Efficient and error-free fluorescent gene tagging in human organoids without double-strand DNA cleavage.

CRISPR-associated nucleases are powerful tools for precise genome editing of model systems, including human organoids. Current methods describing fluorescent gene tagging in organoids rely on the generation of DNA double-strand breaks (DSBs) to stimulate homology-directed repair (HDR) or non-homologous end joining (NHEJ)-mediated integration of the desired knock-in. A major downside associated with DSB-mediated genome editing is the required clonal selection and expansion of candidate organoids to verify the genomic integrity of the targeted locus and to confirm the absence of off-target indels. By contrast, concurrent nicking of the genomic locus and targeting vector, known as in-trans paired nicking (ITPN), stimulates efficient HDR-mediated genome editing to generate large knock-ins without introducing DSBs. Here, we show that ITPN allows for fast, highly efficient, and indel-free fluorescent gene tagging in human normal and cancer organoids. Highlighting the ease and efficiency of ITPN, we generate triple fluorescent knock-in organoids where 3 genomic loci were simultaneously modified in a single round of targeting. In addition, we generated model systems with allele-specific readouts by differentially modifying maternal and paternal alleles in one step. ITPN using our palette of targeting vectors, publicly available from Addgene, is ideally suited for generating error-free heterozygous knock-ins in human organoids.

Introduction

Since the development of efficient genome editing technology, molecular and cell biological research increasingly relies on genetically modified in vitro model systems. In particular, the visualization of endogenous proteins using fluorescent knock-in reporters allows for a precise assessment of their subcellular localization and dynamics during cellular homeostasis and disease [1].

Organoids, in particular of human origin, represent next-generation model systems that recapitulate in vivo tissue architecture and functionality more accurately than 2D cell lines [2]. However, the precise engineering of large knock-in reporters in organoids can be laborious when using conventional CRISPR-mediated strategies to stimulate homology-directed repair (HDR) [3-5] or non-homologous end joining (NHEJ) [6] based editing. While generally effective, these strategies rely on the generation of genomic double-strand breaks (DSBs) by CRISPR-associated nucleases, which often result in both on- and off-target indel mutations as a consequence of error-prone repair by repeated cycles of NHEJ. On-target indels are often generated in the secondary “untargeted allele” that is not carrying the knock-in and may result in missense or nonsense mutations. In addition, while HDR generally results in error-free repair, generating knock-ins via NHEJ-based ligation of a linearized DNA fragment often results in indels within the up- and downstream junctions of the knock-in allele [6,7]. Consequently, existing knock-in protocols inherently require sequence verification of individually picked organoid clones, which is laborious, time consuming, and eliminates genetic heterogeneity in tumor-derived organoid models.

In cell lines, large knock-ins have been generated without introducing DSBs by using the partially inactivated Cas9 D10A nickase variant [8-10], which generates single-strand DNA breaks (nicks) in the genomic strand that hybridizes with the guide RNA [11]. By simultaneously nicking the genomic target locus and the extremities of both homology arms within the targeting vector, a strategy known as in-trans paired nicking (ITPN) [8], efficient knock-in alleles can be generated without double-strand DNA cleavage. Unlike conventional CRISPR/Cas9-mediated genome editing, ITPN modifies target loci with high fidelity, since single genomic nicks are rarely mutagenic [8,12]. By avoiding double-strand DNA cleavage, ITPN enables the insertion of heterozygous reporters or pathogenic (germline) mutations with intact “untargeted” secondary alleles and with minimal risk of off-target indels. Consequently, knock-in cells can be pooled to expedite the expansion and, thus, the generation time of a knock-in line (2 weeks). Pooling successfully targeted organoids is particularly useful for organoid models where clonal selection is laborious. Furthermore, by avoiding clonal selection, preexisting genetic diversity in tumor-derived organoid lines is largely preserved. Here, we investigate the efficiency and fidelity of fluorescent gene tagging via ITPN in human organoids. In addition, we present a palette of easy-to-use targeting vector backbones and protocols for N- or C-terminal fluorescent gene tagging using ITPN.

Results

To probe the efficiency of fluorescent knock-ins in human organoids (Fig 1A), we designed an N-terminal mScarlet knock-in at the human SEC61B locus. We constructed different targeting vectors in order to compare editing efficiencies of various knock-in strategies (Fig 1B). To stimulate editing via NHEJ-mediated ligation of a linearized mScarlet-coding fragment into the Cas9-generated genomic DSB [6,7], we constructed a vector carrying the mScarlet-coding sequence flanked by copies of the genomic Cas9 target site. Alternatively, we included 20 bp microhomology to stimulate genomic integration via the microhomology-mediated end joining (MMEJ) pathway [13]. In addition, we generated vectors with 1 kb homology arms following a traditional targeting vector design that is without flanking Cas9 target sites, or with flanking Cas9 target sites to support genomic integration via ITPN or in-trans paired cleavage (ITPC) [14,15].

Fig 1

Fluorescent gene tagging in human organoids without double-strand DNA cleavage.

(A) Schematic representation of the workflow used to capture fluorescent knock-in efficiencies in human organoids. To ensure optimal outgrowth post-electroporation, organoids are trypsinized to a cell suspension consisting of approximately 5 cell clumps. After electroporation, cells are allowed to expand for 10 days without selecting for cells that received the knock-in constructs. Prior to flow analysis, organoids may consist of partial knock-in populations. To capture the overall knock-in efficiency, organoid cultures are trypsinized to a single-cell suspension and flow analyzed. (B) Schematic representation of the SEC61B targeting strategy. mScarlet was flanked with homology arms matching up- and downstream sequences of the N-terminus of the human SEC61B locus, coding for Protein transport protein Sec61 subunit beta. Cas9 was targeted close to the start of the coding region using a gRNA as indicated (green arrow). Cas9 cleavage sites (triangles) and protospacer adjacent motifs (black bar) are indicated. Up- and downstream homology is represented in blue and red, respectively. Compositions of targeting vectors supporting different knock-in strategies are indicated. (C) Knock-in efficiencies of mScarlet at the human SEC61B locus in a patient-derived tumor organoid model using various knock-in strategies. WT or D10A nickase (Nick) SpCas9 was codelivered with targeting vectors indicated in (B). In all knock-in experiments, targeting vectors were electroporated at equimolar ratios between conditions to correct for differences in vector size. Editing efficiency (% mScarlet+ cells) was determined by single-cell flow analysis 10 days post-electroporation (n = 3 independent experiments). * p < 0.05 in a Ratio paired t test. Error bars indicate SEM. The inset shows representative stills of mScarlet-SEC61B localization in patient-derived tumor organoids (scale bar = 10 μm). (D) As in (C), knock-in efficiency of mScarlet at the human SEC61B locus in tumor organoids using targeting vectors with different homology arm lengths flanked by Cas9 target sites and codelivered with Cas9 D10A nickase to support ITPN (n = 3 independent experiments). Error bars indicate SEM. (E) As in (B), schematic showing the targeting strategy for ITPN-mediated integration of mScarlet (0.7 kb) or mScarlet-P2A-PuromycinR (1.4 kb) at the C-terminus of the human HIST1H2BC locus, coding for Histone H2B type-1C. (F) As in (C), knock-in efficiency of mScarlet (mSC; 0.7 kb) or mScarlet-P2A-PuromycinR (mSC-PR; 1.4 kb) in tumor organoids at the C-terminus of the human HIST1H2BC locus (n = 3 independent experiments). The difference between mSC and mSC-PR was nonsignificant in a Ratio paired t test. Error bars indicate SEM. (G) Knock-in efficiency of an mScarlet knock-in at the SEC61B locus in human colon normal and tumor organoids via ITPN using 1 kb homology arms (n = 3, n = 6 independent experiments for normal and tumor organoids, respectively). In all control conditions, the targeting vector was cotransfected with a guide targeting a different gene. The difference between normal and tumor KI organoids was nonsignificant in a two-sided unpaired t test. Error bars indicate SEM. Underlying data for panels C, D, F, and G are provided in S1 Data. Raw FCS files are available on the FlowRepository (FR-FCM-Z4PJ). ITPC, in-trans paired cleavage; ITPN, in-trans paired nicking; MMEJ, microhomology-mediated end joining; NHEJ, non-homologous end joining; WT, wild-type.

