Julia Joung, Silvana Konermann, Jonathan S Gootenberg, Omar O Abudayyeh, Randall J Platt, Mark D Brigham, Neville E. Sanjana, Feng Zhang

Forward genetic screens are powerful tools for the unbiased discovery and functional characterization of specific genetic elements associated with a phenotype of interest. Recently, the RNA-guided endonuclease Cas9 from the microbial immune system CRISPR (clustered regularly interspaced short palindromic repeats) has been adapted for genome-scale screening by combining Cas9 with guide RNA libraries. Here we describe a protocol for genome-scale knockout and transcriptional activation screening using the CRISPR-Cas9 system. Custom- or ready-made guide RNA libraries are constructed and packaged into lentivirus for delivery into cells for screening. As each screen is unique, we provide guidelines for determining screening parameters and maintaining sufficient coverage. To validate candidate genes identified from the screen, we further describe strategies for confirming the screening phenotype as well as genetic perturbation through analysis of indel rate and transcriptional activation. Beginning with library design, a genome-scale screen can be completed in 6-10 weeks followed by 3-4 weeks of validation.

Rahul Sinha, Geoff Stanley, Gunsagar S. Gulati, Camille Ezran, Kyle J. Travaglini, Eric Wei, Charles K.F. Chan, Ahmad N. Nabhan, Tianying Su, Rachel M. Morganti, Stephanie D Conley, Hassan Chaib, Kristy Red-Horse, Michael T Longaker, Michael P. Snyder, Mark A Krasnow, Irving L. Weissman

Illumina-based next generation sequencing (NGS) has accelerated biomedical discovery through its ability to generate thousands of gigabases of sequencing output per run at a fraction of the time and cost of conventional technologies. The process typically involves four basic steps: library preparation, cluster generation, sequencing, and data analysis. In 2015, a new chemistry of cluster generation was introduced in the newer Illumina machines (HiSeq 3000/4000/X Ten) called exclusion amplification (ExAmp), which was a fundamental shift from the earlier method of random cluster generation by bridge amplification on a non-patterned flow cell. The ExAmp chemistry, in conjunction with patterned flow cells containing nanowells at fixed locations, increases cluster density on the flow cell, thereby reducing the cost per run. It also increases sequence read quality, especially for longer read lengths (up to 150 base pairs). This advance has been widely adopted for genome sequencing because greater sequencing depth can be achieved for lower cost without compromising the quality of longer reads. We show that this promising chemistry is problematic, however, when multiplexing samples. We discovered that up to 5-10% of sequencing reads (or signals) are incorrectly assigned from a given sample to other samples in a multiplexed pool. We provide evidence that this “spreading-of-signals” arises from low levels of free index primers present in the pool. These index primers can prime pooled library fragments at random via complementary 3′ ends, and get extended by DNA polymerase, creating a new library molecule with a new index before binding to the patterned flow cell to generate a cluster for sequencing. This causes the resulting read from that cluster to be assigned to a different sample, causing the spread of signals within multiplexed samples. We show that low levels of free index primers persist after the most common library purification procedure recommended by Illumina, and that the amount of signal spreading among samples is proportional to the level of free index primer present in the library pool. This artifact causes homogenization and misclassification of cells in single cell RNA-seq experiments. Therefore, all data generated in this way must now be carefully re-examined to ensure that “spreading-of-signals” has not compromised data analysis and conclusions. Re-sequencing samples using an older technology that uses conventional bridge amplification for cluster generation, or improved library cleanup strategies to remove free index primers, can minimize or eliminate this signal spreading artifact.

Ryan A. Flynn, Benjamin A. H. Smith, Alex G. Johnson, Kayvon Pedram, Benson M. George, Stacy A. Malaker, Karim Majzoub, Jan E. Carette, Carolyn R. Bertozzi

Glycans modify lipids and proteins to mediate inter- and intramolecular interactions across all domains of life. RNA, another multifaceted biopolymer, is not thought to be a major target of glycosylation. Here, we challenge this view with evidence that mammalian cells use RNA as a third scaffold for glycosylation in the secretory pathway. Using a battery of chemical and biochemical approaches, we find that a select group of small noncoding RNAs including Y RNAs are modified with complex, sialylated N-glycans (glycoRNAs). These glycoRNA are present in multiple cell types and mammalian species, both in cultured cells and in vivo. Finally, we find that RNA glycosylation depends on the canonical N-glycan biosynthetic machinery within the ER/Golgi luminal spaces. Collectively, these findings suggest the existence of a ubiquitous interface of RNA biology and glycobiology suggesting an expanded role for glycosylation beyond canonical lipid and protein scaffolds.

