Multiple applications of genome editing by CRISPR-Cas9 necessitate stringent regulation and Cas9 variants have accordingly been generated whose activity responds to small ligands, temperature or light. However, these approaches are often impracticable, for example in clinical therapeutic genome editing in situ or gene drives in which environmentally-compatible control is paramount. With this in mind, we have developed heritable Cas9-mediated mammalian genome editing that is acutely controlled by the cheap lysine derivative, Lys(Boc) (BOC). Genetic code expansion permitted non-physiological BOC incorporation such that Cas9 (Cas9BOC) was expressed in a full-length, active form in cultured somatic cells only after BOC exposure. Stringently BOC-dependent, heritable editing of transgenic and native genomic loci occurred when Cas9BOC was expressed at the onset of mouse embryonic development from cRNA or Cas9BOC transgenic females. The tightly controlled Cas9 editing system reported here promises to have broad applications and is a first step towards purposed, spatiotemporal gene drive regulation over large geographical ranges.
Many applications of genome editing by CRISPR-Cas91,2,3,4 would benefit from being tightly controllable to minimize unintended, potentially harmful genome cleavage due to leaky nuclease activity. Such regulation would likely impact multiple contexts, including restricting genome editing to given anatomical sites in clinical somatic cell gene therapy5,6 and gene drives4,7. Gene drives duplicate a segment of genomic DNA in vivo independently of selection and in principle work in any sexually reproducing species so that all offspring inherit the gene drive segment7. Given their speed and efficiency, gene drives have the potential to accelerate the dissemination of beneficial genetics in insects, crops and animals; they might streamline the introduction of homozygous mutations to study recessive alleles, eliminate destructive invasive species and agricultural pests, or to improve livestock rapidly and cheaply (for example, to prevent disease transmission), eliminating the need for protracted breeding programs4. Recently, exceptionally high rates of gene drive transmission based on CRISPR-Cas9 genome editing were demonstrated in flying insect populations8,9. However this remarkable efficiency mandates remarkable control.
Previous methods to control Cas9 activity (reviewed in ref.10) have involved modulating light11,12,13 or temperature13,14, but these approaches are difficult to regulate where barriers exist to light penetration (eg in larger organoids in vitro or environmentally) and are limited under conditions of temperature homeostasis (eg in mammals). Small molecule effectors including rapamycin, trimethoprim and (Z)-4-hydroxytamoxifen have also been used to control Cas9 activity in which, although each system differs, in general a domain is fused to Cas9 such that effector binding activates Cas9 through formation of functional protein15,16,17, protein conformational change18 or translocation into the nucleus19. However, these approaches often suffer from high background activities16,18,19. For instance, a rapamycin-inducible system of gene activation caused a ~10-fold target increase in the absence of rapamycin16 and the original demonstration of Cas9 switching by conformational change exhibited background editing of ~10%; refinement to reduce the background significantly reduced on-target editing18. Moreover, rapamycin and trimethoprim are antibiotics with the potential to alter the microbiome, and as potential drivers of antimicrobial resistance, their deployment is likely to be problematic. (Z)-4-Hydroxytamoxifen, which is the major active metabolite of the anticancer agent, tamoxifen, is an antagonist of the estrogen receptor, hemolytic towards human erythrocytes, an endocrine disruptor and aquatic toxin20,21,22. The cost of small molecule regulators in financial and environmental terms is also likely to be prohibitive on large scales.
Clearly, Cas9 regulation by other means will be advantageous or essential for many of its potential applications. To address this, we considered that a cheap amino acid such as the lysine derivative, H-Lys(Boc)-OH (BOC), might be harnessed for Cas9 control. BOC can be incorporated into proteins of interest by genetic code expansion23, suggesting that a novel tier of regulating Cas9 might be achieved by generating a functional Cas9 variant whose expression (translation) and activity depended on the presence (incorporation during translation) of BOC. We therefore set out to evaluate the control of Cas9 by genetic code expansion in mammalian heritable genome alteration as a first step to the stringent control of gene drives and other applications.
Establishing genetic code expansion in mouse embryos
As a starting point, we characterized genetic code expansion to control eGFP expression in transgenic mouse embryos. To this end, an orthogonal pyrrolysyl-tRNA synthetase/tRNA (PylRS/Pyl tRNA) pair was selected to direct incorporation of BOC into an amber stop codon at position 150 of eGFP, giving eGFPN150B 24; we anticipated that in this system, translation of eGFPN150B would be terminated prematurely (at codon 150) in the absence of BOC and functional eGFPN150B protein produced only following BOC exposure (Fig. 1A–D). Because we were interested in modulating protein activity in a mouse model, we designed an eGFPN150B transgene to express each of the components required for ubiquitous, constitutive expression of the eGFPN150B system (PylRS, Pyl tRNA and eGFPN150B; Supplementary Figure S1A) and coinjected it with wild-type (wt) mouse B6D2F1 sperm into B6D2F1 metaphase II (mII) oocytes (Supplementary Figure S1B). This method of transgenesis by intracytoplasmic sperm injection (ICSI) typically yields >80% of transgene-expressing preimplantation embryos after ~4 days of development (E3.5)25,26. Following eGFPN150B transgene injection, some of the resultant embryos (28.9%; n = 44) clearly fluoresced after 4 days of culture in media supplemented with 10 mM BOC for the final 24 h, but only ~2% fluoresced (with a weak or partial signal; n = 46) when BOC was omitted (p < 0.001; Supplementary Figure S1C,D). This suggested that the orthogonal system could enable BOC-inducible eGFPN150B expression in embryos. To enhance spatiotemporally regulated embryonic expression, we synthesized all RNA components of the BOC system (ie PylRS cRNA, eGFPN150B cRNA and Pyl tRNA) in vitro and coinjected them into B6D2F1 mII oocytes followed by exposure to different concentrations of BOC (Supplementary Figure S2A). These injections were analogous to eGFPN150B transgenesis (Supplementary Figure S1A,B) except that RNA was delivered directly rather than relying on transcription of transgene DNA. All injected oocytes continually exposed to 1 mM BOC expressed readily-detectable green fluorescence (ie eGFPN150B) after 5 and 24 h (Supplementary Figure S2B–D). Thus, RNA injection resulted in efficient BOC-inducible eGFPN150B expression in oocytes, paving the way for its application to other proteins.
