The efficacy and the mutation spectrum of genome editing methods can vary substantially depending on the targeted sequence. A simple, quick assay to accurately characterize and quantify the induced mutations is therefore needed. Here we present TIDE, a method for this purpose that requires only a pair of PCR reactions and two standard capillary sequencing runs. The sequence traces are then analyzed by a specially developed decomposition algorithm that identifies the major induced mutations in the projected editing site and accurately determines their frequency in a cell population. This method is cost-effective and quick, and it provides much more detailed information than current enzyme-based assays. An interactive web tool for automated decomposition of the sequence traces is available. TIDE greatly facilitates the testing and rational design of genome editing strategies.

While nuclear compartmentalization is an essential feature of three-dimensional genome organization, no genomic method exists for measuring chromosome distances to defined nuclear structures. In this study, we describe TSA-Seq, a new mapping method capable of providing a "cytological ruler" for estimating mean chromosomal distances from nuclear speckles genome-wide and for predicting several Mbp chromosome trajectories between nuclear compartments without sophisticated computational modeling. Ensemble-averaged results in K562 cells reveal a clear nuclear lamina to speckle axis correlated with a striking spatial gradient in genome activity. This gradient represents a convolution of multiple spatially separated nuclear domains including two types of transcription "hot zones." Transcription hot zones protruding furthest into the nuclear interior and positioning deterministically very close to nuclear speckles have higher numbers of total genes, the most highly expressed genes, housekeeping genes, genes with low transcriptional pausing, and super-enhancers. Our results demonstrate the capability of TSA-Seq for genome-wide mapping of nuclear structure and suggest a new model for spatial organization of transcription and gene expression.

SummaryThe RNA-guided DNA endonuclease Cas9 is a powerful tool for genome editing. Little is known about the kinetics and fidelity of the double-strand break (DSB) repair process that follows a Cas9 cutting event in living cells. Here, we developed a strategy to measure the kinetics of DSB repair for single loci in human cells. Quantitative modeling of repaired DNA in time series after Cas9 activation reveals variable and often slow repair rates, with half-life times up to ∼10 hr. Furthermore, repair of the DSBs tends to be error prone. Both classical and microhomology-mediated end joining pathways contribute to the erroneous repair. Estimation of their individual rate constants indicates that the balance between these two pathways changes over time and can be altered by additional ionizing radiation. Our approach provides quantitative insights into DSB repair kinetics and fidelity in single loci and indicates that Cas9-induced DSBs are repaired in an unusual manner.

Repair of DNA double-strand breaks (DSBs) must be properly orchestrated in diverse chromatin regions to maintain genome stability. The choice between two main DSB repair pathways, nonhomologous end-joining (NHEJ) and homologous recombination (HR), is regulated by the cell cycle as well as chromatin context. Pericentromeric heterochromatin forms a distinct nuclear domain that is enriched for repetitive DNA sequences that pose significant challenges for genome stability. Heterochromatic DSBs display specialized temporal and spatial dynamics that differ from euchromatic DSBs. Although HR is thought to be the main pathway used to repair heterochromatic DSBs, direct tests of this hypothesis are lacking. Here, we developed an in vivo single DSB system for both heterochromatic and euchromatic loci in Drosophila melanogaster. Live imaging of single DSBs in larval imaginal discs recapitulates the spatio-temporal dynamics observed for irradiation (IR)-induced breaks in cell culture. Importantly, live imaging and sequence analysis of repair products reveal that DSBs in euchromatin and heterochromatin are repaired with similar kinetics, employ both NHEJ and HR, and can use homologous chromosomes as an HR template. This direct analysis reveals important insights into heterochromatin DSB repair in animal tissues and provides a foundation for further explorations of repair mechanisms in different chromatin domains.

Summary DNA double-strand break (DSB) repair is mediated by multiple pathways. It is thought that the local chromatin context affects the pathway choice, but the underlying principles are poorly understood. Using a multiplexed reporter assay in combination with Cas9 cutting, we systematically measure the relative activities of three DSB repair pathways as a function of chromatin context in >1,000 genomic locations. This reveals that non-homologous end-joining (NHEJ) is broadly biased toward euchromatin, while the contribution of microhomology-mediated end-joining (MMEJ) is higher in specific heterochromatin contexts. In H3K27me3-marked heterochromatin, inhibition of the H3K27 methyltransferase EZH2 reverts the balance toward NHEJ. Single-stranded template repair (SSTR), often used for precise CRISPR editing, competes with MMEJ and is moderately linked to chromatin context. These results provide insight into the impact of chromatin on DSB repair pathway balance and guidance for the design of Cas9-mediated genome editing experiments.