Yuancheng Lu, Anitha Krishnan, Benedikt Brommer, Xiao Tian, Margarita Meer, Daniel L. Vera, Chen Wang, Qiurui Zeng, Doudou Yu, Michael S. Bonkowski, Jae-Hyun Yang, Emma M. Hoffmann, Songlin Zhou, Ekaterina Korobkina, Noah Davidsohn, Michael B. Schultz, Karolina Chwalek, Luis A. Rajman, George M. Church, Konrad Hochedlinger, Vadim N. Gladyshev, Steve Horvath, Meredith S. Gregory-Ksander, Bruce R. Ksander, Zhigang He, David A. Sinclair

Ageing is a degenerative process leading to tissue dysfunction and death. A proposed cause of ageing is the accumulation of epigenetic noise, which disrupts youthful gene expression patterns that are required for cells to function optimally and recover from damage. Changes to DNA methylation patterns over time form the basis of an 'ageing clock', but whether old individuals retain information to reset the clock and, if so, whether this would improve tissue function is not known. Of all the tissues in the body, the central nervous system (CNS) is one of the first to lose regenerative capacity. Using the eye as a model tissue, we show that expression of Oct4, Sox2, and Klf4 genes (OSK) in mice resets youthful gene expression patterns and the DNA methylation age of retinal ganglion cells, promotes axon regeneration after optic nerve crush injury, and restores vision in a mouse model of glaucoma and in normal old mice. This process, which we call recovery of information via epigenetic reprogramming or REVIVER, requires the DNA demethylases Tet1 and Tet2, indicating that DNA methylation patterns don't just indicate age, they participate in ageing. Thus, old tissues retain a faithful record of youthful epigenetic information that can be accessed for functional age reversal.

Han Li, Kyle A. Beckman, Veronica Pessino, Bo Huang, Jonathan S. Weissman, Manuel D. Leonetti

CRISPR/Cas technologies have transformed our ability to manipulate genomes for research and gene-based therapy. In particular, homology-directed repair after genomic cleavage allows for precise modification of genes using exogenous donor sequences as templates. While both single-stranded DNA (ssDNA) and double-stranded DNA (dsDNA) forms of donors have been used as repair templates, a systematic comparison of the performance and specificity of repair using ssDNA versus dsDNA donors is still lacking. Here, we describe an optimized method for the synthesis of long ssDNA templates and demonstrate that ssDNA donors can drive efficient integration of gene-sized reporters in human cell lines. We next define a set of rules to maximize the efficiency of ssDNA-mediated knock-in by optimizing donor design. Finally, by comparing ssDNA donors with equivalent dsDNA sequences (PCR products or plasmids), we demonstrate that ssDNA templates have a unique advantage in terms of repair specificity while dsDNA donors can lead to a high rate of off-target integration. Our results provide a framework for designing high-fidelity CRISPR-based knock-in experiments, in both research and therapeutic settings. Update: November 12th, 2019 Dear bioRxiv community, The conclusions of this pre-print (originally posted in August 2017) are outdated. While the experiments we present here are accurate, a recent and more systematic analysis revealed that the integration outcomes driven by different forms of HDR donors are more complex than our methods could originally identify. We initially analyzed donor integration only in FACS-selected cells, which under-estimates alleles where the mis-integration of payload leads to non-functional selection markers, and we quantified integration by ddPCR, which is an indirect read-out of sequence properties. These approaches could not capture the full details of donor integration events in our experiments. To address this, we have now developed a new framework based on long-read amplicon sequencing and an integrated computational pipeline to precisely analyze knock-in repair outcomes across a wide range of experimental parameters. Our new data uncover a complex repair landscape in which both single-stranded and double-stranded donors can lead to high rates of imprecise integration in some cell types. Please read our new bioRxiv pre-print entitled “Deep profiling reveals substantial heterogeneity of integration outcomes in CRISPR knock-in experiments” for further information. I hope that this example highlights one of the powers of pre-prints: the ability to update scientific discussions (and set records straight) as new results are obtained, often fueled by the availability of new technologies. Please do not hesitate to contact me directly for any questions or comments.

