Bacillus thuringiensis (Bt) has been successfully used commercially for more than 60 years for biocontrol of insect pests. Since 1996, transgenic plants expressing Bt crystal (Cry) proteins have been used commercially to provide protection against insects that predate on corn and cotton. More recently, Bt Cry proteins that target nematodes have been discovered. One of these, Cry14Ab, has been expressed in transgenic soybean plants and found to provide significant protection against the soybean cyst nematode, Heterodera glycines. However, to date there has been no description of high-level resistance to any Cry14A family protein in nematodes. Here, we describe forward genetic screens to identify such mutants using the nematode Caenorhabditis elegans. Although non-conditional screens failed to identify highly resistant C. elegans, a conditional (temperature-sensitive) genetic screen identified one mutant, bre-6(ye123) (for Bt protein resistant), highly resistant to both Cry14Aa and Cry14Ab. The mutant comes at a high fitness cost, showing significant delays in growth and development and reduced fecundity. bre-6(ye123) hermaphrodites are only weakly resistant to copper intoxication, indicating that the mutant is not highly resistant to all insults. Backcrossing—whole genome sequencing was used to identify the gene mutated in ye123 as the nuclear hormone receptor nhr-31. RNAi, DNA rescue, and CRISPR analyses confirm that resistance to Cry14Aa intoxication in bre-6(ye123) is due to mutation of nhr-31 and was renamed nhr-31(ye123). As predicted for a mutation in this gene, nhr-31(ye123) animals showed significantly reduced expression of most of the subunits of the C. elegans vacuolar ATPase (vATPase). Mutants in the vATPase subunits unc-32 and vha-7 also show resistance to Cry14Aa and/or Cry14Ab. These data demonstrate that nhr-31 and the vATPase play a significant role in the intoxication of C. elegans by Cry14A family proteins, that reduction in vATPase levels result in high resistance to Cry14A family proteins, and that such resistance comes at a high fitness cost. Based on the relative difficulty of finding resistant mutants and the fitness cost associated with the vATPase pathway, our data suggest that transgenic Cry14Ab plants may hold up well to resistance by nematode parasites.

Piwi-interacting RNAs (piRNAs) are genomically encoded small RNAs that engage Piwi Argonaute proteins to direct mRNA surveillance and transposon silencing. Despite advances in understanding piRNA pathways and functions, how the production of piRNA is regulated remains elusive. Here, using a genetic screen, we identify casein kinase II (CK2) as a factor required for piRNA pathway function. We show that CK2 is required for the localization of PRG-1 and for the proper localization of several factors that comprise the 'upstream sequence transcription complex' (USTC), which is required for piRNA transcription. Loss of CK2 impairs piRNA levels suggesting that CK2 promotes USTC function. We identify the USTC component twenty-one-U fouled-up 4 (TOFU-4) as a direct substrate for CK2. Our findings suggest that phosphorylation of TOFU-4 by CK2 promotes the assembly of USTC and piRNA transcription. Notably, during the aging process, CK2 activity declines, resulting in the disassembly of USTC, decreased piRNA production, and defects in piRNA-mediated gene silencing, including transposons silencing. These findings highlight the significance of posttranslational modification in regulating piRNA biogenesis and its implications for the aging process. Overall, our study provides compelling evidence for the involvement of a posttranslational modification mechanism in the regulation of piRNA biogenesis.

Retroviruses and closely related LTR retrotransposons export full-length, unspliced genomic RNA (gRNA) for packaging into virions and to serve as the mRNA encoding GAG and POL polyproteins. Because gRNA often includes splice acceptor and donor sequences used to splice viral mRNAs, retroelements must overcome host mechanisms that retain intron-containing RNAs in the nucleus. Here we examine gRNA expression in Cer1, an LTR retrotransposon in C . elegans which somehow avoids silencing and is highly expressed in germ cells. Newly exported Cer1 gRNA associates rapidly with the Cer1 GAG protein, which has structural similarity with retroviral GAG proteins. gRNA export requires CERV ( C . e legans r egulator of v iral expression), a novel protein encoded by a spliced Cer1 mRNA. CERV phosphorylation at S214 is essential for gRNA export, and phosphorylated CERV colocalizes with nuclear gRNA at presumptive sites of transcription. By electron microscopy, tagged CERV proteins surround clusters of distinct, linear fibrils that likely represent gRNA molecules. Single fibrils, or groups of aligned fibrils, also localize near nuclear pores. During the C . elegans self-fertile period, when hermaphrodites fertilize oocytes with their own sperm, CERV concentrates in two nuclear foci that are coincident with gRNA. However, as hermaphrodites cease self-fertilization, and can only produce cross-progeny, CERV undergoes a remarkable transition to form giant nuclear rods or cylinders that can be up to 5 microns in length. We propose a novel mechanism of rod formation, in which stage-specific changes in the nucleolus induce CERV to localize to the nucleolar periphery in flattened streaks of protein and gRNA; these streaks then roll up into cylinders. The rods are a widespread feature of Cer1 in wild strains of C . elegans , but their function is not known and might be limited to cross-progeny. We speculate that the adaptive strategy Cer1 uses for the identical self-progeny of a host hermaphrodite might differ for heterozygous cross-progeny sired by males. For example, mating introduces male chromosomes which can have different, or no, Cer1 elements.

