Efficient genome editing with chimeric oligonucleotide-directed editing

Domain: neuroscience
Published: 2026-04-13
Editorial outcome: major_revision

Abstract
Prime editing has emerged as a precise and powerful genome editing tool, offering a favorable gene editing profile compared to other Cas9-based approaches. Here we report several nCas9-DNA polymerase fusion proteins and their engineered versions to create a simple and efficient two-component chimeric oligonucleotide-directed editing (CODE) system. CODE contains a derivative of Bst DNA polymerase engineered for increased thermostability and processivity as well as a chimeric pegRNA (cpegRNA) for programmable search and replace genome editing. Additionally, CODEMax(exo+) features a 5’ to 3’ exonuclease activity that promotes effective strand invasion and repair outcomes favoring the incorporation of the desired edit. We demonstrate that CODEs can perform small insertions, deletions, and substitutions with improved efficiency compared to PEMax at many loci in HEK293T cells with plasmid- and RNP-based delivery. We also show that CODEMax can successfully modify mouse and bovine embryos with up to 9.3% precise editing. Further optimization of CODEMax sSystems may enhance editing outcomes in embryos and other challenging contexts. Overall, CODEs complement existing prime editors to expand the toolbox for genome manipulations without double-stranded breaks.

Introduction
Prime editing has gained prominence as a highly effective genome editing tool due to its precision and versatility1-3. Pioneered by Anzalone et al.1, the technology can correct all 12 possible single-base substitutions and make small insertions or deletions without generating double-stranded breaks. Prime editing typically uses a nickase Cas9-reverse transcriptase (nCas9-RT) fusion protein (prime editor) and a prime editing guide RNA (pegRNA), which has a 3’ extension containing a primer binding site (PBS) and reverse transcription template (RTT) encoding the desired edit. The 5’ spacer region of the pegRNA directs the prime editor to the target site, resulting in nicking of the non-target strand by nCas9. Hybridization of the 3’ PBS to the nicked strand then generates an initiation site for reverse transcription to occur. Subsequently, the correct edits are installed into a 3’ edited DNA flap which can be incorporated into the genome1,4.

Although prime editing has been iteratively improved through many rounds at the level of both the prime editor and pegRNA, the system still suffers from variable efficiency across genomic loci5-7. Existing prime editors use M-MLV or other reverse transcriptases, which possess relatively moderate processivity, hindering their capability to perform more complex edits effectively. Additionally, after the prime editor generates a 3’ edited flap, it competes with the 5’ unedited flap for incorporation into the genome, reducing the likelihood of a successful edit. Furthermore, since the pegRNA contains a PBS that is complementary to the spacer sequence, it forms a stable RNA-RNA duplex, which creates a barrier for ribonucleoprotein complexation between the pegRNA and the prime editor8,9. Recent efforts have been made to circumvent the auto-inhibitory interaction within the pegRNA by optimizing the melting temperature of the PBS and by incorporating mismatches into the PBS to reduce misfolded pegRNA interactions which leads to improved prime editing efficiency8,9. Although prime editing can make versatile edits without induction of double-strand breaks in the DNA, the accuracy of prime editors still has room for improvement. One of the most common imprecise edits observed for prime editing is an overextension of the reverse transcriptase past the RTT into the scaffold region of the pegRNA. This readthrough becomes problematic because it results in the incorporation of undesired bases into the genome10, although recent studies have shown methods to lower its occurrence by engineering highly structured regions within the pegRNA4,11,12.

We reasoned that creating chimeric oligonucleotide-directed editing (CODE) systems consisting of a DNA-dependent DNA polymerase paired with a chimeric pegRNA (cpegRNA) containing a DNA primer binding site and a DNA polymerase template may address some of the limitations of current reverse transcriptase-based prime editors. We hypothesized that a cpegRNA could reduce the auto-inhibitory effect observed for traditional pegRNAs, as the DNA-RNA duplex is inherently less stable than the RNA-RNA duplex13. Moreover, DNA polymerases are an abundant family of proteins with diverse molecular properties, rendering them intriguing candidates to achieve chimeric oligonucleotide-directed editing. The use of DNA polymerases with advantageous properties, such as high processivity, proofreading capability, and minimal reverse transcriptase activity has the potential to improve editing efficiency, reduce unintended edits, and enable new types of edits to be performed.

Recent studies have also investigated the use of DNA-dependent DNA polymerases in place of reverse transcriptases for precise search-and-replace genome editing14,15. Liu et al. reported a four-component DNA Polymerase Editing system comprised of nCas9, Phi29 DNA polymerase-MCP fusion protein, sgRNA, and a chimeric DNA-RNA template with an MS2 loop. Ferreira et al. developed Click Editors that utilize nCas9 fused to both a DNA polymerase and a ssDNA tethering domain that localizes the DNA template for polymerization to the edit site. These studies demonstrate the potential of utilizing DNA-dependent DNA polymerases as an additional tool for precision genome editing.

Here we survey a class of 13 CODEs that consist of a cpegRNA and a nickase Cas9-DNA polymerase fusion protein. This simple yet versatile two-component system allows for delivery of the CODE fusion-protein via plasmids, mRNA, or ribonucleoprotein (RNP) complexes for effective and accurate genome editing. We observed that CODE improved gene correction efficiency compared to conventional PE2 and PEMax at several genomic loci in cell culture with low unintended edits. Lastly, we successfully validate CODE in mouse and bovine embryos, achieving precise editing up to 9.3%. Overall, engineered CODEs expand the gene editing toolbox and offer versatility as well as flexibility toward therapeutic applications.

Results
A survey of 13 DNA polymerase-mediated editors for targeted genome modification

To determine whether a nickase Cas9 (nCas9) fused to a DNA polymerase (DNAP) could perform precise genome editing when delivered as a ribonucleoprotein complex, we first constructed nCas9 fusion proteins paired with a variety of wild-type polymerases from viral or bacterial origins (Fig. 1a). A total of 13 DNAPs with diverse properties such as thermostability, proofreading activity, processivity, and size were selected and screened (Fig. 1b). Each construct was expressed in E. coli and purified for ribonucleoprotein delivery. To evaluate the efficiency of CODE candidates, we generated a HEK293T-based reporter cell line containing an open reading frame with a green fluorescence protein (GFP) upstream of a red fluorescence protein (mCherry). We also installed a premature termination codon (PTC) into the mCherry gene so that the cells only display green fluorescence, similar to Liu et al16. Upon successful A-to-G substitution within the PTC, the stop codon is converted to the CAA codon which allows for mCherry translational readthrough.

Next, we designed a cpegRNA to target the constitutively expressed GFP-mCherry reporter gene containing the PTC. The cpegRNA is comprised of a 20-nt RNA targeting sequence, a guide RNA scaffold, and a 3’- end DNA extension sequence containing a primer binding site (PBS) and a DNAP template encoding the desired corrections (Fig. 1c). Each CODE candidate was complexed with the cpegRNA prior to delivery into HEK293T cells via nucleofection and edit efficiency was quantified by the percentage of mCherry positive cells (Fig. 1d)10.

We observed that many CODE candidates were able to induce mCherry activation, albeit with low efficiencies compared to the engineered prime editor PE2. Among the most efficient wild-type CODEs were those that employed polymerases from T4 and T5 bacteriophages, reaching 4.1% and 10.2% mCherry activation, respectively. Additionally, the large fragment Bst DNAP derived from Geobacillus stearothermophilus achieved 4.7% while full length Bst DNAP was even more effective at 11.1% mCherry activation.

Liu et al.14 and Ferreira da Silva et al.15 recently reported prime editing adjacent systems that utilize DNA-dependent DNA polymerases such as Phi29 DNAP and Klenow fragment. Within our editing system, we observed that CODE candidates employing DNA Polymerase I and the Klenow fragment exhibited moderate mCherry activation (3.8% and 4.5%, respectively) while Phi29 DNAP showed low editing activity (2.3%) (Fig. 1e). Although our chimeric editing system differs from these studies, our data support the reported abilities of Phi29 and the Klenow fragment to extend primed templates at a nick in the genomic DNA. Together, these data demonstrate that multiple DNA polymerases can utilize a DNA template within a chimeric pegRNA to perform precise edits in mammalian cells.

Engineering T4 DNA polymerase for improved editing efficiency

For further engineering of CODEs for greater functionality and utility, we selected T4 DNAP and Bst, large fragment DNAP (Bst-LF). T4 DNAP is a mesophilic polymerase that possesses strong 3’-5’ exonuclease activity17,18. We strived to determine how the T4 DNAP location impacted the CODE-T4 editing efficiency by positioning the T4 DNAP either on the C-terminus (CODE-T4v1) or the N-terminus (CODE-T4v2) of nCas9 connected by a 33-amino acid linker (Fig. 2a). We did not observe much difference in the percentage of mCherry activation between the N-terminal and C-terminal fusion constructs. We next aimed to investigate how the 3’-5’ exonuclease activity of T4 DNAP affected the performance of CODE (Fig. 2b). We installed a Y320A mutation on T4 DNAP of CODE-T4v1 editor (CODE-T4v3), which has been shown to diminish the polymerase exonuclease activity by 50-fold19. Notably, this single mutation increased the efficiency of mCherry activation 2.4-fold compared to the wild-type editor (Fig. 2b). We proceeded to install a T4 Gene 32 Protein (gp32) on the N-terminus of T4 DNA polymerase to generate a CODE-T4v4 editor (Fig. 2a,b). T4 gp32 is a single-stranded binding protein (SSB) that is crucial for T4 replication and repair20,21. However, we observed reduced activity compared to CODE-T4v3, possibly due to gp32 SSB being inactive in a fusion format or being sterically hindered by nCas9 and T4 DNAP.