Michael P. Meers, Terri Bryson, Steven Henikoff

We previously described a novel alternative to Chromatin Immunoprecipitation, Cleavage Under Targets & Release Using Nuclease (CUT&RUN), in which unfixed permeabilized cells are incubated with antibody, followed by binding of a Protein A-Micrococcal Nuclease (pA/MNase) fusion protein (1). Upon activation of tethered MNase, the bound complex is excised and released into the supernatant for DNA extraction and sequencing. Here we introduce four enhancements to CUT&RUN: 1) a hybrid Protein A-Protein G-MNase construct that expands antibody compatibility and simplifies purification; 2) a modified digestion protocol that inhibits premature release of the nuclease-bound complex; 3) a calibration strategy based on carry-over of E. coli DNA introduced with the fusion protein; and 4) a novel peak-calling strategy customized for the low-background profiles obtained using CUT&RUN. These new features, coupled with the previously described low-cost, high efficiency, high reproducibility and high- throughput capability of CUT&RUN make it the method of choice for routine epigenomic profiling.

Feng Gao, Yue Sun, Feng Jiang, Xiaoyue Bai, Chunyu Han

RNAs have important and diverse functions. Visualizing an isolated RNA in living cells provide us essential information of its roles. By now, there are two kinds of live RNA imaging systems invented, one is the MS2 system and the other is the Cas13a system. In this study, we show that when fused with split-Fp, CasE can be engineered into a live RNA tracking tool.

Christopher D Go, James D.R. Knight, Archita Rajasekharan, Bhavisha Rathod, Geoffrey G Hesketh, Kento T Abe, Ji-Young Youn, Payman Samavarchi-Tehrani, Hui Zhang, Lucie Y Zhu, Evelyn Popiel, Jean-Philippe Lambert, Étienne Coyaud, Sally W.T. Cheung, Dushyandi Rajendran, Cassandra J Wong, Hana Antonicka, Laurence Pelletier, Brian Raught, Alexander F Palazzo, Eric A Shoubridge, Anne-Claude Gingras

Compartmentalization is an essential characteristic of eukaryotic cells, ensuring that cellular processes are partitioned to defined subcellular locations. High throughput microscopy and biochemical fractionation coupled with mass spectrometry have helped to define the proteomes of multiple organelles and macromolecular structures. However, many compartments have remained refractory to such methods, partly due to lysis and purification artefacts and poor subcompartment resolution. Recently developed proximity-dependent biotinylation approaches such as BioID and APEX provide an alternative avenue for defining the composition of cellular compartments in living cells. Here we report an extensive BioID-based proximity map of a human cell, comprising 192 markers from 32 different compartments that identifies 35,902 unique high confidence proximity interactions and localizes 4,145 proteins expressed in HEK293 cells. The recall of our localization predictions is on par with or better than previous large-scale mass spectrometry and microscopy approaches, but with higher localization specificity. In addition to assigning compartment and subcompartment localization for many previously unlocalized proteins, our data contain fine- grained localization information that, for example, allowed us to identify proteins with novel roles in mitochondrial dynamics. As a community resource, we have created humancellmap.org, a website that allows exploration of our data in detail, and aids with the analysis of BioID experiments.

Pinar Akcakaya, Maggie L. Bobbin, Jimmy A. Guo, Jose M Lopez, M. Kendell Clement, Sara P. Garcia, Mick D. Fellows, Michelle J. Porritt, Mike A. Firth, Alba Carreras, Tania Baccega, Frank Seeliger, Mikael Bjursell, Shengdar Q. Tsai, Nhu T. Nguyen, Roberto Nitsch, Lorenz M Mayr, Luca Pinello, Mohammad Bohlooly-Y, Martin J. Aryee, Marcello Maresca, J. Keith Joung

CRISPR-Cas genome-editing nucleases hold substantial promise for human therapeutics but identifying unwanted off-target mutations remains an important requirement for clinical translation. For ex vivo therapeutic applications, previously published cell-based genome-wide methods provide potentially useful strategies to identify and quantify these off-target mutation sites. However, a well-validated method that can reliably identify off-targets in vivo has not been described to date, leaving the question of whether and how frequently these types of mutations occur. Here we describe Verification of In Vivo Off-targets (VIVO), a highly sensitive, unbiased, and generalizable strategy that we show can robustly identify genome-wide CRISPR-Cas nuclease off-target effects in vivo. To our knowledge, these studies provide the first demonstration that CRISPR-Cas nucleases can induce substantial off-target mutations in vivo, a result we obtained using a deliberately promiscuous guide RNA (gRNA). More importantly, we used VIVO to show that appropriately designed gRNAs can direct efficient in vivo editing without inducing detectable off-target mutations. Our findings provide strong support for and should encourage further development of in vivo genome editing therapeutic strategies.