Argonaute/small RNA pathways and heterochromatin work together to propagate transgenerational gene silencing, but the mechanisms behind their interaction are not well understood. Here, we show that induction of heterochromatin silencing in C. elegans by RNAi or by artificially tethering pathway components to target RNA causes co-localization of target alleles in pachytene nuclei. Tethering the nuclear Argonaute WAGO-9/ HRDE-1 induces heterochromatin formation and independently induces small RNA amplification. Consistent with this finding, HRDE-1, while predominantly nuclear, also localizes to peri-nuclear nuage domains, where amplification is thought to occur. Tethering a heterochromatin-silencing factor, NRDE-2, induces heterochro-matin formation, which subsequently causes de novo synthesis of HRDE-1 guide RNAs. HRDE-1 then acts to further amplify small RNAs that load on downstream Argonautes. These findings suggest that HRDE-1 plays a dual role, acting upstream to initiate heterochromatin silencing and downstream to stimulate a new cycle of small RNA amplification, thus establishing a self-enforcing mechanism that propagates gene silencing to future generations.

Germline Argonautes direct transcriptome surveillance within perinuclear membraneless organelles called nuage. In C. elegans, a family of Vasa-related Germ Line Helicase (GLH) proteins localize in and promote the formation of nuage. Previous studies have implicated GLH proteins in inherited silencing, but direct roles in small-RNA production, Argonaute binding, or mRNA targeting have not been identified. Here we show that GLH proteins compete with each other to control Argonaute pathway specificity, bind directly to Argonaute target mRNAs, and promote the amplification of small RNAs required for transgenerational inheritance. We show that the ATPase cycle of GLH-1 regulates direct binding to the Argonaute WAGO-1, which engages amplified small RNAs. Our findings support a dynamic and direct role for GLH proteins in inherited silencing beyond their role as structural components of nuage.

In animals, Argonaute small-RNA pathways scan germline transcripts to silence self-replicating genetic elements. However, little is known about how endogenous gene expression is recognized and licensed. Here, we show that the presence of introns and, by inference, the process of mRNA splicing prevents default Argonaute-mediated silencing in the C. elegans germline. The silencing of intronless genes is initiated independently of the piRNA pathway but nevertheless engages multiple components of the downstream amplification and maintenance mechanisms that mediate transgenerational silencing, including both nuclear and cytoplasmic members of the worm-specific Argonaute gene family (WAGOs). Small RNAs amplified from intronless mRNAs can trans-silence cognate intron-containing genes. Interestingly, a second, small RNA-independent cis-acting mode of silencing also acts on intronless mRNAs. Our findings suggest that cues put in place during mRNA splicing license germline gene expression and provide evidence for a splicing-dependent and dsRNA- and piRNA-independent mechanism that can program Argonaute silencing.

In Caenorhabditis elegans, targeted genome editing techniques are now routinely used to generate germline edits. The remarkable ease of C. elegans germline editing is attributed to the syncytial nature of the pachytene ovary which is easily accessed by microinjection. This protocol describes the step-by-step details and troubleshooting tips for the entire CRISPR-Cas genome editing procedure, including gRNA design and microinjection of ribonucleoprotein complexes, followed by screening and gen-otyping in C. elegans, to help accessing this powerful genetic animal system. For complete details on the use and execution of this protocol, please refer to Ghanta and Mello (2020).

The authors wish to withdraw this manuscript and apologize for errors in the initial submission. All the original experiments were performed by YY. Unfortunately, JP and members of the Mello lab have not been able to replicate some aspects of the study. JP has failed to independently reproduce the specific results showing RNAi-triggered relocalization of target RNA, and P granule specific accumulation of (the P granule component GLH-1) as reported. The conditions/strains analyzed by JP were as follows: (1)oma-1FISH on WT worms [control, 6 hr and 12 hroma-1(RNAi)]. (2)oma-1FISH on OMA-1:GFP worms [control, 6 hr oma-1(RNAi), or 6 hr gfp(RNAi)]. (3)oma-1FISH on WT worms [control, 4hr, 6hr, 8 hr, and 10 hroma-1(RNAi)]. 10-23 gonads were analyzed per experiment. Fixation conditions were essentially as described, with the only known difference being that gonads were not exposed to detergent prior to fixation. Using YYs reagents and protocol the Mello lab has not observed an obvious relocalization of target RNA to P granules (marked by GFP::GLH-1) after 6 hrsoma-1(RNAi);n=92 gonads. CM, JP and DG consider that the published images accurately represent the image stacks provided by YY as representative, raw data, but JP and CM note configurations of FISH signals in germ nuclei and gonad anatomy that they consider unusual. CM, JP and DG have not detected any evidence of image manipulation.YY states that none of the raw image data were manipulated beyond standard adjustments for brightness and contrast prior to processing images for publication as described. However, YY reports that the images were not representative of the majority of sample gonads, and instead were pre-selected under low magnification for rare examples with asymmetrical, expanded P granules.Efforts to identify conditions that explain the rare gonads imaged by YY continue in the Mello lab, as do efforts to reproduce independently each of the other reported results; we plan to provide an update in the near future.

In several species, Piwi/piRNA genome silencing defects cause immediate sterility that correlates with transposon expression and transposon-induced genomic instability. In C. elegans , mutations in the Piwi-related gene ( prg-1 ) and other piRNA deficient mutants cause a transgenerational decline in fertility over a period of several generations. Here we show that the sterility of late generation piRNA mutants correlates poorly with increases in DNA damage signaling. Instead, sterile individuals consistently exhibit altered perinuclear germ granules. We show that disruption of germ granules does not activate transposon expression but induces multiple phenotypes found in sterile prg-1 pathway mutants. Furthermore, loss of the germ granule component pgl-1 enhances prg-1 mutant infertility. Environmental restoration of germ granule function for sterile pgl-1 mutants restores their fertility. We propose that Piwi mutant sterility is a reproductive arrest phenotype that is characterized by perturbed germ granule structure and is phenocopied by germ granule dysfunction, independent of genomic instability.