Genetic code expansion enables regulation of Cas9 activity via an orthogonal amino acid in tissue culture
We wished to evaluate whether genetic code expansion could be harnessed so that the RNA-guided nuclease activity of Cas91,2,27 could be controlled by adding BOC. Since Cas9 residues can be substituted with a bulky Lys derivative without abolishing endonuclease activity11, we tested Cas9 in which K510 or K742 were replaced with BOC residues. To evaluate this system, HEK293 cells were first transfected with constructs encoding eGFP gRNA, eGFP fused to a destabilization domain (eGFP-DD)28, Pyl tRNA, PylRS and Cas9K510B, Cas9K742B or Cas9K510B,K742B. Transfectants exposed to BOC expressed Cas9 mutants accompanied by a marked reduction of eGFP protein levels, suggestive of BOC-induced Cas9K510B eGFP gene targeting activity (Fig. 2A). Cas9K510B activity was induced by BOC in the micromolar range in a dose-dependent manner (Fig. 2B, n = 3) and Cas9K510B disappearance post-induction was accelerated when it was fused to the destabilization domain (Cas9K510B-DD in Fig. 2C,D; p = 0.002, n = 3). We accordingly evaluated induction of Cas9 mutant activity by BOC in early embryos.
BOC-switchable endogenous transgene editing in one-cell embryos
Oocytes were coinjected with Pyl tRNA and cRNAs encoding PylRS and Cas9K510B, Cas9K742B or Cas9K510B,K742B, incubated for 4~5 h to allow cRNA-encoded protein expression29 and then coinjected with eGFP gRNA plus sperm30 from males homozygous for a ubiquitously-expressed single-copy eGFP transgene31 previously produced by us (Fig. 3A). Mutant Cas9 protein expression was enabled by exposure to BOC immediately after RNA injection for 4~5 or 24 h (Fig. 3A). In these experiments (with Cas9K510B unless stated otherwise), intracytoplasmic sperm injection (ICSI) was expected to produce embryos that fluoresced unless the eGFP gene had successfully been inactivated by editing3. As anticipated, control embryos lacking one or more of the BOC system components all contained brightly fluorescent blastomeres because editing had not taken place and the paternal (sperm-derived) eGFP transgene was therefore expressed (Fig. 3B,C). By contrast, when all BOC components were present and embryos exposed to 1 mM BOC, bright fluorescence was eliminated (Fig. 3B,C). This clearly indicated that the eGFP gene had been inactivated with a high efficiency. Development to the blastocyst stage was reduced following 1 mM BOC exposure for 24 h but it was more efficient following briefer (4~5 h) BOC exposure (Fig. 3D). eGFP mRNAs were intact in fluorescing blastocysts (n = 12) but in all tested non-fluorescing blastocysts, sequencing revealed that transcripts corresponding to eGFP contained one or more mutations corresponding to the targeted region (n = 8; Supplementary Figure S4A). Analogous experiments with cRNA encoding a different Cas9BOC mutant, Cas9K742B, followed by 4~5 h BOC exposure also efficiently yielded non-fluorescing blastocysts (n = 38; Fig. 3E). Induced editing in the system also worked with the double mutant, Cas9K510B,K742B (n = 24), albeit less efficiently; 97.4% for Cas9K742B (Fig. 3E) vs 50.0% for the double mutant (Fig. 3F). These data show that BOC system component levels compatible with embryo development induced efficient editing by Cas9K510B and Cas9K742B. Furthermore, the developmental read-out clearly demonstrates that any truncated Cas9 protein produced in the absence of BOC, or pan-transcriptomic amber-codon read-through of endogenous mRNAs during BOC treatment, had limited cytotoxicity, because even though mouse preimplantation embryos are exquisitely sensitive to developmental perturbation, it did not prevent high rates of development. Moreover, editing was absolutely BOC-dependent, showing that in these experiments the system exhibited little, if any, leakiness.
Heritable, BOC-dependent transgene editing
We next assessed editing heritability in the BOC-inducible system by performing a Cas9K510B series in which Pyl tRNA- and cRNA-injected oocytes were immediately cultured in 1 mM BOC for 4~5 h, injected with eGFP gRNA plus eGFP+ sperm (Fig. 3A) and the resulting 2-cell embryos non-selectively transferred to surrogate mothers. When injection of RNA system components and/or exposure to BOC were omitted, the resulting control offspring fluoresced green (Fig. 4A,B), which was expected given that an intact eGFP allele was present in all fertilizing sperm. However, only 1/18 (5.6%) of pups derived from test embryos following BOC induction were fluorescent, suggestive of efficient editing (Fig. 4A,B). Genomic sequencing of the targeted eGFP region revealed that green fluorescent offspring possessed intact alleles (n = 2), whereas non-all fluorescing offspring harbored at least one of a range of mutations (n = 14; Fig. 4C). Although individuals typically contained a mixture of edited eGFP sequences consistent with mosaicism, we did not detect a mixture of fluorescence and non-fluorescence in offspring, suggesting that editing had occurred efficiently at the 1-cell stage. These data suggest that BOC had efficiently induced, and was a stringent requirement for, Cas9K510B genome editing activity and that the extent of editing in preimplantation embryos provided an accurate meter of editing in offspring.
