conditions. Moreover, CLDN1 expression is up-regulated in several cancers, and the level of CLDN1 expression has been proposed as a prognostic marker of patient survival. Here, we generated and characterised a novel reporter cell line expressing endogenous fluorescent levels of CLDN-1, allowing dynamic monitoring of CLDN-1 expression levels. Specifically, a hepatocellular carcinoma Huh7.5.1 monoclonal cell line was bioengineered using CRISPR/Cas9 to endogenously express a fluorescent TagRFP-T protein fused at the N-terminus of the CLDN1 protein. These cells were proved useful to measure CLDN1 expression and distribution in live cells. However, the cells were resistant to hepatitis C virus (HCV) infection, of which CLDN1 is a viral receptor, while retaining permissiveness to VSV-G-decorated pseudoparticles. Nonetheless, the TagRFP-CLDN1+/+ cell line showed expected CLDN1 protein localisation at TJs and the cell monolayer had similar impermeability and polarisation features as its wild-type counterpart. Finally, using fluorescence recovery after photobleaching (FRAP) approaches, we measured that the majority of endogenous and overexpressed TagRFP-CLDN1 diffuses rapidly within the TJ, whereas half of the overexpressed EGFP-CLDN1 proteins were stalled at TJs. The Huh7.5.1 TagRFP-CLDN1+/+ edited cell line showed physiological features comparable to that of non-edited cells, but became resistant to HCV infection. Our data also highlight the important impact of the fluorescent protein chosen for endogenous tagging. Although HCV-related studies may not be achieved with these cells, our work provides a novel tool to study the cell biology of TJ-associated proteins and a potential screening strategy measuring CLDN1 expression levels.

Double-Check Base Editing for Efficient A to G Conversions.

With the development of CRISPR/Cas9 technology, a new generation of editing methods that convert specific bases has enabled precise single-base mutations. To date, conversion of cytosine to thymidine and adenine to guanine has been achieved using the cytidine deaminase APOBEC1 and adenosine deaminase (TadA), respectively. However, the base editing efficiency can be unacceptably low in some cell types or at certain target loci. One reason might be the lack of a selective pressure against the survival of nonedited cells. Few studies on ABE in prokaryotes have been reported, probably due to the relatively low editing efficiency of TadA. Improving the editing efficiency is the key for establishing base editing techniques and especially the ABE technologies. In this work, a selective pressure against nonedited cells was implemented to increase the base editing efficiency. First, we fused nCas9 or dCas9 with TadA to compare the editing efficiency of nCas9-TadA and dCas9-TadA fusion complexes in the model prokaryote Escherichia coli. While nCas9-TadA was able to achieve A to G base editing (ABE) with a moderate efficiency, dCas9-TadA had a very low efficiency. To enrich for edited cells and increase the base-editing efficiency, we utilized the induction of double-strand breaks by active Cas9, which leads to the death of prokaryotic cells. By introducing an inducible active Cas9 with the same editing gRNA as the nCas9-TadA in the base editing process, the cells with nonedited target bases remained vulnerable to Cas9 and were eliminated. Thus, a double-check base editing (DBE) method was established, which significantly improved the editing efficiency of ABE in E.coli, reaching 99.0% for some sites. By placing a selective pressure against nonedited cells, the DBE strategy might also be applied to various scenarios to increase the efficiency of many different base editing targets or even for epigenetic DNA modification techniques.

Ex Vivo Gene-Edited Cell Therapy for Sickle Cell Disease: Disruption of the BCL11A Erythroid Enhancer with Zinc Finger Nucleases Increases Fetal Hemoglobin in Plerixafor Mobilized Human CD34+ Cells

