Archive for the ‘Crispr’ Category

ICP: an integrated pipeline for classifying CRISPR/Cas9 induced mutant alleles

We developed an integrated bioinformatic tool ICP (Integrated Classifier Pipeline), to parse complex DSB repair outcomes induced by CRISPR/Cas9 and automatically call for experimental errors generated during NGS library preparation and sequencing: 1) a Nucleotide Position Classifier (NPClassifier), and 2) a Single Allele-resolution Classifier (SAClassifier). We employed these two complementary sequence analysis modules in tandem to enable in-depth interpretation of deep sequencing data at single allele resolution (Fig.1ac, see Methods section for detailed description of ICP tools). In line with the unique DNA signatures generated by distinct DSB repair pathways, we categorized the repair products into four major categories. Alleles with a deletion only on the PAM-distal side (PAM-proximal side was protected by Cas9 protein after cleavage), a common category, were termed as PEPPR class mutations (PAM-End Proximal Protected Repair, PEPPR)41,42. While single strand cleavage by the Cas9 RuvC domain can also nick the non-complementary strand at locations beyond the canonical site between the 6th and 7th nucleotide upstream of the PAM sequence, we restrict our analysis here to the majority cases wherein Cas9 cleavage generates blunt DSB ends to simplify the robust classification scheme developed in this study43,44,45. Mutant alleles judged to be generated by directly annealing 2bp microhomology sequences spanning the gRNA cleavage site were assigned into MMEJ class (again acknowledging that such alleles can also be generated with 1bp microhomology sequence, which however, are not readily amenable to the semi-automated analysis we developed)46,47,48, while pure deletion alleles not belonging to either the PEPPR or MMEJ categories were classified as DELET class mutations. Remaining alleles that include insertions-only and indels (deletion plus insertion) were categorized as insertion class (INSRT) mutations (Fig.1b).

The process of DSB repair pattern profiling consists of preparing a NGS library (a), classifying the resulting parsed alleles (b) and displaying processed alleles by rank order and class of mutations (c). a NGS library preparation: Genomic DNA from F1 test flies carrying both Cas9 and gRNA expressing cassettes either maternally (dark blue bars) or paternally (red bars, or progeny from other designated crosses) are subjected for targeted PCR amplification with primers containing Illumina compatible adapters at the 5 terminal to detect somatic indels. The gray rectangle represents a short region of genomic DNA containing a Cas9/gRNA target: purple circle depicts Cas9 protein and sky-blue line is gRNA. b Classification: Raw NGS data are subjected to the NPClassifier to parse alleles into specific primary categories required for building allelic dictionaries used by the SAClassifier. Four major indel groups are categorized: PEPPR (PAM-End Proximal Protected Repair, sky-blue), MMEJ (Microhomology Mediated End-Joining, dark pink), DELET (deletion, any deletions do not belong to PEPPR and MMEJ, orange) and INSRT (insertion, including the alleles only with inserted nucleotides or had deletions and insertions, purple). The 24-nt short PEPPR, MMEJ and DELET dictionaries are used for a more accurate classification and error calling by binning together all alleles with the same seed region that match primary allelic entries in the SAClassifier dictionaries. c DSB repair pattern visualization: intuitive rendering of the processed raw sequence data as an output of rank ordered classes of alleles. Allelic classes derived from NGS sequencing of individual flies or mosquitoes are displayed by their ranked frequency (allele landscape) and repair pattern fingerprints (color-coded by categories).

Briefly, raw reads generated from deep sequencing were subjected to a preliminary categorization using the NPClassifier, which recognizes the relative positions of editing start- and end-points flanking Cas9 cleavage site and then generates a collection of priori alleles for each category. These primary outputs (MMEJ and DELET) were used for building full-length standard comprehensive dictionaries listing all observed mutations and derived 24-nt short dictionaries (with the same seed region flanking the Cas9 cleavage site) as inputs of the SAClassifier. In addition, a synthetic PEPPR dictionary was built by iteratively increasing the length of deletions by a single nucleotide distal to the PAM site, excluding alleles belonging to the MMEJ category. By fishing the raw reads with 24-nt dictionaries, we were able to automatically recognize reads that also contained experimentally generated errors (e.g., from PCR amplification), which usually are located outside of the narrow 24-nt short dictionary window, thereby assigning such composite alleles to correctly matched root alleles (Fig.1b). These dual iteratively employed ICP classification tools provide a robust and precise classification of CRISPR/Cas9 induced DSB repair outcomes. Next, we developed an evocative user-friendly interface to visualize processed allelic category information in the form of rank ordered allelic landscape plots and repair pattern fingerprints (color-coded DSB repair categories), both of which are sorted by read frequency (Fig.1c). These intuitively accessible data outputs are far more informative and discriminating than the unprocessed primary DNA sequence reads (e.g., compare the seemingly idiosyncratic raw lesions depicted in Fig.2a to the obviously unique processed and concordant replicate patterns shown Fig.2b, c). The ICP was thus employed to visualize results in all the following experiments.

a Examples of the top five somatic indels from individual flies derived from split-drive crosses in which the Cas9 transgene is inherited either maternally (Maternal-S, left) or paternally (Paternal-S, right), but separately from a cassette carryingthe gRNAtransmittedby the other parent. Purple stars indicate the color codes for mutation categories (dark pink: MMEJ, sky-blue: PEPPR, orange: DELET, purple: INSRT) and dark green star indicates the separate raw sequence color coded for the four nucleotides A, T, G, and C. The red bar indicates Paternal-S crosses while dark blue bar represents Maternal-S crosses. b Landscapes of top 50 alleles ranked by reads ratio. All six sequenced individual flies are plotted together, with dark blue lines plotting the data from Maternal-S crosses and the red lines from Paternal-S crosses. The y-axis presents the fraction of reads for a given allele and the x-axis depicts the top 50 alleles according to rank order by read frequency. c DSB repair fingerprints for three representative sequenced individual flies from each cross. The x-axis is the same as depicted in panel b. Both panels show the top 50 ranked alleles. d. Bar plots of Class Fraction for top 50 alleles. Color codes for classes are as in panels a and c. Correlation analysis of two out of three replicates from Maternal-S cross (e) or Paternal-S (f) cross. r2 values and p-values are indicated. Source data for panels b, d, e and f are provided as a Source Data file.

Since DSB repair outcomes have been found to vary considerably as a function of Cas9 or gRNA source and level49,50, we employed the ICP platform to parse somatic indels generated by co-expressing Cas9 and gRNAs in somatic cells of fruit flies (Drosophila melanogaster) and mosquitoes (Anopheles stephensi) in various configurations associated with gene-drive systems. We first applied ICP analysis to a split gene-drive system inserted into the Drosophila pale (ple)gene that is designed to detect copying of a gene cassette in somatic cells. This element, referred to as a CopyCatcher (pleCC), carries a gRNA targeting the first intron of Drosophila ple locus49. In this current study, we make use of low-level ectopic somatic Cas9 expression (which is substantial and broad for vasa-Cas9) to analyze DSB repair patterns across diverse cell types in F1 progeny carrying both Cas9 and gRNAs51,52,53. Because cells actively undergoing meiosis make up only a small fraction of dividing cells in an adult fly, the mutational effects of Cas9/gRNA cleavage in such F1 individuals largely reflect the somatic action of these nuclease complexes. We thus conducted several alternative crossingschemes to assess the somatic mutagenic activity of vasa-Cas9 and gRNA components when transmitted to F1 individuals in various configurations from their F0 parents: 1) Maternal Split (Maternal-S, females carrying vasa-Cas9 crossed with males carrying pleCC); 2) Paternal Split (Paternal-S, males carrying vasa-Cas9 crossed with females carrying pleCC); and 3) Maternal Full (Maternal-F, females carrying both the pleCC and vasa-Cas9 transgenes); or Paternal Full (Paternal-F, males carrying both the pleCC and vasa-Cas9 transgenes)49. Comparative ICP analysis revealed several striking and consistent differences between the prevalent somatic mutations generated in individual progeny in each of these different crossing schemes. In the case of Paternal-S crosses, the resulting mutations were dominated by PEPPR alleles (4 out of top 5 alleles in Fig.2a, Fig. S1a, and 70% of the top 50 alleles as rendered in rank ordered allelic landscapes and color coded DSB repair fingerprints in Fig.2c). In contrast, Maternal-S crosses primarily generated MMEJ and INSRT indels (4 out of top 5 alleles were MMEJ, and at least 50% of the top 50 alleles were INSRT mutations, Fig.2a, c, Supplementary Fig. S1a). These differences were also evident in the steeper allelic landscape curves that were generated from the Maternal-S versus Paternal-S crosses (Fig.2b) as characterized by the initial portion of the curve depicting the 5 most frequent alleles (i.e., the dark blue lines in Fig.2b are all above the red lines for the 5 most frequent alleles). We further quantified differences in allelic profiles between crosses by bar plots displaying the summed proportions of the different allelic classes (summing the percentages of all alleles from each category) which we termed as Class Fraction (Fig.2d). This analysis revealed that INSRT alleles were generated at a significantly higher frequency in Maternal-S crosses, while the PEPPR class dominated among the top 50 alleles in the reciprocal Paternal-S crosses (Fig.2d).

A striking feature of the highly divergent DSB repair signatures generated from maternally versus paternally inherited Cas9 sources was the remarkable reproducibility of their DSB repair fingerprints observed across three individual replicates from each cross (Fig.2e, f). We performed a correlation analysis within replicates by extracting 23 common alleles across all six sequenced flies and plotted the resulting allelic profiles together relative to an arbitrarily chosen Paternal-S replicate as reference (bold red line, Supplementary Fig. S1b). We observed that the frequency distributions of these 23 common alleles were much more similar to each other within intra-cross comparisons than between inter-crosses (Supplementary Fig. S1b). This trend was also revealed by higher correlation coefficients for intra-cross comparisons than for inter-cross comparisons based on allelic read ratios (Supplementary Fig. S1cg). Conspicuous defining differences between the Maternal-S and Paternal-S fingerprints were also evident based on the Class Fraction index (Fig.2d). In summary, a variety of differing statistical measurements all underscore the robust consistent similarities shared among allele profiles generated from individual replicates of same cross and clearly distinctive DSB repair pattern fingerprints generated by maternal versus paternal Cas9 inheritance.

We extended our ICP analysis of mutant allele profiles generated in the ple locus to the more extreme Maternal-F (dark blue lines) and Paternal-F (red lines) cross schemes to assess the role of inheritance patterns when both the source of vasa-Cas9 and gRNA originated from a single parent49. Again, we observed highly dominant alleles in the Maternal-F crosses, clearly evident in allelic landscapes, that deviated markedly from those produced by the Paternal-F crosses, which produced more evenly distributed spectra of alleles spread across a broad range of allelic frequencies (Fig.3a, b). As expected based on these large differences, the repair pattern fingerprints generated from different crosses produced clearly distinguishable patterns of mutation classes, which was particularly evident when considering the Class Fraction (Fig.3e). Cumulatively, these data suggest that the developmental timing and/or levels of Cas9 expression (maternal, early zygotic, or late zygotic) are likely to play a key role in determining which particular DSB repair pathway or sub-pathway is engaged in resolving DSBs.

ad Unique DSB repair signatures obtained using different Cas9 sources are displayed with the top 20 alleles (landscapes and DSB repair pattern fingerprints). NGS sequencing was performed on pools of 20 adults. a vasa-Cas9 inserted in the X chromosome and the pleCC element carrying the gRNA were both carried by either female or male parents, mimicking a full-drive configuration (Maternal-F and Paternal-F crosses with vasa-Cas9). b vasa-Cas9 split crosses wherein the Cas9 transgene was transmitted either maternally (Maternal-S) or paternally (Paternal-S) and the pleCC gRNA bearing cassette was carried by the other parent. Same Maternal-S versus Paternal-S crosses as in panel b, but using either actin-Cas9 (c) or nanos-Cas9 (d) sources. e Class Fraction Index for crosses in panels ad. Bars are shaded according to allelic class color codes. f UMAP embedding for visualizing a common set of 59 alleles shared between the four split crosses with actin-Cas9 and vasa-Cas9. Dots represent single alleles, and the colors indicate the allelic category. g Distribution of top 20 alleles generated from single flies derived from across between parents carrying theSpo11 gRNA and vasa-Cas9elements (Paternal-S cross: red lines and Maternal-S cross: dark blue lines). The top plot shows the allelic landscape for the top 20 alleles from all six sequenced single flies and the bottom shows three examples of the classification fingerprints (with all allelic classes condensed into single rows) color coded for the allele categories. h Class Fraction Index for Spo11 gRNA crosses. i, j Correlation analysis between two replicates from each cross. Dark blue is Maternal-S and red is for Paternal-S. r2 values and p-values are indicated. Source data are provided as a Source Data file.

Previous studies have shown that the relative frequencies of NHEJ versus HDR events depend on the source of Cas9 both in terms of timing and level of expression49,50,54. We thus wondered whether ICP analysis would similarly reveal distinct DSB repair outcomes for two additional Cas9 sources (actin-Cas9 and nanos-Cas9, expressing level of Cas9: actin-Cas9>vasa-Cas9>nanos-Cas9) inserted at the same locus with vasa-Cas9 (Fig.3c, d)49.

As was observed for the vasa-Cas9 source, the actin-Cas9 and nanos-Cas9 sources both generated differing allelic landscapes and repair pattern fingerprints when transmitted maternally versus paternally, which also were readily distinguishable from each other (Fig.3bd). Mirroring results with the vasa-Cas9 source, significant differences between the proportions of PEPPR versus MMEJ class among the top 20 alleles were observed in Maternal-S versus Paternal-S crosses for actin-Cas9. For the nanos-Cas9 source, both the MMEJ and INSRT categories were particularly reduced in Paternal-S crosses, although this latter sex-based difference was not as dramatic as for the other Cas9 sources (presumably due to its more germline restricted expression, Fig.3d)55,56. Overall, the general trend once again indicated that maternally inherited Cas9 sources biased somatic DSB repair outcomes in favor of MMEJ and INSRT classes over PEPPR alleles, while paternal transmission of Cas9 generated mutant alleles dominated by PEPPR class alleles (Fig.3e).

Based on the overall similarities of the DSB repair outcomes observed for actin-Cas9 and vasa-Cas9 crosses, we extracted a set of 59 shared alleles that appeared in all sequenced samples and performed UMAP (Uniform Manifold Approximation and Projection) analysis to cluster these common alleles, condensing them into 5 distinct clouds (Fig.3f). Clouds 1, 2, 3, and 4 were dominated by alternative subsets of PEPPR alleles distinguished primarily by the length of deletion (the average deletion sizes were 24bp, 40bp, 31bp for PEPPR Mini, Midi-I and Midi-II cluster, and it was longer than 55bp for PEPPR Maxi cluster), while cloud 5 was predominantly comprised of MMEJ alleles. We reviewed raw sequences for the few trans-cloud assigned alleles and discovered that some of these alleles could be interpreted as having been generated from a second round of repair using one of the core alleles from the same cloud as a repair template. For example, we inferred that allele 58 was actually a PEPPR deletion with several nucleotides potentially having been back-filled. This result is consistent with the previous report that alleles with insertions or complex repair outcomes would be generated from several rounds of synthesis following the generation of a primary deletion event57,58. Assessing the impact of such potential complexities, which we ignore here for simplicity, will require additional future scrutiny. The remainder of these alleles, such as allele 44, could be accounted for variability in the exact Cas9 cleavage site (between the 6th and 7th nucleotidescounting from the PAMside), with an extra nucleotide being deleted on the PAM-proximal side of the gRNA cleavage site (Fig.3f)43,59,60. Since both of these outcomes were rare, we hypothesized second-order origins for such outlier alleles further validate the robust nature of our ICP platform in recognizing core primary categories of DNA repair outcomes. We also analyzed the common 59 alleles by plotting their read frequencies and observed that the differences between the allelic landscapes for the two reciprocal crosses per each Cas9 source mirrored the trend in Fig.3ad described above (Supplementary Fig. S2a, b). Cumulatively, these concordant findings support a key role for theparental origin of Cas9 servingas a major determinant of the DSB repair outcome.

Another obvious determinant of DSB repair outcome is the local genomic DNA context. We assessed the general applicability of theICP by employing it to classify alleles generated by gRNAs targeting four other loci: prosalpha2 (pros2), Rab11, Spo11 and Rab5 using the vasa-Cas9 source61. Paralleling our findings from the ple locus, we observed divergent allelic profiles between Paternal-S and Maternal-S crosses with distinct dominant mutation categories based on the specific target site. For example, the predominant allelic classes generated at the Spo11, pros2 and Rab11 loci were PEPPR and INSRT alleles, while PEPPR and MMEJ alleles were most prevalent for the Rab5 targets (Fig.3g, h, Supplementary Figs. S36). Among these four targets, Spo11 displayed the greatest divergence in the prevalence of top alleles generated from Maternal-S and Paternal-S crosses (reminiscent of the fine distinctions parsed for the ple locus, Fig.3g). We nonetheless still observed high correlation coefficients between two replicates within the same cross and significantly lower correlation coefficients associated with inter-cross comparisons between maternal versus paternal Cas9 inheritance (averaged r2=0.33, Fig.3i, j, Supplementary Fig. S3). We also observed distinctive sex-specific DSB repair patterns for Cas9 transmission at the pros2 and Rab11 gRNAs targeting sites (Supplementary Figs. S4 and S5), although these differences were less pronounced than for ple and Spo11 gRNAs, while for Rab5, the allelic patterns were similar for both maternal and paternal crosses (Supplementary Fig. S6, see Supplementary Discussion Section). In summary, these data support the broad utility of the ICP pipeline to deliver unique discernable locus-specific fingerprints associated with distinct parental inheritance patterns of Cas9 that generalize to other genomic targets.