BOC-dependent heritable editing of native alleles
We next determined whether, in an analogous protocol, BOC could induce Cas9K510B-mediated editing of native alleles of the mouse strain, B6. To this end, we selected genes encoding the testis-determining factor, Sry32, or tyrosinase (Tyr), which confers dark fur pigmentation in B6 mice33. Targeting Sry with control Cas9 (lacking a BOC codon) and a mixture of two gRNAs (Supplementary Table S1) yielded 16 females of which five (31.3%) carried Sry harboring a mutation, implying that Sry inactivation had resulted in sex reversal. When we employed Cas9K510B in the absence of BOC, 0/7 of the resulting females carried Sry, but when BOC was included, 2/10 (20.0%) females were Sry-positive (Fig. 5A) and in both cases Sry had undergone frame-shifting edits (Supplementary Figure S4B). Editing the coat-color determinant, Tyr by control Cas9 and three gRNAs (Supplementary Table S1) had previously resulted in 92.9% (n = 14) of offspring with altered coat-color phenotypes34. A similar efficiency (93.3%; n = 15) of Tyr editing was achieved by BOC-dependent Cas9K510B-mediated targeting (Fig. 5B,C); sequence analysis of coat-color mutants (n = 6) revealed that all contained one or more frame-shifting Tyr gene indels in the targeted region (Supplementary Figure S4C) with a pattern of sequence mosaicism analogous to those observed for eGFP editing (Supplementary Figure S4A). This suggests that BOC-induced editing of endogenous loci by Cas9K510B occurred at comparably high efficiencies to editing mediated by control Cas9.
Inducible genome editing via Cas9BOC supplied by a mouse transgenic line
We extended this principle to evaluate BOC-induced editing by Cas9K510B driven in transgenic mice by the ZP3 promoter, pZP3, which is uniquely expressed in growing oocytes and has become a classical driver of transgene expression in the maternal germline35,36. Comparison of expression between pZP3-Cas9 and pZP3-Cas9K510B transgenic lines is confounded by the differing sites of transgene integration in each, but within each line we noted a marked decline in expression in hemizygotes (containing a single transgene allele) from germinal vesicle stage oocytes, such that pZP3 transgene mRNA was not detectable in mature, fertilizable mII oocytes of the type used in ICSI (Fig. 6A). Thus, the levels of transgene-encoded Cas9 and Cas9K510B transcripts were extremely low.
When mII oocytes from pZP3-Cas9 hemizygous transgenic females were injected with eGFP+ sperm plus gRNA targeting eGFP (but no Cas9 cRNA), 21.2 ± 6.9% of the resulting blastocysts (n = 96) underwent editing as judged by loss of fluorescence (Fig. 6B); editing was confirmed by sequencing (Supplementary Figure S4D). Genome editing had therefore been mediated by endogenous transgene-encoded Cas9 in mII oocytes from hemizygous transgenic females.
We next evaluated whether this might be extended to a BOC-inducible system by injecting mII oocytes from pZP3-Cas9K510B hemizygous transgenic females with Pyl tRNA and cRNA encoding PylRS (but without cRNA encoding Cas9K510B), followed by eGFP+ sperm plus gRNA targeting eGFP. Blastocyst development rates were similar in both groups: 63.3 ± 3.1% of injected oocytes (n = 235) developed in the absence of BOC and 64.4 ± 3.8% (n = 281) with it (p = 0.815). When BOC was included, 7.4 ± 2.3% of the resulting blastocysts (n = 188) lacked fluorescence (including one mosaic) and contained edited eGFP transcripts, whereas all (n = 149) of the blastocysts fluoresced brightly when it was omitted (p = 0.006; Fig. 6B). Given that the control editing rate was 21.2% (see above), BOC-dependent editing by Cas9K510B transgene-expressing females had therefore occurred at ~34.9% of the control rate (7.4% for Cas9K510B vs 21.2% for the control; Fig. 6B).
This work demonstrates controllable, heritable genome editing in mouse embryos by inducing Cas9 activity with the cheap, non-physiological amino acid, Lys(Boc) (BOC) via genetic code expansion. Genetic code expansion has previously been employed for the incorporation of non-physiological amino acids other than BOC, always into the bystander reporter protein, GFP in vivo37,38,39. We here demonstrate that the BOC system applied to Cas9 is compatible with efficient full-term development even when it is active at an early and exquisitely sensitive phase of embryogenesis. This not only provides a developmental indication of limited or surmountable toxicity due to broad genomic amber codon read-through (at least when it occurs during early mammalian embryogenesis) but argues against extensive off-target editing in the system40. In contrast to the higher background activities or restricted applicability of many switchable systems11,12,13,16,18,19, this work accordingly demonstrates low or zero Cas9BOC activity in the absence of BOC; without BOC, Cas9BOC activity was undetectable under the conditions of these experiments, suggesting that the system was subject to little, if any, leakiness.
The BOC-inducible principle should also apply to modified Cas9 activities such as ‘nuclease-dead’ Cas9 (dCas9) derivatives fused to transcriptional modifiers (DNA methyltransferase, histone acetyltransferase, transcription factor activation domains, etc.) to enable reversible, dynamic transcriptional gene control and epigenetic editing (modification) on physiological time-scales (within minutes)41,42,43. BOC would have clear potential advantages over other small molecule effectors, which may either interfere with the fused epigenetic modifier domain or be occluded by it, necessitating case-by-case validation. Moreover, some effectors (eg tamoxifen) inherently contribute to chromatin remodeling, complicating data interpretation44,45.