We then inserted a sso7d DNA binding domain at the C-terminus T4 DNAP of CODE-T4v1 and CODE-T4v3 to create CODE-T4v5 and CODE-T4v6. Sso7d, a DNA binding protein derived from Sulfolobus solfactaricus, is known to greatly enhance the processivity of DNA polymerases22. We sought to understand if this binding domain could improve our CODE-T4 editors. Interestingly, we did not observe any significant mCherry activation of these two CODEs compared to the original CODE-T4v1. The CODE-T4v6 editor, which possesses a Y320A mutation, exhibited a similar editing efficiency to the CODE-T4v1. Since the sso7d DNA binding domain possesses some ribonuclease activity, we reasoned that it could potentially degrade the cpegRNA and have a negative impact on CODE23. We therefore deactivated this ribonuclease activity by introducing two mutations E12L and K35L into the sso7d domain to generate CODE-T4v7. However, this version of CODE performed poorly compared to CODE-T4v3 with a 3.8-fold decrease in efficiency (Fig. 2b).

We next focused on engineering the T4 DNAP itself, which has been well-studied and extensively characterized. Mutations such as L412M and I50L have been shown to increase the processivity, although these substitutions cause a slight increase in replication errors24,25. We therefore generated combinations of these CODE-T4 mutants and tested them in HEK293T cells. Notably, we observed up to 20.7% mCherry activation for the CODE-T4Y320A/L412M mutants, which is a 3.8-fold increase compared to CODE-T4v1. We also noted a boost in efficiency for the CODE-T4I50L/Y320A and CODE-T4G255S/Y320A mutants compared to the original CODE-T4v1 (Fig. 2c).

We hypothesized that the 3’-5’ exonuclease activity of T4 DNA polymerase might have a negative impact on the editing efficiency as it could remove bases at the newly synthesized 3’-flap26,27. To test this hypothesis, we generated a CODE-T4D219A and CODE-T5D138A with deficient 3’-5’ exonuclease activity and compared them against the corresponding wild-type CODE-T4 and CODE-T5. Interestingly, we observed no difference in mCherry activation in HEK293T reporter cells, indicating that this 3’-5’ exonuclease activity might have minimal involvement in the prime editing process (Fig. 2d). We finally optimized the amino acid linker length between the nCas9 and T4 DNAP and found that a 45 amino acid linker exhibited the best mCherry activation efficiency (Fig. 2e).

Engineering Bst DNA polymerase for improved editing efficiency

T4 DNA polymerase variants were engineered to increase processivity at the cost of fidelity yet still showed limited activity at endogenous loci. Guided by these findings, we next explored the inherently thermostable Bst DNA polymerase, which was hypothesized to support efficient cellular editing and motivated the development of the Bst-derived CODEMax and CODEMax (exo⁺) systems.

Bst-LF DNAP is a thermophilic DNA polymerase that has strong strand displacement activity; therefore, Bst-LF is often used in isothermal amplification technologies such as Loop-mediated isothermal amplification (LAMP)28,29. Although wild-type Bst-LF is optimally active at high temperatures in amplification reactions, we observed moderate editing efficiency by Bst-LF and full length Bst DNAP in HEK293T cells. Additionally, prime editor 2 (PE2), which utilizes an engineered M-MSLV variant containing five-point mutations that increase the thermostability and processivity, was shown to achieve dramatic increase in prime editing efficiency compared to prime editor 1 (PE1) that contains a wild-type M-MMLV reverse transcriptase1. Although Bst-LF is naturally thermostable, we hypothesized that additional increases in thermostability could further improve its activity in mammalian cells. Thermostability often correlates with overall stability, and for enzymes like M-MLV and Bst-LF operating in a non-native human cell context, greater stability can help maintain its structure and activity. We therefore explored multiple approaches to engineer Bst-LF mutants within our CODE systems to see if increasing the thermostability of Bst-LF DNAP would increase its activity in cells.

Paik et al.30, used a machine learning approach to generate a Bst DNAP variant with increased thermostability. The Bst-LF DNAP variant, referred to as Br512, consists of a modified 47 amino acid actin-binding protein called villin headpiece fused to the N-terminus of the Bst-LF. The fusion of the villin headpiece to Bst-LF DNAP is hypothesized to improve protein folding and increase processivity via stabilization of the DNA/protein complex. We employed the Br512 DNAP in CODE (referred to as CODE-Bstv3) and observed a nearly 3-fold increase in mCherry-positive cells compared to wild-type Bst-LF. In an attempt to further increase efficiency, we tested additional variants of Br512 engineered to have even stronger thermostability. Br512g3.1 and Br512g3.2 variants differ from Br512 in the villin headpiece, where point mutations were rationally designed to supercharge and stabilize the domain (referred to as SC-vHP47). The Br512g3.1 variant bears 3 mutations (N31R, N39K, E43K) on the SC-vHP47, whereas the Br512g3.2 bears 4 mutations (A20K, N31R, N39K, E43K). Furthermore, these two Bst-LF DNAP variants bear additional mutations within the polymerase domain (T493N, A552G, S371D), resulting in a significant increase in thermostability over Br512 while maintaining high functionality in LAMP reactions up to 74°C30. By incorporating these Br512g3.1 and Br512g3.2 DNAP variants into our chimeric oligonucleotide-directed editors, we generated the CODE-Bstv5 and CODE-Bstv6 systems. Notably, CODE-Bstv5 and CODE-Bstv6 resulted in 23.6% and 33.4% efficiency installing the desired edits, approximately 5-fold and 7-fold increases in mCherry activation compared to the original CODE-Bstv1 system.

In-house synthesis of chimeric pegRNA improves editing efficiency

Traditionally, a pegRNA consists of 100% RNA bases and therefore can be synthesized either by enzymatic or chemical synthesis reactions. This advantage provides flexibility to deliver prime editing systems into cells as well as animal models. Oftentimes, it is convenient to co-deliver plasmids encoding prime editors and pegRNAs driven by a U6 promoter. Since the cpegRNA is a chimeric entity consisting of both RNA and DNA bases, it cannot be synthesized by naturally occurring enzymes. Instead, cpegRNAs must be chemically synthesized, which poses challenges for delivery approaches and associated synthesis costs.

To address this hurdle, we developed an inexpensive ligation-based method to synthesize cpegRNAs. Taking advantage of T4 RNA ligase I, which can ligate a 3’-hydroxyl RNA to a 5’-phosphorylated DNA, we established a synthesis protocol for generating full length cpegRNA (Fig. 3a). A benefit of this ligation reaction is that one can easily make a multitude of edit types at the same genomic locus by modifying single stranded DNA oligos which each utilize the same single-guide RNA. This approach drastically decreases the synthesis cost compared to synthetic cpegRNA. Additionally, we placed four modified 2′-O-methylated uracils at the end of the single guide RNA to improve stability and reduce sgRNA scaffold incorporation in the cells (Fig. 3b). We tested in-house synthesized cpegRNAs with CODE-Bstv6 and noted an improved performance with 43.7% mCherry activation (Fig. 3c,d).

Efficient editing of endogenous gene loci with CODEMax and CODEMax(exo+)

We next aimed to investigate how efficiently CODE performs when targeting endogenous genes. Using nucleofection strategies, we observed a significant reduction in editing efficiency for our CODE-Bstv6, possibly due to the cpegRNA chimeric nature initiating cellular response, leading to its degradation. We then explored alternative methods to deliver CODEs by comparing RNP, mRNA, and plasmid approaches. We proposed that encapsulating the cpegRNA with transfection reagents could provide it with some protection from the cellular environment. After several rounds of optimization, we showed that plasmids encoding CODEs and synthetic cpegRNAs can be co-delivered into mammalian cells via transfection (Fig. 3e-g and Fig. S1).

To enhance editing efficiency, we adopted two additional nCas9 mutations (R221K and N394K) that were developed by Chen et al31. to convert PE2 into PEMax. We called this version of chimeric oligonucleotide-directed editing CODEMax, which is a combination of the engineered CODE-Bstv6 and a mutated nCas9 variant (R221K and N394K). Furthermore, our initial screening of CODE candidates (Fig. 1e) showed better performance of a chimeric oligonucleotide-directed editor using full-length Bst DNAP compared to that with Bst-LF. Interestingly, the difference between the truncated and full-length polymerase is the absence of a 5’-3’ exonuclease domain. We hypothesized that a polymerase that possesses strong 5’-3’ exonuclease activity may improve the editing efficiency (Fig. 4a,b and Fig. S2). In prime editing-like systems, the 3’ flap generated after extension by the polymerase, enters competition for incorporation into the genome with the 5’ unedited flap32. Therefore, having a polymerase with 5’-3’ exonuclease activity that can displace and degrade the 5’ unedited strand during extension may be beneficial (Fig. 4b). In support of this hypothesis, Liang et al.33 has demonstrated that the fusion of a T5 exonuclease at the N-terminus of the M-MLV reverse transcriptase increased efficiency of the PE2 system in plants. Therefore, we incorporated the 5’-3’ exonuclease domain back into CODEMax, hereafter referred to as CODEMax(exo+).

