Cleavage-free human genome editing
Authors: Chenzhong Kuang, Yan Xiao, Dirk Hondmann
Journal: Molecular Therapy (2021)

Most gene editing technologies introduce breaks or nicks into DNA, leading to the generation of mutagenic insertions and deletions by non-homologous end-joining repair. Here, we report a new, cleavage-free gene editing approach based on replication interrupted template-driven DNA modification (RITDM). The RITDM system makes use of sequence-specific DLR fusion molecules that are specifically designed to enable localized, temporary blockage of DNA replication fork progression, thereby exposing single-stranded DNA that can be bound by DNA sequence modification templates for precise editing. We evaluate the use of zinc-finger arrays for sequence recognition. We demonstrate that RITDM can be used for gene editing at endogenous genomic loci in human cells and highlight its safety profile of low indel frequencies and undetectable off-target side effects in RITDM-edited clones and pools of cells. Most gene editing technologies introduce breaks or nicks into DNA, leading to the generation of mutagenic insertions and deletions by non-homologous end-joining repair. Here, we report a new, cleavage-free gene editing approach based on replication interrupted template-driven DNA modification (RITDM). The RITDM system makes use of sequence-specific DLR fusion molecules that are specifically designed to enable localized, temporary blockage of DNA replication fork progression, thereby exposing single-stranded DNA that can be bound by DNA sequence modification templates for precise editing. We evaluate the use of zinc-finger arrays for sequence recognition. We demonstrate that RITDM can be used for gene editing at endogenous genomic loci in human cells and highlight its safety profile of low indel frequencies and undetectable off-target side effects in RITDM-edited clones and pools of cells. IntroductionMany current gene correction systems make use of nuclease or nickase activity to introduce DNA breaks. The generation of DNA breaks has the implicit disadvantage that DNA repair processes, such as non-homologous end-joining (NHEJ), are being triggered, which often results in the generation of mutagenic indels.1Kosicki M. Tomberg K. Bradley A. Repair of double-strand breaks induced by CRISPR-Cas9 leads to large deletions and complex rearrangements.Nat. Biotechnol. 2018; 36: 765-771https://doi.org/10.1038/nbt.4192nbt.4192Crossref PubMed Google Scholar,2Pannunzio N.R. Watanabe G. Lieber M.R. Nonhomologous DNA end-joining for repair of DNA double-strand breaks.J. Biol. Chem. 2018; 293: 10512-10523https://doi.org/10.1074/jbc.TM117.000374TM117.000374Abstract Full Text Full Text PDF PubMed Scopus (258) Google Scholar Minimizing mutational damage is an essential feature that will enable a wider range of genome editing-based therapies to progress to the clinic. In addition to the current gene editing technologies, it would be pragmatically useful to have an alternative gene editing system that does not depend on double-stranded DNA (dsDNA) or single-stranded DNA (ssDNA) breakage. We report the development of a new nuclease-free gene editing system—RITDM (replication interrupted template-driven DNA modification)—based on sequence-specific chimeric molecules that interact with replication forks, which, in combination with DNA modification polynucleotides, enables editing without DNA cleavage.During cell division, the replication fork complex transiently exposes short stretches of ssDNA.3Rickman K. Smogorzewska A. Advances in understanding DNA processing and protection at stalled replication forks.J. Cell Biol. 2019; 218: 1096-1107https://doi.org/10.1083/jcb.201809012jcb.201809012Crossref PubMed Scopus (0) Google Scholar Genome-wide slowing and stalling of replication forks leads to a lingering ssDNA presence within cells and can generate signals equivalent to those of damaged DNA to evoke DNA repair.4Cimprich K.A. Cortez D. ATR: an essential regulator of genome integrity.Nat. Rev. Mol. Cell Biol. 2008; 9: 616-627https://doi.org/10.1038/nrm2450nrm2450Crossref PubMed Scopus (0) Google Scholar Introduction of ssODNs can result in their annealing at DNA replication forks, which can be used for