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Meiotic Cas9 expression mediates gene conversion in the male and female mouse germline

['Alexander J. Weitzel', 'Division Of Biological Sciences', 'Section Of Cell', 'Developmental Biology', 'University Of California San Diego', 'La Jolla', 'California', 'United States Of America', 'Hannah A. Grunwald', 'Ceri Weber']

Date: 2022-01 

Abstract Highly efficient gene conversion systems have the potential to facilitate the study of complex genetic traits using laboratory mice and, if implemented as a “gene drive,” to limit loss of biodiversity and disease transmission caused by wild rodent populations. We previously showed that such a system of gene conversion from heterozygous to homozygous after a sequence targeted CRISPR/Cas9 double-strand DNA break (DSB) is feasible in the female mouse germline. In the male germline, however, all DSBs were instead repaired by end joining (EJ) mechanisms to form an “insertion/deletion” (indel) mutation. These observations suggested that timing Cas9 expression to coincide with meiosis I is critical to favor conditions when homologous chromosomes are aligned and interchromosomal homology-directed repair (HDR) mechanisms predominate. Here, using a Cas9 knock-in allele at the Spo11 locus, we show that meiotic expression of Cas9 does indeed mediate gene conversion in the male as well as in the female germline. However, the low frequency of both HDR and indel mutation in both male and female germlines suggests that Cas9 may be expressed from the Spo11 locus at levels too low for efficient DSB formation. We suggest that more robust Cas9 expression initiated during early meiosis I may improve the efficiency of gene conversion and further increase the rate of “super-mendelian” inheritance from both male and female mice.

Citation: Weitzel AJ, Grunwald HA, Weber C, Levina R, Gantz VM, Hedrick SM, et al. (2021) Meiotic Cas9 expression mediates gene conversion in the male and female mouse germline. PLoS Biol 19(12): e3001478. https://doi.org/10.1371/journal.pbio.3001478 Academic Editor: Bon-Kyoung Koo, IMBA, AUSTRIA Received: April 20, 2021; Accepted: November 10, 2021; Published: December 23, 2021 Copyright: © 2021 Weitzel et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited. Data Availability: All primary Sanger sequence traces that confirm the Spo11Cas9-P2A-eGFP knock-in allele and all genotyping data for offspring of the complementation test crosses are available at Zenodo with the identifier 10.5281/zenodo.5510697. Annotated sequence data for the Spo11Cas9-P2A-eGFP knock-in allele is available in GenBank with the accession number OK067273. Funding: This work was funded by a Packard Fellowship in Science and Engineering, award number 2015-63114, from the David and Lucile Packard Foundation and NIH grant R21GM129448 awarded to KLC. The work was also supported by NIH grant R01AI131081 awarded to SMH and an Allen Frontiers Group Distinguished Investigators Award to EB. Work in the laboratories of KLC and EB was also supported by a gift from the Tata Trusts in India to TIGS-UCSD and TIGS-India. AJW and HAG were supported by NIH training grant T32GM007240, and RL was supported by NIH training grant T32GM133351. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. Competing interests: I have read the journal’s policy and the authors of this manuscript have the following competing interests: KLC, VMG, SMH, and EB serve as science advisory board members for Synbal, Inc. All other authors declare they have no competing interests. Abbreviations: DSB, double-strand DNA break; E16.5, embryonic day 16.5; EJ, end joining; gRNA, guide RNA; HDR, homology-directed repair; IF, immunofluorescence; MMEJ, microhomology-mediated end joining; NHEJ, nonhomologous end joining; OCT, optimal cutting temperature