After validating CODEMax and CODEMax(exo+) activity in a reporter cell line (Fig. S3), we tested CODEMax and CODEMax(exo+) by targeting multiple genomic regions with a variety of edit types such as base conversion and transversion, short insertion, and short deletion. We observed enhanced editing efficiency of CODEMax and CODEMax(exo+) compared to PE2 and PEMax at several loci such as EMX1, FANCF, SRD5A3 and DNMT1 with HEK3 and MECP2 being exceptions. (Fig. 4c-i and Fig. S4-S7). It should be noted that these data were head-to-head comparisons of synthetic pegRNA/cpegRNA and protein-encoding plasmid via transfection. However, we noticed that expression of pegRNA under U6 promoter resulted in higher efficiency for PE2 and PEMax compared to delivery of synthetic pegRNA (Fig. S8). With the addition of a nicking guide, which significantly increases edit efficiency for traditional prime editors by evading mismatch repair (MMR), we also observed improved prime editing for CODE systems, suggesting a conserved presence of heteroduplex intermediates1 (Fig. 4e-i and Fig. S5). Additionally, CODEMax(exo+) outperformed CODEMax at a majority of edit sites. Next, we sought to characterize the unintended edits of CODE systems. We demonstrate that CODEMax and CODEMax(exo+) exhibited low unintended scaffold incorporation, like that seen for PEMax at certain edit sites (Fig. S9). On the other hand, the amount of local unintended edits as well as unintended edits at predicted off target sites was comparable between PE and CODE systems (Fig. S10 and S11)1,34.

CODEMax enables editing in mouse and bovine embryos with lower efficiency than PEMax

To further evaluate the editing efficiency of CODE relative to conventional prime editing (PEMax), we performed genome editing experiments in both mouse and bovine embryos. We selected genomic targets shown to be efficiently edited by PEMax in mouse embryos as well as novel targets35,36. We injected zygote stage embryos with mRNA encoding for PEMax or CODEMax and pegRNA (PEMax) or cpegRNA (CODEMax). mRNA expressing a dominant negative MMR protein was also delivered, as it has been shown to increase prime editing efficiency in mouse embryos37. Individual mouse and bovine embryos were collected at the blastocyst stage and prepared for amplicon sequencing (Fig. 5a, b and S12).

PEMax exhibited high efficiency at the previously targeted loci Tubb5 and Hoxd13 in mouse embryos (Fig.5c). CODEMax also demonstrated editing in mouse embryos albeit at consistently lower efficiency. CODEMax achieved an average of 5.3% precise edits Tubb5, however was unsuccessful at the Hoxd13 locus (Fig. 5d). Similarly, at the 7SK locus, which has not previously been validated as a prime editing target, PEMax achieved a higher editing efficiency (11.1%) compared to CODEMax (1.8%) for a single base pair substitution (Fig. 5c-d). Some PEMax and CODEMax treated mouse embryos arrested during development prior to reaching the blastocyst stage. At the Dnmt1 and Hoxd13 loci, there was minimal difference in development between PEMax and CODEMax treated embryos however, there was a higher percentage of arrested embryos for CODEMax when targeting Tubb5 (Fig. 5e). Next, we attempted editing of the rpl12 locus in bovine embryos, where CODEMax achieved a precise editing efficiency of 9.3% (Fig. 5f). However, PEMax outperformed CODEMax and reached 20.3% efficiency. Importantly, low unintended edits occurred for both CODE and PEMax across all loci tested demonstrating high accuracy around the edit site (Fig. 5c-d, f).

Methods
Ethics Statement

Mouse experiments were performed in accordance to guidelines and protocols approved by the University of Florida Institutional Animal Care and Use Committee (IACUC) under protocol number IACUC202300000558. CF-1 females (Envigo, Indianapolis, IN, USA) were used for oocytes and embryo collection for gene editing experiments. B6SJLF1/J males used for breeding were obtained from Jackson Laboratory.

General cloning methods and plasmid construction

CODE gene fragments were either obtained from Addgene or synthesized by Twist Biosciences. Bacterial and mammalian expression plasmids for CODEs were cloned using In-Fusion® cloning (Takara Bio, Cat# 638948). Q5 high-fidelity polymerase (New England Biolabs, Cat# M0491L) was used to amplify gene fragments and non-lentiviral backbone for cloning as well as genomic DNA for deep sequencing. For In-Fusion® cloning involving the assembly of lentiviral backbones, PrimeSTAR® GXL DNA Polymerase (Takara Bio, Cat# R050A) was used for amplification. For transfection into mammalian cells, plasmids expressing CODEs were prepared using ZymoPure II Plasmid Midiprep kit (Zymo Research, Cat# D4201) and diluted down to 1 mg/μL prior to transfection.

Chimeric pegRNA synthesis via T4 RNA ligase-mediated ligation

All pegRNAs, sgRNAs, and 5’ phosphorylate ssDNAs were purchased from IDT. cpegRNAs were also purchased from IDT unless otherwise indicated that they were produced via ligation of 5’ phosphorylated ssDNA to sgRNAs. Synthetic pegRNA were purchased from IDT with 2’-O-methylation and phosphorothioate linkage on the first and last three bases. Synthetic cpegRNA we re purchased from IDT with 2’-O-methylation and phosphorothioate linkage on the first three bases but only phosphorothioate linkage on the last three bases. Ligations were prepared using T4 RNA Ligase I (New England Biolabs, Cat# M0204) as follows: 3.5 μL T4 RNA Ligase Buffer, 4.5 μL PEG8000, 2 μL T4 RNA Ligase I, 3 μL 10mM ATP, 2 μL 100uM sgRNA, 5 μL 100 mM 5’ phosphorylated ssDNA, 0.25 μL Murine RNase Inhibitor (New England Biolabs, Cat# M0314), and 14.75 μL water. Reaction volumes greater than 35μL led to decreased ligation efficiencies. Reactions were incubated at 16°C for 16 hours and purified with Monarch® RNA Cleanup Kits (New England Biolabs, Cat# T2030) according to manufacturer instructions. Products were analyzed via 10% TBE-Urea PAGE electrophoresis (Biorad, Cat# 3450088).

CODE protein expression and purification

Bacterial expression plasmids carrying CODEs were transformed into homemade competent cells propagated from Rosetta™ 2(DE3)pLysS Singles™ Competent Cells (Millipore Sigma, Cat# 71401). Individual colonies were picked and inoculated in 50 mL Luria Broth (Fisher Scientific, Cat# BP9723-2) overnight at 37°C. The culture was then scaled up to 4-12 liters of Terrific Broth (RPI, T15000-10000.0) and grown until OD = 0.8-1.0. The culture was then quickly cooled on ice for 10-15 minutes and induced with 1 mM isopropyl ß-D-1-thiogalactopyranoside (IPTG) (Gold Biotechnology, Cat# I2481C100). For CODE constructs that were built based on the pET-PE2-His backbone (Addgene, #170103), the culture was induced at 18°C for 5 hours followed by 26°C for 14-18 hours. For CODE constructs that were built based on PE-Max-pET21a backbone (Addgene, #204471), the culture was induced at 18°C for 16-18 hours.

Cell pellets were collected the next day by centrifugation (4000 xg for 10 minutes), suspended in 100-150 mL lysis buffer (500 mM NaCl, 50 mM Tris-HCl pH = 7.5, 1 mM TCEP-HCl, 20 mM imidazole, and 5% glycerol) followed by sonication. The lysate was centrifuged at 40,000 xg for 45 minutes before passing through a 0.45 μm filter. The clarified lysate was then injected into a prepacked Ni-NTA affinity column (EconoFit Nuvia IMAC Column, Biorad #12009287) in a FPLC (NGC Quest Plus, Biorad) pre-equilibrated with lysis buffer. Proteins were eluted from the column with 40 mL of elution buffer (500 mM NaCl, 50 mM Tris-HCl pH = 7.5, 1 mM TCEP-HCl, 300 mM Imidazole, and 5% glycerol). The eluted solution was concentrated in an Amicon® Ultra Centrifugal Filter, 50 kDa MWCO (Millipore Sigma, UFC905024) down to 10-15 mL and equilibrated with 40 mL of Buffer A (200 mM NaCl, 50 mM Tris-HCl pH = 7.5, 1 mM TCEP-HCl, and 5% glycerol). The protein mixture was then passed through a 5 mL Hitrap Heparin HP column (Cytiva, Cat# 17040701) pre-equilibrated with Buffer A. The column then underwent gradient elution from Buffer A to Buffer B (2000 mM NaCl, 50 mM Tris-HCl pH = 7.5, 1 mM TCEP-HCl, and 5% glycerol). The purest fractions of the protein were pooled together and concentrated in an Amicon® Ultra Centrifugal Filter (50 kDa MWCO) in final buffer C (500 mM NaCl, 50 mM Tris-HCl pH = 7.5, 1 mM TCEP-HCl, and 5% glycerol) before storing at -80°C. When in use, the protein was diluted in storage buffer (300 mM NaCl, 10 mM Tris-HCl, 0.1 mM EDTA, 1 mM DTT, 50% glycerol, 0.1% Triton® X-100, final pH = 7.4 at 25°C) down to 50 μM and store at -20°C.

Mammalian cell culture

HEK293T and Lenti-X™ 293T were obtained from ATCC (CRL-3216) and Takara Bio (#632180), respectively. U2OS cells were obtained ATCC (HTB-96). The cells were tested with mycoplasma using MycoAlert® Mycoplasma Detection Kit (Lonza, Cat# LT07-118). The cells were cultured and passaged in D10 medium containing DMEM high glucose with GlutaMAX™ supplement and pyruvate (Gibco, Cat# 10569044), 10% Fetal Bovine Serum (Gibco, Cat# A3160902), 1X Penicillin-Streptomycin (Gibco, Cat# 15140122), and 1X MEM non-essential amino acids (Gibco, 11140035). All cell lines were incubated at 37°C and 5% CO2.