Targeting vectors were coelectroporated with wild-type or D10A nickase SpCas9 expression constructs in a patient-derived tumor organoid model obtained from a colorectal cancer biobank [16]. We visually confirmed the expected localization of mScarlet within knock-in organoids for each condition prior to flow analysis of mScarlet+ cells 10 days post-electroporation (Fig 1C). Flanking homology arms with Cas9 target sites to stimulate ITPN or ITPC resulted in substantially higher editing efficiencies when compared to a traditional targeting vector design with the same homology arm length (Figs 1C and S1A). In addition, NHEJ and MMEJ conditions underperformed when compared to targeting vectors with long homology arms, in particular when combined with nickase Cas9. Notably, ITPN resulted in a similar fraction of knock-in cells when compared to a traditional knock-in strategy that uses wild-type Cas9 and targeting vectors without flanking Cas9 target sites.

To investigate the fidelity of ITPN-mediated fluorescent knock-ins, we performed sequence analyses on polyclonal knock-in lines that were generated according to above-described conditions. To determine the risk for off-target indels, we analyzed the fidelity of the secondary allele that is not carrying the knock-in as a proxy for the likeliest candidates for off-target modifications. Using TIDE analysis [17], we show that wild-type Cas9 conditions result in a high frequency of indels within the secondary allele, whereas knock-in organoids generated via ITPN displayed >99% sequence integrity of their secondary allele (S2 Fig). Next, to investigate the fidelity of ITPN-mediated knock-ins, we generated 11 clonal knock-in lines from the ITPN condition and examined the knock-in alleles via Sanger sequencing. All knock-ins contained intact 5′ and 3′ junctions and no evidence for tandem integration was found (S3A Fig). Moreover, in agreement with previous TIDE analysis on polyclonal cultures, we confirmed the absence of indels in the untargeted allele of heterozygous clones (S3B Fig). Finally, to exclude the presence of off-target editing, we performed whole genome sequencing (WGS) on 3 out of the 11 clonal ITPN-mediated knock-in lines. We investigated the somatic mutation burden of these clones in 166 regions, which were predicted in silico to be likely off-target protospacer loci. No genomic aberrations were identified in the unmodified allele, the predicted off-target protospacer regions or the 200 bases surrounding the predicted sites (S1 Table). The lack of all variants ranging from single base substitutions to structural variation breakpoints confirms the absence of mutations due to incorrectly repaired off-target nuclease activity as well as off-target integrations of the knock-in cassette.

Collectively, these data indicate that ITPN enables highly efficient and indel-free fluorescent gene tagging in human organoids and makes sequence confirmation of clonal lines unnecessary. Consequently, all knock-in organoids can immediately be pooled to expedite the expansion of the edited organoid line and to maintain genetic diversity of patient-derived tumor organoid models.

Traditional design of targeting vectors requires long homology arms to maximize the chance of homologous recombination between the genomic locus and targeting vector. However, vectors with long homology arms are challenging to assemble and are inconvenient for locus-specific genotyping by PCR. To investigate whether efficiency of fluorescent gene tagging is lost when ITPN is mediated by shorter homology arms, we generated a series of targeting vectors with decreasing homology. At the SEC61B locus, the homology demand of ITPN-mediated mScarlet integration peaked at 800 bp (Figs 1D and S1B). While vectors with shorter homology arms were accompanied with lower editing efficiencies, they were sufficient to generate knock-in lines and may be preferred in situations of challenging vector assembly and/or genotyping.

Next, to probe whether knock-in size influences editing efficiency, we designed a C-terminal knock-in at the HIST1H2BC locus and constructed targeting vectors with 500 bp homology to integrate either mScarlet (0.7 kb) or mScarlet-P2A-Puro (1.4 kb) (Fig 1E). Surprisingly, we found no substantial difference in knock-in efficiency between the 2 targeting vector variants, suggesting that a knock-in size in the range of <1.4 kb has no notable influence on editing efficiency via ITPN (Figs 1F and S1C).

Since SEC61B and HIST1H2BC are ubiquitously expressed genes, we decided to investigate if we could knock-in mScarlet-P2A-Blast into normal human colon organoids at the C-terminus of KRT20, which is exclusively expressed in differentiated cells. Following a short pulse of Blasticidin selection, we observed clonal organoids with a subpopulation of cells showing the expected cytoplasmic red fluorescence (S4 Fig). Since differentiated cells do not form organoids as efficiently as stem cells, lines that involve fluorescent knock-ins in differentiation genes such as KRT20 are best generated either using a short pulse of selection or by manually picking and pooling clonal organoids that contain (some) fluorescent cells.

Finally, we compared the efficiency of an N-terminal mScarlet knock-in at the SEC61B locus between tumor and normal colon organoids (Figs 1G and S1D). The knock-in efficiency in tumor organoids was higher (although not significant), which may be attributed to a difference in culture conditions and electroporation efficiency.

A major downside of generating targeting vectors with homology arms flanked by Cas9 target sites at their extremities is the time-intensive molecular cloning. To expedite the cloning of targeting vectors for fluorescent gene tagging at either the N- or C-terminus, we generated a series of minimalistic targeting vector backbones allowing seamless one-step integration of both homology arms using SapI-based Golden Gate assembly [4] (Fig 2A). Targeting vector backbones carrying state-of-the-art monomeric fluorescent proteins are made available from Addgene, including optional P2A-linked selection elements (Fig 2B). Using our optimized vector backbones, targeting vectors can be assembled in the same amount of cloning time as is required for the insertion of gRNA oligos into Cas9 expression vectors. Consequently, when using our vector backbones for ITPN, fluorescent reporter alleles in cell lines and organoid models can be generated in as little as 2 weeks, including molecular cloning procedures for vector assembly (Fig 2C). We summarized our recommendations for knock-in design and one-step targeting vector assembly in a protocol (S1 File). In addition, new variants of targeting vector backbones, e.g., replacing the donor with a different fluorescent protein sequence, can be generated in a short amount of time (S2 File).