Eukaryotic cells use guided search to coordinately control dispersed genetic elements. Argonaute proteins and their small RNA cofactors engage nascent RNAs and chromatin-associated proteins to direct transcriptional silencing. The small ubiquitin-like modifier (SUMO) has been shown to promote the formation and maintenance of silent chromatin (called heterochromatin) in yeast, plants, and animals. Here, we show that Argonaute-directed transcriptional silencing in Caenorhabditis elegans requires SUMOylation of the type 1 histone deacetylase HDA-1. Our findings suggest how SUMOylation promotes the association of HDAC1 with chromatin remodeling factors and with a nuclear Argonaute to initiate de novo heterochromatin silencing.

Germlines shape and balance heredity, integrating and regulating information from both parental and foreign sources. Insights into how germlines handle information have come from the study of factors that specify or maintain the germline fate. In early Caenorhabditis elegans embryos, the CCCH zinc finger protein PIE-1 localizes to the germline where it prevents somatic differentiation programs. Here, we show that PIE-1 also functions in the meiotic ovary where it becomes SUMOylated and engages the small ubiquitin-like modifier (SUMO)-conjugating machinery. Using whole-SUMO-proteome mass spectrometry, we identify HDAC SUMOylation as a target of PIE-1. Our analyses of genetic interactions between pie-1 and SUMO pathway mutants suggest that PIE-1 engages the SUMO machinery both to preserve the germline fate in the embryo and to promote Argonaute-mediated surveillance in the adult germline.