Introduction The mouse remains the single most utilized laboratory model of human physiology and disease. Yet, breeding strategies that combine multiple homozygous transgenic or knock-in alleles are cumbersome in mice due to small litter sizes and long generation times relative to other traditional model systems. Practical challenges therefore limit the full potential for mouse genetic research to model certain complex human genetic disorders and to engineer models that better mimic human metabolism and immunity for therapeutic drug design. We previously demonstrated that super-mendelian inheritance mediated by CRISPR/Cas9 is feasible through the female germline of mice, increasing the transmission frequency of a transgenic allele [1]. Similar systems have been proposed for implementation as a “gene drive” over multiple generations in wild rodents to limit the spread of infectious disease or to reduce populations of invasive mice and rats in sensitive island ecosystems [2–6]. To assess the feasibility of CRISPR/Cas9–mediated gene conversion, we previously engineered a “CopyCat” transgene that disrupts the fourth exon of Tyrosinase, a gene that is required for melanization in mice [7]. The TyrCopyCat transgene expresses the single guide RNA (gRNA) used to target the site of its own insertion and an mCherry fluorescent marker to track its inheritance. We then used available genetic tools to express Cas9 in the early embryo and in the male and female germline. There are two “nonconservative” repair outcomes of CRISPR/Cas9–mediated double-strand DNA breaks (DSBs) that alter the wild-type target sequence in a TyrCopyCat heterozygous animal. First, the two ends may be rejoined by nonhomologous end joining or microhomology mediated end joining [NHEJ or MMEJ, hereafter referred to as “end joining” (EJ)], which can cause insertion or deletion mutations (indels) that disrupt the gRNA target site and prevent subsequent cutting (reviewed in [8]). Alternatively, the DSB can be repaired by homology-directed repair (HDR) using genomic sequences that flank the TyrCopyCat transgene on the homologous chromosome, referred to as “gene conversion” or “homing.” If interchromosomal HDR occurs, then the transgene that was inherited on only one chromosome is copied into the DSB of the homologous locus such that its genotype is converted from heterozygosity to homozygosity [1]. CRISPR/Cas9–mediated gene conversion is highly efficient (>90%) in Dipteran insects and in yeast [9–12]. Using multiple Cre/lox strategies to initiate Cas9 expression in oogonia and spermatogonia of mice [13,14], we previously demonstrated gene conversion in the female but not in the male germline [1]. The frequency of TyrCopyCat allele transmission increased from 50%, predicted by mendelian inheritance, to an average of 72% among offspring of 5 females after CRISPR/Cas9–mediated gene conversion in the germline. We also recombined ultra-tightly linked loci (approximately 9 kb apart) by gene conversion in 22.5% of offspring of females. Although this was an important demonstration of the powerful potential of such approaches, use of two transgenes (Cre and conditional Cas9) to control the timing of Cas9 expression is not an optimal approach to simplify complex crossing schemes. The observation that gene conversion occurred exclusively in the female germline suggested that sex differences in the progression of gonial development might be key. Oogonia rapidly mature into primary oocytes that initiate meiosis during fetal development [15], while spermatogonia are mitotic throughout the life of the animal and sporadically produce cohorts of meiotic primary spermatocytes [16,17]. In contrast to mitosis, homologous chromosomes are aligned during meiosis I, and the molecular machinery that facilitates interchromosomal HDR predominates over EJ [18]; either or both of these conditions might allow gene conversion to occur in meiotic but not in mitotic cells. We therefore interpreted our data to suggest that timing Cas9 expression to coincide with meiosis I may be critical to allow for gene conversion in both sexes. Here, we use a single transgene to evaluate the importance of initiating Cas9 expression during prophase of meiosis I for gene conversion to occur in mice. We engineered a knock-in allele at the Spo11 locus to express Cas9 in conjunction with endogenous SPO11 using a P2A self-cleaving peptide (Fig 1A). Spo11 is the endonuclease and topoisomerase that catalyzes DSB formation required for crossing over during meiotic recombination in prophase of meiosis I [19,20]. For this analysis, the timing but not the catalytic activity of Spo11 expression is important. In mammals, Spo11 is expressed only in mature testes and in embryonic ovaries, and its expression peaks during prophase of meiosis I when homologous chromosomes are aligned [21,22]. PPT PowerPoint slide