Reporter cell line and stably expressed CODE cell line generation

For lentiviral packaging in a T75 flask, 10 μg of transfer plasmid was co-transfected with 5 μg of pMD2.G (Addgene, #12259) and 7.5 μg of psPAX2 (Addgene, #12260) into Lenti-X™ 293T cells with 50 μL of Lipofectamine 3000 and 42 μL P3000 Enhancer reagents (ThermoFisher, Cat# L3000008), which were diluted in Opti-MEM™ I Reduced Serum Medium (Gibco, 31985062) following the manufacturer instructions. The medium was changed to D10 medium six hours later, and the cells were incubated for an additional 48-60 hours. The cells were harvested and pelleted down via centrifugation at 4000 xg for 10 minutes at 4°C. The supernatant was passed through a 0.45 μm filter, aliquoted, and stored at -80°C until use.

Lentiviral particles were titered using the qPCR Lentivirus Titer Kit (ABM, Cat# LV900). For lentiviral transduction, 5 x105 HEK293T cells were infected at an MOI of 5 with viral supernatant via reverse transduction. Briefly, viral supernatant was added to a 6-well plate first. The cells were then counted and resuspended in D10 medium supplemented with 10 μg/mL of TransduceIT™ Transduction Reagent (Mirus Bio, Cat# MIR6620) and transferred to the wells pre-added with viral supernatant. The cells were incubated for 72 hours before flow cytometry sorting and/or antibiotic selection.

Mammalian cell RNP nucleofection and plasmid transfection

Ribonucleoprotein (RNP) complexes were delivered into HEK293T cells via a 4D-Nucleofector® X Unit (Lonza, Cat# AAF-1003X) in a strip format using SF Cell Line 4D-Nucleofector™ X Kit S (Lonza, Cat# V4XC-2032). Purified PEs and CODEs were complexed with either pegRNA or cpegRNA to form RNP at 50 pmol protein: 200 pmol peg/cpegRNA ratio for 15 minutes at room temperature. Around 2x105 cells were resuspended in Lonza SF buffer, mixed with the RNP, and electroporated using the program ED-130. The mixture was then incubated at 37°C and 5% CO2 for 10 minutes before adding to a 48-well plate pre-added with DMEME medium supplemented with 10% Fetal Bovine Serum (no antibiotic). Cells were harvested after 72 hours.

Plasmids encoding prime editors and chimeric editors and synthetic pegRNA/cpegRNA were delivered to HEK293T cells utilizing TransIT-X2® Dynamic Delivery System (Mirus Bio, Cat# MIR6000). 24 hours prior to transfection cells were seeded at of 2x105 cells per well in 24-well plates. Immediately prior to transfection, the media was replaced with antibiotic-free D10 media. 1500 ng of chimeric/prime editor plasmid, 750 ng of synthetic cpegRNA/pegRNA, and 500 ng of ngRNA plasmid (if applicable) were complexed for 20 minutes with 6 μL of TransIT-X2 in 50 μL of Opti-MEM™ Reduced Serum Medium (Gibco, Cat# 31985062). After complexing, reactions were added dropwise to each well and harvested after 72 hours.

Bovine In Vitro Embryo Production

Germinal vesicle stage oocytes (GV oocytes) were collected as cumulus-oocyte complexes (COCs) from ovaries of slaughtered cows. Oocytes were matured for 22–23 hours in BO-IVM medium (IVF Bioscience, Falmouth, UK) at 38.5°C with 6% CO2. Cryopreserved semen from a proven Holstein bull was prepared in BO-SemenPrep medium (IVF Bioscience, Falmouth, UK) to a final sperm concentration of 2 × 10⁶ spermatozoa/ml for in vitro fertilization (IVF). Gametes were co-incubated in BO-IVF medium at 38.5°C with 6% CO2 for 16 hours. After IVF, embryos were denuded from cumulus cells by 5 minutes of vortexing in BO-Wash medium (IVF Bioscience, Falmouth, UK).

Mouse Embryo Isolation and Culture

Female mice (at least 6 weeks old) were superovulated with 5 IU pregnant mare serum gonadotropin (PMSG; Folligon, Merck Animal Health). 46 hours post-administration, mice were injected with 5 IU hCG and mated overnight with B6SJLF1/J males. Zygotes were collected from excised oviducts 18 hours post-mating and cultured in KSOM medium (M8410; Sigma-Aldrich). All animal experiments adhered to guidelines and protocols approved by the Institutional Animal Care and Use Committee (IACUC) at the University of Florida.

In Vitro Transcription and Microinjection

H2b:GFP (provided by Dr. Martin Anger, Laboratory of Cell Division Control, IAPG CAS), PEMax, and CODEMax mRNA was prepared from linearized plasmids using the T3 mMessage mMachine Kit (Ambion). Microinjections were performed between 20 and 24 hours after hCG administration. The microinjection platform was equipped with a microinjector (FemtoJet 4i, Eppendorf), an inverted microscope (Leica, DMi8), and micromanipulators (Narishige). The injection solution contained 20 ng/μL of H2b:GFP mRNA, 300 ng/μL PEMax or CODEMax mRNA, and 5 uM pegRNA or cpegRNA in MEM-PVA medium. Embryos were cultured in a single drop of KSOM medium and covered with mineral oil until reaching the blastocyst stage (day 3.5 and 7.5 for mouse and bovine, respectively).

Genomic DNA library preparation and targeted amplicon deep sequencing

Genomic DNA was extracted from the HEK293T and U2OS cells using the QuickExtract™ DNA Extraction Solution (Biosearch Technologies) system according to the manufacturer’s instructions. Embryos were harvested with the Red Extract-N-Amp kit (Sigma XNAT-100RXN) using 4 μL of extraction buffer, 1 μL of tissue preparation buffer, and 4 μL of neutralization buffer37,41. The DNA was then amplified in the first round of PCR using Q5 DNA Polymerase (NEB), and Illumina barcodes were appended during a second PCR. The products were then gel extracted, pooled together, and loaded on an Illumina MiSeqDx using a MiSeq Reagent Nano Kit v2 (Illumina, Cat# MS-101-1001) according to the manufacturer’s protocol. CRISPResso2 was used to determine the percentage of precise editing and indels42. Quantification window was defined by the parameter “-qwc” spanning 10 base pairs upstream and downstream flanking the targeting sequence in cases where there was no nicking guide and 10 base pairs flanking the targeting sequence and the nicking guide in the case where a nicking guide was used.

Statistics and Reproducibility

All cell culture experiments were performed in triplicates. The number of embryos tested were determined based on the number of embryos produced at the time of the experimentation (n12). No data were excluded from the analyses, and no statistical method was used to predetermine sample size.

Data availability

All data supporting the study are available within the main text, supplementary information, and supplementary data files. Source data are also included in this paper. Sequencing data have been deposited to NCBI database with the accession number PRJNA1266727 [https://www.ncbi.nlm.nih.gov/bioproject/PRJNA1266727].