Fig 2

One-step targeting vector assembly and ITPN expedite fluorescent gene tagging.

(A) Schematic outline of one-step TV assembly via SapI-based golden gate-mediated homology arm ligation. Homology arms can be amplified from genomic DNA or ordered as commercially synthesized DNA fragments. (B) Overview of knock-in backbone constructs available from Addgene. Knock-in backbones contain one of 4 different fluorescent proteins and optional P2A-linked resistance cassettes. Backbone constructs are suitable for knock-ins at either the C- or N-terminus, as indicated. (C) Schematic workflow outlining fluorescent gene tagging in organoids using ITPN. Following electroporation, organoids generally require approximately 10 days of outgrowth before FACS purification of fluorescent knock-in cells. Alternatively, fluorescent clonal organoids can be handpicked and pooled. Sequence verification of individually picked clonal lines is not required when editing via ITPN. ITPN, in-trans paired nicking; TV, targeting vector.

To probe the efficiency of ITPN using our newly designed targeting vectors, we generated triple fluorescent knock-ins by simultaneous targeting of 3 separate genomic loci. Specifically, we targeted the C-terminus of the HIST1H2BC locus to knock-in mTurquoise2-P2A-puromycinR, the C-terminus of the CDH1 locus to knock-in mScarlet, and as a third locus, we included an N-terminal knock-in of mNeongreen at either the LMNA, SEC61B, or MAP4 locus (Fig 3A). DNA cocktails containing different combinations of targeting vectors and their respective Cas9 expression constructs were electroporated into fractionated tumor organoids. Organoids were allowed to form for 10 days without puromycin selection prior to quantification of the raw knock-in efficiencies by single-cell flow analysis. As expected, in all 3 conditions, the knock-in fractions were dominated by cells that carried single knock-ins in either one of the targeted genes. However, we readily detected cells carrying multiple knock-ins, including cells where all 3 genes were edited simultaneously (Fig 3B). The overall knock-in efficiencies for each targeted gene and the fraction of cells carrying multiple knock-ins are summarized in Fig 3C. To confirm the fidelity of the gene fusions, we generated polyclonal triple knock-in lines from each editing condition by manual picking and pooling clonal triple positive organoids. TIDE analysis again confirmed the absence of on-target indels in the untargeted alleles of all edited genes (S5 Fig). In addition, we confirmed the intended integration of the knock-in via Sanger sequencing (S6 Fig). Next, we recorded overnight growth of our TKI-3 knock-in line using live-cell imaging to demonstrate normal growth behavior and phenotype (Fig 3D). Each channel could be recorded without excessive bleaching, allowing a multidimensional dynamic readout of chromosomal instability during mitosis, including chromatin errors (H2B1C), spindle assembly (MAP4), and membrane defects or binucleation (CDH1).

Fig 3

Multiplexed fluorescent gene tagging in human organoids using ITPN.

(A) Multiplexed fluorescent gene tagging in tumor human colon organoids at 3 different genomic loci using ITPN. C-terminal integrations of mScarlet at the CDH1 locus and mTurquoise2-P2A-Puromycin into the HIST1H2BC locus were combined with N-terminal integration of mNeongreen at either the LMNA, SEC61B, or MAP4 locus. In the schematics: Cas9 D10A nick positions (red triangles) and protospacer adjacent motifs (black bars) are indicated for each knock-in design, as well as the gRNA used (green arrow). Organoids were electroporated simultaneously with all 3 targeting vectors to generate one-step multiplexed triple knock-ins. (B) All 3 targeting combinations yielded triple knock-in populations with practical efficiencies, as indicated by flow analysis (numbers indicate frequencies (%) of knock-in cells within the entire targeted cell population). Imaging snapshots show the expected subcellular localization of each fusion protein (scale bar = 10 μm). Raw FCS files are available on the FlowRepository (FR-FCM-Z4PJ). (C) Overview of the multiplexed gene tagging efficiencies as determined by flow cytometry analysis. Raw FCS files are available on the FlowRepository (FR-FCM-Z4PJ). (D) Live-cell imaging of tumor human colon organoids carrying CDH1-mScarlet, HIST1H2BC-mTurquoise2, and mNeongreen-MAP4 knock-ins. The top panel shows representative stills of organoid growth over time (scale bar = 10 μm). For divisions I and II, snapshots of each channel are shown in metaphase and anaphase (scale bar = 5 μm). ITPN, in-trans paired nicking.

Taken together, these results demonstrate that ITPN maintains high levels of fidelity across different genomic loci and allows multiplexed fluorescent gene tagging in human organoids. Using conventional editing protocols, generating organoid lines carrying multiple fluorescent knock-ins is highly laborious. By using ITPN, organoids with multiple edits can be generated within 2 weeks. Alternatively, in case an attempt to multiplex gene targeting fails, cells with a single knock-in can be pooled and retargeted. Moreover, we generated the same combinations of triple knock-ins in 2 rounds of targeting and used intermediate antibiotic selection to enrich for knock-in cells instead of manual picking (S7 Fig).

Since the sequence integrity of the untargeted allele that is not carrying the knock-in is maintained when editing via ITPN, this secondary allele can be retargeted using the same locus-specific targeting vector to obtain homozygous knock-ins. This also enables straightforward differential modification of maternal and paternal alleles by offering 2 different targeting vectors for the same locus. To investigate if ITPN allows the simultaneous generation of biallelic knock-ins carrying different fluorescent tags within each allele, we targeted the SEC61B, MAP4, and HIST1H2BC loci in tumor organoids with both mNeongreen and mScarlet targeting vectors. Flow analysis at 10 days post-electroporation confirmed the presence of a double-positive cell population for each targeted locus (Fig 4A). Genotyping of manually picked lines confirmed correct modification of each allele (S8 Fig). In addition, imaging of biallelic knock-in organoids confirmed the detection of both allele-specific reporters (Fig 4B). Next, we performed live-cell imaging of our HIST1H2BC double knock-in organoids and assessed the biallelic fluorescent output (green versus red) for single cells over time (Fig 4C), as a straightforward showcase how differential allele-specific modifications could be used to study allele-specific expression levels [18,19]. This proof of principle underscores the power of ITPN to create allele-specific readouts that, depending on the knock-in template, can be applied to address many biological questions, ranging from allele-specific expression patterns to differential biochemical properties between wild-type and mutant proteins.

Fig 4

One-step differential fluorescent knock-ins at a single locus.