genome editing in lower eukaryotes.5Barbieri E.M. Muir P. Akhuetie-Oni B.O. Yellman C.M. Isaacs F.J. Precise editing at DNA replication forks enables multiplex genome engineering in eukaryotes.Cell. 2017; 171: 1453-1467.e13https://doi.org/10.1016/j.cell.2017.10.034Abstract Full Text Full Text PDF PubMed Scopus (74) Google Scholar We hypothesized that we could convert these genome-wide effects to a locus-specific approach by blocking replication forks specifically in the vicinity of a target locus, and that this approach could be developed for usage in the larger genomes of mammalian cells. Such a system could enable locus-specific gene editing in human cells.We first developed sequence-specific blocking agents (named DLRs) by designing fusion proteins in which one module (D: DNA binding) binds sequence specifically to one strand of DNA, and, through a linker (L: linker), connects with a second DNA binding module (R: replication fork block) that binds to the opposite strand of DNA. The hypothesis underpinning this design was that such a molecule could (reversibly) bind to both DNA strands and thereby form a temporary “clamp” at the target site. For the sequence-specific D-module, we used zinc-finger arrays, which have been used extensively in the creation of various types of fusion proteins (Figure 1A ).6Klug A. The discovery of zinc fingers and their applications in gene regulation and genome manipulation.Annu. Rev. Biochem. 2010; 79: 213-231https://doi.org/10.1146/annurev-biochem-010909-095056Crossref PubMed Scopus (491) Google Scholar In the R-module, we used a domain that can bind to the phosphate backbone of the opposing DNA strand. When introduced into a cell, DLR fusion proteins can bind to a specific genomic locus and obstruct the progression of a replication fork, thereby exposing single-stranded sequences to which complementary oligonucleotides with (a) desired sequence modification(s) can bind (Figure 1B). After the binding of a DNA modification template, the DNA contains (a) sequence mismatch(es). The cellular DNA repair mechanisms can resolve these mismatches. Depending on which strand is used for correction, part of the cells will be converted to have the desired sequence modification.We demonstrated the direct interaction between DLR fusion molecules and the replication fork, enabling a novel mechanism for gene editing in mammalian cells. Next, we applied RITDM at various disease-relevant loci in the human genome and showed that RITDM can be used to introduce heritable and precise genetic modifications. We achieved precise editing involving (1) single base pair changes, (2) restoration of reading frames by inserting nucleotides, and (3) gene knockouts by the introduction of stop codons and/or restriction enzyme sites. Since RITDM does not require DNA cleavage at the target site, the mutational indel incidence remains at a low level on a par with that of the background. Moreover, both DLR and the donor template must work in concert, implying that off-target effects are of second-order kinetics. This results in a favorable safety profile of RITDM. Taken together, RITDM is a novel DNA cleavage-free editing system that can be used to achieve gene editing while avoiding the generation of indels and/or other off-target effects. This approach thus provides an additional option in the development of gene correction therapies.ResultsDevelopment of sequence-specific blocking DLR fusion moleculesWhen designing a DLR molecule for a specific locus, the D domain must be programmable to recognize the target sequence on one strand of DNA. In principle, any DNA sequence recognition platform can be used, including zinc-finger arrays, TALE effectors, and dCAS9 along with a gRNA. In consideration of keeping the size of DLR constructs as small as possible to also allow the use of in vivo delivery vectors, we chose to use zinc-finger arrays.7Sera T. Uranga C. Rational design of artificial zinc-finger proteins using a nondegenerate recognition code table.Biochemistry. 