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In this manuscript by Nguyen, et al, the authors describe the development of the CODE genome editing system, which uses a prime editing-like mechanism but replaces the reverse transcriptase (RNA- and DNA-dependent DNA polymerase) and RNA template in prime editing systems with non-RT DNA polymerases (T4, Bst) and DNA template. The authors began by testing 13 different DNA polymerase enzymes as fusions to Cas9 nickase, co-delivered with a chimeric pegRNA harboring a DNA template, in a HEK293T reporter cell line with. Several DNA polymerases were able to support editing and activation of the reporter, with full-length Bst DNA polymerase showing the best activity among the wild-type enzymes. T4 and Bst polymerases were chosen for further engineering efforts, which included approaches that diminish 3’-5’ exonuclease activity of the polymerase, optimize the linker length, increase DNA binding and processivity, and increase thermostability. These efforts yielded multiple CODE-T4 and CODE-Bst systems with improved editing activities compared to the starting constructs. The authors next described an approach to efficiently synthesize chimeric pegRNAs using T4 RNA ligase to conjugate chemically synthesized guide RNAs to DNA oligonucleotides. They demonstrated that cpegRNAs generated in this manner were indeed functional for editing. The authors next generated a CODEMax editor from CODE-Bstv6 by incorporation of Cas9 mutations, and a CODEMax(exo+) editor that reintroduces the 5’-3’ exonuclease domain of Bst into the fusion protein. In some cases, CODEMax(exo+) substantially improved editing efficiency, while in other cases, CODEMax was superior. Finally, the authors demonstrated that they could successfully edit mouse and bovine embryos with CODEMax by injecting CODEMax mRNA along with a chimeric pegRNA and an mRNA encoding an MMR suppressing MLH1 dominant negative protein.\nMajor Comments\nThere are two prior studies that have set the precedence for the use of DNA templates and non-RT DNA polymerases for prime editing approaches, both of which are cited and discussed by the authors in the introduction: Liu, et al (PMID 37709915), and da Silva, at al (PMID 37745481). However, these prior publications use distinct components and architectures, and they do not develop the T4 and Bst DNA polymerases used in this study. In some ways, this study represents the simplest design of DNA polymerase/DNA template prime editing system that most resembles the design of the original PE system. The authors also develop a straightforward approach for generating chimeric RNA/DNA pegRNAs based on a simple ligation approach, which makes the system straightforward to use.\nWhile this work does not establish a clear benefit for using the CODE system over the existing RT and RNA-based prime editing systems in all situations, there are potentially many avenues for optimization in the future, as well as specific applications where CODE might be superior. I have just one experimental request below that is necessary to characterize the activity of CODE editing systems in the context of other strategies.\n1. In Liu, et al (PMID, 37709915), the authors demonstrated that MMLV can utilize DNA templates for prime editing. In this CODE manuscript, many of the data panels (e.g., figure 2e, 3d, 3f, 3g, 4c-h) contain PE2 or PEmax editor comparisons, but it is not clear from the figure legends whether the PE2 and PEmax editors are being tested with chimeric pegRNAs or standard synthetic all-RNA pegRNAs. Using chimeric pegRNAs offers the best approach for comparing the DNA polymerase enzymes directly (i.e., MMLV to Bst), but using an all-RNA pegRNA offers the best approach for comparing the standard prime editing system to the CODE systems. In addition, figure S6 generally shows lower prime editing with synthetic pegRNAs compared to U6-transcribed pegRNAs, suggesting synthesis quality or delivery could be influencing editing outcomes. The authors should generate synthetic chimeric pegRNAs and synthetic all-RNA pegRNAs using the same synthetic approach (i.e., T4 RNA ligase) if possible, and at two or more endogenous genomic loci, the authors should characterize the editing efficiency of PE2 or PEmax with both chimeric pegRNAs and all-RNA pegRNAs and compare to the CODE systems. These experiments would help to contextualize the utility of CODE and help to elucidate where the benefits of CODE may come from.\nMinor comments\n1. At the end of the second paragraph of the introduction, in reference to reverse transcription into the scaffold, the authors state that “recent studies have shown methods to lower its occurrence by engineering highly structured regions within the pegRNA.” Suggest adding citation PMID 40204736, “Modified pegRNAs mitigate scaffold-derived prime editing by-products,” to this statement.\n2. Figure 3f-g are missing corresponding descriptions in the figure legend.\n3. For figures containing editing data, including supplementary figures, it would be helpful to note somewhere in the figure or in the legend which cell type the experiments were conducted in.\n4. There is no description in the main text or methods about whether chemical modifications were present in the synthetic pegRNAs/cpegRNAs used. Those details are critical to anyone attempting to reproduce this work, so specific details should be provided.", "reviewer_2": "The article reported a novel genome editing tool, the Chimeric Oligonucleotide Directed Editing (CODE) system, which represents an improvement over existing Prime editing technologies. By combining a DNA-dependent DNA polymerase with a chimeric pegRNA (cpegRNA), it looks that the CODE system partially addresses the limitations of the reverse transcriptase used in traditional Prime editing. The study provides additional toolkit of genome editing as well as offers potential solutions to some of the limitations of existing Prime editing technologies, such as unstable editing efficiency and limited capacity for complex editing. Also it proposes future directions for optimizing the CODE system. There are certain issues:\n1. The connection between the results sections “Engineering T4 DNA polymerase for improved editing efficiency” and “Engineering Bst DNA polymerase for improved editing efficiency” is not clear. The subsequent development of CODEMax and CODEMax (exo+) is also based on variants of Bst DNAP. It is highly suggested to show how the findings from T4 DNAP were applied to the modification of Bst DNAP, as well as a comparison between T4 DNAP and Bst DNAP. This would help explain the rationale for developing CODEMax and CODEMax (exo+) based on Bst DNAP, thereby enhancing the logical coherence of the manuscript.\n2. The data in the results section “Efficient editing of endogenous gene loci with CODEMax and CODEMax (exo+)” show that, compared with PE2 and PEMax, CODEMax and CODEMax (exo+) have improved editing efficiency at several sites in cells, but lower efficiency at some other sites (Figures 4c-i and S3-S5). It is suggested to provide the reasons for these differences, and compare the advantages and limitations of CODEMax and CODEMax (exo+) with PE2 and PEMax.\n3. The data in the results section “CODEMax enables editing in mouse and bovine embryos with lower efficiency than PEMax” show that, despite higher accuracy, CODEMax has much lower editing efficiency than PEMax at multiple sites in embryos, and no editing at all at the Hoxd13 site (Figures 5c-d, f). It doesn’t support the robust application of CODEMax. It may be further discussed, highlighting the advantages of CODEMax as well as its limitations.\n4. The editing sites chosen in the results sections “Efficient editing of endogenous gene loci with CODEMax and CODEMax (exo+)” and “CODEMax enables editing in mouse and bovine embryos with lower efficiency than PEMax” are completely different between cells and embryos, and the results are not consistent. Parallel comparison by selecting the same editing sites in both cells and embryos should be performed for the editing efficiency of CODEMax and PEMax.\n5. The results in Figure 1e show the editing efficiency of CODE-T5 (10.2%) and CODE-T4 (4.1%) in the reporter cell line. Why was T4 DNAP chosen for the futher study rather than the more efficient T5 DNAP?\n6. Bst-LF DNAP is a naturally thermostable enzyme. Is there any reason for enhancing its thermostability could improve its overall performance in cells? Although the subsequent results confirmed this hypothesis, explaining the reasoning process and discussing the relevant mechanisms would enhance the manuscript.\n7. One of the advantages is that CODEMax has fewer unintended edits. However, the observation of unintended edits is limited to the vicinity of the editing sites, and there is a lack of a systematic assessment of the off-target effects of the CODEMax system. It is suggested to provide experimental data on off-target effects, such as whole-genome sequencing (WGS) or RNA sequencing (RNA-seq) results, to evaluate the specificity and safety of the CODEMax system.\n8. The long-term development and physiological functions of the edited embryos is not presented. At least, it should be discussed in the discussion section."}, "author_response": "Authors’ response to the reviewers: We appreciate the reviewers’ feedback on our manuscript titled “Efficient Genome Editing with Chimeric Oligonucleotide-Directed Editing”. We have addressed all the comments below with responses marked in blue and changes to the manuscript highlighted in yellow.\nREVIEWER COMMENTS\nReviewer #1 (R1) (Remarks to the Author):\nIn this manuscript by Nguyen, et al, the authors describe the development of the CODE genome editing system, which uses a prime editing-like mechanism but replaces the reverse transcriptase (RNA- and DNA-dependent DNA polymerase) and RNA template in prime editing systems with non-RT DNA polymerases (T4, Bst) and DNA template. The authors began by testing 13 different DNA polymerase enzymes as fusions to Cas9 nickase, co-delivered with a chimeric pegRNA harboring a DNA template, in a HEK293T reporter cell line with. Several DNA polymerases were able to support editing and activation of the reporter, with full-length Bst DNA polymerase showing the best activity among the wild-type enzymes. T4 and Bst polymerases were chosen for further engineering efforts, which included approaches that diminish 3’-5’ exonuclease activity of the polymerase, optimize the linker length, increase DNA binding and processivity, and increase thermostability. These efforts yielded multiple CODE-T4 and CODE-Bst systems with improved editing activities compared to the starting constructs. The authors next described an approach to efficiently synthesize chimeric pegRNAs using T4 RNA ligase to conjugate chemically synthesized guide RNAs to DNA oligonucleotides. They demonstrated that cpegRNAs generated in this manner were indeed functional for editing. The authors next generated a CODEMax editor from CODE-Bstv6 by incorporation of Cas9 mutations, and a CODEMax(exo+) editor that reintroduces the 5’-3’ exonuclease domain of Bst into the fusion protein. In some cases, CODEMax(exo+) substantially improved editing efficiency, while in other cases, CODEMax was superior. Finally, the authors demonstrated that they could successfully edit mouse and bovine embryos with CODEMax by injecting CODEMax mRNA along with a chimeric pegRNA and an mRNA encoding an MMR suppressing MLH1 dominant negative protein.\nMajor Comments\nThere are two prior studies that have set the precedence for the use of DNA templates and non-RT DNA polymerases for prime editing approaches, both of which are cited and discussed by the authors in the introduction: Liu, et al (PMID 37709915), and da Silva, at al (PMID 37745481). However, these prior publications use distinct components and architectures, and they do not develop the T4 and Bst DNA polymerases used in this study. In some ways, this study represents the simplest design of DNA polymerase/DNA template prime editing system that most resembles the design of the original PE system. The authors also develop a straightforward approach for generating chimeric RNA/DNA pegRNAs based on a simple ligation approach, which makes the system straightforward to use.