(A) The SEC61B, MAP4, and HIST1H2BC loci of a patient-derived tumor organoid model were targeted via ITPN with both mNeongreen and mScarlet targeting vectors according to the design shown in Fig 3A. Flow analysis was performed 10 days post-electroporation. The percentage of cells carrying a single knock-in and cells carrying both knock-ins are indicated. Raw FCS files are available on the FlowRepository (FR-FCM-Z4PJ). (B) Representative stills of tumor organoids carrying biallelic mNeongreen and mScarlet modifications at either the SEC61B, MAP4, and HIST1H2BC loci (scale bar = 10 μm). (C) Allele-specific readout in human colorectal cancer organoids containing differentially tagged HIST1H2BC alleles. Organoids carrying mNeongreen and mScarlet knock-ins at the HIST1H2BC locus were live-cell imaged for 18 hours. Top panels show imaging stills of the green/red composite over time. The left bottom graph shows the changes of allele-specific output over time for a single representative cell. The corrected fluorescent signals of mScarlet, mNeongreen, as well as the ratio mScarlet/mNeongreen are plotted. The right bottom graph shows 7 out of 17 analyzed single cell ratios (mScarlet/mNeongreen). Underlying data are provided in S1 Data. ITPN, in-trans paired nicking.

Discussion

Here, we show that large knock-ins such as fluorescent gene tags can be generated in human organoids with high efficiency and fidelity using ITPN. The strategy is superior to conventional Cas9-mediated genome editing as ITPN is DSB independent, which minimizes the risk of undesired mutations, both at potential off-target sites as well as within the untargeted allele that does not carry the knock-in. This also implies that ITPN is ideally suited for the introduction of heterozygous pathogenic (germline) mutations. An additional advantage is that sequence verification of individually picked clonal lines becomes obsolete and thus that all de novo generated knock-in cells can immediately be pooled. This accelerates expansion of the early culture and reduces the overall generation time of knock-in models. In addition, pooling all de novo generated knock-in cells preserves the genetic heterogeneity present in the original culture, which is important when working with patient-derived tumor organoid models. To circumvent labor-intensive molecular cloning of targeting vectors, we generated a palette of vector backbones that can be locus customized in the same amount of cloning time required for gRNA oligo insertion into Cas9 expression vectors. Moreover, our targeting vector backbone design is modular, so that the donor template itself can easily be adapted to more complex and sophisticated reporter designs [5]. Our vector backbones are available on Addgene, and our protocols (S1 and S2 Files) contain detailed instructions for their application.

While absolute knock-in efficiency depends on many variables, the most practical determinant of a successful strategy is that knock-ins are consistently obtained with each attempt. To showcase the robust efficiency of ITPN in organoids, we generated triple knock-ins at 3 independent genomic loci in a single targeting round, and we accomplished a one-step generation of biallelic knock-ins carrying allele-specific reporters. In our hands, efficiency of ITPN is superior to CRISPR strategies that use conventional donor templates. In agreement with earlier reports [8,9,14,15], an important variable that influences overall efficiency was the ITPN or ITPC of the donor. Although NHEJ-mediated knock-ins by strategies such as CRISPR-HOT [6] minimize the need for molecular cloning, our results indicate that ITPN substantially improves efficiency and fidelity when compared to NHEJ-mediated strategies, while our dedicated vector backbones minimize molecular cloning to a similar extent.

Due to the near exact sequence similarity between maternal and paternal alleles, differences in expression patterns between 2 alleles of a given gene are poorly understood. Likewise, changes in subcellular localization, protein interactions, and/or biochemical properties between wild-type and mutant proteins, such as oncogenes, are rarely examined in the same cells due to the near impossible task of modifying both alleles independently with different tags. Since ITPN generates knock-in alleles without modifying the untargeted allele, the same locus can be retargeted with an alternative donor template. As a proof of principle, we generated a biallelic knock-in with different fluorophores at the HIST1H2BC, SEC61B, and MAP4 loci in a single targeting round. Accurate estimation of allelic imbalance is important to understand genetic and epigenetic mechanisms of gene regulation, and dysregulation during carcinogenesis.

In conclusion, ITPN is a versatile strategy that generates fast and efficient knock-ins in human organoids. We envision that our approach can easily be applied to organoid models derived from other tissues or sources, such as pluripotent stem cell-derived organoid models. Various CRISPR-mediated knock-in strategies have been reported to date that reach sufficient efficiencies to make genetic editing practical in organoid models. ITPN is comparable in terms of efficiency but stands out as being DSB independent and therefore has the highest intrinsic fidelity of precise genome editing. In combination with the seamless one-step generation of targeting vectors, ITPN represents an important technological advance in generating high-fidelity knock-in model systems.

Materials and methods

Vector assembly

Targeting vector backbones were generated by recombinase-based seamless assembly (In-Fusion cloning, Takara Biotech) of a commercially synthesized DNA fragment (IDT or Genscript) carrying mNeongreen [20], mTurquiose2 [21], mScarlet [22], or photoconvertible Dendra2 [23] with optional P2A-selection elements (S2 Table) into a PCR-amplified generic backbone fragment (FWD: tcctcgctcactgactcgct, REV: gcggtattttctccttacgcatctg). See S2 File for a detailed explanation of how to generate new targeting vector backbones. To generate locus-specific targeting vectors, commercially synthesized homology arm fragments (S3 Table) were inserted into targeting vector backbones using SapI-based golden gate assembly as previously described [4]; see S1 File for a more detailed protocol. Cas9 wild-type (addgene #48139) and Cas9 D10A nickase (addgene #48141) locus-specific expression vectors were generated according to published protocols [24].

Organoid culture

Patient-derived tumor organoid with identifier P9T (PDTO-9) was obtained from a previously published colorectal cancer biobank [16]. PDTO-9 was maintained at 37°C with 5% CO2 atmosphere seeded in RGF Basement Membrane Extract (BME), Type 2 (Cultrex). Culture media consisted of advanced DMEM/F12 (Gibco) supplemented with penicillin–streptomycin (Lonza, 10 U ml−1), GlutaMAX (Gibco, 1x), HEPES buffer (Gibco, 10 mM), Noggin-conditioned medium (10%), R-spondin1-conditioned medium (10%), B-27 (Gibco, 1x), nicotinamide (Sigma-Aldrich, 10 mM), N-acetylcysteine (Sigma-Aldrich, 1.25 mM), SB202190 (Gentaur, 10 μM), A83-01 (Tocris, 500 nM), and recombinant human EGF (PeproTech, 50 ng ml−1). PDTO-9 cultures were passaged weekly and maintained below passage 10. Briefly, PDTOs were dissociated using trypsin–EDTA (Sigma-Aldrich) and seeded in BME in a prewarmed 24-well plate. ROCK inhibitor Y-27632 (Gentaur, 10 μM) was added to culture medium upon plating for 2 days.

Normal human colon organoids were maintained at 37°C with 5% CO2 atmosphere seeded in growth factor reduced Matrigel (BD Biosciences). Culture media [25] consisted of advanced DMEM/F12 (Gibco) supplemented with penicillin–streptomycin (Lonza, 10 U ml−1), GlutaMAX (Gibco, 1x), HEPES buffer (Gibco, 10 mM), Noggin-conditioned medium (10%), R-spondin1-conditioned medium (20%), B-27 (Gibco, 1x), N-acetylcysteine (Sigma-Aldrich, 1.25 mM), A83-01 (Tocris, 500 nM), recombinant human EGF (PeproTech, 50 ng ml−1), recombinant human IGF-1 (BioLegend, 100 ng/ml), recombinant human FGF-basic (Peprotech, FGF-2 50 ng/ml), and 0.5 nM Wnt surrogate (U-protein Express). Organoid were passaged as described above.