Article Figures and data Abstract Introduction Results Discussion Materials and methods Data availability References Decision letter Author response Article and author information Metrics Abstract Precise and efficient insertion of large DNA fragments into somatic cells using gene editing technologies to label or modify endogenous proteins remains challenging. Non-specific insertions/deletions (INDELs) resulting from the non-homologous end joining pathway make the process error-prone. Further, the insert is not readily removable. Here, we describe a method called CRISPR-mediated insertion of exon (CRISPIE) that can precisely and reversibly label endogenous proteins using CRISPR/Cas9-based editing. CRISPIE inserts a designer donor module, which consists of an exon encoding the protein sequence flanked by intron sequences, into an intronic location in the target gene. INDELs at the insertion junction will be spliced out, leaving mRNAs nearly error-free. We used CRISPIE to fluorescently label endogenous proteins in mammalian neurons in vivo with previously unachieved efficiency. We demonstrate that this method is broadly applicable, and that the insert can be readily removed later. CRISPIE permits protein sequence insertion with high fidelity, efficiency, and flexibility. Introduction The CRISPR/Cas9 technology has revolutionized genomic editing (Cong et al., 2013; Doudna, 2020; Heidenreich and Zhang, 2016; Jinek et al., 2012; Mali et al., 2013). However, one of the major unresolved challenges is to use CRISPR-based technologies to precisely and efficiently knock in large DNA fragments in somatic cells of living animals (i.e., without generating transgenic animals) to label or modify endogenous proteins. Such technology holds promise for both studying biological mechanisms and gene therapy. Toward this goal, several studies have developed CRISPR/Cas9-based approaches to insert fluorescent protein (FP)-encoding sequences to label and visualize endogenous proteins in somatic cells, such as postmitotic neurons (Artegiani et al., 2020; Gao et al., 2019; Mikuni et al., 2016; Nakade et al., 2014; Nishiyama et al., 2017; Schmid-Burgk et al., 2016; Suzuki et al., 2016; Uemura et al., 2016; Willems et al., 2020). Fluorescent visualization of proteins in living cells is essential for the mechanistic dissection of cellular and organismic processes because many functions of cells and organisms are established and manifested by their constituent proteins. However, conventional methods have fallen short in this regard: immunolabeling is typically incompatible with live imaging, and it usually does not permit sparse labeling for cell-specific contrast in tissues. FP tagging typically involves overexpression, which may result in undesired off-target effects. Knock-ins of FP tags are costly and time-consuming in mammalian species, and, with the exception of certain complex schemes (e.g., Fortin et al., 2014; He and Huang, 2018), are typically associated with poor contrast due to global expression of the FP-tagged protein of interest (e.g., Herzog et al., 2011). In contrast, if successful, CRISPR-based FP labeling of endogenous proteins in somatic cells may overcome these limitations and remove a major bottleneck in studying biological mechanisms. Unfortunately, current CRISPR-based somatic protein labeling technologies are faced with major limitations. First, the in vivo labeling efficiency is moderate, especially for the large DNA insertions required for FP labeling (~15% or less) (Mikuni et al., 2016; Nishiyama et al., 2017; Suzuki et al., 2016; Uemura et al., 2016). Second, and more importantly, these methods, which target protein coding sequences or their immediately adjacent exonic sequences, are error-prone. The precise insertion of short (<50 bp) DNA sequences has started to become possible via prime editing (Anzalone et al., 2019), which only nicks the DNA; however, the insertion of larger DNA fragments still requires the generation of double-stranded DNA breaks (DSBs). At the edited loci involving DSBs, including those loci where the DNA insertion does not occur, the efficient non-homologous end joining (NHEJ) pathway often results in unwanted insertions/deletions (INDELs) that cause mutations and/or frame shifts (Cong et al., 2013; Doudna, 2020; Heidenreich and Zhang, 2016; Mali et al., 2013) (see also Figure 2 and Figure 1—figure supplement 1). This is the case regardless of whether NHEJ or homologous-dependent repair (HDR) is engaged for the insertion process (Roberts et al., 2017). Finally, it has been increasingly recognized that reversible DNA editing may allow for great flexibility in both research and gene therapy (Nakamura et al., 2019); however, the inserted labels are not readily removable at a later time using existing approaches. Here, we present a strategy, called CRISPR-mediated insertion of exon (CRISPIE), which allows for the nearly error-free insertion of FP sequences with high efficiency. Instead of targeting gene exons, CRISPIE targets introns and inserts a designer donor module, which includes an exon encoding the desired protein sequence and the surrounding intronic sequences. INDELs occurring at the insertion junction within the intronic region of DNA will be spliced out (Figure 1A), resulting in very low error rates at the mRNA level (>98% correct). CRISPIE is flexible and broadly compatible with: (1) both N- and C-terminal labeling, (2) proteins with diverse structures and functions, including pre- and post-synaptic proteins and cytoskeletal proteins, (3) all major transfection methods, (4) FPs with diverse colors, and (5) multiple animal species. In part because introns offer ample editing sites to choose from, and because INDELs at the DNA level do not affect the success of editing, a high labeling efficiency (up to 43%) was achieved in cortical neurons of living mice. Importantly, CRISPIE-mediated DNA insertions are erasable. By flanking the donor module with additional designer CRISPR editing sites in the intronic region, the inserted DNA fragment can be erased at a later time. CRISPIE may allow for the routine labeling of proteins at endogenous levels and can be expanded to the insertion of other genetically encoded functional sequences to manipulate protein function. Figure 1 with 4 supplements see all Download asset Open asset The CRISPR-mediated insertion of exon (CRISPIE) strategy and its application to label human β-actin. (A) The conceptual design of the CRISPIE strategy showing that, although insertions/deletions (INDELs) and inverted insertion events may occur at the DNA level during editing, only wild-type mRNAs or mRNAs with the desired precise insertion are produced. Orange boxes: endogenous exon sequences; black lines: endogenous intron sequences; blue lines and green boxes: intron and exon sequences, respectively, of the designer donor module with white arrows showing the correct orientation of the reading frame; red stars: INDELs. (B) Schematics of the single guide RNA/Streptococcus pyogenes Cas9 (sgRNA/SpCas9) plasmid (upper) and the donor plasmid (lower) that are used for panels D–G. Purple triangles: the sgRNA-targeted location and orientation, spl.: splicing; acptr: acceptor. See Figure 1—figure supplement 3 for design details. (C) Schematic of the targeted intron of ACTB with the purple triangle showing the sgRNA targeting site and orientation. Orange and gray boxes: exonic sequences that do and do not encode protein sequences, respectively. (D) Representative two-photon (2 p) images of live U2OS cells with the indicated transfection. Note that the mRuby3 (magenta) expression levels were highly variable across cells due to the variability in plasmid transfection; however, the green label intensities were comparable across cells, as expected for the expression from an endogenous locus. (E) Labeling efficiency (green cell counts over red cell counts) for ACTB in U2OS cells. n = 8 field of views (FOVs), two independent transfections. (F) Representative confocal (Airyscan) and super-resolution structured illumination microscopy (SIM) images of live U2OS cells expressing CRISPIEd β-actin. (G) Representative time-lapse confocal images of live U2OS cells showing the dynamics of actin ruffles (arrows). The dashed yellow line outlines the cell morphology at time zero. (H) Cell growth curves of β-actin-CRISPIEd cells and untransfected cells. n = 6 FOVs. Figure 1—source data 1 Numeric data for Figure 1E. https://cdn.elifesciences.org/articles/64911/elife-64911-fig1-data1-v4.xlsx Download elife-64911-fig1-data1-v4.xlsx Results Design and demonstration of CRISPIE To achieve high-fidelity FP labeling, we developed the CRISPIE method, which inserts a designer exon module that consists of an exogenous exon encoding the FP flanked by intronic sequences, including the splicing acceptor and donor sites, into the intronic region of the target gene via the NHEJ pathway (Figure 1A, Figure 1—figure supplement 1). NHEJ exhibits a higher editing efficiency than HDR in postmitotic somatic cells, such as neurons (Heidenreich and Zhang, 2016; Heyer et al., 2010; Mao et al., 2008; Saleh-Gohari and Helleday, 2004). Because introns are spliced out of mRNAs, they are often poorly conserved and tolerate stochastic mutations better than exons (e.g., see Figure 1—figure supplement 2). INDELs at the joint junction of the insertion therefore will not result in mutations in the mRNA and thus the encoded protein under most circumstances (Figure 1A). In addition, NHEJ-mediated donor DNA insertion can result in unwanted integration with inverted orientation, and INDELs can occur at loci where the donor fails to insert. However, neither of these events will produce disrupted mRNAs when using CRISPIE (Figure 1A). Conceptually, unlike conventional exonic targeting, CRISPIE only results in either wild-type or successfully labeled mRNAs. For