\nResponse: We sincerely thank the reviewer for the careful evaluation and favorable view of our work. The constructive feedback provided has been valuable in helping us further refine the manuscript.\nR1.1. In Liu, et al (PMID, 37709915), the authors demonstrated that MMLV can utilize DNA templates for prime editing. In this CODE manuscript, many of the data panels (e.g., figure 2e, 3d, 3f, 3g, 4c-h) contain PE2 or PEmax editor comparisons, but it is not clear from the figure legends whether the PE2 and PEmax editors are being tested with chimeric pegRNAs or standard synthetic all-RNA pegRNAs. Using chimeric pegRNAs offers the best approach for comparing the DNA polymerase enzymes directly (i.e., MMLV to Bst), but using an all-RNA pegRNA offers the best approach for comparing the standard prime editing system to the CODE systems. In addition, figure S6 generally shows lower prime editing with synthetic pegRNAs compared to U6-transcribed pegRNAs, suggesting synthesis quality or delivery could be influencing editing outcomes. The authors should generate synthetic chimeric pegRNAs and synthetic all-RNA pegRNAs using the same synthetic approach (i.e., T4 RNA ligase) if possible, and at two or more endogenous genomic loci, the authors should characterize the editing efficiency of PE2 or PEMax with both chimeric pegRNAs and all-RNA pegRNAs and compare to the CODE systems.\nThese experiments would help to contextualize the utility of CODE and help to elucidate where the benefits of CODE may come from.\nResponse to R1.1: We thank the reviewer for this thoughtful suggestion. To address this point, we have performed additional experiments using both a fluorescent reporter system (A-to-G conversion described in Figure 1c) and endogenous genomic loci. In these experiments, we compared the editing efficiencies of PE2 and PEMax using chimeric pegRNAs (cpegRNAs) and the editing efficiencies of CODEMax and CODEMax(exo+) using all-RNA pegRNAs. The data are shown here:\nReporter editing:\nWe also wish to clarify that the data panels referenced by the reviewer (e.g., Figures 2e, 3d, 3f, 3g, and 4c–h) present results obtained using PE2 or PEMax editors with all-RNA pegRNAs. To avoid any ambiguity, we have revised the corresponding figure captions to explicitly state the type of pegRNA used in each comparison.\nMinor comments\nR1.2. At the end of the second paragraph of the introduction, in reference to reverse transcription into the scaffold, the authors state that “recent studies have shown methods to lower its occurrence by engineering highly structured regions within the pegRNA.” Suggest adding citation PMID 40204736, “Modified pegRNAs mitigate scaffold-derived prime editing by-products,” to this statement.\nResponse to R1.2: We thank the reviewer for noticing this. The citation has been added to the introduction section.\nR1.3. Figure 3f-g are missing corresponding descriptions in the figure legend.\nResponse to R1.3: Thank you for catching this mistake. We have added descriptions of Figure 3f-g to the figure caption.\nR1.4. For figures containing editing data, including supplementary figures, it would be helpful to note somewhere in the figure or in the legend which cell type the experiments were conducted in.\nResponse to R1.4: We thank the reviewer for the suggestion. We have added the cell lines used in all figures in the corresponding figure captions.\nR1.5. There is no description in the main text or methods about whether chemical modifications were present in the synthetic pegRNAs/cpegRNAs used. Those details are critical to anyone attempting to reproduce this work, so specific details should be provided.\nResponse to R1.5: We thank the reviewer for their feedback. We have added more details in the method section regarding the modifications of the synthetic pegRNAs/cpegRNAs used in this study.\nThe article reported a novel genome editing tool, the Chimeric Oligonucleotide Directed Editing (CODE) system, which represents an improvement over existing Prime editing technologies. By combining a DNA-dependent DNA polymerase with a chimeric pegRNA (cpegRNA), it looks that the CODE system partially addresses the limitations of the reverse transcriptase used in traditional Prime editing. The study provides additional toolkit of genome editing as well as offers potential solutions to some of the limitations of existing Prime editing technologies, such as unstable editing efficiency and limited capacity for complex editing. Also it proposes future directions for optimizing the CODE system. There are certain issues:\nR2.1. The connection between the results sections “Engineering T4 DNA polymerase for improved editing efficiency” and “Engineering Bst DNA polymerase for improved editing efficiency” is not clear. The subsequent development of CODEMax and CODEMax (exo+) is also based on variants of Bst DNAP. It is highly suggested to show how the findings from T4 DNAP were applied to the modification of Bst DNAP, as well as a comparison between T4 DNAP and Bst DNAP. This would help explain the rationale for developing CODEMax and CODEMax (exo+) based on Bst DNAP, thereby enhancing the logical coherence of the manuscript.\nResponse to R2.1: We are grateful for the reviewer’s thoughtful comment and the opportunity to clarify the rationale linking the two engineering efforts. The engineering of T4 DNA polymerase and Bst DNA polymerase was performed using distinct optimization strategies aimed at addressing different biochemical limitations. Specifically, T4 DNA polymerase was engineered to increase processivity and therefore speed up its template engagement, whereas Bst DNA polymerase was optimized to improve thermostability, similar to the rationale used for engineering M-MLV reverse transcriptase in the prime editing systems. T4 DNA polymerase is a mesophilic enzyme while Bst DNA polymerase is a thermophilic enzyme. Therefore, we referred to the previous studies in the literature to select the best approach for each enzyme1-5.\nWe also wanted to clarify that our initial studies with T4 DNA polymerase–based CODE editors yielded promising activity in reporter assays, but these variants exhibited low editing efficiencies at endogenous genomic loci, hinting that higher thermostability and hybrid tolerance were critical for high performance in cells. This observation motivated the transition to Bst DNA polymerase, which demonstrated robust endogenous editing activity even in its initial form. We therefore focused subsequent efforts on Bst-derived variants (CODEMax and CODEMax (exo+)) for further engineering and optimization.\nWe have revised the Results section to clearly articulate this rationale and to highlight the distinct engineering objectives of the T4- and Bst-based systems. The added statements read as follows:\n“T4 DNA polymerase variants were engineered to increase processivity at the cost of fidelity yet still showed limited activity at endogenous loci. Guided by these findings, we next explored the inherently thermostable Bst DNA polymerase, which was hypothesized to support efficient cellular editing and motivated the development of the Bst-derived CODEMax and CODEMax (exo⁺) systems.”\nWhile the optimization of T4 DNA polymerase–based CODE provides valuable mechanistic insights, its detailed development will be pursued in a separate study. In the present work, we focus on the engineering and characterization of the Bst DNA polymerase–based CODE systems, which demonstrated more robust editing activity when targeting endogenous genes.\nR2.2. The data in the results section “Efficient editing of endogenous gene loci with CODEMax and CODEMax (exo+)” show that, compared with PE2 and PEMax, CODEMax and CODEMax (exo+) have improved editing efficiency at several sites in cells, but lower efficiency at some other sites (Figures 4c-i and S3-S5). It is suggested to provide the reasons for these differences, and compare the advantages and limitations of CODEMax and CODEMax (exo+) with PE2 and PEMax.\nResponse to R2.2: We appreciate the reviewer’s insightful feedback. The differences in editing efficiency likely reflect intrinsic distinctions between the reverse transcription–based mechanism of PE2/PEMax and the DNA polymerase–driven priming mechanism of the CODE systems. Because the molecular details of priming and extension by DNA polymerases in the context of chimeric RNA/DNA templates remain poorly understood, it is possible that optimal parameters such as the lengths of the primer binding site (PBS) and the edit template differ between CODE and PE systems.\nIn this study, we designed all cpegRNA for CODE systems using PBS and edit template architectures that had been extensively optimized for PE2 and PEMax to enable a controlled, side-by-side comparison. However, these configurations may not represent the optimal design for DNA polymerase–based systems. One limitation of this work is the practical constraint imposed by the synthesis of chimeric pegRNAs, which limits the number of designs that can be feasibly tested. Future optimization of PBS/template lengths, template chemistry, and polymerase variants may further enhance the efficiency and broaden the applicability of the CODE platform.\nWe have revised the Discussion section to include this explanation and to explicitly discuss the advantages and current limitations of CODEMax and CODEMax (exo⁺) relative to PE2 and PEMax. The added discussion reads as follows:\n“The variable efficiencies observed across loci likely reflect mechanistic differences between reverse transcriptase– and DNA polymerase–driven priming. Because the optimal primer binding site and edit template lengths for DNA polymerases remain uncharacterized, we adopted PBS/template architectures previously optimized for PE2 and PEMax to enable a controlled comparison. These parameters may not be optimal for CODE-based systems, and future studies aimed at tuning primer-template design and polymerase kinetics may further enhance editing efficiency and consistency across targets.”\nResponse to R2.3: We appreciate the reviewer’s suggestion to further discuss the advantages and limitations of CODE. Our results demonstrate that CODE systems enable site-specific editing in HEK293T, U2OS, and mouse and bovine embryos (Figs. 1–4, 5, and S3). However, the performance of PEMax, CODEMax, and CODEMax (exo+) varies depending on the cellular context, target locus, and type of edit. While the variable efficiency of CODE (particularly in embryos) is a limitation, its superior performance over PEMax for certain edits highlights CODE as an important complementary tool. Therefore, when targeting novel loci, users may benefit from designing constructs for both PEMax and CODE to identify the most efficient approach for their specific target. As optimal cpegRNA design parameters remain to be fully established, further improvements could enhance CODE’s potential. Additionally, it is important to note that prime editing has not previously been performed in bovine embryos; therefore, these data represent the first demonstration of prime editing in this system.