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TIFF original image Download: Fig 1. Male and female germ cell characterization of Spo11Cas9-P2A-eGFP knock-in expression. (A) Schematic of targeting the Spo11Cas9-P2A-eGFP knock-in allele. (i) Wild-type Spo11 locus. (ii) Construct for insertion. (iii) Final Spo11Cas9-P2A-eGFP knock-in locus. The genotyping strategy is depicted in S7 Fig. (B–G) Immunofluorescent detection of eGFP (green) and VASA (magenta) plus DAPI (blue) in Spo11+/+ (B, E) and Spo11Cas9-P2A-eGFP/+ (C,D; F,G) mice. (B–D) embryonic ovaries at E18.5 (n = 3) and (E–G) adult testes (n = 3). Seminiferous tubules labeled 1 -> 3 are in different stages of maturation as determined by chromatin compaction (DAPI) and relative level of VASA expression in outer perimeter cells. The inset shows outer perimeter cells of tubule 1 (arrowhead points to spermatogonium) that express low VASA and no eGFP. By contrast, the outer perimeter cells of tubule 2 (meiotic primary spermatocytes) express higher VASA and low eGFP, and outer perimeter cells of tubule 3 express the highest eGFP. Scale bar: 50 μm for (B–G) and 25 μm for the insets in (D) and (G). eGFP channel brightness was adjusted in (B–D) independent of (E–G); equivalent exposures and all biological replicates are shown in S4 Fig. https://doi.org/10.1371/journal.pbio.3001478.g001 Using the same TyrCopyCat allele and crossing scheme to assess gene conversion efficiency as in our prior work [1], we demonstrate that Cas9 expression driven by Spo11 during meiosis I promotes gene conversion in the male as well as female germline. Compared to our most efficient prior strategy, the relative fraction of detectable DSB repair that is mediated by HDR [(HDR/(HDR + EJ)] is increased in the male (from 0% to 11%) and is similar to what we previously observed in the female (71% versus 67%) consistent with our hypothesis that restricting DSB repair to meiosis I favors HDR. However, total rates of DSB repair (HDR + EJ) are lower in both sexes resulting in absolute gene conversion frequencies that are low in males and reduced in females compared to previous strategies. We suggest that the level of Cas9 expression driven from the Spo11 locus may be too low for efficient DSB formation, although the timing of expression overlaps the window when interchromosomal HDR does occur. These results demonstrate that gene conversion is possible in the male mouse germline and emphasize the importance of both timing and levels of Cas9 expression for efficient gene conversion systems in rodents.

Discussion Although moderately efficient gene conversion in one parent might be sufficient to increase the probability of obtaining complex multilocus genotypes, such approaches would be vastly improved by gene conversion in both females and in males using a single Cas9 transgene [28]. A previous study employing direct expression from a single Vasa-Cas9 transgene did not facilitate detectable gene conversion, although the recipient chromosome was not marked [29]. Here, using a Spo11Cas9-P2A-eGFP transgene, we have demonstrated the first reported gene conversion in the male mouse germline. We also show that gene conversion as a proportion of all repair events in the female germline is similar to our prior strategy. However, total repair events are substantially reduced in both sexes. Together with our previous findings, our results suggest that the overall gene conversion efficiency is dependent upon two key factors: the absolute rate of nonconservative DSB repair [(HDR + EJ)/total Tyrch] and the relative rate of repair that is HDR [HDR/(HDR + EJ)] shown in Fig 3. Here, we synthesize all of our findings to suggest a model wherein these two rates are influenced by the cumulative amount of Cas9 expression and by the timing of Cas9 expression initiation, respectively. We