2002; 41: 7074-7081https://doi.org/10.1021/bi020095cCrossref PubMed Scopus (91) Google Scholar, 8Choo Y. Klug A. Selection of DNA binding sites for zinc fingers using rationally randomized DNA reveals coded interactions.Proc. Natl. Acad. Sci. U S A. 1994; 91: 11168-11172https://doi.org/10.1073/pnas.91.23.11168Crossref PubMed Scopus (261) Google Scholar, 9Zhu C. Gupta A. Hall V.L. Rayla A.L. Christensen R.G. Dake B. Lakshmanan A. Kuperwasser C. Stormo G.D. Wolfe S.A. Using defined finger-finger interfaces as units of assembly for constructing zinc-finger nucleases.Nucleic Acids Res. 2013; 41: 2455-2465https://doi.org/10.1093/nar/gks1357 gks1357Crossref PubMed Scopus (0) Google Scholar, 10Choo Y. Klug A. Toward a code for the interactions of zinc fingers with DNA: selection of randomized fingers displayed on phage.Proc. Natl. Acad. Sci. U S A. 1994; 91: 11163-11167https://doi.org/10.1073/pnas.91.23.11163Crossref PubMed Scopus (315) Google Scholar Zinc fingers measure only about 30 amino acids long and can target 3 nucleotides, making it the most compact DNA binding element. The L domain is an orientation enabler for the R domain reaching the opposite DNA strand. For the R unit, we ideally wanted to design a structure that is (1) as compact as possible, (2) able to bind to DNA in a non-sequence-specific manner (e.g., via phosphate backbone and/or major or minor groove interaction), (3) have (a) crystal structure(s) available for modeling, and that is (4) present in a broad range of proteins for non-specific DNA binding, assuming such structures could also serve a similar function in our DLR fusion protein. A structure fulfilling these requirements is the PD-(D/E)xK fold that is present in a superfamily of proteins.11Steczkiewicz K. Muszewska A. Knizewski L. Rychlewski L. Ginalski K. Sequence, structure and functional diversity of PD-(D/E)XK phosphodiesterase superfamily.Nucleic Acids Res. 2012; 40: 7016-7045https://doi.org/10.1093/nar/gks382gks382Crossref PubMed Scopus (0) Google Scholar Members of this protein family are involved in numerous activities that involve DNA interaction.12Wah D.A. Bitinaite J. Schildkraut I. Aggarwal A.K. Structure of FokI has implications for DNA cleavage.Proc. Natl. Acad. Sci. U S A. 1998; 95: 10564-10569https://doi.org/10.1073/pnas.95.18.10564Crossref PubMed Scopus (174) Google Scholar A feature of this fold is that it can bind to the phosphate backbone, thereby providing non-sequence-specific binding. Thus, we set out to design the first DLR fusion protein using a zinc-finger array linked to a PD-(D/E)xK domain.Of the number of zinc-finger crystal structures that have been elucidated, we used the 1MEY structure as a reference.13Kim C.A. Berg J.M. A 2.2 A resolution crystal structure of a designed zinc finger protein bound to DNA.Nat. Struct. Biol. 1996; 3: 940-945https://doi.org/10.1038/nsb1196-940Crossref PubMed Scopus (149) Google Scholar This crystal structure of a designed zinc-finger protein provided useful information on amino acid side-chain interactions with nucleotides as well as details on how its C terminus was positioned in the major DNA groove. For the design of the R unit, we made use of the crystal structure of FokI, as it contains a PD-(D/E)xK fold in its catalytic domain.12Wah D.A. Bitinaite J. Schildkraut I. Aggarwal A.K. Structure of FokI has implications for DNA cleavage.Proc. Natl. Acad. Sci. U S A. 1998; 95: 10564-10569https://doi.org/10.1073/pnas.95.18.10564Crossref PubMed Scopus (174) Google Scholar In addition to looking at the potential suitability of FokI-derived R unit structures, we also included another naturally existing PD-(D/E)xK fold, appearing in the small unit of BtsI, in our analysis.14Xu S.Y. Zhu Z. Zhang P. Chan S.H. Samuelson J.C. Xiao J. Ingalls D. Wilson G.G. Discovery of natural nicking endonucleases Nb.BsrDI and Nb.BtsI and engineering of top-strand nicking variants from BsrDI and BtsI.Nucleic Acids Res. 2007; 35: 4608-4618https://doi.org/10.1093/nar/gkm481Crossref PubMed Scopus (42) Google Scholar Not only does its size and stability make it a good candidate for use in protein engineering, but amino acid sequence alignments also showed similarities between BtsI and FokI (Figure S1A). From BtsI, we identified the PD-(D/E)xK fold located within the β2 and β3 sheets of a three-antiparallel β sheet structure and selected this three-antiparallel β sheet structure for further development of the R unit. Moreover, the crystal