BOC-inducible Cas9 in gametes or embryos near the time of fertilization also promises to mediate tight gene drive regulation. Although potentially powerful, gene drives may have unforeseen and uncontrollable consequences4. The Cas9 system described here addresses this, and because BOC is a cheap amino acid, it could potentially be used environmentally to control gene drive expression. We generally used 1 mM BOC, but even in a non-optimized in vitro system, a 1,000-fold lower BOC concentration worked detectably (Fig. 2B). Lower concentrations are expected to exert fewer co-lateral effects and reduce costs. Thus, BOC (or something analogous) promises to enable gene-drive activity that can be controlled over large geographic ranges, in principle restricting them to areas of BOC exposure. Gene-drive switchability will also reduce the emergence of resistance by periodically removing the selectable advantage of gene drive neutralization; once the gene drive is inactive, it cannot be selected against. There is no reason a priori why this approach should be limited to editing by Cas9, and other nuclease frameworks (eg Cpf1) should be amenable to analogous regulation.
In addition, genome editing is non-trivial in large animals, whose importance in the provision of biomedical models and clinical reagents (eg for xenotransplantation) is increasing46. By streamlining safe target locus homogenotization (ie allelic replacement to homozygosity) in founders generated by nuclear transfer cloning, pronucleus or sperm injection or via ES/iPS cells, gene drive cassettes could reduce breeding programs and add flexibility, particularly impacting species such as pigs, where ES or naïve iPS cells have not been described. Gene drives could be activated by BOC in early embryogenesis following natural mating, obviating the need for microsurgery. The principle of Cas9 regulation by BOC should also apply to anti-Cas9 proteins47, thereby producing a neutralizing effect on Cas9 activity that is itself controllable.
Moreover, BOC-regulated Cas9 (variants) may directly impact human biomedical applications including therapeutics6; they would, for example, enhance spatiotemporal control of targeted viral integration and/or activity in somatic cell therapies (by restricting BOC to a given anatomical site and time) and mitigate against neutralization by pre-existing anti-Cas9 antibodies48. This control principle holds for proteins other than Cas9.
In sum, by extending orthogonal post-transcriptional regulation to functional, non-reporter genes, this work achieves a new tier of Cas9 control with the clear potential for broad utility in stem cell biology and suggests applicability to the control of diverse biological contexts in vitro and in vivo, to plants, insects and mammals, environmentally and in clinical practice.
Materials and Methods
Plasmid pEF1α-FLAG-PylRS-IRES-Neo-4xU6-PylTU25C (a kind gift from Dr. Jason Chin) was used for the expression of pyrrolysyl-tRNA synthetase and pyrrolysyl tRNACUA and to direct the incorporation of BOC. Plasmid pCAG-eGFP has been described previously25. Plasmids p3s-Cas9HC (Addgene plasmid #43945) and pU6-gRNA-GFP(5–10) were generous gifts respectively from Dr. Jin-Soo Kim and Prof. Anton Wutz; pU6-gRNA-GFP(5–10) expresses a gRNA targeting eGFP corresponding to the region between codons 5 and 10. We generated pCAG-eGFPN150B using pCAG-eGFP as template in a PCR reaction with primers F1 and R1 (primer sequences are given in Table S1) to replace the N150 codon of eGFP with TAG (a B codon), as verified by sequencing with primers S1 and S2. To construct pCAG-eGFP-DD (encoding eGFPT308D,S473D), a 906 base pair (bp) fragment containing eGFP-DD (GeneArt, Regensburg, Germany) was introduced by Gibson assembly into EcoRI-digested pCAG-eGFP and verified by sequencing with primers S1 and S2. To generate pEF1α-FLAG-PylRS-CAG-eGFP(150TAG)-4xU6-PylTU25C (designated SP44), the 8,286 bp BamHI-SalGI fragment of pEF1α-FLAG-PylRS-IRES-Neo-4xU6-PylTU25C was joined by Gibson assembly to a 2,748 bp CAG-eGFPN150B PCR amplimer generated with primers F4 and R4 from pCAG-eGFPN150B and confirmed by sequencing with primers S1, S2 and S4. To construct plasmid p3s-Cas9HCK510B (SP81), Gibson assembly was used to join the 5,862 bp KpnI-BsrGI fragment from p3s-Cas9HC to a PCR amplimer from p3s-Cas9HC containing Cas9(6–528) in which the codon for K510 had been replaced by TAG (B) using primers F5 and R5. The sequence of p3s-Cas9HCK510B was verified by sequencing with primers S5, S6 and S7. To construct plasmid p3s-Cas9HCK742B (SP82), the 6152 bp PmlI-BsrGI fragment from p3s-Cas9HC was joined by Gibson assembly to a PCR amplimer from p3s-Cas9HC containing Cas9(514–745) (SP116) in which the codon for K742 had been replaced by TAG (B) using primers F6 and R6. The sequence of p3s-Cas9HCK510B was verified by sequencing with primers S7, S8 and S9. Construct