Organoid electroporation

To generate knock-ins in either normal or colorectal tumor organoids, 1 × 106 cells at approximately 5 cell clumps were coelectroporated with 15 μg DNA, at a 1:1 ratio of Cas9 (Addgene #48139) or Cas9 D10A nickase (Addgene #48141) and targeting vector using the NEPA21 Super Electroporator (Nepagene) following described conditions [26].

Flow analysis

Organoids were trypsinized and filtered through a CellTrics 10 μm sieve (Sysmex) to obtain a single-cell suspension. To quantify the knock-in efficiency, cells were flow analyzed (FACSCelesta, BD) at least 10 days post-electroporation. Gates were set based on a negative control/population.

Genotyping and TIDE analysis

Polyclonal or clonal knock-in cultures were established via manual picking or FACS. Site-specific integrations were confirmed by genotyping PCRs on genomic DNA extract using locus-specific primer sets (S4 Table), followed by Sanger sequencing. TIDE analysis was performed on Sanger sequencing data of secondary “non-knock-in” alleles using the Sanger sequencing read of the parental organoid line as a control sample chromatogram.

Whole genome sequencing and read mapping

The genomic DNA of 3 clonally expanded ITPN-mediated SEC61B-mScarlet organoid lines was isolated using the QIAamp DNA Micro Kit according to the manufacturer’s instructions. Illumina sequencing libraries were generated using 200 ng of genomic DNA according to standard protocols (Illumina). Following WGS to a base coverage of 15x (Illumina NovaSeq 6000, 2 × 150 bp), initial processing of the sequence reads was performed using the complete analysis pipeline available at https://github.com/UMCUGenetics/NF-IAP. Briefly, the Burrows-Wheeler Aligner v0.7.17 mapping tool was used to map sequence reads against human reference genome GRCh38 with settings “bwa mem -c 100 –M” [27]. Next, duplicate reads were flagged with Sambamba v0.6.8 and the Genome Analysis Toolkit (GATK) v4.1.3.0 was used for realignment [28].

Variant calling and filtering

Next, variants were multisample called with the GATK HaplotypeCaller v4.1.3.0 and GATK-Queue v.4.1.3.0, based on default settings and the additional option “EMIT_ALL_CONFIDENT_SITES.” Subsequently, GATK VariantFiltration v4.1.3.0 was used to evaluate the quality of the variant positions, with options -snpFilterName SNP_LowQualityDepth -snpFilterExpression “QD < 2.0” -snpFilterName SNP_MappingQuality -snpFilterExpression “MQ < 40.0” -snpFilterName SNP_StrandBias -snpFilterExpression “FS > 60.0” -snpFilterName SNP_HaplotypeScoreHigh -snpFilterExpression “HaplotypeScore > 13.0” -snpFilterName SNP_MQRankSumLow -snpFilterExpression “MQRankSum < −12.5” -snpFilterName SNP_ReadPosRankSumLow -snpFilterExpression “ReadPosRankSum < −8.0” -snpFilterName SNP_HardToValidate -snpFilterExpression “MQ0 > = 4 && ((MQ0 / (1.0 * DP)) > 0.1)” -snpFilterName SNP_LowCoverage -snpFilterExpression “DP < 5” -snpFilterName SNP_VeryLowQual -snpFilterExpression “QUAL < 30” -snpFilterName SNP_LowQual -snpFilterExpression “QUAL > = 30.0 && QUAL < 50.0” -snpFilterName SNP_SOR -snpFilterExpression “SOR > 4.0” -cluster 3 -window 10 -indelType INDEL -indelType MIXED -indelFilterName INDEL_LowQualityDepth -indelFilterExpression “QD < 2.0” -indelFilterName INDEL_StrandBias -indelFilterExpression “FS > 200.0” -indelFilterName INDEL_ReadPosRankSumLow -indelFilterExpression “ReadPosRankSum < −20.0” -indelFilterName INDEL_HardToValidate -indelFilterExpression “MQ0 > = 4 && ((MQ0 / (1.0 * DP)) > 0.1)” -indelFilterName INDEL_LowCoverage -indelFilterExpression “DP < 5” -indelFilterName INDEL_VeryLowQual -indelFilterExpression “QUAL < 30.0” -indelFilterName INDEL_LowQual -indelFilterExpression “QUAL > = 30.0 && QUAL < 50.0” -indelFilterName INDEL_SOR -indelFilterExpression “SOR > 10.0.”

Low-quality and subclonal mutations accumulated during clonal expansion of the organoid lines were excluded by annotating using SMuRF release 2.1.2 as described [29] (https://github.com/ToolsVanBox/SMuRF). We included all variants in each clone at autosomal or X chromosomes present in less than 3 clonal samples that passed VariantFiltration, with a GATK phred-scaled quality score ≥60; minimum base coverage of 5X, a mapping quality ≥30, and a variant allele frequency of at least 0.15 [29,30]. Structural variation calling was performed with the GRIDSS-purple-linx pipeline v1.3.2, using all paired combinations of the 3 WGS samples as tumor-normal pairs [31].

Analysis of in silico predicted off-target regions

All potential off-target protospacer regions for the guide sequence 5′-GGGGTCGGACCAGGCTGTAG-3′ were predicted using the publicly available CasOFFinder tool [32], using an NGG PAM and allowing up to 4 mismatches. As regions of interest, both the potential off-target protospacer regions as well as the flanking 200 bases were considered. Using BEDtools v2.27.1, all variants that passed filtering by SMuRF were intersected with the regions of interest [33]. In addition, all start and end coordinates of the structural variations called by GRIDSS-purple-linx were intersected with the same potential off-target genomic regions.

Live organoid imaging

To support live-cell microscopy of organoids, PDTO-9 or normal organoid cultures were passaged 5 to 7 days prior to imaging. PDTOs were harvested 24 hours before imaging and resuspended in an ice-cold mix of culture media containing 50% v/v BME or 90% v/v Matrigel. The organoid suspension was then seeded in an ice-cold glass bottom WillCo-dish (WillCo Wells B.V.) coated with a thin film of BME or Matrigel. Organoids were allowed to settle on ice before gel polymerization at 37°C and addition of culture media. Outgrowth was captured overnight on a spinning disk confocal system (Nikon, 15-minute frame rate, z-step 1.4 μm). Imaging data were analyzed with Fiji (ImageJ). For the HIST1H2BC biallelic knock-in organoids, a custom-made analysis macro [34] was used to track single cells and monitor their mNeongreen and mScarlet signals. Bleach correction (per channel, per time point) was performed based on integral fluorescence signals of the corresponding organoids.

Flow cytometry analyses for the measurement of knock-in efficiencies.

Representative flow cytometry plots for the analysis of knock-in efficiencies including nonedited controls are shown. Panel (A), (B), (C), and (D) are related to Fig 1C, 1D, 1F, and 1G, respectively. Raw FCS files are available on the FlowRepository (FR-FCM-Z4PJ). ITPC, in-trans paired cleavage; ITPN, in-trans paired nicking; MMEJ, microhomology-mediated end joining; NHEJ, non-homologous end joining; wt, wild-type.