suggest that both levels and timing of Cas9 dictate the dynamics of Cas9–mediated DSB formation and repair in context of the varied cellular and nuclear environments of male and female mitotic and meiotic germ cells. The kinetics and mechanisms of DNA break repair have been historically described in the context of ionizing radiation and restriction enzyme-induced DSBs that are frequently repaired to preserve the original allele [30–35]. Recent work has shown that Cas9 induced DSBs may be repaired differently, as the Cas9 protein may remain bound after a DSB is induced and can thus prevent canonical repair machinery from accessing exposed DSBs [36–38], although local chromatin environments can affect Cas9 binding kinetics [39,40]. These mechanics may explain observations that Cas9–induced DSB repair is substantially more error prone than repair after other means of DSB induction [38]. Although DSBs can be infrequently repaired by homologous recombination between sister chromatids after mid to late pachytene of meiosis I [41], the very high frequency of conserved alleles from our crosses suggests that most were never cut as opposed to cut and correctly repaired. Therefore, nonconservatively repaired alleles are a reasonable proxy for total rate of DSB generation. These studies inform our interpretation of observed differences in the absolute rate of DSB repair in the context of genetically encoded Cas9. In our previous strategies, after Vasa-Cre mediated removal of the lox-STOP-lox, Cas9 was expressed under regulatory control of the strong constitutive CAG promoter at either of the permissive H11 or Rosa26 loci. These strategies generated a nonconservative allele in 100% of Tyrch male germ cells (Fig 3A, all EJ) and in 60% to 76% of female germ cells [1] (Fig 3B, HDR + EJ). By contrast, the absolute rate of DSB repair in this meiotic Spo11Cas9-P2A-eGFP strategy is substantially lower at 29% (76/263) in male (Fig 3A) and 8% (6/73) in female meiotic germ cells (Fig 3B). Previous studies demonstrated that Cas9–induced DSBs follow the dynamics of mass action; across a fixed window of time, total DSBs increase proportionally to the amount of Cas9 [42], and a higher proportion of DSBs occur earlier in the presence of higher levels of Cas9 [43]. Thus, we hypothesize that the lower absolute rate of DSB repair in the Spo11Cas9-P2A-eGFP strategy may reflect lower Cas9 expression from endogenous Spo11 regulatory sequences than from the constitutive CAG promoter at a permissive locus. We note that the difference in absolute rate of DSB formation in males and females of the Spo11Cas9-P2A-eGFP strategy correlates with a higher level of eGFP detected in male meiotic germ cells (S4 Fig). This observed difference is also consistent with previously reported quantification of Spo11 mRNA expression by RT-qPCR, which shows a higher expression level in males than in females [21]. It is essential to note that the total level of Cas9 protein in a cell is dependent not only on the strength of a regulatory sequence, but also on the duration of expression. The 100% absolute rate of DSB repair in the Vasa-Cre;LSL-Cas9 male germline strategies may have resulted from persistent Cas9 expression over several months in the population of self-renewing mitotic spermatogonia (Fig 3A’, blue clock). Such a long duration of Cas9 expression would provide ample time for prevalent DSB repair by EJ; resulting indels would be inherited by primary spermatocytes, thus blocking any subsequent DSB formation and gene conversion at the onset of meiosis. By contrast, the Spo11Cas9-P2A-eGFP transgene delays Cas9 expression by months to be initiated first in meiotic primary spermatocytes (Fig 1G and Fig 3A’, orange line), and expression termination is controlled by the endogenous Spo11 locus. After rising during meiosis I, Spo11 levels drop precipitously in maturing spermatids [22]. Together, the above considerations suggest that the absolute rate of DSB formation correlates with a cumulative level of Cas9 expression that is likely lower in the Spo11Cas9-P2A-eGFP strategy than in the Vasa-Cre;LSL-Cas9 strategies and lower in Spo11Cas9-P2A-eGFP females than in males. In contrast