pU6-gRNA-GFP(51–56) was prepared from pU6-gRNA-GFP(5–10) and its sequence confirmed with primers S16 and S17. Plasmid pEF1α-FLAG-PylRS-CAG-eGFP-DD-4xU6-PylTU25C-U6-gRNA-GFP(5–10)-U6-gRNA-GFP(51–56) was constructed in four steps. First, Gibson assembly of the 8,286 bp pEF1α-FLAG-PylRS-IRES-Neo-4xU6-PylTU25C BamHI-SalGI fragment to the 683 bp fragment (containing IRES) amplified with primers F2 and R2 from pEF1α-FLAG-PylRS-IRES-Neo-4xU6-PylTU25C, and the 758 bp fragment (containing eGFP) amplified with primers F3 and R3 from pCAG-eGFP generated pEF1α-FLAG-PylRS-IRES-eGFP-4xU6-PylTU25C; this was confirmed by sequencing with primers S1, S2 and S4. Secondly, Gibson assembly of the pEF1α-FLAG-PylRS-IRES-eGFP-4xU6-PylTU25C BglII fragment with the 412 bp PCR product generated with primers F13 and R13 from pU6-gRNA-GFP(5–10) yielded pEF1α-FLAG-PylRS-IRES-eGFP-4xU6-PylTU25C-U6-gRNA-GFP(5–10), as confirmed by sequencing with primer S18. In the third step, Gibson assembly of the pEF1α-FLAG-PylRS-IRES-eGFP-4xU6-PylTU25C-U6-gRNA-GFP(5–10) BglII fragment with the 412 bp PCR product generated with primers F13 and R13 from pU6-gRNA-GFP(51–56) yielded pEF1α-FLAG-PylRS-IRES-eGFP-4xU6-PylTU25C-U6-gRNA-GFP(5–10)-U6-gRNA-GFP, (51–56) which was confirmed by sequencing with primer S18. Finally, Gibson assembly of the 9,020 bp BamHI-SalGI fragment of pEF1α-FLAG-PylRS-IRES-eGFP-4xU6-PylTU25C-U6-gRNA-GFP(5–10)-U6-gRNA-GFP(51–56) and the 2,875 bp PCR amplimer generated using primers F4 and R4 from pCAG-eGFP-DD produced pEF1α-FLAG-PylRS-CAG-eGFP-DD-4xU6-PylTU25C-U6-gRNA-GFP(5–10)-U6-gRNA-GFP, (51–56) as confirmed by sequencing with primers S1, S2 and S4. Plasmids were linearized prior to oocyte injection. In brief, the PylRS cassette was excised and inserted into the pCIneo vector (Promega) at its NheI/EcoRI site and PylRS cRNA synthesized in vitro using a T7 transcript kit (Invitrogen). The Pyl tRNA insert was subcloned as a 4 × (U6-PylT) cassette into the vector, to give T7 promoter -directed expression. Plasmids were expanded and purified using EndoFree Plasmid Maxi Kit (QIAGEN) and digested with SfiI/SapI/SallHF (SP44) or with XhoI for Cas9HCK510B (SP81), Cas9HCK742B (SP82) and Cas9HCK510B,K742B (SP116). Plasmid p3S-Cas9HCK510B-DD, encoding Cas9HCK510B (Cas9K510B containing a C-terminal destabilization domain fusion) was constructed by Gibson assembly of the 7,320 bp p3S-Cas9HCK510B XhoI-KpnI fragment to a 233 bp fragment containing the destabilization domain (GeneArt). Fragments were gel-purified (Promega) and each was injected at 1.0 ng/μl into mII oocytes with sperm25.
Cas9 editing assay in HEK293 cells
HEK293 cells were grown at 37 °C in a 5% (v/v) CO2 atmosphere in DMEM plus GlutaMAX medium (Gibco) supplemented with 10% (v/v) fetal bovine serum for 24 h before transfection. Cells (2 × 105/well in a 24-well plate) were transiently transfected with the appropriate p3s-Cas9HC variant and pEF1α-FLAG-PylRS-CAG-eGFP-DD-4xU6-PylTU25C-U6-gRNA-GFP(5–10)-U6-gRNA-GFP(51–56) using Lipofectamine 2000 (Life Technologies) according to the recommendations of the manufacturer and plated in parallel in media either further supplemented with 1 mM BOC or lacking BOC, or at the BOC concentrations indicated for Fig. 2. After 24 h, cells were washed with 0.5 ml phosphate-buffered saline (PBS) and lysed with RIPA buffer (Sigma, R0278; 50 μl per well) containing protease inhibitor (Sigma, P8340) at 4 °C for 10 min. Lysates were pelleted (20,000 g, 10 min, 4 °C) and the supernatant (45 μl) added to 15 μl LDS sample buffer (Life Technologies). Samples were heated (95 °C, 10 min) and loaded onto a 4–20% Mini-PROTEAN TGX Stain-Free Protein Gel (Bio-Rad) for electrophoresis. The gel was imaged on a ChemiDoc XRS + (Bio-Rad) and transferred to a nitrocellulose membrane using a Trans-Blot Turbo Transfer System (Bio-Rad). The membrane was stained with Ponceau S to confirm protein transfer, blocked with PBST (0.05% [v/v] Tween 20 in PBS) containing 5% (w/v) milk powder at 20 °C for 1 h, and then incubated (4 °C, overnight) with a primary mouse anti-GFP antibody (Thermo Fisher, #MA1-952, 1:500 [v/v] dilution). The membrane was then washed three times (10 ml PBST, 5 min per wash). All subsequent washing steps used this procedure. The membrane was then incubated in secondary anti-mouse antibody (Thermo Fisher, #32430, 1:1,000 [v/v] dilution) for 1 h at 20 °C, 1 h and then washed. The signal was developed by addition of Clarity Max™ Western ECL Substrate (Bio-Rad, #1705062). After imaging on a Bio-Rad ChemiDoc XRS + system, the membrane was washed and incubated (~20 °C, 30 min) with Restore Western Blot Stripping Buffer (Thermo Fisher, #21059) and then washed, incubated (4 °C, overnight) with a primary mouse anti-HA antibody (Thermo Fisher, #26183, 1:2000 [v/v] dilution) before being incubated with the secondary antibody and processed for imaging as described above.