(EPS)

Click here for additional data file.

On-target TIDE analysis of different knock-in strategies at the SEC61B locus.

To perform TIDE analyses, genomic DNA was extracted from polyclonal knock-in populations generated either via MMEJ, traditional targeting, ITPC (with wild-type Cas9), or ITPN (with Cas9 D10A nickase) (see Fig 1C). Polyclonal lines were generated by manually pooling approximately 10 clonal knock-in organoids. TIDE analysis was performed using the Sanger sequencing read of the parental patient-derived tumor organoid model as a control sample chromatogram. The NHEJ condition codelivered with wild-type Cas9 was omitted from this analysis as there were too few knock-in organoids available to generate a polyclonal line. The percentage of reads containing indels is displayed in the left upper corners. Underlying data are provided in S2 Data. ITPC, in-trans paired cleavage; ITPN, in-trans paired nicking; MMEJ, microhomology-mediated end joining; NHEJ, nonhomologous end joining; WT, wild-type.

(EPS)

Click here for additional data file.

Fidelity of ITPN-mediated fluorescent knock-ins at the SEC61B locus.

Sanger sequencing was performed on the SEC61B locus of 11 clonal patient-derived tumor organoid knock-in lines generated via ITPN (see Fig 1C). Knock-in lines were generated by handpicking individual large clonal knock-in organoids. The 5′ and 3′ junctions of the knock-in allele and the target region of the “untargeted” allele is shown for each clone. Clone no.7 carries a homozygous mScarlet knock-in at the SEC61B locus and therefore does not contain an untargeted allele. Underlying data are provided in S2 Data. ITPN, in-trans paired nicking.

(TIF)

Click here for additional data file.

KRT20-mScarlet knock-in in normal human colon organoids.

(A) Schematic representation of the KRT20 targeting strategy to knock in mScarlet-P2A-BlasticidinR. The donor was flanked with homology arms matching up- and downstream sequences of the C-terminus of the human KRT20 locus. Cas9 D10A nickase was targeted close to the stop codon using a gRNA as indicated (green arrow). Cas9 D10A nickase cleavage sites (triangles) and protospacer adjacent motifs (black bar) are indicated. Up- and downstream homology is represented in blue and red, respectively. Knock-in organoids were generated by electroporating normal human colon organoid clumps followed by outgrowth for 10 days. To select for successfully targeted cells, organoids were treated for 7 days with Blasticidin, followed by manual picking of a clonal organoid containing red fluorescent cells. (B) Fluorescent images of representative KRT20-mScarlet-P2A-BlasticidinR knock-in organoids containing both KRT20-positive and KRT20-negative cells (red: KRT20-mScarlet). Scale bar = 50 μm. (C) Sanger sequencing was performed on the KRT20-mScarlet-P2A-BlasticidinR knock-in line. The 5′ and 3′ junctions of the knock-in allele are intact. Underlying data are provided in S2 Data.

(EPS)

Click here for additional data file.

TIDE analysis of multiplexed triple knock-in lines in tumor human colon organoids.

TIDE analysis of the “untargeted” alleles for all targeted genes in the one-step triple knock-in lines (see Fig 3). TIDE was performed using the Sanger sequencing read of the parental tumor colon organoid model as a control sample chromatogram. The SEC61B allele of Triple KI-2 was not analyzed because it failed quality control. Underlying data are provided in S2 Data.

(EPS)

Click here for additional data file.

Sanger sequencing of 5′ and 3′ knock-in allele junctions in triple knock-in tumor colon organoids.

Sanger sequencing was performed on polyclonal triple knock-in tumor colon lines generated via manual picking of triple positive clonal organoids. The 5′ and 3′ junctions of each knock-in allele are shown for all 3 multiplexed knock-in conditions (see Fig 3). Underlying data are provided in S2 Data.

(TIF)

Click here for additional data file.

Two-step triple knock-ins in human colon cancer organoids using antibiotic enrichment.

(A) Schematic representation showing the generation of triple fluorescent knock-ins in human colon cancer organoids in 2 rounds of gene targeting. mScarlet-BlastR was integrated into the CDH1 locus, coding for E-cadherin, followed by Blasticidin selection for 2 weeks to enrich for knock-in organoids. Subsequent handpicking generated a pure line (99.5%, Scarlet+). In a second round of gene targeting, mTurquoise2-PuroR was integrated into the HIST1H2BC locus combined with mNeongreen integrations into either the LMNA, SEC61B, or MAP4 loci. In the schematics: Cas9 (D10A) nick positions (red triangles) and protospacer adjacent motifs (black bar) are indicated for each knock-in design, including gRNA used (green arrow). (B) FACS plots of triple knock-in organoids after the second targeting round and puromycin selection for 10 days. Puromycin enrichment resulted in a near complete selection for HIST1H2B-mTQ2-PuroR positive cells (purple and blue populations combined). Numbers in FACS plots indicate frequencies (%) of knock-in populations within the entire targeted cell population. Raw FCS files are available on the FlowRepository (FR-FCM-Z4PJ).

(EPS)

Click here for additional data file.

Sanger sequencing of differential biallelic modifications in a patient-derived tumor organoid line.

Sanger sequencing was performed on polyclonal biallelic tumor knock-in lines carrying both mScarlet and mNeongreen integrations at either the SEC61B, MAP4, or HIST1H2BC locus. The 5′ and 3′ junctions of each knock-in allele are shown for all 3 biallelic knock-in conditions (see Fig 4A). For HIST1H2BC-mSC, the endogenous stop codon of the HIST1H2BC locus was maintained.

(TIF)

Click here for additional data file.

Presence of off-target effects (small indels and structural variations breakpoints) in in silico predicted off-target regions (spacer and flanking regions) in WGS of 3 clonal ITPN-mediated SEC61B-mScarlet knock-in organoid lines.

ITPN, in-trans paired nicking; WGS, whole genome sequencing.

(XLSX)

Click here for additional data file.

Targeting vector backbone fragments.

(XLSX)

Click here for additional data file.

Targeting vector homology arm sequences.

(XLSX)

Click here for additional data file.

Locus-specific primer sets for genotyping and TIDE.

(XLSX)

Click here for additional data file.

Data underlying Figs 1C, 1D, 1F, 1G, and 4C.

(A-D) Knock-in efficiencies (% of total cell population) as determined by flow cytometry analysis for Fig 1C, 1D, 1F and 1G. (E) Fluorescent signals reported in Fig 4C. Values are corrected as explained in the methods.

(XLSX)

Click here for additional data file.

Raw Sanger and TIDE sequencing results for S2–S6 and S8 Figs.

(ZIP)

Click here for additional data file.

Design considerations and cloning protocol for the one-step targeting vector assembly to obtain fluorescent gene tagging using in-trans paired nicking or cleavage.

(DOCX)

Click here for additional data file.

Personalize your one-step targeting vector backbone by replacing the current donor templates.