Given the strong Cas9 inheritance-dependent distinctions observed for allelic profiles resulting from maternal versus paternal Cas9/gRNA-induced DSBs in Drosophila, we wondered whether similar DSB repair pattern fingerprints could be discerned in mosquitoes carrying a linked full gene-drive in which the Cas9 and gRNA transgenes are carried together in a single cassette62,63,64,65. We examined this possibility using the transgenic An. stephensi Reckh drive,which is inserted into the kynurenine hydroxylase (kh) locus63. Because of the Cas9 and gRNA linkage, the Reckh drive behaves as the Maternal-F and Paternal-F cross configurations described above in which all CRISPR components are carried by a single parental sex63.

Consistent with our observations in flies, the Reckh Maternal-F crosses generated a high proportion of indels that were dominated to a remarkable extent by single mutant alleles with read percentages exceeding 85% for each of the three single mosquitoes sequenced, followed by a long distributed tail of lower frequency alleles. The highly biased nature of the replicate allelic distributions is readily revealed by a virtual step-function in their rank-ordered allelic landscapes (Fig.4a). In striking contrast, over 50% alleles recovered from the Paternal-F crosses were wild-type (WT), which presumably reflects alleles that either remained uncut or DSB ends that were rejoined accurately without further editing. The highly predominant WT allele was followed by a very shallow tail distribution of low frequency mutant alleles in the paternal rank-ordered allelic landscapes (Fig.4a). This dramatic difference in allelic profiles between Maternal-F versus Paternal-F crosses was also clearly displayed by the class-tally bars color coded for the different fractions of each class (black = WT) located beneath each landscape (Fig.4a). Here, the Class Fraction Index measure indicated that Maternal-F crosses generated a greater proportion of INSRT alleles in the first two samples, while Paternal-F crosses produced a high frequency of PEPPR alleles (Fig.4b). As in the case of allelic profiles recovered at the ple and Spo11 loci in flies, common sets of highly correlated mutant DSB repair fingerprints were observed across all three replicates of the Paternal-F Reckh crosses (Supplementary Fig. S7). A similar comparison of allelic distributions in the maternal crosses was precluded by virtue of the single highly dominant alleles and corresponding paucity of lower frequency events, the nature of which varied greatly between replicates. We conclude that the high-resolution performance of the ICP platform in Drosophila can be generalized to other insects such as An. stephensi to robustly discern sex-dependent CRISPR transmission patterns resulting in distinct DSB repair outcomes.

a Rank-ordered landscapes of the top 50 alleles generated from NGS analysis of single mosquitoes. Colored bars with red dots indicate mutated alleles, and black bars with black dots indicate an unmutated WT allele. Middle panels: allelic class fingerprints color coded as in previous figures. Bottom bars: fraction of each allelic class, including WT (black), PEPPR (sky-blue), MMEJ (deep pink), DELET (orange) and INSRT (purple). Numbers indicate the percentage of the corresponding class. b Class Fraction Index for single mosquito sequencing data in panel a. c Developmental time-points for sample collections. d Kinetics of Cas9 mutagenesis generated by the Reckh gRNA. Lines represent the summed fraction of mutant alleles at each time-point. Dark-blue lines indicate maternal (Maternal-F) crosses and red lines paternal (Paternal-F) crosses. e DSB repair fingerprints at different timepoints. Samples were collected at the time points shown in panel c and 20 eggs, larvae, pupae or adults were pooled together for genomic DNA extraction and deep sequencing. The far left and far right panels indicate the Class percentages including WT alleles (black), displaying the proportion of each class at single time-points. Source data are provided as a Source Data file.

Given the dramatic differences we observed in the frequency and nature of somatic alleles generated in maternal versus paternal-sourced Cas9 in both flies and mosquitoes, we wondered whether the developmental timing of Cas9/gRNA expression (maternal=early? and paternal=late?) was the key determinant for these highly reproducible DSB repair fingerprints. We tested this hypothesis by assessing whether DSB repair fingerprints varied as a function of developmental progression using a series of narrowly timed sample collections of F1 mosquitoes produced from crosses of Reckh parents to WT and assayed DSB repair spectra using the ICP pipeline at 12 different developmental stages (Fig.4c. Note: as homozygous Reckh transgenic mosquitoes were crossed to WT, all F1 progeny carried one Reckh allele and one WT receiver allele, the latter of which was amplified for DSB repair analysis). We tracked a diminishing proportion of WT (presumably uncut) alleles and a corresponding increase in mutant alleles of various classes at each of the time points (Fig.4d). Strikingly, nearly half of the target alleles were edited in embryos by 30minutes post-oviposition for both the Maternal-F and Paternal-F Reckh crosses, which corresponds to early pre-blastoderm stages prior to the maternal-to-zygotic transition, suggesting a very early activity of Cas9 in mosquito embryos driven either by maternally inherited Cas9/gRNA complexes or potentially by very early zygotic expression of the Cas9 and gRNA components (Fig.4d)66. We also observed similarly frequent indels being generated as early as 30min in flies expressing Cas9 (either maternally or paternally) with a gRNA targeting the pros2 locus, although the dynamics of Cas9 production are distinct in these two organisms (Supplementary Fig. S8a). Following this initial surge in target cleavage, we observed divergent trajectories in the accumulation of mutant alleles between maternal versus paternal lineages. As an overall trend, mutant alleles accumulated progressively in the Maternal-F lineage until virtually no WT alleles remained, while in Paternal-F lineage, even at the endpoint of adulthood, approximately 60% of WT alleles persisted, in line with our single time point experiments (Fig.4a, d, Supplementary Fig. S8b). As observed in the final distributions of adult alleles, progeny from Maternal-F crosses tended to be enriched for INSRT alleles over the entire developmental time course, while PEPPR alleles were more common in Paternal-F crosses with pronounced accumulation of such alleles during later stages (Fig.4e). A finer scale analysis of the categories of mutant alleles generated over time revealed dynamic patterns of prevalent alleles during mosquito developmental stages (Fig.4e). For example, the proportion of MMEJ alleles peaked at the 2-hour and 4-hour time points (Fig.4e). Similarly, a split-drive expressing a gRNA targeting the Drosophila pros2 locus generated distinct temporal profiles of cleavage patterns in crosses from female versus male parents carrying the drive element (Supplementary Fig. S9).

One unexpected feature of the developmental variations in allelic composition we observed was that the proportion of WT alleles increased at certain time points (e.g., 1-hour in maternal cross and 12-hour - day 1=24h in paternal cross). These temporal fluctuations were also observed in flies expressing Cas9 and a pros2 gRNA at two hours after oviposition (Supplementary Figs. S8a and S9), revealing that this phenomenon might reflect a generally relevant form of clonal selection for WT cells during pre-blastoderm stages. The latter clonal selection might arise if mutant cells experienced negative selection at certain development stages. In the case of paternal transmission, one strong line of evidence supporting this WT clonal selection hypothesis is that in adults, the Reckh element is transmitted to over 99% of F1 progeny, indicating that nearly all target alleles in the germline must be WT. This high frequency of paternal germline transmission is also consistent with the high prevalence of WT alleles tallied at 12h in embryos derived from the paternal crosses (Fig.4e, see Supplementary Discussion Section for more in-depth consideration of this point). We analyzed the developmental distributions of 21 common alleles that were generated at all time-points (Supplementary Fig. S10ae). Most of these common alleles belonged to the PEPPR class, while only five were INSRT alleles, despite the INSRT class overall being the most prevalent for both crosses, again suggesting that INSRT alleles have a higher diversity than other mutation categories (Supplementary Fig. S10a). Overall, this analysis is in line with our previous observation that Maternal-F crosses produced more INSRT alleles while Paternal-F crosses generated a preponderance of PEPPR alleles (Supplementary Fig. S10b).

Given the strong influence of maternal versus paternal origin of Cas9 on the resulting distributions of alleles characterized above by ICP analysis, we wondered whether such allelic signatures could be exploited for lineage tracing in randomly mating multi-generational population cages. We first examined ICP outputs from a controlled crossing scheme carried out over three generations with pleCC and Reckh gRNAs to derive allelic fingerprints distinguishing parents of origin by identifying both somatic alleles in the F1 generation as well as assessment of which of those alleles might be transmitted through the germline to non-fluorescent progeny (i.e., those not inheriting the pleCC or Reckh element) at the F2 generation (Fig.5ad, Supplementary Fig. S11). As anticipated, in both pleCC and Reckh Maternal-F crosses, single dominant somatic alleles were observed in the F1 generation, with the top single allele representing more than 50% of all alleles (Fig.5a, c). Furthermore, all such predominant somatic mutant alleles, which precluded gene-cassette copying of the pleCC or Reckh drive elements in those F1 individuals, were transmitted faithfully through the germline to non-fluorescent F2 progeny with approximately 50% frequency. Furthermore, we observed marked differences in the other half of total reads in F2 progeny depending on the origin of Cas9/gRNA complexes. Thus, a distribution of multiple diverse low frequency mutations were generated when crossing F1 pleCC+ or Reckh+ females with WT males (presumably derived from F1 drive females having deposited Cas9/gRNA complexes maternally that then acted on the paternally sourced WT allele somatically in F2 individuals). In the reciprocal male cross, however, approximately 50% of all alleles remained WT (Fig.5b, d, Supplementary Fig. S12af). These findings support the hypothesis that the top somatic indels derived from maternal Cas9 sources were generated at very early developmental stages (possibly at the point of fertilization or shortly thereafter during the first somatic cell division), resulting in a single mutant allele being initially produced and then transmitted to every descendent cell including all germline progenitor cells49. With the paternal-sourced Cas9 and gRNA, arrays of variable somatic mutations were recovered with the most prominent alleles accounting for fewer than 10% of the total alleles in F1 progeny (Fig.5b). Accordingly, paternally generated F1 somatic alleles were more randomly transmitted via the germline of individuals that failed to copy the gene cassette for either the pleCC or Reckh elements. As a result of this diversity of somatic F1 alleles, only occasionally were the most prevalent alleles also transmitted through germline (e.g., individuals 1, 4 and 5 in Fig.5b, Supplementary Fig. S12gl).

Primary DNA sequences of top single alleles and their percentages of the total alleles from six individual sequenced flies derived from ple gRNA Maternal-F (a) and Paternal-F (b) crosses. Gray bars indicate the location of the gRNA protospacer and red arrowheads are the associated PAM sites. The first row depicts the reference sequence covering the expected DSB cleavage site. Colored squares in the right column indicate the class to which a given allele belongs to. The tables shown on the right of each allele show its frequency among all reads. Left columns of the table indicate frequencies of the somatic allele, and the right columns are the top germline mutant allele frequency obtained by sequencing F2 non-fluorescence progeny derived from same F1 individuals whose top somatic allele is displayed in the left column (excluding WT alleles). Colored dots indicate different alleles with the same color shared between two columns indicating that the same allele appeared as both top 1 somatic and germline indels from the same F0 founders. c, d Allele profiles generated by Reckh parents and progeny generated with the same crossing scheme as for the pleCC. c Tabulation of the Maternal-F cross. d Tabulation of the Paternal-F cross. e Crossing scheme forthe Reckh cage trials. Three individual cages were seeded with 10 homozygous Reckh females, 90 WT females and 100 WT males for the maternally initiated lineage, while the paternally initiated cages were seeded with 10 homozygous Reckh males, 90 WT males and 100 WT females. At each of the following three generations, 10 Reckh+ females and 10 Reckh+ males were randomly collected for single mosquito deep sequencing. f Biased inheritance of Reckh was observed in the maternally seeded cages at generations 2 and 3, but not for the paternally seeded cages. Pink bars denote the fraction of sequenced individual mosquitoes inheriting Reckh from female parents, and cyan colored bars represent Reckh inheritance from the males. Source data are provided as a Source Data file.

The Reckh element in mosquitoes performed similarly to the fly pleCC, however, Reckh F1 individuals displayed less frequent zygotic cleavage and a corresponding reduction in the diversity of resulting somatically generated mutations (>50% WT alleles remained, Paternal-F cross). Consistent with this limited number and array of somatic mutations in the F1 generation from Paternal-F cross, NHEJ mutations were only rarely transmitted to the F2 generation, probably due to more germline-restricted expression of vasa-Cas9 in mosquitoes as compared to flies (Fig.5c, d). These results again suggest that cleavage and repair events were generated later during development in paternal crosses resulting in a stochastic transmission of F1 somatic alleles to the germline, which were largely uncorrelated with the most prevalent allele present somatically in the F1 parent49. Taken together, these highly divergent sex-dependent DSB repair signatures suggested that such genetic fingerprints could be used to track parental history in the context of randomly mating multi-generation population cages.

Based on the highly dominant mutant indels (Maternal-F) versus WT (Paternal-F) alleles generated by Reckh genetic element described above, we evaluated inheritance patterns of indels in multi-generational cages initiated by a 5% introduction of Reckh into WT populations either through maternal or paternal lineages in the F0 generation (Fig.5e). We randomly selected at least 20 fluorescence marker-positive mosquitoes (10 females and 10 males) for NGS analysis at generations 2 and 3, when the Reckh allele was still present at relatively low frequencies in the population and random mating was more likely to have taken place between Reckh/+ heterozygous and WT mosquitoes. Thus, we envisioned that the source of Reckh allele could be tracked back to a male versus female parent of origin by examining whether a dominant WT allele was present (inherited paternally) or not (inherited maternally) (Fig.5e, f). Following this reasoning, we inferred a strong bias for progeny inheriting the Reckh element from a Reckh+ males mating with WT females during generations 2 and 3 than the reverse (i.e., female transmission of Reckh alleles) in the maternally seeded lineage. Indeed, in one maternally seeded replicate (cage 2, generation 3), 100% of the progeny had inherited the Reckh element from their fathers (Fig.5f). In contrast to the striking sex-specific transmission bias observed in maternally seeded cages, progeny from paternally seeded cages displayed more evenly distributed stochastic parental inheritance patterns (Fig.5f). These highly reproducible parent of origin signatures demonstrate the utility of ICP in allelic lineage tracking, which could be of great potential utility in evaluating alternative initial release strategies for gene-drive mosquitoes as well as post-release surveillance of gene-drives as they spread through wild target populations (see Discussion).

Another important challenge for deciphering DSB repair outcomes is to track both NHEJ and gene-cassette mediated HDRevents within the same sample. Such a comprehensive genetic detection tool could have broad impactful applications (see Discussion). For example, one important and non-trivial application is to follow the progress of gene-drives in a marker free fashion as they spread through insect populations. Such dual tracking capability would address the potential concern that mutations eliminating a dominant marker for the gene-drive element could evade phenotype-based assessments of the drive process. Accordingly, we devised a three-step short-amplicon based deep sequencing (200400bp) strategy based on tightly linked colony-specific nucleotide polymorphisms distinguishing donor versus receiver chromosomes to detect copying of two CopyCatcher elements, pleCC and hthCC, from their chromosomes of origin (donor chromosome) to WT homologous (receiver chromosome) targets (Fig.6a)49. Notably, this strategy only amplified the inserted gene cassette on the donor chromosome and or the cassette if it copied onto the receiver chromosome. Thus, the measured allelic frequencies indicate the relative proportions of gene cassettes copied to the receiver chromosome versus those residing on the donor chromosome (Fig.6b displays the inferred somatic HDR frequency quantified from the three-step NGS sequencing protocol as well as Indels quantified by our standard 2-step NGS sequencing protocol - see Methods section for additional details).

a Scheme for tracking gene-drive copying using NGS. Gray bars: genomic DNA, pink oval: Cas9 protein, sky-blue line: gRNA, colored asterisks: polymorphisms. Color coded rectangles represent four nucleotides. Four possible recombinants listed are generated by resolving Holliday junctions at different sites marked with black crosses. b NGS sequencing-based quantification of somatic HDR generated by pleCC in F1 progeny. Areas delineated by dotted lines indicate patches of cells in which somatic HDR copying events have taken place either under bright field (upper) or RFP fluorescent filed (middle). Bottom bars are the summary of the inferred frequency for the somatic HDR (orange), indels (green) and WT alleles (black) derived from the deep sequencing data using the same samples photographed above. More than three flies from each cross were imaged and used for analysis. Scale bars indicate 200 pixels. c Somatic HDR profile with ple gRNA. The red line is for Maternal-F cross and dark blue line for the Paternal-F cross. d Diagram of the hthCC. Black double arrow: recoded hth cDNA, blue rectangles: exon 1, light green rectangles: exons 2-14, and colored lines underneath represent probes used for detection. e In situ images with embryos laid from hthCC-vasa-Cas9 females crossed with WT males. Blue=exon 1, green=WT exons 2-14, red=recoded cDNA for exons 2-14. Insets are magnified single nuclei indicated by colored arrows. This experiment has been repeated at least three times. Scale bars stand for 10m. f Temporal profiles for somatic HDR-mediated copying of the hthCC element assessed by NGS as described for the pleCC in panels c and f. Y-axis tabulates the percentage of HDR at a given time point. Table at the bottom quantifies the HDR fraction at given time points for both the Paternal-F and Maternal-F crosses. Source data are provided as a Source Data file.

In our first set of experiments, we analyzed editing outcomes by examining F1 progeny derived from Maternal-S and Paternal-S pleCC crosses. We compared the rates of somatic HDR measured by NGS analysis to those evaluated by image-based phenotypes associated with copying of the CopyCatcher element. As summarized previously, CopyCatchers such as the pleCC are designed to permit quantification of concordant homozygous mutant clonal phenotypes (e.g., pale patches of thoracic cuticle and embedded sectors ofcolorless bristles), with underlying DsRed+ fluorescent cell phenotypes49. Individual flies in which imaging-based analysis had been conducted were then subject toseparate NGS HDR-fingerprinting and INDELs-fingerprinting resulting in a comprehensive quantification of HDR, NHEJ, and WT alleles within the same sample (Fig.6b, libraries for HDR-fingerprinting and INDELs-fingerprinting were prepared from the same individual fly, but with different DNA preparation and sequencing protocols as detailed description in Methods). For these experiments, F1 flies were genotyped and those carrying both Cas9 and pleCC gRNA were used for NGS analysis (data shown here are the inferred frequencies of somatic HDR, NHEJ events, and WT alleles). This dual integrated analysis revealed that HDR in the Maternal-S crosses resulted in ~15% somatic HDR-mediated cassette copying events on average based on sequencing, and that such cassette copying was yet more frequent in Paternal-S crosses, producing ~25% somatic HDR. The nearly two-fold greater HDR-mediated copying efficiency detected by sequencing in Paternal-S crosses mirrors phenotypic outcomes wherein maternally inherited Cas9 similarly results in a lower frequency of cassette copying detected by fluorescence image analysis in somatic cells than for paternally inherited Cas9 (Fig.6b)49.