to the absolute rate of DSB repair, we hypothesize that the relative rate of HDR [HDR/(HDR + EJ)] is primarily a function of the timing of Cas9 activity rather than the amount of Cas9 protein. In meiotic cells, particularly in early prophase I (leptotene through mid-pachytene), EJ-specific machinery is not detected; homologous recombination becomes the exclusive repair pathway and predominantly occurs between homologous chromosomes [44,45]. Later in prophase I, after mid-pachytene, EJ again becomes the primary DSB repair choice [18]. Second, the single homologous chromosome that is the template for gene conversion is far more likely to be aligned and accessible during meiosis I, and machinery is expressed that limits recombination to occur between homologous chromosomes [18,46]. From these data, we hypothesize that a permissive window for gene conversion by interchromosomal HDR opens in leptotene and closes in mid-pachytene of meiosis I (Fig 3A’ and 3B’, red shading), whereas DSBs induced before or after this window may be more likely repaired by EJ. Since SPO11 is necessary for meiotic homologous recombination, our knock-in allele that appears to unintentionally reduce its function could cause lower overall rates of HDR, including gene conversion, during this window. However, we note that Spo11 is haplosufficient in mice [23]. The similar relative rate of HDR in females of both genetic strategies may reflect a narrow temporal difference between the onset of Cas9 expression in the Spo11Cas9-P2A-eGFP strategy compared to the Vasa-Cre;LSL-Cas9 strategies, since Vasa-Cre is expressed in oogonia that rapidly mature into meiotic oocytes. In the male Vasa-Cre;LSL-Cas9 strategies, by contrast, DSBs may have been formed and repaired by EJ in most or all mitotic spermatogonia prior to opening the permissive window. Using the Spo11Cas9-P2A-eGFP that initiates expression in meiotic primary spermatocytes may bring the onset of Cas9 expression closer to the permissive window. Therefore, temporally shifting Cas9 to the onset of meiosis in both sexes likely resulted in disproportionate improvement of the relative rate of HDR in males versus females. Our results mark a significant advance in the feasibility of super-mendelian inheritance strategies in rodents by achieving germline gene conversion in both sexes. We propose that adjusting the onset and increasing the level of Cas9 expression can further improve the absolute rate of DSB repair and the proportion of repair that is gene conversion by HDR in both males and females. This may be achieved by identifying promoters that express at a high level and that fall within or immediately prior to the permissive window for HDR during meiosis I and engineering transgenes that cannot disrupt endogenous gene function. An alternative approach might be to use a promoter that drives strong expression with less temporal specificity and to chemically control the timing of Cas9 activity (e.g., with tamoxifen or trimethoprim [47]). Both wild release applications to constitute a “gene drive” and laboratory applications to facilitate breeding complex genotypes will benefit from further increasing the efficiency of gene conversion by improved genetic strategies. However, improvements that require drug delivery would not be practical for implementation in the wild. Increasing the absolute rate of DSB repair versus the relative rate of HDR will have different effects on improving gene drive versus laboratory applications. Gene drive efficiency can tolerate a lower absolute rate of repair as long as the proportion of repair that is HDR is very high. Otherwise systems must be designed to select against indels that would be resistant to subsequent gene conversion (reviewed in [48]). In a laboratory setting, where the researcher can genotype to choose animals, a higher rate of EJ is more tolerable so long as the rate of HDR is sufficient to obtain genotypes of interest. While showing that CRISPR/Cas9–mediated gene conversion is possible in both male and female mouse germlines, our work reveals nuances that differ from insects and that must be considered for further refinement and implementation in rodents.