convenience, after a gene is labeled by CRISPIE, we say it has been 'CRISPIEd'. As a proof of principle, we first labeled human β-actin (ACTB) at its N-terminus with monomeric EGFP (mEGFP) in human U2OS cells by co-transfecting a plasmid that expresses a single guide RNA (sgRNA) targeting the ACTB gene and Streptococcus pyogenes Cas9 (SpCas9), a plasmid carrying the donor module (Figure 1B and Figure 1—figure supplement 3B1), and a transfection marker plasmid that expresses mRuby3. The sgRNA/SpCas9 complex cuts an editing site at the first intron of ACTB (Figure 1C), as selected using the BROAD Institute sgRNA designer (Doench et al., 2016), and releases the donor module from the plasmid by cutting sgRNA-targeted sequences flanking the module (Figure 1B, lower panel). The released module is then inserted into the cut site at the ACTB intron via the NHEJ pathway. The module-flanking sgRNA-targeted sequences are in the reverse orientation to facilitate the insertion of the module in the forward orientation (Suzuki et al., 2016). The donor module includes an exon in the appropriate translational phase (0–0 for intron 1 of ACTB) encoding mEGFP and peptide linkers (Figure 1—figure supplement 3B1). At the exon-intron junctions, ~100 bp of intronic sequence and ~10 bp of adjacent exonic sequence were taken from an obligatory intron (i.e., no reported alternative splicing events) and its adjacent exons of mouse CaMKIIα (Camk2a), a highly expressed protein in the brain, to include the splicing donor or acceptor site (Wang and Burge, 2008). mEGFP labeling was observed at 3–4 days post-transfection (Figure 1D) and required the presence of both sgRNA/SpCas9 and the donor module. Over 35% (37 ± 5%) of transfected cells, as identified by the expression of mRuby3 (red), were mEGFP (green)-labeled (Figure 1E). This likely represents a lower bound of labeling efficiency, since the co-transfection rate of all three plasmids is unlikely to reach 100%. Confocal and super-resolution structured illumination microscopy (SIM) revealed that the mEGFP signals exhibited the characteristic distribution of actin protein (Fischer et al., 1998; Planchon et al., 2011; Figure 1F). Time-lapse confocal microscopy revealed dynamic actin ruffles at the edges of cells (Figure 1G), which is consistent with the current knowledge of actin dynamics (Fischer et al., 1998; Planchon et al., 2011). Furthermore, cells with CRISPIEd actin grew at a rate indistinguishable from that of unlabeled cells in the same dish (Figure 1H and Figure 1—figure supplement 4). These results are consistent with the predictions for a functional, FP-labeled endogenous actin protein. CRISPIE-mediated protein insertion is nearly error-free at the mRNA level To characterize the insertion events and INDEL rates at the insertion site, we sorted transfected cells into red-only (transfected, but not labeled) and green (transfected and labeled) cells. The actin genomic DNA and mRNA at the editing locus were analyzed using PCR and RT-PCR, respectively (Figure 2A and B; see Figure 2—figure supplement 1 and Supplementary files 1 and 2 for primers, and anticipated PCR products and sizes). As expected, bands corresponding to the successful insertion (i.e., insertion in the forward orientation) at the targeted site are observed in green-labeled cells for both the genomic DNA and the mRNA (Figure 2A). Systematic analyses of green-labeled cells detected all possible editing events at the genomic DNA level, including no label insertion, forward insertion, and inverted insertion (Figure 2B), indicating that insertion events were typically non-homologous (i.e., both chromosomes differentially edited). Had the coding sequence been targeted, an inverted label insertion would inevitably cause disruptive mutations. However, under our conditions, because the donor only contained the splicing signals for exon inclusion in the forward orientation, only wild-type and forwardly inserted mRNAs were detected (Figure 2B), demonstrating the advantage of CRISPIE. Figure 2 with 2 supplements see all Download asset Open asset CRISPR-mediated insertion of exon (CRISPIE)-mediated β-actin labeling is resistant to inverted insertion events and insertions/deletions (INDELs). (A) Gel images of PCR analyses for genomic DNA and RT-PCR analyses for mRNA for untransfected (untr.) and transfected U2OS cells with (transf. +label) and without (transf. -label) successful actin labeling. As detailed in Figure 2—figure supplement 1, specific primers were used for the targeting site at the ACTB gene and mRNA with and without the desired label insertion, and for a control site (TUBA1B) that is not edited in this experiment. (B) Gel images of PCR and RT-PCR analyses for the genomic DNA and mRNA, respectively, of successfully labeled cells to detect non-labeled events, and both 5' and 3' junctions of forward and inverted label insertion events. (C) Representative next-generation sequencing (NGS) results of the genomic DNA and mRNA prepared from transfected U2OS cells without successful actin labeling corresponding to bands ii and ii', respectively. INDELs are marked in red. (D) Pie graphs of the relative proportions of INDEL events based on NGS results of the PCR/RT-PCR products from panels A and B, as marked by the Roman numerals. Statistical tests were performed using a χ2 test. From top to bottom, n = 54216, 1181, 1589, 2767, and 6036 for genomic DNA, and 5771, 2936, 4166, 3613, and 5949 for mRNA. Figure 2—source data 1 Numeric data for Figure 2D. https://cdn.elifesciences.org/articles/64911/elife-64911-fig2-data1-v4.xlsx Download elife-64911-fig2-data1-v4.xlsx To evaluate how INDELs may affect a targeted gene, the PCR amplicons corresponding to no label insertion events in untransfected, red-only, and green cells, as well as those corresponding to forward insertion events in green cells were subjected to next-generation sequencing (NGS) (Figure 2C and D; sequenced amplicons correspond to gel bands in Figure 2A and B, as indicated by Roman numerals). Remarkably, at the genomic DNA level in red-only cells, over 70% (834/1181 reads) of PCR amplicon sequences contained INDELs (Figure 2C, and second column of Figure 2D), confirming the predicted high rate of INDELs during gene editing. In contrast, at the mRNA level in these cells, 99.7% (2930/2936 reads) of the sequences did not contain INDELs (p<0.001, cf. genomic DNA) (Figure 2D, second column), which was no different from control, untransfected cells (99.7%, 5761/5771 reads) (Figure 2D, first column). Similarly, undesired INDELs were frequently detected in all sequenced amplicons from green cells at the genomic DNA level (Figure 2D). Similar high rates of INDELs were also observed at the genomic DNA level in other insertion locations of ACTB and in another gene (TUBA1B) across two different cell lines (Figure 2—figure supplement 2). However, few errors were detected at the mRNA level (INDELs 24–87% for DNA vs. 0.3–2% for mRNA; Figure 2D). These numbers correspond to correct-to-mutation ratios of 50- to 300-fold for mRNA, which are around two orders of magnitude higher than those for DNA. These results demonstrate that, although inverted insertion and INDEL events may occur at the DNA level—which would have caused mutations or frame shifts when using existing strategies to target coding sequences—CRISPIE is resistant to these disruptive events at the mRNA level. CRISPIE can be optimized to achieve high labeling efficiency For insertion strategies that target coding sequences, the options for targeting sites are limited, and not all potential targeting sites allow for high-efficiency editing (Doench et al., 2014). Furthermore, as discussed above, INDELs cause mutations or frame shifts, thereby reducing the likelihood of successful labeling. In contrast, CRISPIE is insensitive to INDELs, and the lengths of introns usually offer ample choices for potential targeting sites. CRISPIE may therefore have the potential to achieve higher labeling efficiency than strategies that target coding sequences. To test this, we first asked whether different intronic insertion sites may exhibit different labeling efficiencies. Five different editing sites (i1 to i5 in Figure 3A) were selected using the BROAD Institute sgRNA designer (with Azimuth v2.0 scores of 0.57–0.69) (Doench et al., 2016). By using a generic donor that was released from the plasmid by a designed sgRNA (called DRS-2; see Figure 1—figure supplement 3B7 for donor plasmid design) (Gao et al., 2019), we found that all five sites permitted the successful labeling of ACTB; however, the labeling efficiencies varied significantly (Figure 3B), with the highest labeling efficiency corresponding to around 20-fold of the lowest. Figure 3 Download asset Open asset CRISPR-mediated insertion of exon (CRISPIE) is optimizable and achieves higher efficiency than exon labeling. (A) Schematic positions of the tested editing sites in the intron 1 and exon 2 of ACTB that are targeted in panels B and C. The asterisk marks the site used for ACTB in subsequent experiments. (B) Comparison of the editing efficiency of five different intronic locations. For comparison under identical conditions, a generic donor excised using an independent single guide RNA (sgRNA) (DRS-2) was used (Figure 1—figure supplement 3B7). n = 18 FOVs (field of views) from two independent transfections. (C) Representative images and quantification of successful editing rates at the intronic location i4 vs. the only two possible exonic locations for N-terminally labeled ACTB, each using their specifically designed donors. n = 18 FOVs from two independent transfections. Figure 3—source data 1 Numeric data for Figure 3B and C. https://cdn.elifesciences.org/articles/64911/elife-64911-fig3-data1-v4.xlsx Download elife-64911-fig3-data1-v4.xlsx We also examined FP labeling that directly targets the coding sequence for