Susannah J Salter, Michael J. Cox, Elena M Turek, Szymon T Calus, William O Cookson, Miriam F. Moffatt, Paul Turner, Julian Parkhill, Nick Loman, Alan W Walker

The study of microbial communities has been revolutionised in recent years by the widespread adoption of culture independent analytical techniques such as 16S rRNA gene sequencing and metagenomics. One potential confounder of these sequence-based approaches is the presence of contamination in DNA extraction kits and other laboratory reagents. In this study we demonstrate that contaminating DNA is ubiquitous in commonly used DNA extraction kits, varies greatly in composition between different kits and kit batches, and that this contamination critically impacts results obtained from samples containing a low microbial biomass. Contamination impacts both PCR-based 16S rRNA gene surveys and shotgun metagenomics. These results suggest that caution should be advised when applying sequence-based techniques to the study of microbiota present in low biomass environments. We provide an extensive list of potential contaminating genera, and guidelines on how to mitigate the effects of contamination. Concurrent sequencing of negative control samples is strongly advised.

Gali Housman, Igor Ulitsky

Long noncoding RNAs (lncRNAs) are a diverse class of RNAs with increasingly appreciated functions in vertebrates, yet much of their biology remains poorly understood. In particular, it is unclear to what extent the current catalog of over 10,000 distinct annotated lncRNAs is indeed devoid of genes coding for proteins. Here we review the available computational and experimental schemes for distinguishing between recent genome-wide applications. We conclude that the model most consistent with available data is that a large number of mammalian lncRNAs undergo translation, but only a very small minority of such translation events result in stable and functional peptides. The outcome of the majority of the translation events and their potential biological purposes remain an intriguing topic for future investigation.

Omar O Abudayyeh, Jonathan S Gootenberg, Silvana Konermann, Julia Joung, Ian M Slaymaker, David B.T. Cox, Sergey Shmakov, Kira S. Makarova, Ekaterina Semenova, Leonid Minakhin, Konstantin Severinov, Aviv Regev, Eric S Lander, Eugene V. Koonin, Feng Zhang

The CRISPR-Cas adaptive immune system defends microbes against foreign genetic elements via DNA or RNA-DNA interference. We characterize the Class 2 type VI-A CRISPR-Cas effector C2c2 and demonstrate its RNA-guided RNase function. C2c2 from the bacterium Leptotrichia shahii provides interference against RNA phage. In vitro biochemical analysis show that C2c2 is guided by a single crRNA and can be programmed to cleave ssRNA targets carrying complementary protospacers. In bacteria, C2c2 can be programmed to knock down specific mRNAs. Cleavage is mediated by catalytic residues in the two conserved HEPN domains, mutations in which generate catalytically inactive RNA-binding proteins. These results broaden our understanding of CRISPR-Cas systems and suggest that C2c2 can be used to develop new RNA-targeting tools.

Bernd Zetsche, Matthias Heidenreich, Prarthana Mohanraju, Iana Fedorova, Jeroen Kneppers, Ellen M DeGennaro, Nerges Winblad, Sourav R Choudhury, Omar O Abudayyeh, Jonathan S Gootenberg, Wen Y Wu, David A Scott, Konstantin Severinov, John van der Oost, Feng Zhang

Microbial CRISPR-Cas defense systems have been adapted as a platform for genome editing applications built around the RNA-guided effector nucleases, such as Cas9. We recently reported the characterization of Cpf1, the effector nuclease of a novel type V-A CRISPR system, and demonstrated that it can be adapted for genome editing in mammalian cells. Unlike Cas9, which utilizes a trans-activating crRNA (tracrRNA) as well as the endogenous RNaseIII for maturation of its dual crRNA:tracrRNA guides, guide processing of the Cpf1 system proceeds in the absence of tracrRNA or other Cas (CRISPR associated) genes, suggesting that Cpf1 is sufficient for pre-crRNA maturation. This has important implications for genome editing, as it would provide a simple route to multiplex targeting. Here, we show for two Cpf1 orthologs that no other factors are required for array processing and demonstrate multiplex gene editing in mammalian cells as well as in the mouse brain by using a designed single CRISPR array.