(DOCX)

Click here for additional data file.

18 May 2021

Dear Dr Snippert,

Thank you for submitting your manuscript entitled "Efficient fluorescent gene tagging in human organoids without double-strand DNA cleavage" for consideration as a Methods and Resources article by PLOS Biology. Please accept my apologies for the delay in getting back to you as we consulted with an academic editor about your submission.

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17 Jun 2021

Dear Dr Snippert,

Thank you very much for submitting your manuscript "Efficient fluorescent gene tagging in human organoids without double-strand DNA cleavage" for consideration as a Methods and Resources paper at PLOS Biology. I'm handling your paper temporarily while my colleague Richard Hodge is out of the office. Your manuscript has been evaluated by the PLOS Biology editors, an Academic Editor with relevant expertise, and by three independent reviewers.

The reviews are attached below. You'll see that the reviewers find your manuscript interesting and of potential use to the field, but note that the methodology is not yet fully validated. Specifically, Reviewer #1 and #2 raise overlapping concerns with the characterization of the targeting efficiency and ask that this is calculated in the primary organoids. Reviewer #2 also asks that additional biological replicates are provided for the colonic organoids. In addition, Reviewer #3 asks for additional sequencing validation data to address potential on- and off-target mutations that could occur using the strategy. Please address all of the reviewer's concerns.

In light of the reviews (below), we will not be able to accept the current version of the manuscript, but we would welcome re-submission of a much-revised version that takes into account the reviewers' comments. We cannot make any decision about publication until we have seen the revised manuscript and your response to the reviewers' comments. Your revised manuscript is also likely to be sent for further evaluation by the reviewers.

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*****************************************************

REVIEWERS' COMMENTS:

Reviewer #1:

In this manuscript, Bollen et al reported a double-strand DNA-free system to allow seamless one-step targeting vector assembly to support both N- and C-terminal fluorescent knock-ins. Three genomic loci were simultaneously modified in a single round of targeting in colon cancer organoids. Similar approaches were applied on healthy colon organoids. This can be a potential very useful tool for organoid field. However, some concerns need to be addressed before considered for publication.

Major concerns:

1. The targeting efficiency needs to be better characterized. Based on Figure 3b. All cells in one organoid express the knock-in proteins. Is this true for all organoids? If yes, what is the percentage of organoids that are triple positive, what is the percentage of organoids that express two fluorescent proteins, what is the percentage of organoids that express mScarlet, mTurquoise2, or Neongreen?

2. Is there any difference of targeting efficiency between normal and cancer organoids? The targeting efficiency should be quantified for both normal and cancer organoids as Question 1.

3. Figure S2C, sequencing of isolated clones should be performed to validate the heterozygous KI .

Minor concerns:

1. Scale bars are missing for most figure panels, which should be added and described in the legends.

2. Gating control should be added for all flow cytometry analysis.

3. Statistical analysis is missing in Figure 1e.

Reviewer #2:

[identifies herself as Joo-Hyeon Lee]

Summary

Efficient CRISPR based gene targeting and generation of knock-ins have been challenging for 3D organoids derived from primary tissue. This manuscript describes a method to generate fluorescent reporter organoid lines by introducing homology directed repair (HDR) using Cas9 D10A nickase, which instead of creating a double stranded break (DSB) nicks only the allele targeted by the gRNA. The protocol would be beneficial to the field for efficient generation of human organoid reporter lines enabling the long-term tracing of cells. The authors showed the way to knock-in fluorescent proteins into multiple locus of the same cells, and further generate homologues lines by sequential targeting of both gene alleles. The classical antibiotics-based selection strategy could also be used in conjunction with florescence. These various plasmids used for fluorescent knock-in are also available with different fluorescent proteins to the community, proving that this protocol can be broadly useful.

General Comments

Originality and significance: The strategy of in-trans paired nicking using modified Cas9 nicking variant has been reported in 2D culture systems such as HEK293T cells, as cited in the manuscript (Chen et al., 2017). This protocol adapted the in-trans paired nicking strategy to generate florescence-based selection in primary tissue-derived organoids, which can also be combined with antibiotics-based selection, allowing the efficient genetic engineering of primary human organoid culture models.

Data and methodology: A major concern in this manuscript is that there is no description of biological replicates and validations in Fig. 3 and 4. For this system to be widely applicable, the authors should test multiple 'colonic' organoids and provide the detailed information of variation and efficiency of this strategy across biological repeats.

Conclusions: The authors used HEK293T cells to calculate the editing efficiency of their strategy in Fig. 1. Given that this manuscript focuses on 3D human primary organoids, the editing efficiency of in-trans paired/cleavage method should be provided in human primary organoid culture systems with appropriate statistics.

Suggested improvements:

1. Fig. 1: As stated above, it is crucial to provide the editing efficiency and the determination of homology arm length in human primary organoids, instead of HEK293T cells. Further, in Fig. 1b, did the authors observe the off-target effects with WT Cas9? TIDE analysis with sanger sequencing data would be useful to determine off-target effects in both WT Cas9 and nickase Cas9. In Fig. 1c, homozygous knock-in by FACS analysis must be supported by locus sequencing data either in the figure or as supplementary dataset.

2. Fig. 3: The locus that were targeted are actively transcribing/constitutively active (LMNA, HIST1H2BC, CDH1…). Can inactive locus be targeted with similar efficiency? For wider application of this protocol, both actively transcribing and inactive genomic regions must be targeted. Further, can an inactive locus in the colonic organoids targeted with a fluorescent reporter be turned-on by in vitro culture modification? OR can a florescent knock-in into an active gene be turned-off transiently? This will determine how well the florescence reports the transcriptional activity of the gene.

3. How does the size of knock-in construct influence on the efficiency of knock-in? Is the efficiency of knock-in at CDH1 locus similar to "mScarlet" (Fig. 3) and "mScarlet-P2A-Blasticidin" (Sup Fig. 1)? How much does insert size influence on the efficiency of knock-in with this strategy in this manuscript?

4. Fig. 3 and Sup Fig. 1: How many biological repeats have been conducted for this strategy? How much efficiency variations have been observed?

5. Have the authors tested whether organoid lines generated by the multiplex fluorescent gene tagging strategy can be stably maintained over multiple passages? Can organoid lines with 3x fluorescent reporters be frozen (liqN2), recovered and subculture for long-term tracing? It would be informative to provide these further validations for wider utility.

6. It would be informative to provide the sanger sequencing results for the targeted locus for Fig. 3, Fig. 4, and Sup. Fig. 1.

Clarity and context: The manuscript has been well written with a logical flow and a clear abstract, introduction, and conclusion.

Reviewer #3:

[identifies himself as Andrew Bassett]

Summary

Bollen et al. demonstrate that large knock-ins in human organoids can be achieved with good efficiency and precision through use of in-trans paired nicking (nicking of both the genomic locus and targeting vector). The combination of a nicked targeting vector and nickase enzyme shows comparable overall rates of editing compared with WT Cas9, highlighting the potential of generating knock-ins without inducing a double-stranded break. The potential of this technique is further demonstrated through the generation of a triple KI in human colon organoids and the integration of different fluorescent markers on each allele at a single locus, the latter of which establishes a method for the study of allele-specific expression dynamics. Finally, to address the lengthy cloning process associated with generating targeting vectors with homology arms, the authors demonstrate a one-step targeting vector assembly strategy utilising Golden-gate cloning, resulting in a targeting vector with the specific upstream and downstream homology regions on either side of a fluorescent tag. The targeting vector backbone variants developed during this work are available through Addgene.