structure of FokI indicated that this structure can orient perpendicularly on the DNA phosphate backbone.12Wah D.A. Bitinaite J. Schildkraut I. Aggarwal A.K. Structure of FokI has implications for DNA cleavage.Proc. Natl. Acad. Sci. U S A. 1998; 95: 10564-10569https://doi.org/10.1073/pnas.95.18.10564Crossref PubMed Scopus (174) Google Scholar In FokI, the loop between β1 and β2 in the PD-(D/E)xK fold participates in DNA major groove binding,12Wah D.A. Bitinaite J. Schildkraut I. Aggarwal A.K. Structure of FokI has implications for DNA cleavage.Proc. Natl. Acad. Sci. U S A. 1998; 95: 10564-10569https://doi.org/10.1073/pnas.95.18.10564Crossref PubMed Scopus (174) Google Scholar which would support the desired orientation of the R unit versus that of the zinc-finger DNA binding D domain. To enable this desired orientation, we replaced the loop of BtsI with that of FokI to create an R unit that combines the major groove-binding loop as found in FokI with the compact β sheet structure (β1, β2, and β3) of BtsI (Figures 1A and S1A).When considering how we could link the N-terminal zinc finger to the C-terminal R domain, we again looked at the crystal structures. The C terminus (in MEY1) of the zinc finger bound to DNA is positioned in the major groove of the DNA it is bound to. The crystal structure of the antiparallel β sheet structure (as in Fok1), when it is oriented on top of the phosphate backbone of the opposing strand, has its N terminus in close proximity to the major groove. The N terminus of the R unit could thus be fused to the C terminus of the zinc-finger array using a relatively short linker. We then used a four-amino-acid LRGS linker, which is a prototypical linker used in synthetic zinc-finger biology.15Wright D.A. Thibodeau-Beganny S. Sander J.D. Winfrey R.J. Hirsh A.S. Eichtinger M. Fu F. Porteus M.H. Dobbs D. Voytas D.F. Joung J.K. Standardized reagents and protocols for engineering zinc finger nucleases by modular assembly.Nat. Protoc. 2006; 1: 1637-1652Crossref PubMed Scopus (162) Google ScholarTo avoid the risk of residual nuclease activity, amino acid residues with a negatively charged side chain, an aspartic acid (D) residue from the β2 sheet, or glutamic acid (E) from the β3 at the catalytically active sites in the PD-(D/E)xK fold were replaced by amino acids either with an uncharged side chain, such as serine (S), glutamine (Q), asparagine (N), or threonine (T), a hydrophobic chain, such as alanine (A), valine (V), leucine (L), isoleucine (I), or methionine (M), or with a positively charged chain, such as histamine (H), arginine (R), or lysine (K), which, in native FokI and in BtsI, eliminates nuclease activity (Figure S1B).11Steczkiewicz K. Muszewska A. Knizewski L. Rychlewski L. Ginalski K. Sequence, structure and functional diversity of PD-(D/E)XK phosphodiesterase superfamily.Nucleic Acids Res. 2012; 40: 7016-7045https://doi.org/10.1093/nar/gks382gks382Crossref PubMed Scopus (0) Google ScholarMutant EGFP reporter gene editing by RITDMWe developed a gene correction reporter system to test if our approach using DLR fusion proteins in combination with a modification template could be used to perform gene editing. For this proof-of-principle validation we developed a cell line with a GFP reporter (Figure S2A). The integrated mutant EGFP (EGFPDP2) gene bears a frameshift causing a deletion (G) in combination with a G-to-C point mutation, rendering the expressed protein nonfunctional (Figures 2A and S2B). We obtained a clone with two inserted copies of the EGFPDP2 construct with the idea that, if one copy was corrected, the other copy could serve as an indicator for potential off-target effects. As a first attempt, a DLR fusion protein with a 5-zinc-finger array, recognizing a unique 15-nucleotide sequence in EGFPDP2, was developed (Figure 2B). A 142-nucleotide ssDNA correction template was designed that incorporated the corrections for a “G insertion” and a “C-to-G conversion” (Figure S2B).Figure 2Genome editing by RITDMShow full caption(A) Diagram of a mutated EGFP (EGFPDP2) reporter gene in the HEK293 reporter cell line. The EGFDP2 reporter has a deletion of single nucleotide (G) and a G-to-C point mutation. Restoration of EGFP reading frame after RITDM