Template-directed CRISPR/Cas9 editing is a powerful tool for introducing subtle mutations in genomes. However, the success rate of incorporation of the desired mutations at the target site is difficult to predict and therefore must be empirically determined. Here, we adapted the widely used TIDE method for quantification of templated editing events, including point mutations. The resulting TIDER method is a rapid, cheap and accessible tool for testing and optimization of template-directed genome editing strategies. A free web tool for TIDER data analysis is available at http://tide.nki.nl.

Current genome editing tools enable targeted mutagenesis of selected DNA sequences in many species. However, the efficiency and the type of introduced mutations by the genome editing method are largely dependent on the target site. As a consequence, the outcome of the editing operation is difficult to predict. Therefore, a quick assay to quantify the frequency of mutations is vital for a proper assessment of genome editing actions. We developed two methods that are rapid, cost-effective, and readily applicable: (1) TIDE, which can accurately identify and quantify insertions and deletions (indels) that arise after introduction of double strand breaks (DSBs); (2) TIDER, which is suited for template-mediated editing events including point mutations. Both methods only require a set of PCR reactions and standard Sanger sequencing runs. The sequence traces are analyzed by the TIDE or TIDER algorithm (available at https://tide.nki.nl or https://deskgen.com). The routine is easy, fast, and provides much more detailed information than current enzyme-based assays. TIDE and TIDER accelerate testing and designing of DSB-based genome editing strategies.

Template-directed CRISPR/Cas9 editing is a powerful tool for introducing subtle mutations ingenomes. However, the success rate of incorporation of the desired mutations at the target site is difficult to predict and therefore must be empirically determined. Here, we adapted the widely used TIDE method for quantification of templated editing events, including point mutations. The resulting TIDER method is a rapid, cheap and accessible tool for testing and optimization of template-directed genome editing strategies.. CC-BY-NC-ND 4.0 International license peer-reviewed) is the author/funder. It is made available under aThe copyright holder for this preprint (which was not . http://dx.doi.org/10.1101/218156 doi: bioRxiv preprint first posted online Nov. 13, 2017; 2 The CRISPR system for genome editing has become one of the most popular techniques in molecular biology. CRISPR endonucleases such as Cas9 can cleave genomic DNA with high precision, and due to error-prone repair mechanisms this can result in small insertions or deletions (indels) [1][2][3] .Alternatively, precisely designed small nucleotide changes can be incorporated near the break site by providing a donor template 4,5 , such as a single-stranded oligodeoxynucleotide (ssODN) 4,6 . By homology-directed repair (HDR), the DNA of the donor template is exchanged with the genomic DNA, and thereby the desired mutations are introduced 7,8 . Such precise editing offers the possibility to create and study specific mutations, or to correct disease-causing nucleotide variants 5,9 .A current limitation of this template-directed strategy is that the efficacy is unpredictable and often low. Because error-prone non-templated repair pathways are active besides HDR, various indels are often introduced at the target site instead of the desired mutation. Moreover, a substantial fraction of the target sequence may remain unaltered. Thus, exposing a pool of cells to CRISPR and a donor template yields a complex mixture of cells with wild-type DNA, indels and the designed mutation, with unpredictable ratios [10][11][12] . A quick and easy assay to determine these ratios is of key importance, particularly if one wants to estimate how many cells are to be cloned from the pool in order to obtain at least one clonal line with the desired mutation.High throughput sequencing of DNA around the induced break site is a powerful tool to analyze the mutation spectrum 13 , but is also expensive and requires substantial computational analysis. The copyright holder for this preprint (which was not . http://dx.doi.org/10.1101/218156 doi: bioRxiv preprint first posted online Nov. 13, 2017; 3 the sgRNA is used as input to determine the expected break site. Indels are then quantified by computational decomposition of the mixture of sequences in the test sequence trace, using the control sequence trace for comparison 14 .TIDER takes a similar decomposition approach, but it requires one additional capillary sequencing trace. This "reference" trace is derived from a pure DNA sample that c...