Methods Animals Production of the TyrCopyCat transgenic mice was previously described in [1]. Tyrchinchilla mice are strain FVB.129P2-Pde6b+ Tyrc-ch/AntJ, stock number 4828, from the Jackson Laboratory (ME, USA). Tyrnull mice are strain CD-1, stock number 022, from Charles River Laboratories (MA, USA). The Spo11Cas9-P2A-eGFP knock-in mice were produced by Biocytogen (MA, USA) using CRISPR/Cas9 to insert P2A-Cas9-P2A-GFP flanked by 1.5-kb arms of homology to Spo11 exon 13 and the 3′ UTR. Mice were housed in accordance with UCSD Institutional Animal Care and Use Committee protocols and fed on a standard breeders diet. All animal research was conducted under UCSD Animal Welfare Assurance number D16-00020 and approved by Institutional Animal Care and Use Committee protocol number S14014. Humane euthanasia was carried out by low-flow exposure to CO 2 until loss of consciousness followed by cervical dislocation as a secondary method. Sample collection, immunofluorescence, histology, imaging, and western blotting We collected testes from adult male Spo11Cas9-P2A-eGFP/+ and wild-type mice. To obtain meiotic ovaries, we crossed Spo11Cas9-P2A-eGFP/+ males to CD-1 females. Once fertilization was inferred by detection of a copulatory plug, the female was separated from the breeding male. At the desired embryonic stage, we dissected embryos from the uterus, confirmed the embryonic stage by limb morphological stage [49], bisected the embryo in transverse anterior to the ovaries, and collected the tail tip for PCR genotyping to establish sex and presence of the Spo11Cas9-P2A-eGFP transgene. Embryos were fixed in 4% PFA overnight. The following day, they were transferred to 20% sucrose solution until saturated. Embryos were embedded in optimal cutting temperature (OCT) blocks and sectioned at 6-μm thickness. Slides were frozen at -80°C until processing. Slides were removed from −80°C and dried. We performed antigen retrieval for 10′ in 5 μg/ml Proteinase K at room temperature. We then fixed in 4%PFA 1xPBS for 5′ and washed 3× for 10′ in 1x PBS. Slides were then blocked (3% BSA, 5% NGS, 0.1% Triton, 0.02% SDS, in 1X PBS) for 1 to 2 hours at room temperature before addition of primary antibodies diluted in block [chicken anti-eGFP, 1:500, Thermo Fisher catalog # A10262; rabbit anti-human Ddx4 (VASA), 1:250, Abcam catalog # AB13840] and incubated at 4°C for 18 hours. Slides were then washed 3× for 30′ in PBST (1X PBS, 0.1% Triton) prior to incubation in secondary antibodies [Invitrogen (MA, USA), Alexa-488 goat anti-chicken and Life Technologies (CA, USA), 647 goat anti-rabbit, 1:250 each] in block with 300 nM DAPI [Cell Signaling Technology (MA, USA), catalog # 4083S] and incubated at 4°C for 18 hours. We then washed 3× for 30′ in PBST and mounted under coverslips in Vectashield (Vector Labs, CA, USA). Images were collected on Leica TCS SP8 multiphoton confocal microscope. Excitation frequency GFP (488 nm): emission range 505 to 550 nm. Excitation frequency Vasa (647 nm): emission range 655 to 685 nm. Excitation frequency DAPI (405 nm): emission range 415 to 470 nm. The pixel intensity was determined in FIJI by ZProjection “average intensity” for each channel. The brightness of eGFP signal intensity was adjusted equivalently across all samples (Fig 1, S4 Fig) with the exception of the testis in Fig 1F and 1G, which was reduced for clarity. For hematoxylin and eosin staining of slides, the staining agent was Harris hematoxylin solution. Samples were differentiated in 1% acid alcohol. The bluing agent was saturated lithium carbonate and slides were counterstained in a solution of 0.1% eosin Y and 0.1% phloxine B in acid alcohol. The samples were then dehydrated to 100% EtOH and cleared in xylenes. The slides were mounted under coverslips in Permount and cured for 24 hours. Samples