editing. (B) Schematic depicting that a mutated EGFP (EGFPDP2) reporter gene was targeted and repaired using a specifically designed RITDM system. The DLR molecule recognizes 15-nucleotide sequences, specifically. (C) EGFP was restored to express functionally in the reporter cells, as shown in forms of positive cells (left), cellular cluster (middle), and enrichment of positive cells (right). (D) Representative flow plots from flow cytometric analysis of alive EGFP cells 7 days post-genomic editing with the indicated conditions of control, donor alone, RITDM genome editing. Quantification of GFP (+) cells (per million) from the indicated conditions. Bar graphs represent two independent experiments with standard error shown. (E) The endogenous ApoE gene was genetically modified at codon 112 by RITDM in HEK293 cells. (F) The DLR molecule 27 nucleotide sequences at the target. (G) Single-nucleotide T-to-C conversion was detected by ddPCR. The bar graph represents the editing frequencies at codon 112 of the ApoE gene achieved by RITDM in HEK293 cells measured and calculated by ddPCR. (H) Enhanced genome editing frequencies were detected by ddPCR, while ssODN alone did not induce T-to-C conversion. The bar graph represented the editing frequencies at codon 112 of the ApoE gene achieved by RITDM in HEK293 cells measured and calculated by ddPCR.View Large Image Figure ViewerDownload Hi-res image Download (PPT)Five days after the transfection of thymidine synchronized cells, fluorescent green cells and cellular clusters could be observed using fluorescent microscopy (Figure 2C, left and middle). Following reseedings, we could obtain pure green cell populations (Figure 2C, right). Cells were also analyzed by flow cytometry 7 days after transfection. This showed that, under these initial conditions, RITDM editing resulted in 300 ± 60 cells per million with bright green fluorescence in a normal cellular morphology, whereas the “ssODN alone” control showed a frequency as low as 19 ± 7 per million cells (Figure 2D), which is in line with published observations.16Igoucheva O. Alexeev V. Yoon K. Targeted gene correction by small single-stranded oligonucleotides in mammalian cells.Gene Ther. 2001; 8: 391-399https://doi.org/10.1038/sj.gt.3301414Crossref PubMed Scopus (145) Google Scholar,17Urnov F.D. Miller J.C. Lee Y.L. Beausejour C.M. Rock J.M. Augustus S. Jamieson A.C. Porteus M.H. Gregory P.D. Holmes M.C. Highly efficient endogenous human gene correction using designed zinc-finger nucleases.Nature. 2005; 435: 646-651https://doi.org/10.1038/nature03556Crossref PubMed Scopus (1316) Google Scholar These observations confirmed that RITDM could be used for gene editing.To further evaluate the nature of the genetic conversion at the GFP reporter locus, we extracted genomic DNA from a pure green cellular population that was obtained after repeated reseeding. We used Sanger sequencing to confirm the genetic modifications (Figure S2C) followed by next-generation sequencing. Approximately 59.4% of the reads had the anticipated G insertion and C-to-G conversion (Figures S2D–S2F), indicating that one allele of EGFPDP2 was corrected into EGFP by RITDM gene editing.We tested a series of donors with lengths between 60 and 200 nucleotides that also varied the position of the editing sequences. All donors with a length between 90 and 200 nucleotides yielded green cells or green clusters. We did not observe a meaningful pattern relevant to the positions of genetic modification within the donor either symmetrically or asymmetrically. These observations were in line with results obtained when we started using RITDM to edit codon 112 of the ApoE gene. When using a correction oligonucleotide of 60 nucleotides, we only observed a weak ddPCR signal, while a 71-nucleotide oligonucleotide allowed an increase in signal (Figure S15A). Therefore, we subsequently designed donor templates with an approximate length of 150 nucleotides with the modification sequence in its middle.Conversion of codon 112 of human ApoE by RITDM gene editingTo confirm that RITDM could also modify an endogenous human gene, we selected the ApoE gene and aimed to convert a “T” to “C” at codon 112 in ApoE in HEK293 cells. ApoE has three major alleles (E2, E3, and E4) that differ at codons 112 and 158. E4 is a risk allele for late-onset Alzheimer disease (LOAD), E3 is the dominant allele in the human population, and E2 reduces the likelihood and age of onset of LOAD.18Corder E.H. Saunders A.M. Strittmatter W.J. Schmechel D.E. Gaskell P.C. Small G.W. Roses A.D. Haines J.L. Pericak-Vance M.A. Gene dose of apolipoprotein E type 4 allele and the risk of Alzheimer’s disease in late onset families.Science. 