Increases in fetal hemoglobin (HbF) are associated with improved clinical outcomes in the inherited hemoglobinopathies. We are developing a novel gene-edited cell therapy to treat patients with sickle cell disease (SCD) and beta-thalassemia (BT) using autologous hematopoietic stem and progenitor cells (HSPCs) genetically modified with zinc finger nucleases (ZFNs) to restore high levels of HbF expression. The ZFNs have been designed to specifically target the GATA motif within an intronic erythroid-specific enhancer (ESE) of BCL11A, the gene encoding a transcriptional regulator of the fetal-to-adult hemoglobin switch. Previously, we reported successful ZFN-mediated, ex vivoBCL11A gene editing in dual mobilized HSPCs, i.e. peripheral stem cells mobilized with a combination of granulocyte colony-stimulating factor (G-CSF) and plerixafor1. The editing procedure was optimized for high on-target/low off-target modification levels and increases in HbF in erythroid progeny. This drug product, ST-400, passed extensive safety testing and is currently in a phase 1/2a clinical trial for transfusion-dependent beta-thalassemia (ClinicalTrials.gov number NCT03432364).An analogous drug product, BIVV003, is being developed as a therapeutic for SCD. In patients with SCD, the use of G-CSF for stem cell mobilization is not recommended due to the risk of clinical complications. Therefore, peripheral HSPCs are obtained from SCD patients via single agent, plerixafor mobilization and apheresis2-5. Understanding the effect of this change in mobilization strategy on ZFN editing efficiency and specificity is a key element in preparing for SCD gene-edited cell therapy. In the present studies, we demonstrate comparability of ZFN editing outcomes in single and dual mobilized HSPCs obtained from healthy donors. In plerixafor mobilized HSPCs from five healthy donors at research scale, ZFN-mediated gene editing induced an efficient modification at the BCL11A ESE target site (>75% of alleles modified, as measured by MiSeq deep sequencing) with high post-editing viability (77%). Similar gene editing efficiencies (>70%) were obtained in HSPCs at clinical manufacturing scale (n=2). Further, in vitro HbF protein levels and HbF+ cell frequencies within erythroid progeny of edited cells were increased by >4 and 3-fold respectively - compared to non-edited cells in the same culture conditions, using reverse phase high-performance liquid chromatography and flow cytometry (n=4 healthy donors at research scale). Single cell clone analysis revealed that ZFN-mediated gene editing targeted both alleles of BCL11A at high frequency (91-94% of edited cells within erythroid progeny) with high levels of replicable GATA-disrupting indel patterns. On average, each edited allele contributed an additional 17.6% increase in HbF production in vitro, with a statistically-significant increase in HbF level for biallelic edited vs. unedited controls (3.4 fold).Critical to BIVV003 use in clinical trials, ZFN-mediated gene editing did not impair single agent mobilized HSPC function in vitro based on measurements of colony forming unit (CFU) production and frequencies of long-term HSC (LT-HSC) and common myeloid progenitor (CMP) cells by flow cytometry. In agreement with this data, injection of BIVV003 into immune-deficient NBSGW mice resulted in robust long-term engraftment (21 weeks) without any impact on the number of HSPCs and their differentiated progeny. Overall, these data demonstrate potential efficacy of ZFN-edited HSPCs (BIVV003) as a novel cell therapy for SCD patients.Holmes et al., 2017 (ASH abstract)Fitzhugh et al., 2009Lagresle-Peyrou et al., 2018Hsieh and Tisdale, 2018Yannaki et al., 2012 DisclosuresMoran:Bioverativ, a Sanofi Company: Employment. Ling:Bioverativ, a Sanofi Company: Employment. Lessard:Bioverativ, a Sanofi Company: Employment. Vieira:Bioverativ a Sanofi Company: Employment. Hong:Bioverativ, a Sanofi Company: Employment. Holmes:Sangamo Therapeutics: Employment. Reik:Sangamo Therapeutics: Employment. Dang:Sangamo Therapeutics: Employment. Gray:Sangamo Therapeutics: Employment. Levasseur:Bioverativ, a Sanofi Company: Employment. Rimmele:Bioverativ, a Sanofi Company: Employment.