Target gRNA synthesis
Target gRNAs were produced as previously described49. Briefly, T7-gRNA-scaffold products were amplified by PCR using Pfx polymerase (Invitrogen) with Scaffold-Fwd and Scaffold-Rev primers and the T7 gRNA vector as template with PCR reaction parameters: 94 °C, 2 min followed by 25 cycles of 94 °C, 15 sec; 55 °C, 30 sec; 68 °C, 20 sec; final extension at 68 °C, 7 min. gRNA scaffold PCR product were gel purified (Promega) and used as a template using T7-target oligo and the amplified gRNA scaffold. gRNA was synthesized by PCR with Pfx using T7-target oligo primers (Supplementary Table S1), T7-fwd primer and scaffold-Rev primer with the parameters: 94 °C, 5 min followed by 25 cycles of 94 °C, 15 sec; 55 °C, 30 sec; 68 °C, 20 sec; final extension at 68 °C, 7 min. The amplification of single 130 bp PCR products was in each case confirmed by 2% (w/v) agarose gel electrophoresis and products purified (Promega) for following synthesis of gRNA in vitro3 using the MEGAshortscript T7 Transcription Kit (Invitrogen) as recommended by the manufacturer.
Collection and culture of oocytes and embryos
Animal procedures complied with the Animals (Scientific Procedures) Act, 1986 and experimental protocols were approved by the University of Bath Ethical Review Board. Wild-type mouse (Mus musculus) strains were bred from C57BL/6J (B6) and DBA/2 stocks in-house or supplied by Charles River (L’Arbresle, France). A 129SvJ line containing a single-copy eGFP transgene under the control of the pCAG promoter-enhancer has been reported previously31. Oocytes were collected from 8- to 12-week-old females that had been super-ovulated by standard serial intraperitoneal injection of 5 IU pregnant mare serum gonadotropin (PMSG) followed 48 h later by 5 IU human chorionic gonadotropin (hCG) and held under mineral oil in humidified 5% CO2 (v/v air) at 37 °C, until required29,31. Embryo culture in vitro was in KSOM under mineral oil in humidified 5% CO2 (v/v in air) at 37 °C.
Sperm preparation and microinjection
Cauda epididymidal sperm from 8- to 12-week-old males were prepared as described previously3,30,31. Sperm were resuspended in ice-cold nuclear isolation medium (NIM; ~0.5 ml per epididymis) and stored on ice or at 4 °C. Immediately prior to injection, ~50 μl of freshly-prepared sperm suspension was mixed 20 μl of polyvinylpyrrolidone (PVP, average Mr ≈ 360,000; Sigma-Aldrich) solution (15% [w/v]). DNA or gRNA solutions were immediately mixed with the sperm/PVP suspension to give each at a final concentration of 1 ng/μl and 200 ng/μl respectively in the suspension. For endogenous gene editing of Tyr and Sry loci in the BOC system, the gRNA working concentration was 800 ng/μl. Approximately 2.0~2.5 μl of suspension containing a single sperm head was injected (ICSI) within ~60 min of DNA/gRNA mixing into mII oocytes held in droplets of M2 under mineral oil, essentially as described25,30,31. Where indicated, mII oocytes that had been injected with RNA were injected with sperm heads (ICSI) 3~4 h later.
Synthesis of RNA in vitro and microinjection into mII oocytes
The preparation of 5′-capped and polyadenylated in vitro transcripts was from cRNA synthesized from linearized plasmid template DNA in a T7 mScript™ Standard mRNA Production System (Cellscript, USA) as previously described29,31. Synthesis in vitro of gRNA and tRNA was with the MEGAshortscript T7 RNA synthesis kit (Invitrogen) according to the instructions of the vendor. Each took place at 37 °C for 3~4 h in a 20 μl total reaction volume containing DNA template and T7 RNA polymerase. RNAs were precipitated, dissolved in nuclease-free water, quantified on a Nanophotometer and stored in aliquots at −80 °C until required. RNA solutions were diluted as appropriate with sterile water and injected within 1 h of thawing via a piezo-actuated micropipette into mII oocytes, or in the case of gRNA, in the presence of sperm. Working concentrations of RNA injected into mII oocytes were: 600 ng/μl for gRNA, 100 ng/μl for cRNA encoding non-mutant Cas9, 600 ng/μl for cRNA encoding PylRS, eGFPN150B or mutant Cas9, and 1,200 ng/μl for PylTU25C tRNA (Pyl tRNA). For BOC-induced editing of endogenous genomic Sry and Tyr sequences, the working concentrations of cRNA encoding mutants was 3,300 ng/μl and for gRNA, 800 ng/μl. We estimate that 5~10 pl were typically injected per mII oocyte.
Reverse transcriptase PCR
Total RNA from embryos and tissues was extracted using the TRI reagent (Sigma) according to the instructions provided. From each total RNA sample, 800 ng were used for first strand cDNA synthesis by Superscript III Reverse transcriptase (Invitrogen). Following synthesis, cDNA was treated with RNase H (Invitrogen) for 20 min at 37 °C and 1 μl of resultant cDNA used to charge PCR reactions with the appropriate target primers (Supplementary Table S1).
Embryonic day (E1.5) 2-cell embryos were transferred to the oviducts of pseudo-pregnant CD-1 females at day 0.5 (ie plugged females that had been mated with vasectomized males the previous night). Where appropriate, pups were delivered by Cesarian section and fostered by CD-1 females.