1993; 261: 921-923https://doi.org/10.1126/science.8346443Crossref PubMed Scopus (7163) Google Scholar,19Chartier-Harlin M.C. Parfitt M. Legrain S. Pérez-Tur J. Brousseau T. Evans A. Berr C. Vidal O. Roques P. Gourlet V. et al.Apolipoprotein E, epsilon 4 allele as a major risk factor for sporadic early and late-onset forms of Alzheimer’s disease: analysis of the 19q13.2 chromosomal region.Hum. Mol. Genet. 1994; 3: 569-574https://doi.org/10.1093/hmg/3.4.569Crossref PubMed Scopus (366) Google Scholar A DLR fusion protein was developed with an array of 9 zinc fingers recognizing a 27-nucleotide sequence on the leading strand of human ApoE gene (Figures 2E, 2F, and S3A). We evaluated a number of potential binding sites close to the target site and found this 27-nucleotide sequence to be unique in the human genome, even when we allowed for multiple mismatches in BLAST searches. For the donor template, a 129-nucleotide ssDNA with a T-to-C substitution located in its middle was used.Detection of T-to-C conversion was performed by droplet digital PCR (ddPCR). One common primer was located inside this ssDNA template sequence while the other was located outside. Allele-specific probes conjugated with fluorophores FAM and HEX were designed to distinguish between C and T, respectively (Figure S3A). Following transfection of HEK293 cells with the DLR and correction template, cells were allowed to recover and grow on complete culture medium for 7 days, after which the genomic DNA was isolated and used in ddPCR analysis. Raw droplet data are shown in Figure 2G, where C droplets are displayed in the top panel and T droplets are in the lower panel. “No DNA” was used as negative control, showing neither C nor T droplets. Wild-type fibroblast was used as a positive control because of its heterozygous T/C genotype for codon 112 of human ApoE, showing both C and T droplets. The untreated HEK293 only had T droplets, demonstrating a homozygous T/T genotype. After RITDM editing, C droplets appeared, demonstrating successful T-to-C genetic conversion at codon 112 of human ApoE with an average DNA editing frequency of 1.37% ± 0.09% (Figure 2G; Table S3), obtained from three independent experiments. We also confirmed the successful T-to-C conversion by next-generation sequencing. Cells edited by RITDM showed a T-to-C conversion at the expected nucleotide position with a frequency of 1.6%. Since HEK293 has five to six copies of chromosome 19, a DNA conversion rate of 1.5%–1.6% implies that the frequency of cells containing at least one corrected copy will be higher and may be in the 7.5%–9.6% range. Compared with untransfected cells, no obvious unwanted single-nucleotide polymorphisms (SNPs) were detected (Figure S3B). We further optimized the transfection protocol by increasing the amount of DNA plasmid and ssODN, which resulted in an increase of the correction frequency to 9.88% ± 4.19%, as measured by ddPCR (Figure 2H; Table S3).Following these experiments, we used the human U937 cell line, which is homozygous ApoE4/E4.20Schaffer S. Lam V.Y. Ernst I.M. Huebbe P. Rimbach G. Halliwell B. Variability in APOE genotype status in human-derived cell lines: a cause for concern in cell culture studies?.Genes Nutr. 2014; 9: 364https://doi.org/10.1007/s12263-013-0364-4Crossref PubMed Scopus (10) Google Scholar We aimed to convert ApoE4 to E3 by a C-to-T conversion at codon 112 by co-transfection of the same DLR used in the experiments above. For a DNA modification template we used a 150-nucleotide ssODN with a desired C-to-T substitution (Figures 3A and S5A ). As detected by ddPCR, RITDM targeting resulted in C-to-T gene conversion, which was verified by next-generation sequencing (Figures 3A, S5B, and S5C). We repeated these experiments multiple times and