In This Issue

HomeCirculation ResearchVol. 121, No. 10In This Issue Free AccessIn BriefPDF/EPUBAboutView PDFView EPUBSections ToolsAdd to favoritesDownload citationsTrack citationsPermissions ShareShare onFacebookTwitterLinked InMendeleyReddit Jump toFree AccessIn BriefPDF/EPUBIn This Issue Ruth Williams Ruth WilliamsRuth Williams Search for more papers by this author Originally published27 Oct 2017https://doi.org/10.1161/RES.0000000000000183Circulation Research. 2017;121:1103Monocyte PKCδ Deletion Accelerated Atherosclerosis (p 1153)Li et al uncover an unexpected effect of protein kinase C deletion on atherosclerosis.Download figureDownload PowerPointExcess lipids in the blood can activate monocytes and enhance their uptake into artery walls, thus promoting the development of atherosclerotic plaques. This lipid-induced activation of monocytes involves the signal transduction factor protein kinase C (PKC); but PKC has a number of isoforms, so exactly which enzyme is involved in the process, and how, remains unclear. To find out, Li and colleagues examined rats with hyperlipidemia. They showed that PKC levels were indeed increased in the animals’ monocytes, and that most of this was accounted for by an increase in PKCδ. To determine the physiological significance of this increase, the team generated transgenic atherosclerosis-prone mice with macrophages lacking PKCδ. To their surprise, the researchers found that mice lacking PKCδ in macrophages exhibited accelerated atherosclerosis compared with controls, their plaques were larger, and contained more macrophages. Further investigations revealed that the numbers of plaque macrophages increased due to reduced apoptosis and increased proliferation of these cells, as well as to increased phosphorylation of pro-survival factors. The authors conclude that specific activation of the δ isoform of PKC in monocytes may be a way to limit atherosclerosis progression.CRISPR/Cas9 Postnatal Cardiac Gene Editing (p 1168)Gene editing of mouse hearts is not yet a reliable approach for functional genetic analyses, say Johansen et al.Download figureDownload PowerPointEngineering mice to possess specific genetic deletions is a powerful tool for studying gene function. But generating transgenic mice is time-consuming, expensive, and requires large numbers of animals. Gene editing is a successful approach for studying gene functions in cultured cells and has been used in live animals to correct genetic defects. So Johansen and colleagues considered that it might be a cheaper, quicker, and less animal-intensive approach for studying gene functions in vivo. The team introduced gene-editing vectors targeting 3 genes of known function (Myh6, Tbx20, and Sav1) into the hearts of newborn mice. However, only 1 of the genes (Myh6) resulted in the expected phenotype. Further examinations revealed that only 5% to 15% of gene copies had been disrupted by the editing, resulting in a mosaic of edited and nonedited cells, which, in the case of Sav1 and Tbx20, was clearly insufficient to produce a phenotype. While the team could boost the editing efficiency of Sav1 by targeting 2 sites in the gene simultaneously, the authors conclude that considerable technical improvements would be necessary for in vivo gene editing to be a reliable tool for gene function studies.Umbilical Cord MSCs for Heart Failure (p 1192)Umbilical cord stem cells could be a promising therapy for heart failure, say Bartolucci et al.Download figureDownload PowerPointStem cell treatment for heart failure has shown promise in animal models of the disease, but in clinical trials, the benefits have been moderate, at best. Most clinical trials have used bone marrow–derived mononuclear cells or mesenchymal stem cells (BM-MSCs), but MSCs derived from human umbilical cords offer several advantages over their bone marrow counterparts, say Bartolucci and colleagues. For one thing, these cells have more proliferative and differentiation potential than adult cells, and they are very easy to obtain. The team now reports on a first-of-its-kind randomized placebo-controlled trial of umbilical cord MSCs (UM-MSCs) for heart failure. Fifteen patients aged between 18 and 75 received intravenous injections of UC-MSCs, while another 15 received a placebo. None of the UC-MSC recipients suffered adverse effects as a result of the treatment. In comparison with control patients, the treatment recipients exhibited a significant and sustained improvement in heart function—including increased left ventricle ejection fraction—and an improved quality of life. These encouraging results now pave the way for further larger-scale trials of UC-MSCs to examine how dosage, administration methods, and other factors might further improve outcomes. Previous Back to top Next FiguresReferencesRelatedDetails October 27, 2017Vol 121, Issue 10 Advertisement Article InformationMetrics © 2017 American Heart Association, Inc.https://doi.org/10.1161/RES.0000000000000183 Originally publishedOctober 27, 2017 PDF download Advertisement

Efficient Genome Editing of a Facultative Thermophile Using Mesophilic spCas9.

Well-developed genetic tools for thermophilic microorganisms are scarce, despite their industrial and scientific relevance. Whereas highly efficient CRISPR/Cas9-based genome editing is on the rise in prokaryotes, it has never been employed in a thermophile. Here, we apply Streptococcus pyogenes Cas9 (spCas9)-based genome editing to a moderate thermophile, i.e., Bacillus smithii, including a gene deletion, gene knockout via insertion of premature stop codons, and gene insertion. We show that spCas9 is inactive in vivo above 42 °C, and we employ the wide temperature growth range of B. smithii as an induction system for spCas9 expression. Homologous recombination with plasmid-borne editing templates is performed at 45–55 °C, when spCas9 is inactive. Subsequent transfer to 37 °C allows for counterselection through production of active spCas9, which introduces lethal double-stranded DNA breaks to the nonedited cells. The developed method takes 4 days with 90, 100, and 20% efficiencies for gene deletion, knockout, and insertion, respectively. The major advantage of our system is the limited requirement for genetic parts: only one plasmid, one selectable marker, and a promoter are needed, and the promoter does not need to be inducible or well-characterized. Hence, it can be easily applied for genome editing purposes in both mesophilic and thermophilic nonmodel organisms with a limited genetic toolbox and ability to grow at, or tolerate, temperatures of 37 and at or above 42 °C.

RNA-guided editing of bacterial genomes using CRISPR-Cas systems.

The targeting of nucleases to specific DNA sequences facilitates genome editing. Recent work demonstrated that the CRISPR-associated (Cas) nuclease Cas9 can be targeted to sequences in vitro simply by modifying a short7 CRISPR RNA (crRNA) guide. Here we use this CRISPR-Cas system to introduce marker-free mutations in Streptococcus pneumoniae and Escherichia coli. The approach involves re-programming Cas9 by using a crRNA complementary to a target chromosomal locus and introducing a template DNA harboring a desired mutation and an altered crRNA recognition site for recombination with the target locus. We exhaustively analyze Cas9 target requirements to define the range of targetable sequences and show strategies for editing sites that do not meet these requirements. Alone or together with recombineering, CRISPR assisted editing induces recombination at the targeted locus and kills non-edited cells leading to a recovery of close to a 100% of edited cells. Multiple crRNA can be used to modify several loci simultaneously. Our results show that CRISPR-mediated genome editing only requires programming of the crRNA and template sequences and thus constitutes a useful tool for genetic engineering.