\nTo better reflect these points, we have updated the discussion section accordingly:\n“The engineered CODE systems make advancements upon a new class of genome editors utilizing DNA-dependent DNA polymerases. The abundance and diversity of DNA polymerases across all domains of life offer a rich resource for tool development in the future. Key properties of wild-type DNA polymerases can be harnessed for enhanced editing performance such as thermostability, processivity, proofreading ability, and 5’ to 3’ exonuclease activity. Traditional prime editors have been extensively engineered and refined over many years to improve their performance; similarly, this study demonstrates that CODE systems have great potential for continued optimization. In addition, future efforts can customize polymerases for specific applications, as exemplified by the specialization of prime editors by Doman et al6. Importantly, the relative performance of CODE and PEMax varies with the target locus and edit type, with each system showing advantages in different contexts, thereby providing complementary tools for diverse editing applications. Expanding the genome editing toolbox to include systems driven by reverse transcriptases and DNA-dependent DNA polymerases will open new opportunities for precision genome engineering.”\nR2.4. The editing sites chosen in the results sections “Efficient editing of endogenous gene loci with CODEMax and CODEMax (exo+)” and “CODEMax enables editing in mouse and bovine embryos with lower efficiency than PEMax” are completely different between cells and embryos, and the results are not consistent. Parallel comparison by selecting the same editing sites in both cells and embryos should be performed for the editing efficiency of CODEMax and PEMax.\nWe selected the more closely conserved rpl12 locus to directly assess editing performance on similar sequences across distinct biological contexts. We tested the mismatched bovine peg/cpegRNA and a redesigned human-matched peg/cpegRNA with PEMax, CODEMax, and CODEMax (exo+) in HEK293T cells. PEMax tolerated the mismatches and achieved 5.8% editing using the bovine pegRNA, whereas neither CODEMax nor CODEMax (exo+) showed detectable activity. Interestingly, none of the systems edited when the fully matched human peg/cpegRNA was used. These results highlight a broader challenge in peg/cpegRNA design, where in some cases a mismatched pegRNA can outperform a sequence-matched version7. The new experimental data supporting this analysis is below to demonstrate cross-system comparison and illustrate these sequence-dependent and context-dependent effects.\nResponse to R2.5: We appreciate the reviewer’s thoughtful question. We were also encouraged by the strong reporter activity observed with T5 DNA polymerase. However, we selected T4 DNA polymerase for subsequent development for two main reasons. First, T4 DNAP is one of the most extensively characterized DNA polymerases with a substantial literature describing the effect of various mutations on processivity and fidelity, providing a strong foundation for rational engineering and structure-guided mutagenesis1-3. Second, although T5 DNAP initially showed higher reporter editing activity, we encountered consistent difficulties in purifying the nCas9–T5 DNAP fusion protein. It was challenging to achieve sufficient yield and quality for follow-up experiments during our initial screening in which purified proteins were required for RNP delivery, despite multiple optimization attempts. Given these constraints, we prioritized T4 DNAP as a tractable and well-studied platform for initial proof-of-concept engineering of the CODE system.\nWe also added a statement on the major reason why T5 DNA polymerase was not explored further in the Discussion section. Hopefully, it helps clarify why we didn’t choose T5 DNA polymerase for optimization.\n“Although T5 DNA polymerase exhibited higher reporter editing activity, we selected T4 DNA polymerase for subsequent optimization because its structure and biochemical properties have been extensively characterized, facilitating rational engineering.”\nR2.6. Bst-LF DNAP is a naturally thermostable enzyme. Is there any reason for enhancing its thermostability could improve its overall performance in cells? Although the subsequent results confirmed this hypothesis, explaining the reasoning process and discussing the relevant mechanisms would enhance the manuscript.\nResponse to R2.6: We thank the reviewer for the question. Although Bst DNA polymerase is intrinsically thermostable, further enhancing its stability can improve the robustness and efficiency of the Cas9–DNAP fusion within the cellular environment. First, higher thermostability often correlates with improved protein folding and resistance to proteolytic degradation under crowded intracellular conditions, thereby increasing the effective concentration and half-life of the enzyme in cells. Second, thermostable variants tend to maintain stronger and longer-lived interactions with nucleic acid substrates; in the context of prime-editing–like reactions, this can enhance processivity and reduce premature dissociation from RNA/DNA hybrids8. Third, previous optimization of M-MLV reverse transcriptase for PE2 and PEMax demonstrated that mutations conferring thermostability also improved editing efficiency at physiological temperatures, suggesting that structural rigidity, rather than heat tolerance, drives better catalytic performance9. Together, these factors provide a mechanistic basis for why improving thermostability in\nTo better explain the reasoning and potential mechanism behind the correlation between thermostability and cellular activity, we made the below changes to the section “Engineering Bst DNA polymerase for improved editing efficiency”:\n“Bst-LF DNAP is a thermophilic DNA polymerase that has strong strand displacement activity; therefore, Bst-LF is often used in isothermal amplification technologies such as Loop-mediated isothermal amplification (LAMP)27,28. Although wild-type Bst-LF is optimally active at high temperatures in amplification reactions, we observed moderate editing efficiency by Bst-LF and full length Bst DNAP in HEK293T cells. Additionally, prime editor 2 (PE2), which utilizes an engineered M-MLV variant containing five-point mutations that increase the thermostability and processivity, was shown to achieve dramatic increase in prime editing efficiency compared to prime editor 1 (PE1) that contains a wild-type M-MMLV reverse transcriptase9. Although Bst-LF is naturally thermostable, we hypothesized that additional increases in thermostability could further improve its activity in mammalian cells. Thermostability often correlates with overall stability, and for enzymes like M-MLV and Bst-LF operating in a non-native human cell context, greater stability can help maintain its structure and activity. We therefore explored multiple approaches to engineer Bst-LF mutants within our CODE systems to see if increasing the thermostability of Bst-LF DNAP would increase its activity in cells.”\nR2.7. One of the advantages is that CODEMax has fewer unintended edits. However, the observation of unintended edits is limited to the vicinity of the editing sites, and there is a lack of a systematic assessment of the off-target effects of the CODEMax system. It is suggested to provide experimental data on off-target effects, such as whole-genome sequencing (WGS) or RNA sequencing (RNA-seq) results, to evaluate the specificity and safety of the CODEMax system.\nResponse to R2.7: We thank the reviewer for this important comment regarding off-target assessment. We recognize that methods such as WGS or RNA-Seq could offer additional insights, but targeted deep sequencing of the most likely off-target sites is more aligned with the scope and resources of the current study. Additionally, distant off-target effects have been shown to be very infrequent for prime editing (less than 1% of reads), and therefore unlikely to be captured with the read depth achieved during WGS (which has only10-30x read depth and therefore cannot accurately quantify off-target edits below XX% frequency). For these reasons, we selected a more focused analysis method that enables detection even of low abundance off-target edits. Previously used for prime editing off-target characterization, this method detects genome modification at the most common off-target sites for Cas9, which have been experimentally determined9-11.\nWe analyzed the gDNA from two previous editing experiments (HindIII insertion at the HEK3 locus and +6 G deletion at the FancF locus) to quantify all modifications at the four most likely off-target sites reported for each locus.\nWe have revised the results section to include the following:\n“Next, we sought to characterize the unintended edits of our CODE systems. We demonstrate that CODEMax and CODEMax(exo+) exhibited low unintended scaffold incorporation like that seen for PEMax at certain edit sites (Fig. S7). On the other hand, the amount of local unintended edits as well as unintended edits at predicted off target sites was comparable between PE and CODE systems (Fig. S8 and S9).\nR2.8. The long-term development and physiological functions of the edited embryos is not presented. At least, it should be discussed in the discussion section.\nResponse to R2.8: We thank the reviewer for the feedback. Of the genes that we tested, successful editing is not expected to result in physiological changes up to the blastocyst stage13. Interestingly, one of the genes we edited in bovine embryos, rpl12, was found to be essential for embryonic development in another study that used prime editing14. Although our edits were not designed to result in a change in phenotype, the data presented in Figure 5e shows that the process of microinjection can result in the arrest of embryo development. Arrest post-microinjection is most commonly due to damage of the cellular and nuclear membrane, however PE and CODE may also have more direct effects on development that were not explored in this study.\n“Furthermore, PEMax and CODE systems had comparable effects on blastocyst development when targeting Hoxd13 and Dnmt1; however CODEMax treatment targeting Tubb5 resulted in more frequent arrested development than PEMax. As embryos were prepared for sequencing at the blastocyst stage to assess editing efficiency, long-term developmental or physiological effects were not characterized in this study.\n15"}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors have provided new data in their rebuttal comparing the activity of PE2, PEMax, CODEMax and CODEMax(exo+) with either pegRNAs or cpegRNAs. However, these data don’t appear to be included in any of the main text or supplementary figures. Did the authors intend to include these data in the revised manuscript, or were these data intentionally left out? As an aside, the data is not labeled clearly in the graphs—some conditions have no editing and as such no visible bars with colors to indicate whether a pegRNA or cpegRNA was used.\nFurthermore, for these experiments, it was not made clear whether the authors produced the all-RNA pegRNAs with the same ligation approach that was used for the production of cpegRNAs (as requested in my initial comments to the authors), or if full-length pegRNAs were produced by standard linear synthesis. Have the authors tried to generate pegRNAs with the same ligation approach that is used to produce cpegRNAs?\nAs synthesis quality and purity of pegRNAs is known to influence editing outcomes, and since smaller fragment syntheses for ligation substrates can be achieved with higher purities than for longer full-length guides, this is an important control to determine if the production method and/or related impurities influenced editing efficiencies. If they have not already done so, the authors should produce all-RNA pegRNAs by ligation for comparison to cpegRNAs and CODE editors. They are encouraged to test edits that have unexpectedly low or inconsistent PE2/PEMax efficiency, such as the DNMT1 +1 TCA insertion (this edit had notably higher editing with a pol-III transcribed pU6-pegRNA, as shown in Figure S6a).