Genotyping was performed on ear-punch biopsies collected at weaning as described previously3 and digested at 55 °C for 5 h in 100 μl of a lysis buffer containing 10% (w/v) sodium dodecyl sulfate and 2 mg/ml proteinase K (Sigma). One microliter of a 1:10 (v/v) dilution of each genomic DNA sample was used for genotyping by PCR in a 10 μl reaction volume. For single blastocyst genomic PCR, individual blastocysts (E4.5–E5.5) were each collected into a PCR tube containing 0.5% (w/v) sarcosyl and flash-frozen in liquid nitrogen before being subjected to target PCR amplification with primer sequences as given in Supplementary Table S1. Where appropriate, PCR products were gel-purified for sequence analysis.
Following oocyte injection, fluorescence of embryos was visualized on an IX71 (Olympus, Japan) microscope equipped with an AndroZyla cMOS camera and OptoLEP illumination system (Cairn Research, UK) as previously described31. Images were processed with ImageJ (imagej.nih.gov/ij/) or MetaMorph (Molecular Devices, USA) analysis software. Quantitative analyses subtracted background from subject area fluorescence intensities, which can produce negative results in beads experiments in which background levels from latex are lower than those of mII oocytes.
All experiments were performed on at least two days. The number of samples (n) per experiment reflects oocyte or embryo survival after manipulation. All samples were randomly collected; that is, we did not knowingly select different classes of healthy oocytes or embryos except where stated and no data were selectively excluded. Data analysis was performed with or without blinding. Statistical differences between pairs of data sets were analyzed by chi-squared or two-tailed unpaired t-test. Values of p < 0.05 were considered statistically significant.
Mali, P. et al. RNA-guided human genome engineering via Cas9. Science 339, 823–826 (2013).
Cong, L. et al. Multiplex genome engineering using CRISPR/Cas systems. Science 339, 819–823 (2013).
Suzuki, T., Asami, M. & Perry, A. C. F. Asymmetric parental genome engineering by Cas9 during mouse meiotic exit. Sci. Rep. 4, 7621 (2014).
Esvelt, K. M., Smidler, A. L., Catteruccia, F. & Church, G. M. Concerning RNA-guided gene drives for the alteration of wild populations. eLife 3, e03401 (2014).
Pineda, M., Moghadam, F., Ebrahimkhani, M. R. & Kiani, S. Engineered CRISPR systems for next generation gene therapies. ACS Synth. Biol. 6, 1614–1626 (2017).
Dai, W. J. et al. CRISPR-Cas9 for in vivo Gene Therapy: Promise and Hurdles. Mol. Ther. Nucleic Acids 5, e349 (2016).
Burt, A. Site-specific selfish genes as tools for the control and genetic engineering of natural populations. Proc. R. Soc. Lond. B 270, 921–928 (2003).
Gantz, V. M. & Bier, E. Genome editing. The mutagenic chain reaction: a method for converting heterozygous to homozygous mutations. Science 348, 442–444 (2015).
Hammond, A. et al. A CRISPR-Cas9 gene drive system targeting female reproduction in the malaria mosquito vector Anopheles gambiae. Nat. Biotechnol. 34, 78–83 (2016).
Nihongaki, Y., Otabe, T. & Sato, M. Emerging approaches for spatiotemporal control of targeted genome with inducible CRISPR-Cas9. Anal. Chem. https://doi.org/10.1021/acs.analchem.7b04757 (2016).
Hemphill, J., Borchardt, E. K., Brown, K., Asokan, A. & Deiters, A. Optical Control of CRISPR/Cas9 Gene Editing. J. Am. Chem. Soc. 137, 5642–5645 (2015).
Nihongaki, Y., Kawano, F., Nakajima, T. & Sato, M. Photoactivatable CRISPR-Cas9 for optogenetic genome editing. Nat. Biotechnol. 33, 755–760 (2015).
Richter, F. et al. Engineering of temperature- and light-switchable Cas9 variants. Nucleic Acids Res. 44, 10003–10014 (2016).
Moreno-Mateos, M. A. et al. CRISPR-Cpf1 mediates efficient homology-directed repair and temperature-controlled genome editing. Nat. Commun. 8, 2024 (2017).
Davis, K. M., Pattanayak, V., Thompson, D. B., Zuris, J. A. & Liu, D. R. Small molecule-triggered Cas9 protein with improved genome-editing specificity. Nat. Chem. Biol. 11, 316–318 (2015).
Zetsche, B., Volz, S. E. & Zhang, F. A split-Cas9 architecture for inducible genome editing and transcription modulation. Nat. Biotechnol. 33, 139–142 (2015).
Nguyen, D. P. et al. Ligand-binding domains of nuclear receptors facilitate tight control of split CRISPR activity. Nat. Commun. 7, 12009 (2016).
Oakes, B. L. et al. Profiling of engineering hotspots identifies an allosteric CRISPR-Cas9 switch. Nat. Biotechnol. 34, 646–651 (2016).
Liu, K. I. et al. A chemical-inducible CRISPR-Cas9 system for rapid control of genome editing. Nat. Chem. Biol. 12, 980–987 (2016).
Cruz Silva, M. M., Madeira, V. M., Almeida, L. M. & Custódio, J. B. Hemolysis of human erythrocytes induced by tamoxifen is related to disruption of membrane structure. Biochim. Biophys. Acta 1464, 49–61 (2000).
Brauch, H., Mürdter, T. E., Eichelbaum, M. & Schwab, M. Pharmacogenomics of tamoxifen therapy. Clin. Chem. 55, 1770–1782 (2009).