Our genetic analysis of stage-dependent differences in DSB repair pathway activity in this study is consistent with a commonly held view in the gene-drive field based on a variety of indirect genetic transmission data that HDR-mediated cassette copying does not occur efficiently during early embryonic stages50,51,63,67,68,69,70. This inference, however, has not yet been verified experimentally. We thus sought to provide direct evidence supporting this key supposition using NGS-based HDR-fingerprinting to track the somatic HDR events across a range of developmental stages in both Maternal-F and Paternal-F crosses in which the Cas9 and gRNA transgenes are transmitted together either maternally or paternally using our validated NGS sequencing protocol. Notably, we collected samples at 9 timepoints and pooled 20 F1 progeny together for pooled sequencing to prime the developmental profile of somatic HDR with pleCC (samples were thus collected without genotyping since it is impractical to genotype individual embryos and young larvae). Because of the limitations imposed by embryo pooling we were unable to use the same samples collected here for also quantifying the generation of somatic NHEJ alleles (i.e., only half of the F1 progeny carried the vasa-Cas9 transgene on the X chromosome and those embryos lacking this transgene were not suitable for generating mutations - note that such an analysis was possible in the case of the viable Reckh drive shown in Fig.4e as well as for a viable split-drive allele inserted into the essential prosalpha2 locus shown in Supplementary Fig. S9). Indeed, NGS analysis detected only very rare examples of somatic HDR events in early embryos derived from both crosses (Fig.6c). Notably, HDR in the Paternal-F cross detected by this sequencing protocol increased substantially to 35.9% during adult stages, a period coinciding with the temporal peak of the pale expression profile (note that in this experiment we employed the actin-Cas9 rather than vasa-Cas9 source, which has higher level of Cas9 expression in somatic cells and generates a correspondingly higher frequency of somatic HDR)49.

We extended our sequencing-based strategy to quantify somatic HDR using a second CopyCatcher element (hthCC) designed specifically to identify even rare copying events in early blastoderm-stage embryos. The hthCC is inserted into the homothorax (hth) gene and was engineered to visualize HDR-mediated copying of the gene cassette by fluorescence in situ hybridization (FISH) using discriminating fluorescent RNA probes complementary to specific endogenous versus recoded cDNA sequences (Fig.6d, e). In this system, copying of the transgene from the donor chromosome to the receiver chromosome would be indicated by the presence of two nuclear dots of red fluorescence detected by the hth recoded cDNA-specific probe (indicating two copies of recoded hth cDNA). In contrast, cells in which no copying occurred should contain only a single nuclear red dot signal (from the donor allele). Such in situ analysis detected no clear case of gene cassette copying in any of the ~5000 blastoderm stage cells examined across ~500 embryos (with the caveat that some mitotic nuclei generate ambiguous signals depending on their orientation). This qualified negative result assessed by in situ analysis was consistent with the very low estimates of HDR frequency during the same early blastoderm-stage developmental window based on NGS analysis in staged time-course experiments, although the latter sequencing method did detect very low levels of somatic HDR at ~3hours after egg laying from the Paternal-F crosses (and no copying until day three of larvae with the maternal cross Fig.6df). The very low levels of somatic HDR observed in early embryos for the hthCC construct either by in situ hybridization or by NGS sequencing parallel the results summarized above for the pleCC element (Fig.6c, f). The maximal somatic HDR frequency observed for the hthCC Maternal-F crosses (0.06% at day 3 after egg laying) was somewhat lower than that for the similar cross for pleCC (0.35% at adult stage), consistent with the predominance of single mutant alleles being generated at very early stages following fertilization in Maternal-F crosses. In contrast to the exceedingly rare copying of the hthCC element detected in early embryos for either the Maternal-F or Paternal-F crosses, the same element frequently copied to the homologous chromosome during later developmental stages in Paternal-F crosses as assessed by NGS sequencing. The hthCC elementagain copied with somewhat lower efficiency than the pleCC element (e.g., 15.2% for hthCC versus 35.9% for pleCC tabulated in adults), presumably reflecting differing genomic cleavage rates or gene conversion efficiencies generated by their respective gRNAs (including total cleavage levels and temporal features). In aggregate, these two examples of quantitative analysis of copying frequencies based on both NGS and in situ analysis demonstrate that ICP and NGS-based quantification of gene conversion events can be successfully integrated for a comprehensive analysis of DSB repair outcomes, including both NHEJ and HDR events as a function of developmental stage. These powerful tools also could be applied for following gene-drive spread through freely mating populations in a marker-free manner as well as for a variety of other applications including gene therapy (see Discussion).

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Developmental progression of DNA double-strand break repair deciphered by a single-allele resolution mutation ... - Nature.com

Abstract

Genome editing is the modification of genomic DNA at a specific target site in a wide variety of cell types and organisms, including insertion, deletion and replacement of DNA, resulting in inactivation of target genes, acquisition of novel genetic traits and correction of pathogenic gene mutations. Due to the advantages of simple design, low cost, high efficiency, good repeatability and short-cycle, CRISPR-Cas systems have become the most widely used genome editing technology in molecular biology laboratories all around the world. In this review, an overview of the CRISPR-Cas systems will be introduced, including the innovations, the applications in human disease research and gene therapy, as well as the challenges and opportunities that will be faced in the practical application of CRISPR-Cas systems.

Keywords: CRISPR, Cas9, Genome editing, Human disease models, Rabbit, Gene therapy, Off target effects

Genome editing is the modification of genomic DNA at a specific target site in a wide variety of cell types and organisms, including insertion, deletion and replacement of DNA, resulting in inactivation of target genes, acquisition of novel genetic traits and correction of pathogenic gene mutations [1], [2], [3]. In recent years, with the rapid development of life sciences, genome editing technology has become the most efficient method to study gene function, explore the pathogenesis of hereditary diseases, develop novel targets for gene therapy, breed crop varieties, and so on [4], [5], [6], [7].

At present, there are three mainstream genome editing tools in the world, zinc finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs) and the RNA-guided CRISPR (clustered regularly interspaced short palindromic repeats)-Cas (CRISPR-associated) nucleases systems [8], [9], [10]. Due to the advantages of simple design, low cost, high efficiency, good repeatability and short-cycle, CRISPR-Cas systems have become the most widely used genome editing technology in molecular biology laboratories all around the world [11], [12]. In this review, an overview of the CRISPR-Cas systems will be introduced, including the innovations and applications in human disease research and gene therapy, as well as the challenges and opportunities that will be faced in the practical application of CRISPR-Cas systems.

CRISPR-Cas is an adaptive immune system existing in most bacteria and archaea, preventing them from being infected by phages, viruses and other foreign genetic elements [13], [14]. It is composed of CRISPR repeat-spacer arrays, which can be further transcribed into CRISPR RNA (crRNA) and trans-activating CRISPR RNA (tracrRNA), and a set of CRISPR-associated (cas) genes which encode Cas proteins with endonuclease activity [15]. When the prokaryotes are invaded by foreign genetic elements, the foreign DNA can be cut into short fragments by Cas proteins, then the DNA fragments will be integrated into the CRISPR array as new spacers [16]. Once the same invader invades again, crRNA will quickly recognize and pair with the foreign DNA, which guides Cas protein to cleave target sequences of foreign DNA, thereby protecting the host [16].

CRISPR-Cas systems can be classified into 2 classes (Class 1 and Class 2), 6 types (I to VI) and several subtypes, with multi-Cas protein effector complexes in Class 1 systems (Type I, III, and IV) and a single effector protein in Class 2 systems (Type II, V, and VI) [17], [18]. The classification, representative members, and typical characteristics of each CRISPR-Cas system are summarized in [10], [12], [15], [16], [17], [18].

Summary of CRISPR-Cas systems.

Type II CRISPR-Cas9 system derived from Streptococcus pyogenes (SpCas9) is one of the best characterized and most commonly used category in numerous CRISPR-Cas systems [18], [19]. The main components of CRISPR-Cas9 system are RNA-guided Cas9 endonuclease and a single-guide RNA (sgRNA) [20]. The Cas9 protein possesses two nuclease domains, named HNH and RuvC, and each cleaves one strand of the target double-stranded DNA [21]. A single-guide RNA (sgRNA) is a simplified combination of crRNA and tracrRNA [22]. The Cas9 nuclease and sgRNA form a Cas9 ribonucleoprotein (RNP), which can bind and cleave the specific DNA target [23]. Furthermore, a protospacer adjacent motif (PAM) sequence is required for Cas9 proteins binding to the target DNA [20].

During genome editing process, sgRNA recruits Cas9 endonuclease to a specific site in the genome to generate a double-stranded break (DSB), which can be repaired by two endogenous self-repair mechanisms, the error-prone non-homologous end joining (NHEJ) pathway or the homology-directed repair (HDR) pathway [24]. Under most conditions, NHEJ is more efficient than HDR, for it is active in about 90% of the cell cycle and not dependent on nearby homology donor [25]. NHEJ can introduce random insertions or deletions (indels) into the cleavage sites, leading to the generation of frameshift mutations or premature stop codons within the open reading frame (ORF) of the target genes, finally inactivating the target genes [26], [27]. Alternatively, HDR can introduce precise genomic modifications at the target site by using a homologous DNA repair template [28], [29] (). Furthermore, large fragment deletions and simultaneous knockout of multiple genes could be achieved by using multiple sgRNAs targeting one single gene or more [30], [31].

Mechanism of genome editing. Double-strand break (DSB) induced by nucleases can be repaired by non-homologous end joining (NHEJ) or homology-directed repair (HDR) pathways. NHEJ can introduce random insertions or deletions (indels) of varying length at the site of the DSB. Alternatively, HDR can introduce precise genomic modifications at the target site by using a homologous DNA donor template.

CRISPR-Cas systems have become the most favorite genome editing tool in the molecular biology laboratory since they were confirmed to have genome editing capabilities in 2012 [23]. They have made numerous achievements in the field of correcting pathogenic mutations, searching for essential genes for cancer immunotherapy, and solving key problems in organ xenotransplantation [5], [32], [33]. Unfortunately, there are still some limitations which need to solve in CRISPR-Cas systems, such as potential off-target effects, limited genome-targeting scope restricted by PAM sequences, and low efficiency and specificity [34], [35]. Therefore, many research teams have been trying to improve this tool.

By introducing two point mutations, H840A and D10A, into HNH and RuvC nuclease domain, researchers have obtained a nuclease dead Cas9 (dCas9) [36]. The dCas9 lacks DNA cleavage activity, but DNA binding activity is not affected. Then, by fusing transcriptional activators or repressors to dCas9, the CRISPR-dCas9 system can be used to activate (CRISPRa) or inhibit (CRISPRi) transcription of target genes [37], [38]. Additionally, dCas9 can be fused to various effector domains, which enables sequence-specific recruitment of fluorescent proteins for genome imaging and epigenetic modifiers for epigenetic modification [39], [40]. Furthermore, this system is easy to operate and allows simultaneous manipulation of multiple genes within a cell [38].

In order to improve the efficiency of site-directed mutagenesis, base editing systems containing dCas9 coupled with cytosine deaminase (cytidine base editor, CBE) or adenosine deaminase (adenine base editor, ABE) have been developed [41], [42]. It can introduce CG to TA or AT to GC point mutations into the editing window of the sgRNA target sites without double-stranded DNA cleavage [41], [42]. Since base editing systems avoid the generation of random insertions or deletions to a great extent, the results of gene mutation are more predictive. However, owing to the restriction of base editing window, base editing systems are not suitable for any target sequence in the genome. Accordingly, C-rich sequences, for example, would produce a lot of off-target mutations [43]. Therefore, researchers have always been trying to develop and optimize novel base editing systems to overcome this drawback [44]. At present, base editing systems have been widely used in various cell lines, human embryos, bacteria, plants and animals for efficient site-directed mutagenesis, which may have broad application prospects in basic research, biotechnology and gene therapy [45], [46], [47]. In theory, 3956 gene variants existing in Clin var database could be repaired by base substitution of C-T or G-A [42], [48].

An NGG PAM at the 3 end of the target DNA site is essential for the recognization and cleavage of the target gene by Cas9 protein [20]. Besides classical NGG PAM sites, other PAM sites such as NGA and NAG also exist, but their efficiency of genome editing is not high [49]. However, such PAM sites only exist in about one-sixteenth of the human genome, thereby largely restricting the targetable genomic loci. For this purpose, several Cas9 variants have been developed to expand PAM compatibility.

In 2018, David Liu et al.[50] developed xCas9 by phage-assisted continuous evolution (PACE), which can recognize multiple PAMs (NG, GAA, GAT, etc.). In the latter half of the same year, Nishimasu et al. developed SpCas9-NG, which can recognize relaxed NG PAMs [51]. In 2020, Miller et al. developed three new SpCas9 variants recognizing non-G PAMs, such as NRRH, NRCH and NRTH PAMs [52]. Later in the same year, Walton et al. developed a SpCas9 variant named SpG, which is capable of targeting an expanded set of NGN PAMs [53]. Subsequently, they optimized the SpG system and developed a near-PAMless variant named SpRY, which is capable of editing nearly all PAMs (NRN and NYN PAMs) [53].

By using these Cas9 variants, researchers have repaired some previously inaccessible disease-relevant genetic variants [51], [52], [53]. However, there are still some drawbacks in these variants, such as low efficiency and cleavage activity [50], [51]. Therefore, they should be further improved by molecular engineering in order to expand the applications of SpCas9 in disease-relevant genome editing.

In addition to editing DNA, CRISPR-Cas systems can also edit RNA. Class 2 Type VI CRISPR-Cas13 systems contain a single RNA-guided Cas13 protein with ribonuclease activity, which can bind to target single-stranded RNA (ssRNA) and specifically cleave the target [54]. To date, four Cas13 proteins have been identified: Cas13a (also known as C2c2), Cas13b, Cas13c and Cas13d [55]. They have successfully been applied in RNA knockdown, transcript labeling, splicing regulation and virus detection [56], [57], [58]. Later, Feng Zhang et al. developed two RNA base edting systems (REPAIR system, enables A-to-I (G) replacement; RESCUE system, enables C-to-U replacement) by fusing catalytically inactivated Cas13 (dCas13) with the adenine/cytidine deaminase domain of ADAR2 (adenosine deaminase acting on RNA type 2) [59], [60].

Compared with DNA editing, RNA editing has the advantages of high efficiency and high specificity. Furthermore, it can make temporary, reversible genetic edits to the genome, avoiding the potential risks and ethical issues caused by permanent genome editing [61], [62]. At present, RNA editing has been widely used for pre-clinical studies of various diseases, which opens a new era for RNA level research, diagnosis and treatment.

Recently, Anzalone et al. developed a novel genome editing technology, named prime editing, which can mediate targeted insertions, deletions and all 12 types of base substitutions without double-strand breaks or donor DNA templates [63]. This system contains a catalytically impaired Cas9 fused to a reverse transcriptase and a prime editing guide RNA (pegRNA) with functions of specifying the target site and encoding the desired edit [63]. After Cas9 cleaves the target site, the reverse transcriptase uses pegRNA as a template for reverse transcription, and then, new genetic information can be written into the target site [63]. Prime editing can effectively improve the efficiency and accuracy of genome editing, and significantly expand the scope of genome editing in biological and therapeutic research. In theory, it is possible to correct up to 89% known disease-causing gene mutations [63]. Nevertheless, as a novel genome editing technique, more research is still needed to further understand and improve prime editing system.

So far, as a rapid and efficient genome editing tool, CRISPR-Cas systems have been extensively used in a variety of species, including bacteria, yeast, tobacco, Arabidopsis, sorghum, rice, Caenorhabditis elegans, Drosophila, zebrafish, Xenopus laevis, mouse, rat, rabbit, dog, sheep, pig and monkey [64], [65], [66], [67], [68], [69], [70], [71], [72], [73], [74], [75], [76], [77], [78], as well as various human cell lines, such as tumor cells, adult cells and stem cells [79], [80]. In medical field, the most important application of CRISPR-Cas systems is to establish genetically modified animal and cell models of many human diseases, including gene knockout models, exogenous gene knock-in models, and site directed mutagenesis models [80], [81].

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Establishing animal models of human diseases

Animal models are crucial tools for understanding gene function, exploring pathogenesis of human diseases and developing new drugs. However, traditional methods for generating animal models are complex, costly and time-consuming, which severely limit the application of animal models in basic medical research and preclinical studies [82]. Since the discovery of CRISPR-Cas systems, a series of genetically modified animal models have successfully been generated in a highly efficient manner [72], [73], [74], [75], [76], [77], [78].

Among numerous model animals, mice are widely used for scientific studies and recognized as the most important model animals in human disease research [83]. So far, researchers have successfully generated many genetically modified mouse models, such as cancer, cardiovascular disease, cardiomyopathy, Huntington's disease, albino, deafness, hemophilia B, obesity, urea cycle disorder and muscular dystrophy [84], [85], [86], [87], [88], [89], [90], [91], [92], [93]. Nevertheless, owing to the great species differences between humans and rodents, they cant provide effective assessment and long-term follow-up for research and treatment of human diseases [94]. Therefore, the application of larger model animals, such as rabbits, pigs and non-human primates, is becoming more and more widespread [74], [77], [78]. With the development of CRISPR-Cas systems, generating larger animal models for human diseases has become a reality, which greatly enriches the disease model resource bank.