Søeren M. Karst, Ryan M. Ziels, Rasmus H. Kirkegaard, Emil A. Sørensen, Daniel McDonald, Qiyun Zhu, Rob Knight, Mads Albertsen

High-throughput amplicon sequencing of large genomic regions remains challenging for short-read technologies. Here, we report a high-throughput amplicon sequencing approach combining unique molecular identifiers (UMIs) with Oxford Nanopore Technologies or Pacific Biosciences CCS sequencing, yielding high accuracy single-molecule consensus sequences of large genomic regions. Our approach generates amplicon and genomic sequences of >10,000 bp in length with a mean error-rate of 0.0049-0.0006% and chimera rate <0.022%.

Lucas Sinclair, Omneya Ahmed Osman, Stefan Bertilsson, Alexander Eiler

As new sequencing technologies become cheaper and older ones disappear, laboratories switch vendors and platforms. Validating the new setups is a crucial part of conducting rigorous scientific research. Here we report on the reliability and biases of performing bacterial 16S rRNA gene amplicon paired-end sequencing on the MiSeq Illumina platform. We designed a protocol using 50 barcode pairs to run samples in parallel and coded a pipeline to process the data. Sequencing the same sediment sample in 248 replicates as well as 70 samples from alkaline soda lakes, we evaluated the performance of the method with regards to estimates of alpha and beta diversity. Using different purification and DNA quantification procedures we always found up to 5-fold differences in the yield of sequences between individually barcodes samples. Using either a one-step or a two-step PCR preparation resulted in significantly different estimates in both alpha and beta diversity. Comparing with a previous method based on 454 pyrosequencing, we found that our Illumina protocol performed in a similar manner -- with the exception for evenness estimates where correspondence between the methods was low. We further quantified the data loss at every processing step eventually accumulating to 50\% of the raw reads. When evaluating different OTU clustering methods, we observed a stark contrast between the results of QIIME with default settings and the more recent UPARSE algorithm when it comes to the number of OTUs generated. Still, overall trends in alpha and beta diversity corresponded highly using both clustering methods. Our procedure performed well considering the precisions of alpha and beta diversity estimates, with insignificant effects of individual barcodes. Comparative analyses suggest that 454 and Illumina sequence data can be combined if the same PCR protocol and bioinformatic workflows are used for describing patterns in richness, beta-diversity and taxonomic composition. (version 1.1 resubmitted to PLOS one 2014-Sept-08)

Andie C. Hall

Nucleic acid stains are necessary for Agarose Gel Electrophoresis (AGE). The commonly used but mutagenic Ethidium Bromide is being usurped by a range of safer but more expensive alternatives. These safe stains vary in cost, sensitivity and the impedance of DNA as it migrates through the gel. Modified protocols developed to reduce cost increase this variability. In this study, five Gel stains (GelRed™, GelGreen™, SYBR™ safe, SafeView and EZ-Vision®In-Gel Solution) two premixed loading dyes (SafeWhite, EZ-Vision®One) and four methods (pre-loading at 100x, pre-loading at 10x, precasting and post-staining) are evaluated for sensitivity and effect on DNA migration. GelRed™ was found to be the most sensitive while the EZ-Vision® dyes and SafeWhite had no discernible effect on DNA migration. Homemade loading dyes were as effective as readymade ones at less than 4% of the price. This method used less than 1% of the dye needed for the manufacturer recommended protocols. Thus, with careful consideration of stain and method, Gel stain expenditure can be reduced by over 99%.

Erika C Urdaneta, Carlos H. Vieira-Vieira, Timon Hick, Hans-Herrmann Wessels, Davide Figini, Rebecca Moschall, Jan Medenbach, Uwe Ohler, Sander Granneman, Matthias Selbach, Benedikt M Beckmann

Recent methodological advances allowed the identification of an increasing number of RNA-binding proteins (RBPs) and their RNA-binding sites. Most of those methods rely, however, on capturing proteins associated to polyadenylated RNAs which neglects RBPs bound to non-adenylate RNA classes (tRNA, rRNA, pre-mRNA) as well as the vast majority of species that lack poly-A tails in their mRNAs (including all archea and bacteria). To overcome these limitations, we have developed a novel protocol, Phenol Toluol extraction (PTex), that does not rely on a specific RNA sequence or motif for isolation of cross-linked ribonucleoproteins (RNPs), but rather purifies them based entirely on their physicochemical properties. PTex captures RBPs that bind to RNA as short as 30 nt, RNPs directly from animal tissue and can be used to simplify complex workflows such as PAR-CLIP. Finally, we provide a first global RNA-bound proteome of human HEK293 cells and Salmonella Typhimurium as a bacterial species.