were imaged at 20× magnification under brightfield on an Olympus (Tokyo, Japan), BX61 microscope. Mouse testis lysate was acquired from dissections of mice at approximately 6 months of age (n = 3 wild type, n = 3 Spo11-Cas9-eGFP heterozygotes) and placed into a lysis solution (8 M Urea, 75 mM NaCl, 50 mM Tris pH 8.0, 1 mM NaF, 1 mM beta-Gly). Each sample was dounce homogenized at 4°C, sonicated 3× 10 seconds at 4°C, and centrifuged at 4°C for 10 minutes at 15,000 g. Protein quantification was achieved via Pierce BCA Protein Assay Kit (Thermo Fisher (MA, USA), #23225). For western blotting, three 4% to 20% MP TGX Stain-Free Gel (Bio-Rad # 4568093) were loaded with Precision Plus (Bio-Rad (CA, USA), # 1610374), MagicMark (Thermo Fisher #LC5602), and 30 ug of testis lysate. Each gel underwent elecrophoresis for 1.5 hours at 30 mA, 100 V, and transferred to a 0.2 uM nitrocellulose membrane at 60 V with variable ampere at 4°C for 1.5 hours. Membranes were Ponceau stained to verify transfer and rinsed 2x in ddH2O. For blocking, membranes were washed for 5 minutes at RT with 1X TBS, incubated in block (1X TBS, 0.1% Tween, 5% nonfat dry milk) for 1 hour at RT and washed 3× in TBST (1X TBS, 0.1% Tween) for 5 minutes. For detection using mouse anti-Cas9 in mouse tissues, the block included 20 ug/mL Affinipure Fab Fragment (Jackson ImmunoResearch (PA, USA), catalog # 115-007-003) to reduce background detection of mouse IgG. Primary antibodies were diluted [rabbit anti-Spo11, 1:500, Sigma Aldrich catalog # MABE1167; chicken anti-GFP 1:1,000, Thermo Fisher catalog # A10262; mouse anti-Cas9 1:1,000, Cell Signaling catalog # 14697] in TBST with 5% nonfat dry milk and incubated with membranes for 18 hours at 4°C followed by washes 3× for 5 minutes in TBST. HRP-conjugated secondary antibodies were diluted [anti-rabbit 1:1,000, Cell Signaling catalog # 7074; anti-chicken 1:500, Thermo Fisher catalog # A16054; anti-mouse 1:1,000, Cell Signaling catalog # 7076] in TBST, and membranes were incubated for 1 hour at RT. Detection was accomplished using the Clarity Max ECL Substrate (Bio-Rad #1705060), and membranes were imaged on a Bio-Rad ChemiDoc XRS+ Gel Imaging System. Genotyping The Spo11Cas9-P2A-eGFP knock-in allele was detected as in S7 Fig using primers and conditions in S1 Table. We detected inheritance of the TyrCopyCat transgene by observing red fluorescence of mCherry (S5 Fig). We then genotyped all animals for the Tyrch allele by PCR amplification of a 392-bp region containing the SNP followed by Sanger Sequencing (GeneWiz, CA, USA). Finally, we confirmed the TyrCopyCat genotype in each Tyrch animal using PCR primers that bind inside and outside of the transgene to amplify an 838-bp fragment only from transgenic animals (S6 Fig). PCR genotypes were performed according to conditions set in S1 Table. The Spo11Cas9-P2A-eGFP knock-in allele was confirmed to be the correct sequence and integrated into the correct genomic location by 9 tiled PCR amplifications (S1 Fig) using primers and conditions in S1 Table. The flanking amplicons, LHom and RHom, were each amplified using a primer that annealed inside the transgenic element (either the body of Cas9 or the body of eGFP) and a primer outside the homology arm in the genomic locus to confirm proper site-specific integration. Each of the 9 amplicons were then Sanger sequenced for verification (GeneWiz). All raw Sanger sequence traces are available at the associated Zenodo archive.

Acknowledgments We thank H. Cook-Andersen, M. Wilkinson, and R. Knight for discussions of strategy and meiotic drivers. V. Ruthig provided advice on germline immunofluorescence and staging of ovaries and testes. We thank D. Yelon for use of the Leica TCS SP8 multiphoton confocal microscope.

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