Our research focuses on the generation of genetically modified rabbit models using CRISPR-Cas systems. Compared with mice, rabbits are closer to humans in physiology, anatomy and evolution [95]. In addition, rabbits have a short gestation period and less breeding cost. All these make them suitable for studies of the cardiovascular, pulmonary and metabolism diseases [95], [96]. Nowadays, we have generated a series of rabbit models for simulating human diseases, including congenital cataracts, duchenne muscular dystrophy (DMD), X-linked hypophosphatemia (XLH), etc (summarized in ) [97], [98], [99], [100], [101], [102], [103], [104], [105], [106], [107], [108], [109], [110], [111], [112], [113], [114]. Take the generation of PAX4 gene knockout rabbits as an example, the procedure we used to establish genetically modified rabbit models is summarized in and .

CRISPR-Cas system mediated rabbit models of human diseases.

Generation of PAX4 gene knockout (KO) rabbits using CRISPR-Cas9 system. (A) Schematic diagram of the sgRNA target sites located in the rabbit PAX4 locus. PAX4 exons are indicated by yellow rectangles; target sites of the two sgRNA sequences, sgRNA1 and sgRNA2, are highlighted in green; protospacer-adjacent motif (PAM) sequence is highlighted in red. Primers F and R are used for mutation detection in pups. (B) Microinjection and embryo transfer. First a mixture of Cas9 mRNA and sgRNA is microinjected into the cytoplasm of the zygote at the pronuclear stage. Then the injected embryos are transferred into the oviduct of recipient rabbits. After 30days gestation, PAX4 KO rabbits are born. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

Summary of the PAX4 KO rabbits generated by CRISPR-Cas9 system.

In addition, the pig is an important model animal extensively used in biomedical research. Compared with mice, their body/organ size, lifespan, anatomy, physiology, metabolic profile and immune characteristics are more similar to those of humans, which makes the pig an ideal model for studying human cardiovascular diseases and xenotransplantation [115]. At present, several genetically modified pig models have been successfully generated, including neurodegenerative diseases, cardiovascular diseases, cancer, immunodeficiency and xenotransplantation model [116], [117], [118], [119], [120], [121], [122].

To date, non-human primates are recognized as the best human disease models. Their advantage is that their genome has 98% homology with the human genome; also, they are highly similar to humans in tissue structure, immunity, physiology and metabolism [123]. Whats more, they can be infected by human specific viruses, which makes them very important models in infectious disease research [124]. Nowadays, researchers have generated many genetically modified monkey models, such as cancer, muscular dystrophy, developmental retardation, adrenal hypoplasia congenita and Oct4-hrGFP knockin monkeys [125], [126], [127], [128], [129].

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Establishing cell models of human diseases

It was found that the efficiency of CRISPR-Cas mediated genome editing is higher in vitro than in vivo, thus the use of genetically modified cell models can greatly shorten the research time in medical research [130]. Until now, researchers have used CRISPR-Cas systems to perform genetic manipulations on various cell lines, such as tumor cells, adult cells and stem cells, in order to simulate a variety of human diseases [79], [80].

Fuchs et al. generated the RPS25-deficient Hela cell line by knocking out ribosomal protein eS25 (RPS25) gene using CRISPR-Cas9 system [131]. Drost et al. edited four common colorectal cancer-related genes (APC, P53, KRAS and SMAD4) in human intestinal stem cells (hISCs) by CRISPR-Cas9 technology [132]. The genetically modified hISCs with 4 gene mutations possessed the biological characteristics of intestinal tumors and could simulate the occurrence of human colorectal cancer [132]. Jiang et al. induced site-specific chromosome translocation in mouse embryonic stem cells by CRISPR-Cas9, in order to establish a cell and animal model for subsequent research on congenital genetic diseases, infertility, and cancer related to chromosomal translocation [133].

In addition, induced pluripotent stem cells (iPSCs) have shown great application prospect in disease model establishment, drug discovery and patient-specific cellular therapy development [134]. iPSCs have the ability of self-renewal and multiple differentiation potential, which are of great significance in disease model establishment and regenerative medicine research [135]. In recent years, by combining CRISPR-Cas systems with iPSC technology, researchers have generated numerous novel and reliable disease models with isogenic backgrounds and provided new solutions for cell replacement therapy and precise therapy in a variety of human diseases, including neurodegenerative diseases, acquired immunodeficiency syndrome (AIDS), -thalassemia, etc [134], [135], [136].

With the development of CRISPR-Cas systems and the discovery of novel Cas enzymes (Cas12, Cas13, etc.), CRISPR-based molecular diagnostic technology is rapidly developing and has been selected as one of the world's top ten science and technology advancements in 2018 [137].

Unlike Cas9, Cas13 enzymes possess a collateral cleavage activity, which can induce cleavage of nearby non-target RNAs after cleavage of target sequence [54]. Based on the collateral cleavage activity of Cas13, Feng Zhang et al.[138] developed a Cas13a-based in vitro nucleic acid detection platform, named SHERLOCK (Specific High Sensitivity Enzymatic Reporter UnLOCKing). It is composed of Cas13a, sgRNA targeting specific RNA sequences and fluorescent RNA reporters. After Cas13a protein recognizes and cleaves the target RNA, it will cut the report RNA and release the detectable fluorescence signal, so as to achieve the purpose of diagnosis [138]. Researchers have used this method to detect viruses, distinguish pathogenic bacteria, genotype human DNA and identify tumor DNA mutations [137], [138]. Later, Feng Zhang et al. improved SHERLOCK system and renamed it as SHERLOCKv2, which can detect four virus at the same time [139].

In addition to Cas13, Cas12 enzymes are also found to possess collateral cleavage activity [140]. Doudna et al.[141] developed a nucleic acid detection system based on Cas12a (also known as Cpf1), named DETECTR (DNA endonuclease-targeted CRISPR trans reporter). DETECTR has been used to detect cervical cancer associated HPV subtypes (HPV16 and HPV18) in either virus-infected human cell lines or clinical patient samples [141]. Furthermore, Doudna et al. are trying to use the newly discovered Cas14 and CasX proteins in molecular diagnosis, which may further enrich the relevant techniques of CRISPR-based molecular diagnosis [142], [143].

CRISPR-based molecular diagnostic technology has incomparable advantages over traditional molecular diagnostic methods, such as high sensitivity and single-base specificity, which is suitable for early screening of cancer, detection of cancer susceptibility genes and pathogenic genes [137], [144]. Meanwhile, CRISPR diagnostics is inexpensive, simple, fast, without special instrument, and is suitable for field quick detection and detection in less-developed areas [137], [144]. At present, many companies are trying to develop CRISPR diagnostic kits for family use, to detect HIV, rabies, Toxoplasma gondi, etc.

CRISPR-Cas9 system enables genome-wide high-throughput screening, making it a powerful tool for functional genomic screening [145]. The high efficiency of genome editing with CRISPR-Cas9 system makes it possible to edit multiple targets in parallel, thus a mixed cell population with gene mutation can be produced, and the relationship between genotypes and phenotypes could be confirmed by these mutant cells [146]. CRISPR-Cas9 library screening can be divided into two categories: positive selection and negative selection [147]. It has been utilized to identify genes associated with cancer cell survival, drug resistance and virus infection in various models [148], [149], [150]. Compared with RNAi-based screening, high-throughput CRISPR-Cas9 library screening has the advantages of higher transfection efficiency, minimal off-target effects and higher data reproducibility [151]. At present, scientists have constructed human and mouse genome-wide sgRNA libraries, and they have been increasingly improved according to different requirements [152], [153]. In the future, CRISPR-Cas9-based high-throughput screening technology will definitely get unprecedented development and application.

Gene therapy refers to the introduction of foreign genes into target cells to treat specific diseases caused by mutated or defective genes [154]. Target cells of gene therapy are mainly divided into two categories: somatic cells and germ line cells. However, since germ line gene therapy is complicated in technique as well as involves ethical and security issues, today gene therapy is limited to somatic cell gene therapy [155]. Traditional gene therapy is usually carried out by homologous recombination or lentiviral delivery. Nevertheless, the efficiency of homologous recombination is low, and lentiviral vectors are randomly inserted into the recipient genome, which may bring potential security risks to clinical applications [156]. Currently, with the rapid development of CRISPR-Cas systems, they have been widely applied in gene therapy for treating various of human diseases, monogenic diseases, infectious diseases, cancer, etc [155], [156], [157]. Furthermore, some CRISPR-mediated genome-editing therapies have already reached the stage of clinical testing. briefly summarizes the ongoing clinical trials of gene therapy using genome-editing technology, including ZFN, TALEN and CRISPR-Cas systems.

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Monogenic diseases

Monogenic diseases refer to the genetic diseases caused by mutations of a single allele or a pair of alleles on a pair of homologous chromosomes [158]. There are more than 6600 known monogenic diseases around the world, -thalassaemia, sickle cell disease (SCD), hemophilia B (HB), retinitis pigmentosa (RP), leber congenital amaurosis type 10 (LCA10), duchenne muscular dystrophy (DMD), hutchinson-gilford progeria syndrome (HGPS), hereditary tyrosinemia (HT), cystic fibrosis (CF), etc [159]. Most of the monogenic diseases are rare diseases lacking of effective treatment, which will greatly affect the life quality of patients. Nowadays, many animal models of monogenic diseases have been treated with CRISPR-mediated gene therapy. Furthermore, even some CRISPR clinical trials for monogenic diseases are going on [160].

Summary of clinical trials of gene therapy using genome-editing technology.

-Thalassaemia, a hereditary hemolytic anemia disease, is one of the most common and health-threatening monogenic diseases in the world. It is characterized by mutations in the -globin (HBB) gene, leading to severe anemia caused by decreased hemoglobin (Hb) level [161]. For the moment, the only way to cure -thalassemia is hematopoietic stem cell transplantation (HSCT). Yet, high cost of treatment and shortage of donors limit its clinical application [162]. Other therapy, for example, blood transfusion, can only sustain the life of patients but cant cure the disease [161]. To better treat -thalassemia, researchers have turned their attention to gene therapy. A major technical idea is to repair the defective -globin gene of iPSCs from patients with -thalassemia by CRISPR-Cas9 technology, then red blood cells can be produced normally and the disease could be cured [163], [164]. Besides, reactivating fetal hemoglobin (HbF) expression has also been proposed to be an effective method to treat -thalassemia through knockout of BCL11A gene, which suppresses the expression of fetal hemoglobin [165], [166].

Additionally, CRISPR-Cas systems have also been used for the treatment of other hematologic diseases, such as sickle cell disease (SCD) and hemophilia B (HB). SCD is a monogenic disease caused by a single-nucleotide mutation in human -globin gene, leading to a substitution of glutamic acid by valine and the production of an abnormal version of -globin, which is known as hemoglobin S (HbS) [167]. CRISPR-Cas9 system has been used to treat SCD by repairing the -globin gene mutation or reactivating HbF expression [168], [169]. HB is an X-linked hereditary bleeding disorder caused by deficiency of coagulation factor IX, and the most common treatment for hemophilia B is supplement blood coagulation factor [170], [171]. Huai et al. injected naked Cas9-sgRNA plasmid and donor DNA into the adult mice of F9 mutation HB mouse model for gene correction [172]. Meanwhile, Cas9/sgRNA were also microinjected into germline cells of this HB mouse model for gene correction. Both in vivo and ex vivo experiment were sufficient to remit the coagulation deficiency [172]. Guan et al. corrected the F9 Y371D mutation in HB mice using CRISPR-Cas9 mediated in situ genome editing, which greatly improved the hemostatic efficiency and increased the survival of HB mice [173].

Duchenne muscular dystrophy (DMD) is an X-chromosome recessive hereditary disease, with clinical manifestations of muscle weakness or muscle atrophy due to a progressive deterioration of skeletal muscle function [174]. It is usually caused by mutations in the DMD gene, a gene encoding dystrophin protein [174]. Deletions of one or more exons of the DMD gene will result in frameshift mutations or premature termination of translation, thereby normal dystrophin protein can not be synthesized [175]. Currently, there is no effective treatment for DMD. Conventional drug treatment can only control the disease to a certain extent, but can not cure it. It was found that a functional truncated dystrophin protein can be obtained by removing the mutated transcripts with CRISPR-Cas9 system [176], [177], [178]. In addition, base editing systems can also be applied in DMD treatment by repairing single base mutation or inducing exon skipping by introducing premature termination codons (PTCs) [179].

Retinitis pigmentosa (RP) is a group of hereditary retinal degenerative diseases characterized by progressive loss of photoreceptor cells and retinal pigment epithelium (RPE) function [180]. RP has obvious genetic heterogeneity, and the inheritance patterns include autosomal dominant, autosomal recessive, and X-linked recessive inheritance [180]. To date, there is still no cure for RP. In recent years, with the rapid development of gene editing technology, there has been some progress in the treatment of RP. Several gene mutations causing RP have been corrected by CRISPR-Cas9 in mouse models to prevent retinal degeneration and improve visual function, for example, RHO gene, PRPF31 gene and RP1 gene [181], [182].

Leber Congenital Amaurosis type 10 (LCA10) is an autosomal retinal dystrophy with severe vision loss at an early age. The most common gene mutation found in patients with LCA10 is IVS26 mutation in the CEP290 gene, which disrupts the coding sequence by generating an aberrant splice site [183]. Ruan et al. used CRISPR-Cas9 system to knock out the intronic region of the CEP290 gene and restored normal CEP290 expression [184]. In addition, subretinal injection of EDIT-101 in humanized CEP290 mice showed rapid and sustained CEP290 gene editing [185], [186].

Hutchinson-Gilford Progeria Syndrome (HGPS) is a rare lethal genetic disorder with the characteristic of accelerated aging [187]. A point mutation within exon 11 of lamin A gene activates a cryptic splice site, leading to the production of a truncated lamin A called progerin [188]. However, CRISPR-Cas based gene therapy has opened up a broad prospect in HGPS treatment. Administration of AAV-delivered CRISPR-Cas9 components into HGPS mice can reduce the expression of progerin, thereby improved the health condition and prolonged the lifespan of HGPS mice [189], [190]. In addition, Suzuki et al. repaired G609G mutation in a HGPS mouse model via single homology arm donor mediated intron-targeting gene integration (SATI), which ameliorated aging-associated phenotypes and extended the lifespan of HGPS mice [191].

CRISPR-Cas systems have also showed their advantages in gene therapy of hereditary tyrosinemia (HT) and cystic fibrosis (CF). HT is a disorder of tyrosine metabolism caused by deficiency of fuarylacetoacetate hydrolase (Fah) [192]. Yin et al. corrected a Fah mutation in a HT mouse model by injecting CRISPR-Cas9 components into the liver of the mice [193]. Then, the wild-type Fah protein in the liver cells began to express and the body weight loss phenotype was rescued [193]. CF, an autosomal recessive inherited disease with severe respiratory problems and infections, has a high mortality rate at an early age [194]. It is caused by mutations in the CFTR gene, which encodes an epithelial chloride anion channel, the cystic fibrosis transmembrane conductance regulator (CFTR) [194]. Until now, genome editing strategies have been carried out in cell models to correct CFTR mutations. In cultured intestinal stem cells and induced pluripotent stem cells from cystic fbrosis patients, the CFTR homozygous 508 mutation has been corrected by CRISPR-Cas9 technology, leading to recovery of normal CFTR expression and function in differentiated mature airway epithelial cells and intestinal organoids [195], [196].

(2)

Infectious diseases

In recent years, gene therapy has gradually been applied to the treatment of viral infectious diseases. Transforming host cells to avoid viral infection or preventing viral proliferation and transmission are two main strategies for gene therapy of viral infectious diseases [197].

Human immunodeficiency virus (HIV), a kind of retrovirus, mainly attacks the human immune system, especially the CD4 T lymphocytes. When human cells are invaded by HIV, the viral sequences can be integrated into the host genome, blocking cellular and humoral immunity while causing acquired immunodeficiency syndrome (AIDS) [198]. There is still no known cure for AIDS but it could be treated. Although antiretroviral therapy can inhibit HIV-1 replication, the viral sequences still exist in the host genome, and they could be reactivated at any time [199]. CRISPR-Cas9 system can target long terminal repeat (LTR) and destruct HIV-1 proviruses, thus it is possible to completely eliminate HIV-1 from genome of infected host cells [200], [201]. In addition, resistance to HIV-1 infection could be induced by knockout of the HIV co-receptor CCR5 gene in CD4 T cells [202], [203].

Cervical cancer is the second most common gynecologic malignant tumor. The incidence is increasing year by year and young people are especially prone to this disease. It was found that the occurrence of cervical cancer is closely related to HPV (human papillomavirus) infection [204]. HPV is a double-stranded cyclic DNA virus, E6 and E7 genes located in HPV16 early regions are carcinogenic genes [205]. Researchers designed sgRNAs targeting E6 and E7 genes to block the expression of E6 and E7 protein, subsequently the expression of p53 and pRb was restored to normal, finally increasing tumor cells apoptosis and suppressing subcutaneous tumor growth in in vivo experiments [206], [207], [208]. Moreover, HPV virus proliferation was blocked through cutting off E6/E7 genes, and the virus in the bodies could be eliminated [206], [207], [208].

(3)

Cancer

Cancer is the second leading cause of death worldwide after cardiovascular diseases, and it is also a medical problem that needs to be solved urgently. A variety of genetic or epigenetic mutations have been accumulated in the cancer genome, which can activate proto-oncogenes, inactivate tumor suppressors and produce drug resistance [209], [210]. So far, CRISPR-Cas systems have been used to correct the oncogenic genome/epigenome mutations in tumor cells and animal models, resulting in inhibition of tumor cell growth and promotion of cell apoptosis, thereby inhibiting tumor growth [211], [212], [213].