comparison. There are only two possible SpCas9 editing sites around the start codon of ACTB (referred to as e1 and e2; Figure 3A). Despite using their optimal insertion donors (for both, see Figure 1—figure supplement 3B6), both exonic sites exhibited significantly lower labeling efficiencies compared to the optimized CRISPIE editing (greater than fivefold; Figure 3C). Between the e1 and e2 sites, the one with the higher Azimuth score was in fact less efficient (Azimuth scores = 0.55 and 0.62, respectively), indicating that the labeling efficiency is not yet entirely predictable. Nevertheless, CRISPIE allows for systematic optimization by screening for highly efficient editing sites and, when optimized, can achieve higher labeling efficiency than exonic editing. CRISPIE is broadly applicable in dividing cells The CRISPIE method should be broadly applicable for use with different FPs or with other functional domains, different protein targets, and different animal species. To test different FP labels for potential multiplex imaging studies, we successfully labeled ACTB in U2OS cells using cyan (mTurquoise2), green-yellow (mNeonGreen), and red (mRuby3) FPs (Figure 4A, donor plasmids illustrated in Figure 1—figure supplement 3B2-4). Notably, when both mTurquoise2 and mNeonGreen donors were included simultaneously, double-labeled cells could be found, which appeared to be capable of dividing (Figure 4—figure supplement 1), indicating that FP labeling does not affect cell viability in diploid CRISPIEd cells. To test the applicability of CRISPIE to different protein targets, we labeled several additional important subcellular structures in U2OS cells, namely microtubules, mitochondria, focal adhesion complexes, and the endoplasmic reticulum, by labeling tubulin alpha 1B (TUBA1B), translocase of outer mitochondrial membrane 20 (TOMM20), vinculin (VCL), and calreticulin (CALR), respectively (Figure 4B and Figure 1—figure supplement 3A and B; Lam et al., 2012; Rizzo et al., 2009; Roberts et al., 2017; Shroff et al., 2008). In a growth test, cells with CRISPIEd TUBA1B grew at a rate indistinguishable from that of unlabeled cells in the same dish (Figure 1—figure supplement 4), suggesting that such labeled TUBA1B is functional. Notably, the TOMM20 and VCL genes were labeled at their respective last introns (Figure 1—figure supplement 3A) because their encoded proteins are best tagged at their C-termini (Rizzo et al., 2009; Roberts et al., 2017; Shroff et al., 2008). This demonstrates that CRISPIE is also readily compatible with C-terminal labeling. For some genes, such as VCL, there are coding sequences of significant lengths present at the last exon, downstream of the possible CRISPIE labeling site. To prevent potential disruption of protein function caused by an insertion in the middle of the protein, the coding sequence of the last exon of VCL was included in the insertion donor sequence, and a stop codon was introduced at the end of the mEGFP coding sequence (Figure 1—figure supplement 3B). To test the applicability in different animal species, we further labeled β-actin (Actb) in mouse Neuro 2A cells. A mouse-specific sgRNA was used because introns are degenerative across species (e.g., see Figure 1—figure supplement 2). The labeling was successful and was dependent on the correct combination of the species and the corresponding sgRNA (Figure 4C). CRISPIE labeling was also successful in rat neurons (Figure 5A, see below). Overall, these results indicate that CRISPIE is versatile and readily applicable to many different FP labels, protein species, and animal species, and can be used to tag proteins at their N- and C-termini. Figure 4 with 1 supplement see all Download asset Open asset CRISPR-mediated insertion of exon (CRISPIE) is applicable to labeling with different colors, different proteins, and different animal species. (A) Representative images of ACTB labeled with each of the three indicated fluorescent proteins (FPs) of different colors in U2OS cells. (B) Representative images of four different proteins, as indicated, labeled with monomeric EGFP (mEGFP) in U2OS cells. (C) Representative images of human and mouse β-actin labeling in human U2OS cells and mouse Neuro 2A cells, respectively, under the indicated conditions. Figure 5 with 3 supplements see all Download asset Open asset CRISPR-mediated insertion of exon (CRISPIE) is applicable to different proteins in postmitotic neurons using various transfection methods. (A) Representative images of the indicated proteins labeled in rat cultured hippocampal slices transfected using the biolistic method. (B) Representative images of the indicated proteins labeled in cultured hippocampal slices from SpCas9 ± mice transfected using the biolistic method. Arrowheads indicate spines where each of the three proteins is differentially enriched or excluded. (C) Spine enrichment index (SEI) of the three proteins in panel B. From left to right, n (spines/neurons)=55/5, 26/3, and 22/2. (D, E) Representative images (D) and labeling efficiency quantifications (E) of β-actin or α-tubulin 1B-labeled neurons that are labeled in vivo via in utero electroporation (IUE) in the mouse somatosensory cortex and imaged in acute brain slices. n = 10 FOVs (field of views) from three mice for the quantification of both proteins. (F) Representative side view (x-z, left), top view (x-y, middle), and zoomed-in (x-y, right) images of labeled Actb (green) in cortical neurons that were transfected via IUE and were imaged in living mice via a cranial window. (G) Representative images of β-actin-CRISPIEd neurons in acute brain slices labeled via adeno-associated virus (AAV) injections into the cortex of SpCas9 ± mice. Figure 5—source data 1 Numeric data for Figure 5C and E. https://cdn.elifesciences.org/articles/64911/elife-64911-fig5-data1-v4.xlsx Download elife-64911-fig5-data1-v4.xlsx CRISPIE is broadly applicable to postmitotic neurons using various transfection methods in vitro and in vivo To determine whether CRISPIE can be used in postmitotic cells, such as neurons, and whether it is compatible with different transfection methods, we applied it to label proteins in neurons in cultured slices and in vivo using three common transfection methods. First, we used the biolistic method (i.e., 'gene gun') to deliver the sgRNA and donor for rat β-actin (rActb), and the cytosolic marker mRuby3, to cultured rat hippocampal slices. Notably, although the biolistic transfection method is widely used in studying the cellular functions of neurons, CRISPR-based labeling has not yet been demonstrated using this method, likely due in part to the relatively low labeling efficiencies. For this same reason, high-contrast double labeling—for example, the target protein together with a cell morphology marker—in brain tissue remains challenging. In our experiments, 6–45 neurons were transfected per brain slice, as determined by their red mRuby3 fluorescence. Among them, on average 15% (15 ± 4%, n = 6 slices) exhibited successful labeling of actin (Figure 5A). In some slices, as many as six to seven successfully labeled neurons of different neuronal types, including dentate gyrus neurons, CA1 and CA3 pyramidal neurons, and interneurons, could be detected (Figure 5—figure supplement 1). Glial cells could also be labeled (Figure 5—figure supplement 1B). The labeled neurons exhibited normal miniature excitatory postsynaptic currents (mEPSCs) in frequency and amplitude when compared to adjacent untransfected neurons (Figure 5—figure supplement 2). These results indicate that CRISPIE provides a viable option for protein labeling using gene gun in cultured slices. Similar results were obtained in cultured mouse hippocampal slices from an SpCas9 heterozygous genetic background, which constitutively expresses SpCas9 and thereby simplified the experimental design (Figure 5B). Other synaptic proteins, including synapsin (SYN1), tubulin (TUBA1B), and CaMKIIα, were also successfully labeled (Figure 5A and B). In many labeled cells, both the mRuby3 cytosolic marker and mEGFP were readily detectible, allowing us to compare the subcellular distribution of three proteins—β-actin, α-tubulin 1B, and CaMKIIα—in neuronal dendrites (Figure 5C). By quantifying their distribution between the dendrites and spines using a previously described spine enrichment index (SEI) (Zhong et al., 2009), we found that the three proteins exhibited distinct subcellular distributions: actin was highly enriched in dendritic spines, and CaMKIIα was moderately enriched, whereas tubulin was excluded from spines (Figure 5C; see Materials and methods for quantification details). To determine whether CRISPIE can be used to label endogenous proteins in vivo, in utero electroporation (IUE) was used to introduce the sgRNA, the donor, and the cytosolic marker mRuby3 into the somatosensory cortex of SpCas9 mice to label β-actin or tubulin in separate experiments (Figure 5D). Among the transfected red cells, the labeling efficiencies were 43 ± 5% for actin and 30 ± 5% for tubulin (Figure 5E), both of which are significantly higher than previously reported for CRISPR-based endogenous labeling of any protein in vivo (Mikuni et al., 2016; Nishiyama et al., 2017; Suzuki et al., 2016; Uemura et al., 2016). This high level of labeling efficiency allowed us to routinely perform in vivo imaging in the brains of living mice (Figure 5F and Figure 5—figure supplement 3). Finally, as viruses have become an increasingly prevalent method of transfection, we generated an adeno-associated virus (AAV) harboring the sgRNA targeting β-actin, the donor, and mRuby3 (Figure 1—figure supplement 3C). When injected into the cortex of SpCas9 mice, this virus successfully transduced neurons in vivo and labeled the actin cytoskeleton (Figure 5G). Overall, CRISPIE can be readily applied using a variety of widely used methods to