Orias, F. et al. Tamoxifen ecotoxicity and resulting risks for aquatic ecosystems. Chemosphere 128, 79–84 (2015).
Chin, J. W. Expanding and Reprogramming the Genetic Code of Cells and Animals. Annu. Rev. Biochem. 83, 379–408 (2014).
Tsai, Y. H., Essig, S., James, J. J., Lang, K. & Chin, J. W. Selective rapid and optically switchable regulation of protein function in live mammalian cells. Nat. Chem. 7, 554–561 (2015).
Perry, A. C. F. et al. Mammalian transgenesis by intracytoplasmic sperm injection. Science 284, 1180–1183 (1999).
Perry, A. C. F. et al. Efficient metaphase II transgenesis with different transgene archetypes. Nat. Biotechnol. 19, 1071–1073 (2001).
Jinek, M. et al. A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity. Science 337, 816–821 (2012).
Li, X. et al. Generation of destabilized green fluorescent protein as a transcription reporter. J. Biol. Chem. 273, 34970–34975 (1998).
Suzuki, T., Yoshida, N., Suzuki, E., Okuda, E. & Perry, A. C. F. Full-term mouse development by abolishing Zn2+-dependent metaphase II arrest without Ca2+ release. Development 137, 2659–2669 (2010).
Yoshida, N. & Perry, A. C. F. Piezo-actuated mouse intracytoplasmic sperm injection (ICSI). Nat. Protoc. 2, 296–304 (2007).
Suzuki, T. et al. Mice produced by mitotic reprogramming of sperm injected into haploid parthenogenotes. Nat. Commun. 7, 12676 (2016).
Berta, P. et al. Genetic evidence equating SRY and the testis-determining factor. Nature 348, 448–450 (1990).
Müller, G., Ruppert, S., Schmid, E. & Schutz, G. Functional analysis of alternatively spliced tyrosinase gene transcripts. EMBO J. 7, 2723–2730 (1988).
Sunagawa, G. A. et al. Mammalian Reverse Genetics without Crossing Reveals Nr3a as a Short-Sleeper Gene. Cell Rep. 14, 662–677 (2016).
Philpott, C. C., Ringuette, M. J. & Dean, J. Oocyte-specific expression and developmental regulation of ZP3, the sperm receptor of the mouse zona pellucida. Dev. Biol. 121, 568–575 (1987).
Lewandoski, M., Wassarman, K. M. & Martin, G. R. Zp3-cre, a transgenic mouse line for the activation or inactivation of loxP-flanked target genes specifically in the female germ line. Curr. Biol. 7, 148–151 (1997).
Ernst, R. J. et al. Genetic code expansion in the mouse brain. Nat. Chem. Biol. 12, 776–778 (2016).
Chen, Y. T. et al. Heritable expansion of the genetic code in mouse and zebrafish. Cell Res. 27, 294–297 (2017).
Han, S. et al. Expanding the genetic code of Mus musculus. Nat. Commun. 8, 14568 (2017).
Schaefer, K. A. et al. Unexpected mutations after CRISPR-Cas9 editing in vivo. Nat. Methods 14, 547–548 (2017).
Konermann, S. et al. Genome-scale transcriptional activation by an engineered CRISPR-Cas9 complex. Nature 517, 583–588 (2015).
Vora, S., Tuttle, M., Cheng, J. & Church, G. Next stop for the CRISPR revolution: RNA-guided epigenetic regulators. FEBS J. 283, 3181–3193 (2016).
Liao, H. K. et al. In Vivo Target Gene Activation via CRISPR/Cas9-Mediated Trans-epigenetic Modulation. Cell, https://doi.org/10.1016/j.cell.2017.10.025 (2017).
Liu, X. F. & Bagchi, M. K. Recruitment of distinct chromatin-modifying complexes by tamoxifen-complexed estrogen receptor at natural target gene promoters in vivo. J. Biol. Chem. 279, 15050–15058 (2004).
Sharma, D., Saxena, N. K., Davidson, N. E. & Vertino, P. M. Restoration of tamoxifen sensitivity in estrogen receptor-negative breast cancer cells: tamoxifen-bound reactivated ER recruits distinctive corepressor complexes. Cancer Res. 66, 6370–6378 (2006).
Rogers, C. S. Genetically engineered livestock for biomedical models. Transgenic Res. 25, 345–359 (2016).
Pawluk, A. et al. Naturally occurring off-switches for CRISPR-Cas9. Cell 167, 1829–1838.e9 (2016).
Charlesworth, C. T. et al. Identification of Pre-Existing Adaptive Immunity to Cas9 Proteins in Humans. bioRxiv, https://doi.org/10.1101/243345 (2018).
Shao, Y. et al. CRISPR/Cas-mediated genome editing in the rat via direct injection of one-cell embryos. Nat. Protoc. 9, 2493–2512 (2014).
The authors gratefully acknowledge support to A.C.F.P. from the Medical Research Council, UK (MR/N000080/1 and MR/N020294/1), to A.C.F.P. and Y.-H.T. from the Biology and Biological Science Research Council, UK (BB/P009506/1) and to Y.-H.T. from the Wellcome Trust (200730/Z/16/Z). We thank Professor C. Tickle and Dr. M. VerMilyea for incisive comments during manuscript preparation, animal care laboratory staff at the University of Bath, and Professor M. Szczelkun for introducing the Tsai and Perry labs.
The authors declare no competing interests.
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Suzuki, T., Asami, M., Patel, S.G. et al. Switchable genome editing via genetic code expansion. Sci Rep 8, 10051 (2018). https://doi.org/10.1038/s41598-018-28178-3
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