\nAfter another review of this manuscript, I have the following additional comments:\n1. In the last paragraph of the introduction, the sentence beginning “This simple yet versatile two-component system allows for delivery via plasmids, mRNA, or ribonucleoprotein (RNP) complexes…” is a bit misleading, as the cpegRNA cannot be expressed from a plasmid. The authors should revise to make this statement more accurate by saying that the editor protein can be expressed from a plasmid, while the cpegRNA must be delivered as RNA.\n2. In the first paragraph of the results section, for the sentence beginning “We also installed a premature termination codon (PTC) into the mCherry gene” cites ref 16, but that appears to be incorrect (or at least isn’t matched to the text).\n3. In the methods section for the HEK293T fluorescent reporter cell line production, it would be helpful to state the multiplicity of infection used for cell line generation and approximate copy number of integrated lentivirus reporters per cell.\n4. In both Figure 3g and Figure 4g, DNMT1 +1 TCA insertion appears to be tested with PE2 without a nicking sgRNA, yet in Figure 3g there is ~7% editing with PE2 and in Figure 4g there is essentially no editing without a nicking sgRNA. What explains this discrepancy? MECP2 +5 G>C is tested both Figure 3 and Figure 4 and looks more consistent. These experiments may require repeating to determine which result it more representative of outcomes.\n5. Please discuss the possibility for RNaseH-mediated degradation due to hybridization between the cpegRNA RNA spacer and DNA PBS/template sequence.\n6. The description of the methods for how the mRNA was produced is incomplete. Was the IVT template a digested plasmid, or a PCR product? Either further description or reference to another procedure is required.", "reviewer_2": "The concerns have been well addressed."}, "author_response": "Authors’ response to the reviewers: We appreciate the reviewers’ feedback on our manuscript titled “Efficient Genome Editing with Chimeric Oligonucleotide-Directed Editing”. We have addressed all the comments below with responses marked in blue and changes to the manuscript highlighted in yellow.\nREVIEWER COMMENTS\nReviewer #1 (R1, Remarks to the Author):\nR1 | The authors have provided new data in their rebuttal comparing the activity of PE2, PEMax, CODEMax and CODEMax(exo+) with either pegRNAs or cpegRNAs. However, these data don’t appear to be included in any of the main text or supplementary figures. Did the authors intend to include these data in the revised manuscript, or were these data intentionally left out? As an aside, the data is not labeled clearly in the graphs—some conditions have no editing and as such no visible bars with colors to indicate whether a pegRNA or cpegRNA was used.\nFurthermore, for these experiments, it was not made clear whether the authors produced the all-RNA pegRNAs with the same ligation approach that was used for the production of cpegRNAs (as requested in my initial comments to the authors), or if full-length pegRNAs were produced by standard linear synthesis. Have the authors tried to generate pegRNAs with the same ligation approach that is used to produce cpegRNAs?\nAs synthesis quality and purity of pegRNAs is known to influence editing outcomes, and since smaller fragment syntheses for ligation substrates can be achieved with higher purities than for longer full-length guides, this is an important control to determine if the production method and/or related impurities influenced editing efficiencies. If they have not already done so, the authors should produce all-RNA pegRNAs by ligation for comparison to cpegRNAs and CODE editors. They are encouraged to test edits that have unexpectedly low or inconsistent PE2/PEMax efficiency, such as the DNMT1 +1 TCA insertion (this edit had notably higher editing with a pol-III transcribed pU6-pegRNA, as shown in Figure S6a).\nResponse to R1 | We thank the reviewer for the additional feedback. We initially considered whether incorporating all additional data generated during the rebuttal process could potentially be confusing for readers. However, the reviewer raised a valid concern, and we agree that inclusion of the rebuttal data will be helpful to readers. Accordingly, we have added these data as new supplementary figures (Figs. S3 and S7).These figures compare the activity of PE2, PEMax, CODEMax and CODEMax(exo+) with either pegRNAs or cpegRNAs in our reporter cell line as well as at three endogenous genes. To aid in the visual interpretation of the graph, we have changed the formatting of the graphs.\nReviewer #1 has raised an excellent point regarding whether the pegRNAs and cpegRNAs were generated using the same strategy. To develop the ligation-based approach for cpegRNAs, we tested multiple ligases and optimized reaction conditions to establish an efficient DNA-RNA ligation strategy. While an alternative ligation workflow for pegRNAs could be implemented using different ligases and purification steps, the resulting pegRNAs would not be directly comparable due to differences in ligation efficiency, yield, and purity. Furthermore, the unreacted ssRNA vs. ssDNA components of the pegRNAs and cpegRNAs may have different effects after transfection to cells, so it would not be a fair comparison.\nThe data in Figure 3 compared PE2 using a synthetic pegRNA (ordered from IDT) and CODE2 using a ligated cpegRNA. To ensure a fair and robust comparison between pegRNAs and cpegRNAs, we used synthetic versions of both RNAs, each produced by IDT using the same synthesis method, in all figures except for Figure 3. We apologize for the confusion caused by not clearly specifying whether ligated or synthetic cpegRNA was used in each experiment. We included data using both versions to provide readers with flexibility, allowing them to either purchase cpegRNA commercially or generate it in-house. To prevent any confusion, we have now updated all relevant figures and figure legends to clearly state how the pegRNAs and cpegRNAs were obtained.\nR1 | After another review of this manuscript, I have the following additional comments:\nR1.1 | In the last paragraph of the introduction, the sentence beginning “This simple yet versatile two-component system allows for delivery via plasmids, mRNA, or ribonucleoprotein (RNP) complexes…” is a bit misleading, as the cpegRNA cannot be expressed from a plasmid. The authors should revise to make this statement more accurate by saying that the editor protein can be expressed from a plasmid, while the cpegRNA must be delivered as RNA.\nResponse to R1.1 | We appreciate the reviewer’s point. We made the following change to make it clear the various delivery formats were referring to the CODE fusion-protein:\n“This simple yet versatile two-component system allows for delivery of the CODE fusion-protein via plasmids, mRNA, or ribonucleoprotein (RNP) complexes for effective and accurate genome editing.”\nR1.2 | In the first paragraph of the results section, for the sentence beginning “We also installed a premature termination codon (PTC) into the mCherry gene” cites ref 16, but that appears to be incorrect (or at least isn’t matched to the text).\nResponse to R1.2 | We thank the reviewer for bringing this to our attention. The author that should be cited for this reference is Liu. The following change has been made:\n“We also installed a premature termination codon (PTC) into the mCherry gene so that the cells only display green fluorescence, similar to Liu et al16.”\nR1.3 | In the methods section for the HEK293T fluorescent reporter cell line production, it would be helpful to state the multiplicity of infection used for cell line generation and approximate copy number of integrated lentivirus reporters per cell.\nResponse to R1.3 | We thank the reviewer for this recommendation. We made the following change to the Methods:\n“Lentivral particles were titered using the qPCR Lentivirus Titer Kit (ABM, Cat# LV900). For lentiviral transduction, 5 x105 HEK293T cells were infected at an MOI of 5 with viral supernatant via reverse transduction. Briefly, viral supernatant was added to a 6-well plate first. The cells were then counted and resuspended in D10 medium supplemented with 10 μg/mL of TransduceIT™ Transduction Reagent (Mirus Bio, Cat# MIR6620) and transferred to the wells pre-added with viral supernatant. The cells were incubated for 72 hours before flow cytometry sorting and/or antibiotic selection.”\nR1.4 | In both Figure 3g and Figure 4g, DNMT1 +1 TCA insertion appears to be tested with PE2 without a nicking sgRNA, yet in Figure 3g there is ~7% editing with PE2 and in Figure 4g there is essentially no editing without a nicking sgRNA. What explains this discrepancy? MECP2 +5 G>C is tested both Figure 3 and Figure 4 and looks more consistent. These experiments may require repeating to determine which result it more representative of outcomes.\nResponse to R1.1 | We thank the reviewer for the attention to detail. As suggested, we have repeated the PE2 experiment at the DNMT1 site, and the data aligns more closely with the data seen in Figure 4g. Therefore, we have replaced the data in Figure 3g with the most recent, more consistent values.\nSome variation among data performed within different biological experiments is to be expected, because cell conditions can vary across the two collaborating universities (UF and Princeton). However, our new results show consistent results across experiments at both labs.\nR1.5 | Please discuss the possibility for RNaseH-mediated degradation due to hybridization between the cpegRNA RNA spacer and DNA PBS/template sequence.\nResponse to R1.5 | We thank the reviewer for this insight. There is a possibility that the intramolecular hybridization of the cpegRNA can occur before complexing with the CODE protein and trigger RNaseH-mediated degradation of the cpegRNA. Although we include chemical modifications on both the 5’ and 3’ ends of the cpegRNA to minimize exonuclease degradation, this does not eliminate the endonuclease activity of RNaseH. Therefore, we agree with the reviewer that this is something that should be mentioned as a possible limitation to the efficiency of CODE systems. The following sentence was added to the discussion:\n“The variable efficiencies observed across loci likely reflect mechanistic differences between reverse transcriptase– and DNA polymerase–driven priming. Because the optimal primer binding site and edit template lengths for DNA polymerases remain uncharacterized, we utilized PBS/template architectures previously optimized for PEMax or designed with DeepPrime, a tool trained exclusively on prime editing data, to enable a controlled comparison36. These parameters may not be optimal for CODE-based systems, and future studies aimed at tuning primer-template design and polymerase kinetics may further enhance editing efficiency and consistency across targets. Additionally, RNase H-mediated degradation of the cpegRNA in cells may contribute to variable CODE efficiency due to the hybridization of the RNA spacer with the DNA template and PBS.”\nR1.6 | The description of the methods for how the mRNA was produced is incomplete. Was the IVT template a digested plasmid, or a PCR product? Either further description or reference to another procedure is required.\nResponse to R1.6 | We appreciate the reviewer’s comment. We made the following change to the Material and Methods section:\n“H2b:GFP (provided by Dr. Martin Anger, Laboratory of Cell Division Control, IAPG CAS), PEMax, and CODEMax mRNA was prepared from linearized plasmid using the T3 mMessage mMachine Kit (Ambion).”"}, {"round": 3, "reviewer_comments": {"reviewer_1": "I thank the authors for their revisions to the manuscript. I appreciate that there would be the potential need for optimization of the method for making split-synthesis all-RNA pegRNAs to obtain representative results (though I suspect it should be possible with T4 RNA ligase I, the same enzyme used for preparing cpegRNAs). The authors have therefore adequately addressed all of my concerns, and I have no further comments."}, "author_response": ""}]

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