I find the paper to be well written, clear and easy to understand, and the supplementary files provide a clear overview of the strategy and combined with the availability of vectors would enable others to use this methodology. Experiments were well thought out and performed, and do show the versatility and precision of in-trans paired nicking over conventional CRISPR-mediated knock-in methods.

I would recommend acceptance in the methods section of PlOS Biology once experiments to address on and off target mutations and integrations listed below have been addressed, and the minor comments have been corrected.

Major Issues

1. As referenced in this manuscript, the technique of in-trans paired nicking has already been established in other systems such as human cells. Thus, the principal novelty of this study is the application of this technique to human organoids

2. Integrity of non-targeted allele. One of the major benefits of the in trans paired nicking system is that the non-targeted allele should not accumulate indel mutations, allowing for instance multiple rounds of targeting without sequence validation and cloning in-between and meaning that e.g. "…sequence verification of individually picked clones is no longer required since all knock-in cells can immediately be pooled." However, the authors do not directly demonstrate this precision. They should perform experiments to address the frequency of mutations at the non-targeted allele for instance by high throughput sequencing of the WT alleles after a targeting experiment in comparison to the DSB-mediated methodologies.

3. On target precision. Similarly, it has been shown that DSB-mediated genome engineering can introduce undesired and unexpected mutations on target such as long deletions (Kosicki et al. NatBiotech 2018, "Repair of double-strand breaks induced by CRISPR-Cas9 leads to large deletions and complex rearrangements"), and tandem integrations of multiple copies of the transgene, etc (Canaj et al. BioRxIV 2019 "Deep profiling reveals substantial heterogeneity of integration outcomes in CRISPR knock-in experiments"). It would be important to assess the frequency of these on target mutations with the different strategies. This could be done by targeted long read sequencing (e.g. ONT), or long-range PCR and Sanger sequencing.

4. Off-target mutation. Another advantage of using Cas9 nickase is the reduction of off-target mutations. I feel that this is not addressed experimentally and should be done through sequencing potential off-target sites (in comparison to nucleases). A key reference that should be included on this topic is Chen et al. NAR 2020 "Expanding the editable genome and CRISPR-Cas9 versatility using DNA cutting-free gene targeting based on in trans paired nicking."

5. Off-target integration. dsDNA molecules, especially when linearized, are able to integrate randomly (as well as at CRISPR off-target sites) in the genome. The authors should address the frequency at which this occurs by performing analysis of random integration of transgenes (e.g. by ddPCR copy number counting or inverse PCR).

Minor Issues

1. "By contrast, flanking homology arms with Cas9 target sites to facilitate in-trans paired cleavage saturates the homology demand at around ~600bp [10]" I'm not sure the paper they cite really demonstrates this difference, since it seems locus dependent (for instance, Fig 2C is quite similar between the paired cleavage and circular donors). They should adapt the text accordingly

2. The authors should cite additional papers describing the in trans paired nicking approach such as (but not restricted to) the following:

a. Rees et al. 2019 NatComms "Expanding the editable genome and CRISPR-Cas9 versatility using DNA cutting-free gene targeting based on in trans paired nicking " that uses a Cas9 nickase fusion to Rad51 to improve editing outcomes.

b. Hyodo et al. 2020 CellRep "Tandem Paired Nicking Promotes Precise Genome Editing with Scarce Interference by p53"

3. Figure 1 - 5' and 3' annotation on DNA incorrect.

4. Supplementary Figure 1 - Incorrect spelling of 'triple' in diagram title.

17 Nov 2021

Submitted filename: Rebuttal PLOS 211115.docx

Click here for additional data file.

7 Dec 2021

Dear Dr Snippert,

Thank you for submitting your revised Methods and Resources article entitled "Efficient and error-free fluorescent gene tagging in human organoids without double-strand DNA cleavage" for publication in PLOS Biology. I have now obtained advice from two of the original reviewers and have discussed their comments with the Academic Editor.

As you can see, the reviewers appreciated the substantial amount of additional data included in the revised manuscript to address their comments. Based on the reviews, we will probably accept this manuscript for publication. Please make sure to address the following data and other policy-related requests that I have provided below:

(A) We note that flow cytometry data is presented in Figures 3B, 4A, S1 and S7B. We ask that you please provide the raw FCS files of these data and recommend depositing the files in the FlowRepository database due to the potential size of the files (https://flowrepository.org/). If you do use the FlowRepository, please ensure that the files are publicly available at this stage and please provide the accession number/URL for the deposition.

(B) Please deposit the raw Sanger and TIDE sequencing data (Figure S3-S5, S6 and S8) in a public database, such as the GEO. As before, please ensure that the data is made publicly available at this stage and that you provide the accession number/URL for the deposition.

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-------------------------

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Please do not hesitate to contact me should you have any questions.

Sincerely,

Richard

Richard Hodge, PhD

Associate Editor, PLOS Biology

rhodge@plos.org

------------------------------------------------------------------------

Reviewer remarks:

Reviewer #2: The revised manuscript represents a significant improvement over the initial submission. I have no further comment, and appreciate the authors' effort to address the raised questions and concerns.

Reviewer #3 (Andrew Bassett, signs his review): The authors have substantially revised their manuscript, and it now addresses most of my concerns through incorporation of additional sequence analysis of on and off-target mutagenesis, and whole genome characterisation. The changes made have substantially improved the manuscript, which demonstrates application of this methodology to organoid systems, and also provides a toolkit of reagents for others to use to employ this. I am happy for it to be published in Plos Biology.

5 Jan 2022

Dear Dr Snippert,

On behalf of my colleagues and the Academic Editor, Madeline Lancaster, I am pleased to say that we can in principle accept your Methods and Resources "Efficient and error-free fluorescent gene tagging in human organoids without double-strand DNA cleavage" for publication in PLOS Biology, provided you address any remaining formatting and reporting issues. These will be detailed in an email that will follow this letter and that you will usually receive within 2-3 business days, during which time no action is required from you. Please note that we will not be able to formally accept your manuscript and schedule it for publication until you have any requested changes.

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PRESS

We frequently collaborate with press offices. If your institution or institutions have a press office, please notify them about your upcoming paper at this point, to enable them to help maximise its impact. If the press office is planning to promote your findings, we would be grateful if they could coordinate with biologypress@plos.org. If you have not yet opted out of the early version process, we ask that you notify us immediately of any press plans so that we may do so on your behalf.

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Thank you again for choosing PLOS Biology for publication and supporting Open Access publishing. We look forward to publishing your study.

Sincerely,

Richard

Richard Hodge, PhD

Associate Editor, PLOS Biology

rhodge@plos.org

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