Huanle Liu, Oguzhan Begik, Morghan C Lucas, Lindsay F. Rizzardi, Schraga Schwartz, John S Mattick, Martin A. Smith, Eva Maria Novoa

The field of epitranscriptomics has undergone an enormous expansion in the last few years; however, a major limitation is the lack of generic methods to map RNA modifications transcriptome-wide. Here we show that using Oxford Nanopore Technologies, N6-methyladenosine (m6A) RNA modifications can be detected with high accuracy, in the form of systematic errors and decreased base-calling qualities. Our results open new avenues to investigate the universe of RNA modifications with single nucleotide resolution, in individual RNA molecules.

Birgit Koch, Bianca Nijmeijer, Moritz Kueblbeck, Yin Cai, Nike Walther, Jan Ellenberg

Gene tagging with fluorescent proteins (FPs) is essential to investigate the dynamic properties of cellular proteins. Clustered Regularly Interspaced Short Palindromic Repeats/Cas9 technology (CRISPR/Cas9) technology is a powerful tool for inserting fluorescent markers into all alleles of the gene of interest and permits functionality and physiological expression of the fusion protein. It is essential to evaluate such genome-edited cell lines carefully in order to preclude off-target effects caused by either (i) incorrect insertion of the FP, (ii) perturbation of the fusion protein by the FP or (iii) non-specific genomic DNA damage by CRISPR/Cas9. In this protocol, we provide a step-by-step description of our systematic pipeline to generate and validate homozygous fluorescent knock-in cell lines. We have used the paired Cas9D10A nickase approach to efficiently insert tags into specific genomic loci via homology-directed repair (HDR) with minimal off-target effects. It is not only time- and cost-effective to perform whole genome sequencing of each cell clone, but also there are spontaneous genetic variations occurring in mammalian cell lines. Therefore we have developed an efficient validation pipeline of the generated cell lines consisting of junction PCR, Southern Blot analysis, Sanger sequencing, microscopy, Western blot analysis and live cell imaging for cell cycle dynamics which takes between 6-9 weeks. Using this pipeline, 70% of the targeted genes could be tagged homozygously with FPs and resulted in physiological levels and phenotypically functional expression of the fusion proteins. In contrast to a study that systematically tagged genes using CRISPR/Cas9 in human stem cells1, our approach resulted in homozygously tagged proteins of interests.

Caleb A Lareau, Kendell Clement, Jonathan Y Hsu, Vikram Pattanayak, J. Keith Joung, Martin J. Aryee, Luca Pinello

Schaefer et al. recently advanced the provocative conclusion that CRISPR-Cas9 nuclease can induce off-target alterations at genomic loci that do not resemble the intended on-target site. Using high-coverage whole genome sequencing (WGS), these authors reported finding SNPs and indels in two CRISPR-Cas9-treated mice that were not present in a single untreated control mouse. On the basis of this association, Schaefer et al. concluded that these sequence variants were caused by CRISPR-Cas9. This new proposed CRISPR-Cas9 off-target activity runs contrary to previously published work and, if the authors are correct, could have profound implications for research and therapeutic applications. Here we demonstrate that the simplest interpretation of Schaefer et al.'s data is that the two CRISPR-Cas9-treated mice are actually more closely related genetically to each other than to the control mouse. This strongly suggests that the so-called “unexpected mutations” simply represent SNPs and indels shared in common by these mice prior to nuclease treatment. In addition, given the genomic and sequence distribution profiles of these variants, we show that it is challenging to explain how CRISPR-Cas9 might be expected to induce such changes. Finally, we argue that the lack of appropriate controls in Schaefer et al.'s experimental design precludes assignment of causality to CRISPR-Cas9. Given these substantial issues, we urge Schaefer et al. to revise or re-state the original conclusions of their published work so as to avoid leaving misleading and unsupported statements to persist in the literature.