Germlines shape and balance heredity, integrating and regulating information from both parental and foreign sources. Insights into how the germline handles information have come from the identification of factors that specify or maintain the germline fate. In early embryos, the CCCH zinc-finger protein PIE-1 localizes to the germline where it prevents somatic differentiation programs. Here we show that PIE-1 also functions in the meiotic ovary where it becomes SUMOylated and engages the SUMO-conjugating machinery. Using whole-proteome mass spectrometry to detect SUMO-conjugated proteins, we identify HDAC SUMOylation as a target of PIE-1. Our findings suggest that SUMOylation activates HDAC, lowering histone acetylation and enhancing Argonaute-mediated surveillance in the germline.

ABSTRACTCRISPR genome editing has revolutionized genetics in many organisms. In the nematodeCaenorhabditis elegansone injection into each of the two gonad arms of an adult hermaphrodite exposes hundreds of meiotic germ cells to editing mixtures, permitting the recovery of multiple indels or small precision edits from each successfully injected animal. Unfortunately, particularly for long insertions, editing efficiencies can vary widely, necessitating multiple injections, and often requiring co-selection strategies. Here we show that melting double stranded DNA (dsDNA) donor molecules prior to injection increases the frequency of precise homology-directed repair (HDR) by several fold for longer edits. We describe troubleshooting strategies that enable consistently high editing efficiencies resulting, for example, in up to 100 independent GFP knock-ins from a single injected animal. These efficiencies makeC. elegansby far the easiest metazoan to genome edit, removing barriers to the use and adoption of this facile system as a model for understanding animal biology.