In addition, immunotherapy is considered to be a major breakthrough in cancer treatment, especially chimeric antigen receptor-T (CAR-T) cell therapy, which has a significantly therapeutic effect on leukemia, lymphoma and certain types of solid tumors [214], [215], [216]. CAR-T cells are genetically manipulated, patient-specific T cells, which express receptors targeting antigens specially expressed on tumor cells, for example, CD19 CAR-T cells for B cell malignancies. Then these cells will be transfused back to patients to fight against cancer [217]. However, CAR-T cell therapy is complex, time-consuming and expensive, and it is greatly limited by the quality and quantity of autologous T cells. Therefore, researchers have used CRISPR-Cas9 system to develop universal CAR-T cells, such as simultaneously removing endogenous T cell receptor gene and HLA class I encoding gene on T cells of healthy donors and introducing CAR sequence [218], [219], [220]. Thereby, it could be used in multiple patients without causing graft versus host reaction (GVHR). In addition, CRISPR-Cas mediated genome editing has also been used to enhance the function of CAR-T cells by knocking out genes encoding signaling molecules or T cell inhibitory receptors, such as programmed cell death protein 1 (PD-1) and cytotoxic T lymphocyte antigen 4 (CTLA-4) [221], [222].

Though CRISPR-Cas mediated efficient genome editing technologies have been broadly applied in a variety of species and different types of cells, there are still some important issues needed to be addressed during the process of application, such as off-target effects, delivery methods, immunogenicity and potential risk of cancer.

It was found that designed sgRNAs will mismatch with non-target DNA sequences and introduce unexpected gene mutations, called off-target effects [223]. Off-target effects seriously restrict the widespread application of CRISPR-Cas mediated genome editing in gene therapy, for it might lead to genomic instability and increase the risk of certain diseases by introducing unwanted mutations at off-target sites [224]. At present, several strategies have been used to predict and detect off-target effects, online prediction software, whole genome sequencing (WGS), genome-wide, unbiased identification of DSBs enabled by sequencing (GUIDE-seq), discovery of in situ cas off-targets and verification by sequencing (DISCOVER-Seq), etc [225]. Furthermore, to minimize off-target effects, researchers have systematically studied the factors affecting off-target effects and developed a number of effective approaches.

(1)

Rational design and modification of sgRNAs

The specific binding of sgRNA with the target sequence is the key factor in CRISPR-Cas mediated genome editing. Rational design of highly specific sgRNAs might minimize off-target effects [224]. The length and GC content of sgRNAs, and mismatches between sgRNA and its off-target site will all affect the frequency of off-target effects [226]. In addition, on the basis of rational design of sgRNAs, the specificity of CRISPR-Cas systems can be further improved by modifying sgRNAs, such as engineered hairpin sgRNAs and chemical modifications of sgRNAs [227], [228].

(2)

Modification of Cas9 protein

As we know, the interaction between Cas9 and DNA affects the stability of DNA-Cas9/sgRNA complex as well as tolerance to mismatch [229]. Therefore, high-fidelity SpCas9 variants have been developed by introducing amino substitution(s) into Cas9 protein in order to destabilize the function structure of the CRISPR complex [230]. Researchers have developed several highly effective Cas9 mutants, high-fidelity Cas9 (SpCas9-HF1), enhanced specificity Cas9 (eSpCas9), hyper-accurate Cas9 (HypaCas9), etc [231], [232], [233]. All of them can significantly reduce off-target effects while retain robust target cleavage activity.

(3)

Adoption of double nicking strategy

Recently, a double-nicking strategy has been developed to minimize off-target effects, which employs two catalytic mutant Cas9-D10A nickases and a pair of sgRNAs to produce a cleavage on each strand of the target DNA, thus forming a functional double strand break [234]. Additionally, it was proven that the fusion protein generated by combining dCas9 with Fok nuclease can also reduce off-target effects [235]. Only when the two fusion protein monomers are close to each other to form dimers, can they perform the cleavage function [235]. This strategy could greatly reduce DNA cleavage at non-target sites.

(4)

Anti-CRISPRs

Off switches for CRISPR-Cas9 system was first discovered by Pawluk et al. in 2016. They identified three naturally existing protein families, named as anti-CRISPRs, which can specifically inhibit the CRISPR-Cas9 system of Neisseria meningitidis[236]. Later, Rauch et al. discovered four unique type IIA CRISPR-Cas9 inhibitor proteins encoded by Listeria monocytogenes prophages, and two of them (AcrllA2 and AcrllA4) can block SpCas9 when assayed in Escherichia coli and human cells [237]. Recently, Doudna et al. discovered two broad-spectrum inhibitors of CRISPR-Cas9 system (AcrllC1 and AcrllC3) [238]. Therefore, in order to reduce off-target effects, the anti-CRISPRs could be used to prevent the continuous expression of Cas9 protein in cells to be edited.

(5)

Others

The concentration of Cas9/sgRNA can also affect the frequency of off-target mutations [239]. Thus, the optimal concentration of Cas9 and sgRNA needs to be determined by pre-experiment. Besides, the formulation of CRISPR-Cas9 can affect the frequency of off-target mutations as well. Cas9 nucleases can be delivered into target cells in 3 different forms: DNA expression plasmid, mRNA or recombination protein [240]. Currently, the use of Cas9/sgRNA ribonucleoprotein complexes (Cas9-RNPs), which are composed of purified Cas9 proteins in combination with sgRNA, is becoming more and more widespread. It was found that delivery as plasmid usually produces more off-targets than delivery as RNPs, since the CRISPR-Cas system is active for a shorter time without Cas9 transcription and translation stages [241], [242].

Nowadays, how to effectively deliver CRISPR-Cas components to specific cells, tissues and organs for precisely directed genome editing is still a major problem in gene therapy. Ideal delivery vectors should have the advantages of non-toxicity, well targeting property, high efficiency, low cost, and biodegradability [35], [156]. At present, three main delivery methods have been employed in delivering CRISPR-Cas components, including physical, viral and non-viral methods [243]. Physical methods are the simplest way to deliver CRISPR-Cas components, including electroporation, microinjection and mechanical cell deformation. They are simple and efficient, which can also improve the expression of genes, and being widely applied in in vitro experiments [243], [244]. In addition, viral vectors, such as adenovirus, adeno-associated virus (AAV) and lentivirus viral vectors, are being widely used for both in vitro/ex vivo and in vivo delivery due to their high delivery efficiency. They are commonly used for gene delivery in gene therapy, and some of them have been approved for clinical use [245], [246]. However, safety issue of viral vectors is still a major problem needed to be solved in pre-clinical trials. Therefore, researchers have turned their attention to non-viral vectors, for instance, liposomes, polymers and nanoparticles [247]. Based on the advantages of safety, availability and cost-effectiveness, they are becoming a hotspot for the delivery of CRISPR-Cas components [248].

Since all these delivery methods have both advantages and disadvantages, its necessary to design a complex of viral vectors and non-viral vectors, which combines the advantages of both vectors. Along with the deepening of research, various carriers could be modified by different methods to increase the delivery efficiency and reduce the toxicity [249]. In addition, more novel vectors, such as graphene and carbon nanomaterials (CNMs), could also be applied in the delivery of CRISPR-Cas components [250], [251].

Since the components of CRISPR-Cas systems are derived from bacteria, host immune response to Cas gene and Cas protein is regarded as one of the most important challenges in the clinical trials of CRISPR-Cas system [156], [252]. It was found that in vivo delivery of CRISPR-Cas components can elicit immune responses against the Cas protein [252], [253]. Furthermore, researchers also found that there were anti-Cas9 antibodies and anti-Cas9 T cells existing in healthy humans, suggesting the pre-existing of humoral and celluar immune responses to Cas9 protein in humans [254]. Therefore, how to detect and reduce the immunogenicity of Cas proteins is a major challenge will be faced in clinical application of CRISPR-Cas systems. Researchers are trying to handle this problem by modifying Cas9 protein or using Cas9 homologues [255].

Recently, two independent research groups found that CRISPR-Cas mediated double-stranded breaks (DSBs) can activate the p53 signaling pathway [256], [257]. This means that genetically edited cells are likely to become potential cancer initiating cells, and clinical treatment with CRISPR-Cas systems might inadvertently increase the risk of cancer [256], [257], [258]. Although there is still no direct evidence to confirm the relationship between CRISPR-Cas mediated genome editing and carcinogenesis, these studies once again give a warning on the application of CRISPR-Cas systems in gene therapy. It reminds us that there is still a long way to go before CRISPR-Cas systems could be successfully applied to humans.

CRISPR-Cas mediated genome editing has attracted much attention since its advent in 2012. In theory, each gene can be edited by CRISPR-Cas systems, even genes in human germ cells [259]. However, germline gene editing is forbidden in many countries including China, for it could have unintended consequences and bring ethical and safety concerns [260].

However, in March 2015, a Chinese scientist, Junjiu Huang, published a paper about gene editing in human tripronuclear zygotes in the journal Protein & Cell, which brings the ethical controversy of human embryo gene editing to a climax [261]. Since then, genome editing has been challenged by ethics and morality, and legal regulation of genome editing has triggered a heated discussion all around the world.

Then, on Nov. 28, 2018, the day before the opening of the second international human genome editing summit, Jiankui He, a Chinese scientist from the Southern University of Science and Technology, announced that a pair of gene-edited babies, named Lulu and Nana, were born healthy in China this month. They are the worlds first gene-edited babies, whose CCR5 gene has been modified, making them naturally resistant to HIV infection after birth [262]. The announcement has provoked shock, even outrage among scientists around the world, causing widespread controversy in the application of genome editing.

The society was shocked by this breaking news, for it involves genome editing in human embryos and propagating into future generations, triggering a chorus of criticism from the scientific community and bringing concerns about ethics and security in the use of genome editing. Therefore, scientists call on Chinese government to investigate the matter fully and establish strict regulations on human genome editing. Global supervisory system is also needed to ensure genome editing of human embryos moving ahead safely and ethically [263].

Since CRISPR-Cas mediated genome editing technologies have provided an accessible and adaptable means to alter, regulate, and visualize genomes, they are thought to be a major milestone for molecular biology in the 21st century. So far, CRISPR-Cas systems have been broadly applied in gene function analysis, human gene therapy, targeted drug development, animal model construction and livestock breeding, which fully prove their great potential for further development. However, there are still some limitations to overcome in the practical applications of CRISPR-Cas systems, and great efforts still need to be made to evaluate their long-term safety and effectiveness.

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In 2013, two biochemists published a paper proclaiming theyd discovered a potentially game-changing method of manipulating genes. CRISPR which sounds like a veggie-forward gastro pub won them each a Nobel Prize.

In the years since, CRISPR (or Clustered Regularly Interspaced Short Palindromic Repeats) has lived up to the hype. Its altered the global scientific landscape and raised questions about what kinds of revolutionary changes scientists and healthcare providers could and should pursue.

What if we could make foods allergy-free and crops drought-resistant? What if we could eliminate invasive species and protect against infectious diseases like malaria? What if we could revive extinct species? What if we could remove or repair mutations that cause inherited conditions? Or create custom immunotherapies to treat an individuals cancer?

The prospects are that exciting.

If your understanding of genetics starts and ends with high school biology or the (very fictional) Jurassic Park movies youre not alone. This stuff is complicated. Thats why we asked genomics and immunotherapy expert Timothy Chan, MD, PhD, to break CRISPR down for us, so we can better understand why, over a decade later, its still got researchers so excited.

Before we jump into CRISPR, lets start with the concept of gene editing.

Gene editing is the process of altering genetic material (DNA). That could mean changing a few individual genes or an entire sequence. Research has been ongoing for more than a decade thats looking at using gene editing on mutations that cause serious health conditions in people. The goal of this gene editing research is to eliminate or correct the mutation thats causing the health condition, or has the potential to cause one, such as certain cancers. In other research studies, gene editing is being explored so a mutation isnt passed down to children at birth.

For example, the U.S. Food and Drug Administration (FDA) approved a gene therapy in late 2022 that introduces a gene needed for blood clotting into people with hemophilia B. Its one of several cellular and gene therapy products currently in use today.

There are many different techniques and applications for gene editing. CRISPR is one approach to gene editing thats showing promise in ongoing clinical trials.

Now that were clear on what gene editing is, lets focus on a specific approach: CRISPR.

Clustered Regularly Interspaced Short Palindromic Repeats, otherwise known as CRISPR, was originally identified in bacteria, as a bacterial defense system, says Dr. Chan.

Thats right. Bacteria have immune systems, too.

CRISPR contains spacers sequences of DNA left over from unfriendly viruses or other entities as well as repeating sections of genetic material. Those sequences provide acquired immunity, and form the building blocks of the gene editing system or process. It creates a sort of blueprint that allows enzymes in genetic material to make changes to sequences of DNA in living cells. One of the best-known enzymes used for this purpose is called Cas9, which is why youll sometimes hear people talk about CRISPR-Cas9.

Over the years, people have discovered that specific enzymes that allow CRISPR to work Cas9 is one of them.But there are other ones, and they can be tailored to target sequences of interest in the DNA for specific cuts to be made, Dr. Chan explains.

You can think of the underlying mechanism of CRISPR gene editing as being similar to the way magnetic shapes are drawn to each other or the way Lego blocks fit together.

The segments in CRISPR are transcribed into RNA. This RNA includes a guide sequence, which is a match to existing DNA in a persons body.

That guide sequence can be tailored to whatever you want, Dr. Chan says. And as a result, you can make specific alterations or mutations in a part of the genome that you are targeting with a high degree of accuracy.

Along for the ride with this guide sequence is an enzyme like Cas9.

When the guide sequence and enzyme find the desired DNA to edit, the enzyme can then get down to business. It attaches itself to this DNA and makes changes, whether thats a cut or alteration.

CRISPR technology has come a long way, Dr. Chan says. The first generation of CRISPR was a great way to inactivate genes. It only made a break in genes. Then, the DNA would get filled up with natural repair enzymes.

But new versions of CRISPR like CRISPR prime or CRISPR HD are more advanced.

These can allow actual replacements to occur, Dr. Chan continues. You can even very accurately replace one sequence one of the letters in the genome with another letter. And you can make specific mutations.

CRISPRs ability to make very specific, very small cuts has the potential to transform how healthcare providers can address certain genetic diseases.

Dr. Chan is optimistic about the future of CRISPR based on the success of ongoing clinical trials in human subjects. For any type of genetic diseases caused by a single mutated gene, you can use CRISPR to mutate it and make it normal. Thats why its useful. Its a way for us to change errors in the genome.

Right now, CRISPR is geared toward correcting a single change in genes, he adds. While combinations may be possible in the future, were just not there yet.

While gene editing is already in use, CRISPR is still in the clinical trials phase, Dr. Chan says. Its used all the time in research laboratories and industries, he notes. Many clinical trials are testing CRISPR in the setting of genetic diseases and cancer.

Interestingly, CRISPR can be used to detect certain diseases. The best-known example is the Sherlock CRISPR SARS-CoV-2 Kit: A COVID-19 test that received emergency use authorization (EAU) from the FDA in 2020.

But theres no FDA-approved CRISPR therapy right now. The clinical trials are ongoing, he says.

These include trials looking at CRISPR to correct genetic diseases such as cystic fibrosis, Huntingtons disease and muscular dystrophy.

Dr. Chan adds that there are also major clinical trials in process for blood disorders, where CRISPR is being used to correct the gene alteration that causes the condition. As one example, he cites a promising trial looking at CRISPR-Cas9 gene editing for sickle cell disease and -thalassemia, written about in an early 2021 issue of the New England Journal of Medicine. -thalassemia is an inherited blood disorder that impacts the bodys ability to create hemoglobin an iron-dense protein that serves as the primary ingredient in red blood cells.

There are also clinical trials looking to see if CRISPR can be used to treat certain cancers. Dr. Chan notes that chimeric antigen receptor (CAR)T-cell therapy is one of the first gene therapies approved for leukemias. Current research is looking at whether CRISPR technology can make this treatment even more effective.

In CAR T-cell therapy, you take out T-cells from someone and put in a receptor a new way for these cells to target something on cancer cells and then put these cells back in the patient, he explains. Researchers are running trials now where they use CRISPR to alter those T-cells to make them even more active.

CRISPR therapies can take on many different forms. CRISPR has been inserted directly into the body before. It was famously injected into the eyes of seven people with a rare hereditary blindness disorder in 2020, two of whom later told NPR that they regained some ability to see colors. There are human trials in process right now that deliver CRISPR through gels and creams, through food or drink, skin grafts or injections. Ex-vivo delivery is also common: Thats when CRISPR is used to modify a cell outside the body. The cells are then re-inserted into the body using a harmless virus.

The results have been promising so far. I do believe in the next three to five years possibly even sooner were going to see approval to treat some diseases, Dr. Chan states.

With any type of CRISPR therapy, Dr. Chan says theres a risk of getting off-target effects or unexpected side effects.

Whenever youre altering something as fundamental as DNA, you just dont know what might happen he explains. Theres always a chance for the unexpected. You can potentially have effects on your DNA that were not intended.

At the moment, he doesnt have any specific examples of what these effects might be and he notes that existing research suggests the risk is pretty low. Still, data from future research might tell a different story.

Dr. Chan nevertheless sees a lot of potential for CRISPR in the coming years.

The field is moving very quickly, he says. Were seeing continual improvement of the actual CRISPR tools being used.

Its getting more accurate and more flexible in terms of what you can do. There are various engineered modified variants of CRISPR now that are allowing very specific, very accurate changes with fewer off-target effects. So, I think the future is very bright.

View original post here:
What Is CRISPR Gene Editing and How Does It Work?

In 2013, two biochemists published a paper proclaiming theyd discovered a potentially game-changing method of manipulating genes. CRISPR which sounds like a veggie-forward gastro pub won them each a Nobel Prize.

Cleveland Clinic is a non-profit academic medical center. Advertising on our site helps support our mission. We do not endorse non-Cleveland Clinic products or services. Policy

In the years since, CRISPR (or Clustered Regularly Interspaced Short Palindromic Repeats) has lived up to the hype. Its altered the global scientific landscape and raised questions about what kinds of revolutionary changes scientists and healthcare providers could and should pursue.

What if we could make foods allergy-free and crops drought-resistant? What if we could eliminate invasive species and protect against infectious diseases like malaria? What if we could revive extinct species? What if we could remove or repair mutations that cause inherited conditions? Or create custom immunotherapies to treat an individuals cancer?