obtained an average 16.67% ± 7.92% of DNA correction frequencies (Figure S5B).Figure 3Illustrations of the scope of genome editing by RITDMShow full captionA number of disease-related genomic loci were modified by RITDM. For each case, the genomic target sequence is shown in blue and the sequence to be changed in red. The donor template with the desired sequence modification (in green) is shown. For each case the type of editing and the intended consequences are indicated. A representative ddPCR plot illustrates typical experimental data obtained, while the bar charts show the percentage of cells without indels, the percentage of cells edited as intended, respectively, the percentage of cells with indels, as determined by deep sequencing. Experimental results are shown for ApoE (A), Bcl11a (B), and PDCD1 (C). Bar graphs represent at least two independent experiments with standard errors shown.View Large Image Figure ViewerDownload Hi-res image Download (PPT)We performed a number of controls, including one that contained the zinc-finger array without the R unit (Figure S6C). This was done to evaluate if the zinc-finger array by itself could block replication fork progression and elicit gene editing. Blocking of replication fork progression by deadCAS9 (dCAS9) has been described.21Whinn K.S. Kaur G. Lewis J.S. Schauer G.D. Mueller S.H. Jergic S. Maynard H. Gan Z.Y. Naganbabu M. Bruchez M.P. et al.Nuclease dead Cas9 is a programmable roadblock for DNA replication.Sci. Rep. 2019; 9: 13292https://doi.org/10.1038/s41598-019-49837-zCrossref PubMed Scopus (18) Google Scholar,22Doi G. Okada S. Yasukawa T. Sugiyama Y. Bala S. Miyazaki S. Kang D. Ito T. Catalytically inactive Cas9 impairs DNA replication fork progression to induce focal genomic instability.Nucleic Acids Res. 2021; 49: 954-968https://doi.org/10.1093/nar/gkaa12416062770Crossref PubMed Google Scholar Performing this control, we did not observe gene conversion, indicating that the R unit is needed to achieve the intended effect. In addition, the donor alone did not induce genome conversion at significant levels. Also, in subsequent experiments, we never observed gene conversion using zinc-finger array controls (plus modification oligonucleotide) without an R unit (Figures S7C, S8C, and S9C). This is in line with gene editing experiments in which dCAS9 is used as a control where, typically, very low or no gene editing frequencies are observed.23Wu X. Scott D.A. Kriz A.J. Chiu A.C. Hsu P.D. Dadon D.B. Cheng A.W. Trevino A.E. Konermann S. Chen S. et al.Genome-wide binding of the CRISPR endonuclease Cas9 in mammalian cells.Nat. Biotechnol. 2014; 32: 670-676https://doi.org/10.1038/nbt.2889nbt.2889Crossref PubMed Scopus (0) Google Scholar,24Guilinger J.P. Thompson D.B. Liu D.R. Fusion of catalytically inactive Cas9 to FokI nuclease improves the specificity of genome modification.Nat. Biotechnol. 2014; 32: 577-582https://doi.org/10.1038/nbt.2909Crossref PubMed Scopus (617) Google ScholarIn addition, we further ruled out the possibility of false positivity stemming from residual donor template surviving in cells and being co-isolated with chromosomal DNA. Shown in Figure S6A, we performed ddPCR assay using only one end primer. If there was donor template left in the genomic DNA extraction, the donor template then could pair with one end primer to amplify the target region, leading to false-positive droplets that could be detected by the highly sensitive ddPCR assay. Clearly, we did not find any positive droplets in any condition tested that involved donor template alone, chromosomal DNA from mixed positive pooled cells, and positive single cellular clones (Figure S6B).Generation and analysis of ApoE-converted clonesHaving established that RITDM can be used to convert the ApoE gene at codon 112, we wanted to generate clones to enable more detailed genetic and functional characterization of the gene editing effects. In subsequent transfections, single clones were obtained by seeding 0.5 transfected cell/well and continued culturing for 2–3 weeks. Positive clones were identified by ddPCR (Figures S4A and S4B). Positive clone frequencies obtained averaged 11.6% ± 5.3% in three independent experiments (Figure S4C). Sanger sequencing re