Melting and fast cooling double stranded DNA donor molecules prior to injection dramatically increases the frequency of homology-directed repair for edits such as insertions of fluorescent protein markers in Caenorhabditis elegans. Strategies described here enable consistently ..... CRISPR genome editing has revolutionized genetics in many organisms. In the nematode Caenorhabditis elegans, one injection into each of the two gonad arms of an adult hermaphrodite exposes hundreds of meiotic germ cells to editing mixtures, permitting the recovery of multiple indels or small precision edits from each successfully injected animal. Unfortunately, particularly for long insertions, editing efficiencies can vary widely, necessitating multiple injections, and often requiring coselection strategies. Here, we show that melting double-stranded DNA (dsDNA) donor molecules prior to injection increases the frequency of precise homology-directed repair (HDR) by several fold for longer edits. We describe troubleshooting strategies that enable consistently high editing efficiencies resulting, for example, in up to 100 independent GFP knock-ins from a single injected animal. These efficiencies make C. elegans by far the easiest metazoan to genome edit, removing barriers to the use and adoption of this facile system as a model for understanding animal biology.

SUMMARY Eukaryotic cells regulate 5’ triphosphorylated (ppp-) RNAs to promote cellular functions and prevent recognition by antiviral RNA sensors. For example, RNA capping enzymes possess triphosphatase domains that remove the γ phosphates of ppp-RNAs during RNA capping. Members of the closely related PIR1 family of RNA polyphosphatases remove both the β and γ phosphates from ppp-RNAs. Here we show that C. elegans PIR-1 dephosphorylates ppp-RNAs made by cellular RdRPs and is required for the maturation of 26G-RNAs, Dicer-dependent small RNAs that regulate thousands of genes during spermatogenesis and embryogenesis. PIR-1 also regulates the CSR-1 22G-RNA pathway and has critical functions in both somatic and germline development. Our findings suggest that PIR-1 modulates both Dicer-dependent and - independent Argonaute pathways, and provide insight into how cells and viruses use a conserved RNA phosphatase to regulate and respond to ppp-RNA species.

Summary Germlines shape and balance heredity, integrating and regulating information from both parental and foreign sources. Insights into how the germline handles information have come from the identification of factors that specify or maintain the germline fate. In early C. elegans embryos, the CCCH zinc-finger protein PIE-1 localizes to the germline where it prevents somatic differentiation programs. Here we show that PIE-1 also functions in the meiotic ovary where it becomes SUMOylated and engages the SUMO-conjugating machinery. Using whole-proteome mass spectrometry to detect SUMO-conjugated proteins, we identify HDAC SUMOylation as a target of PIE-1. Our findings suggest that SUMOylation activates HDAC, lowering histone acetylation and enhancing Argonaute-mediated surveillance in the germline.

GCNA proteins are expressed across eukarya in pluripotent cells and have conserved functions in fertility. GCNA homologs Spartan (DVC-1) and Wss1 resolve DNA-protein crosslinks (DPCs), including Topoisomerase-DNA adducts, during DNA replication. Here, we show that GCNA mutants in mouse and C. elegans display defects in genome maintenance including DNA damage, aberrant chromosome condensation, and crossover defects in mouse spermatocytes and spontaneous genomic rearrangements in C. elegans. We show that GCNA and topoisomerase II (TOP2) physically interact in both mice and worms and colocalize on condensed chromosomes during mitosis in C. elegans embryos. Moreover, C. elegans gcna-1 mutants are hypersensitive to TOP2 poison. Together, our findings support a model in which GCNA provides genome maintenance functions in the germline and may do so, in part, by promoting the resolution of TOP2 DPCs.

piRNAs (Piwi-interacting small RNAs) engage Piwi Argonautes to silence transposons and promote fertility in animal germlines. Genetic and computational studies have suggested that C. elegans piRNAs tolerate mismatched pairing and in principle could target every transcript. Here we employ in vivo cross-linking to identify transcriptome-wide interactions between piRNAs and target RNAs. We show that piRNAs engage all germline mRNAs and that piRNA binding follows microRNA-like pairing rules. Targeting correlates better with binding energy than with piRNA abundance, suggesting that piRNA concentration does not limit targeting. In mRNAs silenced by piRNAs, secondary small RNAs accumulate at the center and ends of piRNA binding sites. In germline-expressed mRNAs, however, targeting by the CSR-1 Argonaute correlates with reduced piRNA binding density and suppression of piRNA-associated secondary small RNAs. Our findings reveal physiologically important and nuanced regulation of individual piRNA targets and provide evidence for a comprehensive post-transcriptional regulatory step in germline gene expression.