The prospects are that exciting.

If your understanding of genetics starts and ends with high school biology or the (very fictional) Jurassic Park movies youre not alone. This stuff is complicated. Thats why we asked genomics and immunotherapy expert Timothy Chan, MD, PhD, to break CRISPR down for us, so we can better understand why, over a decade later, its still got researchers so excited.

Before we jump into CRISPR, lets start with the concept of gene editing.

Gene editing is the process of altering genetic material (DNA). That could mean changing a few individual genes or an entire sequence. Research has been ongoing for more than a decade thats looking at using gene editing on mutations that cause serious health conditions in people. The goal of this gene editing research is to eliminate or correct the mutation thats causing the health condition, or has the potential to cause one, such as certain cancers. In other research studies, gene editing is being explored so a mutation isnt passed down to children at birth.

For example, the U.S. Food and Drug Administration (FDA) approved a gene therapy in late 2022 that introduces a gene needed for blood clotting into people with hemophilia B. Its one of several cellular and gene therapy products currently in use today.

There are many different techniques and applications for gene editing. CRISPR is one approach to gene editing thats showing promise in ongoing clinical trials.

Now that were clear on what gene editing is, lets focus on a specific approach: CRISPR.

Clustered Regularly Interspaced Short Palindromic Repeats, otherwise known as CRISPR, was originally identified in bacteria, as a bacterial defense system, says Dr. Chan.

Thats right. Bacteria have immune systems, too.

CRISPR contains spacers sequences of DNA left over from unfriendly viruses or other entities as well as repeating sections of genetic material. Those sequences provide acquired immunity, and form the building blocks of the gene editing system or process. It creates a sort of blueprint that allows enzymes in genetic material to make changes to sequences of DNA in living cells. One of the best-known enzymes used for this purpose is called Cas9, which is why youll sometimes hear people talk about CRISPR-Cas9.

Over the years, people have discovered that specific enzymes that allow CRISPR to work Cas9 is one of them.But there are other ones, and they can be tailored to target sequences of interest in the DNA for specific cuts to be made, Dr. Chan explains.

You can think of the underlying mechanism of CRISPR gene editing as being similar to the way magnetic shapes are drawn to each other or the way Lego blocks fit together.

The segments in CRISPR are transcribed into RNA. This RNA includes a guide sequence, which is a match to existing DNA in a persons body.

That guide sequence can be tailored to whatever you want, Dr. Chan says. And as a result, you can make specific alterations or mutations in a part of the genome that you are targeting with a high degree of accuracy.

Along for the ride with this guide sequence is an enzyme like Cas9.

When the guide sequence and enzyme find the desired DNA to edit, the enzyme can then get down to business. It attaches itself to this DNA and makes changes, whether thats a cut or alteration.

CRISPR technology has come a long way, Dr. Chan says. The first generation of CRISPR was a great way to inactivate genes. It only made a break in genes. Then, the DNA would get filled up with natural repair enzymes.

But new versions of CRISPR like CRISPR prime or CRISPR HD are more advanced.

These can allow actual replacements to occur, Dr. Chan continues. You can even very accurately replace one sequence one of the letters in the genome with another letter. And you can make specific mutations.

CRISPRs ability to make very specific, very small cuts has the potential to transform how healthcare providers can address certain genetic diseases.

Dr. Chan is optimistic about the future of CRISPR based on the success of ongoing clinical trials in human subjects. For any type of genetic diseases caused by a single mutated gene, you can use CRISPR to mutate it and make it normal. Thats why its useful. Its a way for us to change errors in the genome.

Right now, CRISPR is geared toward correcting a single change in genes, he adds. While combinations may be possible in the future, were just not there yet.

While gene editing is already in use, CRISPR is still in the clinical trials phase, Dr. Chan says. Its used all the time in research laboratories and industries, he notes. Many clinical trials are testing CRISPR in the setting of genetic diseases and cancer.

Interestingly, CRISPR can be used to detect certain diseases. The best-known example is the Sherlock CRISPR SARS-CoV-2 Kit: A COVID-19 test that received emergency use authorization (EAU) from the FDA in 2020.

But theres no FDA-approved CRISPR therapy right now. The clinical trials are ongoing, he says.

These include trials looking at CRISPR to correct genetic diseases such as cystic fibrosis, Huntingtons disease and muscular dystrophy.

Dr. Chan adds that there are also major clinical trials in process for blood disorders, where CRISPR is being used to correct the gene alteration that causes the condition. As one example, he cites a promising trial looking at CRISPR-Cas9 gene editing for sickle cell disease and -thalassemia, written about in an early 2021 issue of the New England Journal of Medicine. -thalassemia is an inherited blood disorder that impacts the bodys ability to create hemoglobin an iron-dense protein that serves as the primary ingredient in red blood cells.

There are also clinical trials looking to see if CRISPR can be used to treat certain cancers. Dr. Chan notes that chimeric antigen receptor (CAR)T-cell therapy is one of the first gene therapies approved for leukemias. Current research is looking at whether CRISPR technology can make this treatment even more effective.

In CAR T-cell therapy, you take out T-cells from someone and put in a receptor a new way for these cells to target something on cancer cells and then put these cells back in the patient, he explains. Researchers are running trials now where they use CRISPR to alter those T-cells to make them even more active.

CRISPR therapies can take on many different forms. CRISPR has been inserted directly into the body before. It was famously injected into the eyes of seven people with a rare hereditary blindness disorder in 2020, two of whom later told NPR that they regained some ability to see colors. There are human trials in process right now that deliver CRISPR through gels and creams, through food or drink, skin grafts or injections. Ex-vivo delivery is also common: Thats when CRISPR is used to modify a cell outside the body. The cells are then re-inserted into the body using a harmless virus.

The results have been promising so far. I do believe in the next three to five years possibly even sooner were going to see approval to treat some diseases, Dr. Chan states.

With any type of CRISPR therapy, Dr. Chan says theres a risk of getting off-target effects or unexpected side effects.

Whenever youre altering something as fundamental as DNA, you just dont know what might happen he explains. Theres always a chance for the unexpected. You can potentially have effects on your DNA that were not intended.

At the moment, he doesnt have any specific examples of what these effects might be and he notes that existing research suggests the risk is pretty low. Still, data from future research might tell a different story.

Dr. Chan nevertheless sees a lot of potential for CRISPR in the coming years.

The field is moving very quickly, he says. Were seeing continual improvement of the actual CRISPR tools being used.

Its getting more accurate and more flexible in terms of what you can do. There are various engineered modified variants of CRISPR now that are allowing very specific, very accurate changes with fewer off-target effects. So, I think the future is very bright.

Continue reading here:
The Basics of CRISPR Gene Editing - Cleveland Clinic Health Essentials

Abstract

Clustered regularly interspaced short palindromic repeat (CRISPR) and their associated protein (Cas-9) is the most effective, efficient, and accurate method of genome editing tool in all living cells and utilized in many applied disciplines. Guide RNA (gRNA) and CRISPR-associated (Cas-9) proteins are the two essential components in CRISPR/Cas-9 system. The mechanism of CRISPR/Cas-9 genome editing contains three steps, recognition, cleavage, and repair. The designed sgRNA recognizes the target sequence in the gene of interest through a complementary base pair. While the Cas-9 nuclease makes double-stranded breaks at a site 3 base pair upstream to protospacer adjacent motif, then the double-stranded break is repaired by either non-homologous end joining or homology-directed repair cellular mechanisms. The CRISPR/Cas-9 genome-editing tool has a wide number of applications in many areas including medicine, agriculture, and biotechnology. In agriculture, it could help in the design of new grains to improve their nutritional value. In medicine, it is being investigated for cancers, HIV, and gene therapy such as sickle cell disease, cystic fibrosis, and Duchenne muscular dystrophy. The technology is also being utilized in the regulation of specific genes through the advanced modification of Cas-9 protein. However, immunogenicity, effective delivery systems, off-target effect, and ethical issues have been the major barriers to extend the technology in clinical applications. Although CRISPR/Cas-9 becomes a new era in molecular biology and has countless roles ranging from basic molecular researches to clinical applications, there are still challenges to rub in the practical applications and various improvements are needed to overcome obstacles.

Keywords: CRISPR, Cas-9, sgRNA, gene-editing, mechanism, applications

Genome editing is a type of genetic engineering in which DNA is deliberately inserted, removed, or modified in living cells.1 The name CRISPR (Clustered Regularly Interspaced Short Palindromic Repeat) refers to the unique organization of short, partially repeated DNA sequences found in the genomes of prokaryotes. CRISPR and its associated protein (Cas-9) is a method of adaptive immunity in prokaryotes to defend themselves against viruses or bacteriophages.2 Japanese scientist Ishino and his team accidentally found unusual repetitive palindromic DNA sequences interrupted by spacers in Escherichia coli while analyzing a gene for alkaline phosphatase first discovered CRISPR in 1987. However, they did not ascertain its biological function. In 1990, Francisco Mojica identifies similar sequences in other prokaryotes and he named CRISPR, yet the functions of these sequences were a mystery.3 Later on in 2007, a CRISPR was experimentally conferred as a key element in the adaptive immune system of prokaryotes against viruses. During the adaptation process, bacterial cells become immunized by the insertion of short fragments of viral DNA (spacers) into a genomic region called the CRISPR array. Hence, spacers serve as a genetic memory of previous viral infections.4 The CRISPR defense mechanism protects bacteria from repeated viral attacks via three basic stages: adaptation (spacer acquisition), crRNA synthesis (expression), and target interference. CRISPR loci are an array of short repeated sequences found in chromosomal or plasmid DNA of prokaryotes. Cas gene is usually found adjacent to CRISPR that codes for nuclease protein (Cas protein) responsible to destroy or cleave viral nucleic acid.5

Before the discovery of CRISPR/Cas-9, scientists were relied on two gene-editing techniques using restriction enzymes, zinc finger nucleases (ZFN) and Transcription activator-like effector nucleases (TALENs).6 ZFN has a zinc finger DNA binding domain used to bind a specific target DNA sequence and a restriction endonuclease domain used to cleave the DNA at the target site. TALENs are also composed of DNA binding domain and restriction domain like ZFN but their DNA binding domain has more potential target sequence than the ZFN gene-editing tool. In both cases, the difficulty of protein engineering, being expensive, and time-consuming were the major challenges for researchers and manufacturers.6,7 The development of a reliable and efficient method of a gene-editing tool in living cells has been a long-standing goal for biomedical researchers. After figuring out the CRISPR mechanism in prokaryotes, scientists understood that it could have beneficial use in humans, plants, and other microbes. It was in 2012 that Doudna, J, and Charpentier, E discovered CRISPR/Cas-9 could be used to edit any desired DNA by just providing the right template.8 Since then, CRISPR/Cas-9 becomes the most effective, efficient, and accurate method of genome editing tool in all living cells and utilized in many applied disciplines.9 Thus, this review aims to discuss the mechanisms of genome editing mediated by CRISPR/Cas-9 and to highlight its recent applications as one of the most important scientific discoveries of this century, as well as the current barriers to the transformation of this technology.

Based on the structure and functions of Cas-proteins, CRISPR/Cas system can be divided into Class I (type I, III, and IV) and Class II (type II, V, and VI). The class I systems consist of multi-subunit Cas-protein complexes, while the class II systems utilize a single Cas-protein. Since the structure of type II CRISPR/Cas-9 is relatively simple, it has been well studied and extensively used in genetic engineering.10 Guide RNA (gRNA) and CRISPR-associated (Cas-9) proteins are the two essential components in CRISPR/Cas-9 system. The Cas-9 protein, the first Cas protein used in genome editing was extracted from Streptococcus pyogenes (SpCas-9). It is a large (1368 amino acids) multi-domain DNA endonuclease responsible for cleaving the target DNA to form a double-stranded break and is called a genetic scissor.11 Cas-9 consists of two regions, called the recognition (REC) lobe and the nuclease (NUC) lobe. The REC lobe consists of REC1 and REC2 domains responsible for binding guide RNA, whereas the NUC lobe is composed of RuvC, HNH, and Protospacer Adjacent Motif (PAM) interacting domains. The RuvC and HNH domains are used to cut each single-stranded DNA, while PAM interacting domain confers PAM specificity and is responsible for initiating binding to target DNA.12 Guide RNA is made up of two parts, CRISPR RNA (crRNA) and trans-activating CRISPR RNA (tracrRNA). The crRNA is an 1820 base pair in length that specifies the target DNA by pairing with the target sequence, whereas tracrRNA is a long stretch of loops that serve as a binding scaffold for Cas-9 nuclease. In prokaryotes, the guide RNA is used to target viral DNA, but in the gene-editing tool, it can be synthetically designed by combining crRNA and tracrRNA to form a single guide RNA (sgRNA) in order to target almost any gene sequence supposed to be edited.11

The mechanism of CRISPR/Cas-9 genome editing can be generally divided into three steps: recognition, cleavage, and repair.13 The designed sgRNA directs Cas-9 and recognizes the target sequence in the gene of interest through its 5crRNA complementary base pair component. The Cas-9 protein remains inactive in the absence of sgRNA. The Cas-9 nuclease makes double-stranded breaks (DSBs) at a site 3 base pair upstream to PAM.14 PAM sequence is a short (25 base-pair length) conserved DNA sequence downstream to the cut site and its size varies depending on the bacterial species. The most commonly used nuclease in the genome-editing tool, Cas-9 protein recognizes the PAM sequence at 5-NGG-3 (N can be any nucleotide base). Once Cas-9 has found a target site with the appropriate PAM, it triggers local DNA melting followed by the formation of RNA-DNA hybrid, but the mechanism of how Cas-9 enzyme melts target DNA sequence was not clearly understood yet. Then, the Cas-9 protein is activated for DNA cleavage. HNH domain cleaves the complementary strand, while the RuvC domain cleaves the non-complementary strand of target DNA to produce predominantly blunt-ended DSBs. Finally, the DSB is repaired by the host cellular machinery.11,15

Non-homologous end joining (NHEJ), and homology-directed repair (HDR) pathways are the two mechanisms to repair DSBs created by Cas-9 protein in CRISPR/Cas-9 mechanism.16 NHEJ facilitates the repair of DSBs by joining DNA fragments through an enzymatic process in the absence of exogenous homologous DNA and is active in all phases of the cell cycle. It is the predominant and efficient cellular repair mechanism that is most active in the cells, but it is an error-prone mechanism that may result in small random insertion or deletion (indels) at the cleavage site leading to the generation of frameshift mutation or premature stop codon.17 HDR is highly precise and requires the use of a homologous DNA template. It is most active in the late S and G2 phases of the cell cycle. In CRISPR-gene editing, HDR requires a large amount of donor (exogenous) DNA templates containing a sequence of interest. HDR executes the precise gene insertion or replacement by adding a donor DNA template with sequence homology at the predicted DSB site.16,17

In just a few years of its discovery, the CRISPR/Cas-9 genome editing tool has already being explored for a wide number of applications and had a massive impact on the world in many areas including medicine, agriculture, and biotechnology. In the future, researchers hope that this technology will continue to advance for treating and curing diseases, develop more nutritious crops, and eradicating infectious diseases.18 Highlights for some of the recent CRISPR/Cas-9 applications and clinical trials being investigated are discussed below.

More than 6000 genetic disorders have been known so far. But the majority of the diseases lack effective treatment strategies.19 Gene therapy is the process of replacing the defective gene with exogenous DNA and editing the mutated gene at its native location. It is the latest development in the revolution of medical biotechnology. From 1998 to August 2019, 22 gene therapies including the novel CRISPR/Cas-9 have been approved for the treatment of human diseases.20

Since its discovery in 2012, CRISPR/Cas-9 gene editing has held the promise of curing most of the known genetic diseases such as sickle cell disease, -thalassemia, cystic fibrosis, and muscular dystrophy.21,22 CRISPR/Cas-9 for targeted sickle cell disease (SCD) therapy and -thalassemia have been also applied in clinical trials.23 SCD is an autosomal recessive genetic disease of red blood cells, which occurs due to point mutation in the -globin chain of hemoglobin leading to sickle hemoglobin (HbS). During the deoxygenation process, HbS polymerization leads to severe clinical complications like hemolytic anemia.24 Either direct repairing the gene of hemoglobin S or boosting fetal -globin are the two main approaches that CRISPR/Cas-9 is being used to treat SCD.25 However, the most common method used in a clinical trial is based on the approach of boosting fetal hemoglobin. First bone marrow cells are removed from patients and the gene that turns off fetal hemoglobin production, called B-cell Lymphoma 11A (BCL11A) is disabled with CRISPR/Cas-9. Then, the gene-edited cells are infused back into the body.26 BCL11A is a 200 base pair gene found on chromosome 2 and its product is responsible to switch -globin into the -globin chain by repressing -globin gene expression.27 Once this gene is disabled using CRISPR/Cas-9, the production of fetal hemoglobin containing -globin in the red blood cells will increase, thereby alleviating the severity and manifestations of SCD.28

Scientists have been also investigating CRISPR/Cas-9 for the treatment of cystic fibrosis. The genetic mutation of the cystic fibrosis transmembrane conductance regulator (CFTR) gene decreases the structural stability and function of CFTR protein leading to cystic fibrosis.29 CFTR protein is an anion channel protein regulated by protein kinase-A, located at the apical surface of epithelial cells of the lung, intestine, pancreas, and reproductive tract.30 Although there is no cure for cystic fibrosis, symptom-based therapies (such as antibiotics, bronchodilators, and mucus thinning medications) and CFTR modulating drugs have become the first-line treatments to relieve symptoms and reduce the risk of complications.31 Currently, gene manipulation technologies and molecular targets are also being explored. The use of CRISPR/Cas-9 technology for genome editing has great potential, although it is in the early stages of development.32 In 2013, researchers culture intestinal stem cells from two cystic fibrosis patients and corrected the mutation at the CFTR locus resulting in the expression of the correct gene and full function of the protein. Since then, the potential utility of the application of CRISPR/Cas-9 for cystic fibrosis was established.33 Furthermore, Duchenne muscular dystrophy (DMD), which is caused by a mutation in the dystrophin gene and characterized by muscle weakness, has been successfully corrected by CRISPR/Cas-9 in patient-induced pluripotent stem cells.34 Despite considerable efforts, the treatment available for DMD remains supportive rather than curative. Currently, several therapeutic approaches (gene therapy, cell therapy, and exon skipping) have been investigated to restore the expression of dystrophin in DMD muscles.35,36 Deletion/excision of intragenic DNA and removing the duplicated exon by CRISPR/Cas-9 are the new and promising approaches in correcting the DMD gene, which restores the expression of dystrophin protein.37

Moreover, the latest researches show that the CRISPR/Cas-mediated single-base editing and prime editing systems can directly install mutations in cellular DNA without the need for a donor template. The CRISPR/Cas-base editor and prime editor system do not produce DSB, which reduces the possibility of indels that are different from conventional Cas-9.38 So far, two types of base editors have been developed: cytosine base editor (CBE) and adenine base editor (ABE).39 The CBE is a type of base editor composed of cytidine deaminase fused with catalytically deficient or dead Cas-9 (dCas-9). It is one of the novel gene therapy strategies that can produce precise base changes from cytidine (C) to thymidine (T).40 However, the target range of the CBE base editor is still restricted by PAM sequences containing G, T, or A bases. Recently, a more advanced fidelity and efficiency base editor called nNme2-CBE (discovered from Neisseria meningitides) with expanded PAM compatibility for cytidine dinucleotide has been developed in both human cells and rabbits embryos.41 The ABE uses adenosine deaminase fused to dCas-9 to correct the base-pair change from adenosine (A) to guanosine (G).38 Overall, single-base editing through the fusion of dCas-9 to cytidine deaminase or adenosine deaminase is a safe and efficient method to edit point mutations. But both base editors can only fix four-transition mutations (purine to purine or pyrimidine to pyrimidine).42 To overcome this shortcoming, the most recent member of the CRISPR genome editing toolkit called Prime Editor (PE) has been developed to extend the scope of DNA editing beyond the four types of transition mutations.43 PE contains Cas-9 nickase fused with engineered reverse transcriptase and multifunctional primer editing guide RNA (pegRNA). The pegRNA recognizes the target nucleotide sequence; the Cas-9 nickase cuts the non-complementary strand of DNA three bases upstream from the PAM site, exposing a 3-OH nick of genomic DNA. The reverse transcriptase then extends the 3 nick by copying the edit sequence of pegRNA. Hence, PE not only corrects all 12 possible base-to-base transitions, and transversion mutations but also small insertion and deletion mutations in genetic disorders.44

The first CRISPR-based therapy in the human trial was conducted to treat patients with refractory lung cancer. Researchers first extract T-cells from three patients blood and they engineered them in the lab through CRISPR/Cas-9 to delete genes (TRAC, TRBC, and PD-1) that would interfere to fight cancer cells. Then, they infused the modified T-cells back into the patients. The modified T-cells can target specific antigens and kill cancer cells. Finally, no side effects were observed and engineered T-cells can be detected up to 9 months of post-infusion.45 CRISPR/Cas-9 gene-editing technology could also be used to treat infectious diseases caused by microorganisms.46 One focus area for the researchers is treating HIV, the virus that leads to AIDS. In May 2017, a team of researchers from Temple University demonstrated that HIV-1 replication can be completely shut down and the virus eliminated from infected cells through excision of HIV-1 genome using CRISPR/Cas-9 in animal models.47 In addition to the approach of targeting the HIV-genome, CRISPR/Cas-9 technology can also be used to block HIV entry into host cells by editing chemokine co-receptor type-5 (CCR5) genes in the host cells. For instance, an in vitro trial conducted in China reported that genome editing of CCR5 by CRISPR/Cas-9 showed no evidence of toxicity (infection) on cells and they concluded that edited cells could effectively be protected from HIV infection than unmodified cells.48

As the world population continues to grow, the risk of shortage in agricultural resources is real. Hence, there is a need for new technologies for increasing and improving natural food production. CRISPR/Cas-9 is an existing addition to the field since it has been used to genetically modify foods to improve their nutritional value, increase their shelf life, make them drought-tolerant, and enhance disease resistance.18 There are generally three ways that CRISPR is solving the worlds food crisis. It can restore food supplies, help plants to survive in hostile conditions, and could improve the overall health of the plants.49

Beyond genome editing activity, CRISPR/Cas-9 can be used to artificially regulate (activate or repress) a certain target of a gene through advanced modification of Cas-9 protein.15 Researchers had performed an advanced modified Cas-9 endonuclease called dCas-9 nuclease by inactivating its HNH and RuvC domains. The dCas-9 nuclease lacks DNA cleavage activity, but its DNA binding activity is not affected. Then, transcriptional activators or inhibitors can be fused with dCas-9 to form the CRISPR/dCas-9 complex. Therefore, catalytically inactive dCas-9 can be used to activate (CRISPRa) or silence (CRISPRi) the expression of a specific gene of interest.50 Moreover, the CRISPR/dCas-9 can be also used to visualize and pinpoint where specifically the gene of interest is located inside the cell (subcellular localization) by fusing a marker such as Green Fluorescent Proteins (GFP) with dCas-9 enzyme. This enables site-specific labeling and imaging of endogenous loci in living cells for further utilization.51

Despite its great promise as a genome-editing system CRISPR/Cas-9 technology had hampered by several challenges that should be addressed during the process of application. Immunogenicity, lack of a safe and efficient delivery system to the target, off-target effect, and ethical issues have been the major barriers to extend the technology in clinical applications.52 Since the components of the CRISPR/Cas-9 system are derived from bacteria, host immunity can elicit an immune response against these components. Researchers also found that there were both pre-existing humoral (anti-Cas-9 antibody) and cellular (anti-Cas-9 T cells) immune responses to Cas-9 protein in healthy humans. Therefore, how to detect and reduce the immunogenicity of Cas-9 protein is still one of the most important challenges in the clinical trial of the system.53

Safe and effective delivery of the components into the cell is essential in CRISPR/Cas-9 gene editing. Currently, there are three methods of delivering the CRISPR/Cas-9 complex into cells, physical, chemical, and viral vectors. Non-viral (physical and chemical) methods are more suitable for ex vivo CRISPR/Cas-9-based gene editing therapy.54 The physical methods of delivering CRISPR/Cas-9 can include electroporation, microinjection, hydrodynamic injection, and so on. Electroporation applies a strong electric field to the cell membrane to temporarily increase the permeability of the membrane, allowing the CRISPR/Cas-9 complex to enter the cytoplasm of the target cell. However, the main limitation of this method is that it causes significant cell death.55 Microinjection involves injecting the CRISPR/Cas-9 complex directly into cells at the microscopic level for rapid gene editing of a single cell. Nevertheless, this method also has several disadvantages such as cell damage, which is technically challenging and is only suitable for a limited number of cells.56 The hydrodynamic injection is the rapid injection of a large amount of high-pressure liquid into the bloodstream of animals, usually using the tail vein of mice. Although this method is simple, fast, efficient, and versatile, it has not yet been used in clinical applications due to possible complications.57 The chemical methods of CRISPR/Cas-9 delivery involves lipid and polymer-based nanoparticles.58 Lipid nanoparticles/liposomes are spherical structures composed of lipid bilayers membrane and are synthesized in aqueous solutions using Lipofectamine-based reagents. The positively charged liposomes encapsulated with negatively charged nucleic acids thereby facilitate the fusion of the complex across the cell membrane into cells.59 Polymeric nanoparticles, such as polyethyleneimine and poly-L-lysine, are the most widely used carriers of CRISPR/Cas-9 components. Like lipid nanoparticles, polymer-based nanoparticles can also transverse the complex in the membrane through endocytosis.60

Viral vectors are the natural experts for in vivo CRISPR/Cas-9 delivery.61 Vectors, such as adenoviral vectors (AVs), adeno-associated viruses (AAVs), and lentivirus vectors (LVs) are currently being widely used as delivery methods due to their higher delivery efficiency relative to physical and chemical methods. Among them, AAVs are the most commonly used vectors due to their low immunogenicity and non-integration into the host cell genome compared to other viral vectors.62 However, the limited virus cloning capacity and the large size of the Cas-9 protein remain a major problem. One strategy to tackle this hurdle is to package sgRNA and Cas-9 into separate AAVs and then co-transfect them into cells. Recent methods employ a smaller strain of Cas-9 from Staphylococcus aureus (SaCas-9) instead of the more commonly used SpCas-9 to allow packaging of sgRNA and Cas-9 in the same AAVs.54,61 Lately, the development of extracellular vesicles (EVs), for the in vivo delivery of CRISPR/Cas-9 to avoid some of the limitations of viral and non-viral methods has shown a great potential.63

The designed sgRNA will mismatch to the non-target DNA and can result in nonspecific, unexpected genetic modification, which is called the off-target effect.57 The CRISPR/Cas-9 target efficiency is determined by the 20-nucleotide sequences of sgRNA and the PAM sequences adjacent to the target genome. It has been shown that more than three mismatches between the target sequence and the 20-nucleotide sgRNA can result in off-target effects.64 The off-target effect can possibly cause harmful events such as sequence mutation, deletion, rearrangement, immune response, and oncogene activation, which limits the application of the CRISPR/Cas-9 editing system for therapeutic purposes.65 To mitigate the possibility of CRISPR/Cas-9 off-target effect, several strategies have been developed, such as optimization of sgRNA, modification of Cas-9 nuclease, utilization of other Cas-variants, and the use of anti-CRISPR proteins.66 Selecting and designing an appropriate sgRNA for the targeted DNA sequence is an important first step to reduce the off-target effect.67 When designing sgRNA, strategies such as GC content, sgRNA length, and chemical modifications of sgRNA must be considered. Generally speaking, studies revealed that GC content of between 40% and 60%, truncated (short length of sgRNA), and incorporation of 2-O-methyl-3-phosphonoacetate in the sgRNA ribose-phosphate backbone are the preferred methods to increase genome editing efficiency of CRISPR/Cas-9.67,68 Modifying the Cas-9 protein to optimize its nuclease specificity is another way to reduce off-target effects. For instance, mutating either one of the catalytic residues of Cas-9 nuclease (HNH and RuvC) will convert the Cas-9 into nickase that could only generate a single-stranded break instead of a blunt cleavage.69 It has been reported that the use of the inactivated RuvC domain of Cas-9 with sgRNA can reduce the off-target effect by 100 to 1500 times.70 Moreover, the nuclease Cas-12a (previously known as Cpf1) is a type V CRISPR/Cas system that provides high genome editing efficiency.71 Unlike the CRISPR/Cas-9 system, CRISPR/Cas-12a can process pre-crRNA into mature crRNA without tracrRNA, thereby reducing the size of plasmid constructs. The Cas-12a protein recognizes a T-rich (5-TTTN) PAM sequence instead of 5-NGG and provides high accuracy at the target gene loci than Cas-9.69 Recently, the use of multicomponent Class I CRISPR proteins, such as CRISPR/Cas-3 and CRISPR/Cas-10 provides better genome editing efficiency than Cas-9.72 The Cas-3 is an ATP-dependent nuclease/helicase that can delete a large part of DNA from the target site without prominent off-target effect. For instance, the DMD gene were repaired by Cas-3-mediated system in induced pluripotent stem cell.73 The Cas-10 protein does not require the PAM sequence and can identify sequences even in the presence of point mutation.72 Anti-CRISPR (Acr) proteins are phage derived small proteins that inhibit the activity of CRISPR/Cas system. They are a recently discovered method to reduce off-target effects of CRISPR/Cas-9.74 From Acr proteins, AcrIIA4 specifically targets Cas-9 nuclease. AcrIIA4 mimics DNA and binds to the Cas-9 site, making impossible to perform further cleavage in area outside the target region.75 Furthermore, CRISPR/Cas-9 gene editing has been challenged by ethics and safety all over the world. Since the technology is still in its infancy and knowledge about the genome is limited, many scientists restrain that it still needs a lot of work to increase its accuracy and make sure that changes made in one part of the genome do not have unforeseen consequences, especially in the application towards human trials.52

AAVs, adeno-associated viral vectors; ABE, adenine base editor; Acr, anti-CRSPR; AVs, adeno-viral vectors; ATP, adenosine tri-phosphate; BCL11A, B-cell lymphoma 11 A; CAS-9, CRISPR-associated protein-9; CBE, cytidine base editor; CCR5, chemokine receptor type 5; CFTR, cystic fibrosis conductance transmembrane receptor; CRISPR, clustered regularly interspaced short palindromic repeat; CrRNA, CRISPR ribonucleic acid; DMD, Duchenne muscular dystrophy; DNA, deoxyribonucleic acid; DSBs, double-stranded breaks; HDR, homology-directed repair; LVs, lentivirus vectors; NHEJ, non-homologous end Jjining; PAM, protospacer adjacent motif; PD-1, programmed cell death-1; RNA, ribonucleic acid; TALENs, transcriptionactivator like effector nucleases; TRAC, T-cell receptor alpha; TRBC, T-cell receptor beta; TracrRNA, trans-activating CRISPR ribonucleic acid; ZFNs, zinc finger nucleases.

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Mechanism and Applications of CRISPR/Cas-9-Mediated Genome Editing

Youve probably read stories about new research using the gene editing technique CRISPR, also called CRISPR/Cas9. The scientific world is captivated by this revolutionary technology, since it is easier, cheaper and more efficient than previous strategies for modifying DNA.

The term CRISPR/Cas9 stands for Clustered Regularly Interspaced Short Palindromic Repeats/CRISPR associated protein 9. The names reflect important features identified during its discovery, but dont tell us much about how it works, as they were coined before anyone understood what it was.

CRISPR/Cas9 is a system found in bacteria and involved in immune defence. Bacteria use CRISPR/Cas9 to cut up the DNA of invading bacterial viruses that might otherwise kill them.

Today weve adapted this molecular machinery for an entirely different purpose to change any chosen letter(s) in an organisms DNA code.

We might want to correct a disease-causing error that was inherited or crept into our DNA when it replicated. Or, in some cases, we may want to enhance the genetic code of crops, livestock or perhaps even people.

So do we just snip the unwanted gene out and replace it with a good one?

Read more: Explainer: what is genome editing?

We first have to remember that animals and plants are composed of millions of cells, and each cell contains the same DNA. There is no point editing just one cell: we would have to edit the same gene in every single cell. Wed have to snip out millions of genes and paste in millions of new ones.

And not all cells are easy to get to how could we reach cells buried in our bones or deep within a brain?

A better approach is to start at the beginning and edit the genome while there is only one cell a very early embryo.

So, all we need is a giant microscope and a tiny pair of scissors. And that is basically what we use.

Cas9 is the technical name for the virus-destroying scissors that evolved in bacteria. The CRISPR part of the name comes from repeat DNA sequences that were part of a complex system telling the scissors which part of the DNA to cut.

In order to target our Cas9 scissors, we link them to an artificial guide that directs them to the matching segment of DNA.

Remember, DNA comes in two strands, with one strand fitting alongside the other. We make a guide with a code that will line up with only one part of our 3 billion base pair long genome its like a Google search. Its truly possible for our guide to comb through vast amounts of genetic material to find the one section it matches exactly. Then our scissors can make the cut in exactly the right place.

Once the Cas9 scissors cut the DNA just where we intend, the cell will try to repair the break using any available DNA it can find. So, we also inject the new gene we want to insert.

Read more: Now we can edit life itself, we need to ask how we should use such technology

You can use a microscope and a tiny needle to inject the CRISPR/Cas9 together with the guide and the donor DNA, the new gene. Or, you can punch holes in cells with electric currents and let these things just float in, use guns to shoot them in stuck-on tiny bullets, or introduce them encapsulated in bubbles of fat that fuse with the cell membrane and release their contents inside.

But how does the new gene find the right place to embed itself? Imagine you wanted to put in the last piece of a jigsaw puzzle with 3 billion pieces, and its inside a cell, filled with goop like a passionfruit.

What youd do is fabricate a jigsaw piece of precisely the right shape and inject it into the passionfruit. Then its just a case of jiggling around until eventually the piece finds its way to the correct part of the puzzle and slots into the only place it fits.

You dont need to be able to see the DNA in our genome through the microscope its too small. And you dont really have to jiggle either random diffusion (called Brownian motion) will always deliver the jigsaw piece to the place where it fits in the end.

First, the guide will jiggle along and find the right place for the scissors to cut, and then the new donor DNA will similarly line up where it fits and will be permanently stitched into the DNA strand via natural DNA repair mechanisms.

Recently, though, new CRISPR editing systems have been created that dont even require a cut through the DNA. In this case, the CRIPSR/Cas and guide system can deliver an enzyme to a particular gene and alter it, changing perhaps an A to a G or a C to a T, rather than cutting anything out or putting anything in.

Most experiments use mouse embryos or cells grown in petri dishes in artificial liquid designed to be like blood. Other researchers are modifying stem cells that may then be re-injected into patients to repopulate damaged organs.

Only a few labs around the world are actually working with early human embryos. This research is highly regulated and carefully watched. Others work on plant cells, as whole plants can be grown from a few cells.

As we learn more, the scope of what we can do with CRISPR/Cas9 will improve. We can do a lot, but every organism and every cell is different. Whats more, everything in the body is connected, so we must think about unexpected side effects and consider the ethics of changing genes. Most of all we, as a society, should discuss and agree what we wish to achieve.

Read more: Why we can trust scientists with the power of new gene-editing technology

Read the other articles in our precision medicine series here.

Read more here:
What is CRISPR gene editing, and how does it work? - The Conversation

Arch Dis Child Educ Pract Ed. 2016 Aug; 101(4): 213215.

1HYMS Centre for Education Development (CED), Hull, York Medical School, University of York, York, UK

2Molecular Haematology Unit, MRC Weatherall Institute of Molecular Medicine, University of Oxford, John Radcliffe Hospital, Oxford, UK

3Department of Haematology, Oxford University NHS Foundation Trust, Churchill Hospital, Oxford, UK

4Academic Unit of Child Health, Sheffield Children's Hospital, Sheffield, UK

1HYMS Centre for Education Development (CED), Hull, York Medical School, University of York, York, UK

2Molecular Haematology Unit, MRC Weatherall Institute of Molecular Medicine, University of Oxford, John Radcliffe Hospital, Oxford, UK

3Department of Haematology, Oxford University NHS Foundation Trust, Churchill Hospital, Oxford, UK

4Academic Unit of Child Health, Sheffield Children's Hospital, Sheffield, UK

Received 2016 Jan 5; Revised 2016 Feb 18; Accepted 2016 Feb 19.

Keywords: CRISPR/cas9, gene editing, children, genome engineering

Clustered regularly interspaced palindromic repeats (CRISPR)/Cas9 is a gene-editing technology causing a major upheaval in biomedical research. It makes it possible to correct errors in the genome and turn on or off genes in cells and organisms quickly, cheaply and with relative ease. It has a number of laboratory applications including rapid generation of cellular and animal models, functional genomic screens and live imaging of the cellular genome.1 It has already been demonstrated that it can be used to repair defective DNA in mice curing them of genetic disorders,2 and it has been reported that human embryos can be similarly modified.3 Other potential clinical applications include gene therapy, treating infectious diseases such as HIV and engineering autologous patient material to treat cancer and other diseases. In this review we will give an overview of CRISPR/Cas9 with an emphasis on how it may impact on the specialty of paediatrics. Although it is likely to have a significant effect on paediatrics through its impact in the laboratory, here we will concentrate on its potential clinical applications. We will also describe some of the difficulties and ethical controversies associated with this novel technology.

CRISPR/Cas9 is a gene-editing technology which involves two essential components: a guide RNA to match a desired target gene, and Cas9 (CRISPR-associated protein 9)an endonuclease which causes a double-stranded DNA break, allowing modifications to the genome (see ).

The CRISPR/Cas9 system.1 Clustered regularly interspaced palindromic repeats (CRISPR) refers to sequences in the bacterial genome. They afford protection against invading viruses, when combined with a series of CRISPR-associated (Cas) proteins. Cas9, one of the associated proteins, is an endonuclease that cuts both strands of DNA. Cas9 is directed to its target by a section of RNA. This can be synthesised as a single strand called a synthetic single guide RNA (sgRNA); the section of RNA which binds to the genomic DNA is 1820 nucleotides. In order to cut, a specific sequence of DNA of between 2 and 5 nucleotides (the exact sequence depends upon the bacteria which produces the Cas9) must lie at the 3 end of the guide RNA: this is called the protospacer adjacent motif (PAM). Repair after the DNA cut may occur via two pathways: non-homologous end joining, typically leading to a random insertion/deletion of DNA, or homology directed repair where a homologous piece of DNA is used as a repair template. It is the latter which allows precise genome editing: the homologous section of DNA with the required sequence change may be delivered with the Cas9 nuclease and sgRNA, theoretically allowing changes as precise as a single base-pair.

One of the most exciting applications of CRISPR/Cas9 is its potential use to treat genetic disorders caused by single gene mutations. Examples of such diseases include cystic fibrosis (CF), Duchenne's muscular dystrophy (DMD) and haemoglobinopathies. The approach so far has currently only been validated in preclinical models, but there is hope it can soon be translated to clinical practice.

Schwank et al used CRISPR/Cas9 to investigate the treatment of CF. Using adult intestinal stem cells obtained from two patients with CF, they successfully corrected the most common mutation causing CF in intestinal organoids. They demonstrated that once the mutation had been corrected, the function of the CF transmembrane conductor receptor (CFTR) was restored.4

Another disease in which CRISPR/Cas9 has been investigated is DMD. Tabebordbar et al recently used adeno-associated virus (AAV) delivery of CRISPR/Cas9 endonucleases to recover dystrophin expression in a mouse model of DMD, by deletion of the exon containing the original mutation. This produces a truncated, but still functional protein. Treated mice were shown to partially recover muscle functional deficiencies.5 Significantly, it was demonstrated that the dystrophin gene was edited in muscle stem cells which replenish mature muscle tissue. This is important to ensure any therapeutic effects of CRISPR/Cas9 do not fade over time. Two similar studies have described using the CRISPR/Cas9 system in vivo to increase expression of the dystrophin gene and improve muscle function in mouse models of DMD.67 Other studies have used CRISPR/Cas9 to target duplication of exons in the human dystrophin gene in vitro and have shown that this approach can lead to production of full-length dystrophin in the myotubules of an individual with DMD.8

CRISPR/Cas9 could also be used to treat haemoglobinopathies. Canver et al9 recently showed BCL11A enhancer disruption by CRISPR/Cas9 could induce fetal haemoglobin in both mice and primary human erythroblast cells. In the future such an approach could allow fetal haemoglobin to be expressed in patients with abnormal adult haemoglobin. This would represent a novel therapeutic strategy in patients with diseases such as sickle cell disease or thalassaemias. Knock-in of a fully functional -globin gene is much more challenging, which is the reason for this somewhat unusual approach.

Another potential clinical application of CRISPR/Cas9 is to treat infectious diseases, such as HIV. Although antiretroviral therapy provides an effective treatment for HIV, no cure currently exists due to permanent integration of the virus into the host genome. Hu et al showed the CRISPR/Cas9 system could be used to target HIV-1 genome activity. This inactivated HIV gene expression and replication in a variety of cells which can be latently infected with HIV, without any toxic effects. Furthermore, cells could also be immunised against HIV-1 infection. This is a potential therapeutic advance in overcoming the current problem of how to eliminate HIV from infected individuals. After further refinement, the authors suggest their findings may enable gene therapies or transplantation of genetically altered bone marrow stem cells or inducible pluripotent stem cells to eradicate HIV infection.10

There has been increasing interest in the possibility of using CRISPR/Cas9 to modify patient-derived T-cells and stem/progenitor cells which can then be reintroduced into patients to treat disease. This approach may overcome some of the issues associated with how to efficiently deliver gene editing to the right cells.

T-cell genome engineering has already shown success in treating haematological malignancies and has the potential to treat solid cancers, primary immune deficiencies and autoimmune diseases. Genetic manipulation of T-cells has previously been inefficient. However, Schumann et al recently reported a more effective approach in human CD4+ T-cells based on the CRISPR/Cas9 system. Their technique allowed experimental and therapeutic knock-out and knock-in editing of the genome in primary human T-cells. They demonstrated T-cells could be manipulated to prevent expression of the protein PD-1, which other work has shown may allow T-cells to target solid cancers.11

There is also interest in using CRISPR/Cas9-mediated genome editing in pluripotent stem cells or primary somatic stem cells to treat disease. For example Xie et al12 showed the mutation causing -thalassaemia could be corrected in human induced pluripotent stem cells ex vivo. They suggest that in the future such an approach could provide a source of cells for bone marrow transplantation to treat -thalassaemia and other similar monogenic diseases.

A number of challenges remain before the potential of CRISPR/Cas9 can be translated to effective treatments at the bedside. A particular issue is how to deliver gene editing to the right cells, especially if the treatment is to be delivered in vivo. To safely deliver Cas9-nuclease encoding genes and guide RNAs in vivo without any associated toxicity, a suitable vector is needed. AAV has previously been a favoured option for gene delivery.1 However, this delivery system may be too small to allow efficient transduction of the Cas9 gene.1 A smaller Cas9 gene could be used, but this has additional implications on efficacy.1 A number of other non-viral delivery systems are under investigation and this process requires further optimisation.

Another significant concern is the possibility of off-target effects on parts of the genome separate from the region being targeted. Unintentional edits of the genome could have profound long-term complications for patients, including malignancy. The concentration of the Cas9 nuclease enzyme and the length of time Cas9 is expressed are both important when limiting off-target activity.1 Although recent modifications in the nuclease have increased specificity, further work is required to minimise off-target effects and to establish the long-term safety of any treatment.

The therapeutic applications of CRISPR/Cas9 considered in this article have predominantly been directed at somatic cells. A particularly controversial issue surrounding CRISPR/Cas9 is that of gene editing in embryos. It has already been shown that CRISPR/Cas9 technology can alter the genome of human embryos3 which theoretically could prove useful in the preimplantation treatment of genetic diseases. However, any genetic modification of the germline would be permanent and the long-term consequences are unclear. Many oppose germline modification under any circumstances, reasoning that an eventual consequence could be non-therapeutic genetic enhancement.13 It is clear that the ethical boundaries, within which CRISPR/Cas9 can be used, remain to be fully determined.

Clinical bottom line

CRISPR/Cas9 technology has the potential to revolutionise the treatment of many paediatric conditions.

A number of practical and ethical challenges must be overcome before this potential can be realised at the bedside.

Contributors: DK conceived the idea for this article. All authors were involved in writing and reviewing the final manuscript.

Funding: AK is supported by a Wellcome Trust Fellowship (108785/Z/15/Z).

Competing interests: None declared.

Provenance and peer review: Not commissioned; externally peer reviewed.

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What is CRISPR/Cas9? - PMC - National Center for Biotechnology Information

Ten years ago this week, Jennifer Doudna and her colleagues published the results of a test-tube experiment on bacterial genes. When the study came out in the journal Science on June 28, 2012, it did not make headline news. In fact, over the next few weeks, it did not make any news at all.

Looking back, Dr. Doudna wondered if the oversight had something to do with the wonky title she and her colleagues had chosen for the study: A Programmable Dual RNA-Guided DNA Endonuclease in Adaptive Bacterial Immunity.

I suppose if I were writing the paper today, I would have chosen a different title, Dr. Doudna, a biochemist at the University of California, Berkeley, said in an interview.

Far from an esoteric finding, the discovery pointed to a new method for editing DNA, one that might even make it possible to change human genes.

I remember thinking very clearly, when we publish this paper, its like firing the starting gun at a race, she said.

In just a decade, CRISPR has become one of the most celebrated inventions in modern biology. It is swiftly changing how medical researchers study diseases: Cancer biologists are using the method to discover hidden vulnerabilities of tumor cells. Doctors are using CRISPR to edit genes that cause hereditary diseases.

The era of human gene editing isnt coming, said David Liu, a biologist at Harvard University. Its here.

But CRISPRs influence extends far beyond medicine. Evolutionary biologists are using the technology to study Neanderthal brains and to investigate how our ape ancestors lost their tails. Plant biologists have edited seeds to produce crops with new vitamins or with the ability to withstand diseases. Some of them may reach supermarket shelves in the next few years.

CRISPR has had such a quick impact that Dr. Doudna and her collaborator, Emmanuelle Charpentier of the Max Planck Unit for the Science of Pathogens in Berlin, won the 2020 Nobel Prize for chemistry. The award committee hailed their 2012 study as an epoch-making experiment.

Dr. Doudna recognized early on that CRISPR would pose a number of thorny ethical questions, and after a decade of its development, those questions are more urgent than ever.

Will the coming wave of CRISPR-altered crops feed the world and help poor farmers or only enrich agribusiness giants that invest in the technology? Will CRISPR-based medicine improve health for vulnerable people across the world, or come with a million-dollar price tag?

The most profound ethical question about CRISPR is how future generations might use the technology to alter human embryos. This notion was simply a thought experiment until 2018, when He Jiankui, a biophysicist in China, edited a gene in human embryos to confer resistance to H.I.V. Three of the modified embryos were implanted in women in the Chinese city of Shenzhen.

In 2019, a court sentenced Dr. He to prison for illegal medical practices. MIT Technology Review reported in April that he had recently been released. Little is known about the health of the three children, who are now toddlers.

Scientists dont know of anyone else who has followed Dr. Hes example yet. But as CRISPR continues to improve, editing human embryos may eventually become a safe and effective treatment for a variety of diseases.

Will it then become acceptable, or even routine, to repair disease-causing genes in an embryo in the lab? What if parents wanted to insert traits that they found more desirable like those related to height, eye color or intelligence?

Franoise Baylis, a bioethicist at Dalhousie University in Nova Scotia, worries that the public is still not ready to grapple with such questions.

Im skeptical about the depth of understanding about whats at issue there, she said. Theres a difference between making people better and making better people.

Dr. Doudna and Dr. Charpentier did not invent their gene-editing method from scratch. They borrowed their molecular tools from bacteria.

In the 1980s, microbiologists discovered puzzling stretches of DNA in bacteria, later called Clustered Regularly Interspaced Short Palindromic Repeats. Further research revealed that bacteria used these CRISPR sequences as weapons against invading viruses.

The bacteria turned these sequences into genetic material, called RNA, that could stick precisely to a short stretch of an invading viruss genes. These RNA molecules carry proteins with them that act like molecular scissors, slicing the viral genes and halting the infection.

As Dr. Doudna and Dr. Charpentier investigated CRISPR, they realized that the system might allow them to cut a sequence of DNA of their own choosing. All they needed to do was make a matching piece of RNA.

To test this revolutionary idea, they created a batch of identical pieces of DNA. They then crafted another batch of RNA molecules, programming all of them to home in on the same spot on the DNA. Finally, they mixed the DNA, the RNA and molecular scissors together in test tubes. They discovered that many of the DNA molecules had been cut at precisely the right spot.

For months Dr. Doudna oversaw a series of round-the-clock experiments to see if CRISPR might work not only in a test tube, but also in living cells. She pushed her team hard, suspecting that many other scientists were also on the chase. That hunch soon proved correct.

In January 2013, five teams of scientists published studies in which they successfully used CRISPR in living animal or human cells. Dr. Doudna did not win that race; the first two published papers came from two labs in Cambridge, Mass. one at the Broad Institute of M.I.T. and Harvard, and the other at Harvard.

Lukas Dow, a cancer biologist at Weill Cornell Medicine, vividly remembers learning about CRISPRs potential. Reading the papers, it looked amazing, he recalled.

Dr. Dow and his colleagues soon found that the method reliably snipped out pieces of DNA in human cancer cells.

It became a verb to drop, Dr. Dow said. A lot of people would say, Did you CRISPR that?

Cancer biologists began systematically altering every gene in cancer cells to see which ones mattered to the disease. Researchers at KSQ Therapeutics, also in Cambridge, used CRISPR to discover a gene that is essential for the growth of certain tumors, for example, and last year, they began a clinical trial of a drug that blocks the gene.

Caribou Biosciences, co-founded by Dr. Doudna, and CRISPR Therapeutics, co-founded by Dr. Charpentier, are both running clinical trials for CRISPR treatments that fight cancer in another way: by editing immune cells to more aggressively attack tumors.

Those companies and several others are also using CRISPR to try to reverse hereditary diseases. On June 12, researchers from CRISPR Therapeutics and Vertex, a Boston-based biotech firm, presented at a scientific meeting new results from their clinical trial involving 75 volunteers who had sickle-cell anemia or beta thalassemia. These diseases impair hemoglobin, a protein in red blood cells that carries oxygen.

The researchers took advantage of the fact that humans have more than one hemoglobin gene. One copy, called fetal hemoglobin, is typically active only in fetuses, shutting down within a few months after birth.

The researchers extracted immature blood cells from the bone marrow of the volunteers. They then used CRISPR to snip out the switch that would typically turn off the fetal hemoglobin gene. When the edited cells were returned to patients, they could develop into red blood cells rife with hemoglobin.

Speaking at a hematology conference, the researchers reported that out of 44 treated patients with beta thalassemia, 42 no longer needed regular blood transfusions. None of the 31 sickle cell patients experienced painful drops in oxygen that would have normally sent them to the hospital.

CRISPR Therapeutics and Vertex expect to ask government regulators by the end of year to approve the treatment.

Other companies are injecting CRISPR molecules directly into the body. Intellia Therapeutics, based in Cambridge and also co-founded by Dr. Doudna, has teamed up with Regeneron, based in Westchester County, N.Y., to begin a clinical trial to treat transthyretin amyloidosis, a rare disease in which a damaged liver protein becomes lethal as it builds up in the blood.

Doctors injected CRISPR molecules into the volunteers livers to shut down the defective gene. Speaking at a scientific conference last Friday, Intellia researchers reported that a single dose of the treatment produced a significant drop in the protein level in volunteers blood for as long as a year thus far.

The same technology that allows medical researchers to tinker with human cells is letting agricultural scientists alter crop genes. When the first wave of CRISPR studies came out, Catherine Feuillet, an expert on wheat, who was then at the French National Institute for Agricultural Research, immediately saw its potential for her own work.

I said, Oh my God, we have a tool, she said. We can put breeding on steroids.

At Inari Agriculture, a company in Cambridge, Dr. Feuillet is overseeing efforts to use CRISPR to make breeds of soybeans and other crops that use less water and fertilizer. Outside of the United States, British researchers have used CRISPR to breed a tomato that can produce vitamin D.

Kevin Pixley, a plant scientist at the International Maize and Wheat Improvement Center in Mexico City, said that CRISPR is important to plant breeding not only because its powerful, but because its relatively cheap. Even small labs can create disease-resistant cassavas or drought-resistant bananas, which could benefit poor nations but would not interest companies looking for hefty financial returns.

Because of CRISPRs use for so many different industries, its patent has been the subject of a long-running dispute. Groups led by the Broad Institute and the University of California both filed patents for the original version of gene editing based on CRISPR-Cas9 in living cells. The Broad Institute won a patent in 2014, and the University of California responded with a court challenge.

In February of this year, the U.S. Patent Trial and Appeal Board issued what is most likely the final word on this dispute. They ruled in favor of the Broad Institute.

Jacob Sherkow, an expert on biotech patents at the University of Illinois College of Law, predicted that companies that have licensed the CRISPR technology from the University of California will need to honor the Broad Institute patent.

The big-ticket CRISPR companies, the ones that are farthest along in clinical trials, are almost certainly going to need to write the Broad Institute a really big check, he said.

The original CRISPR system, known as CRISPR-Cas9, leaves plenty of room for improvement. The molecules are good at snipping out DNA, but theyre not as good at inserting new pieces in their place. Sometimes CRISPR-Cas9 misses its target, cutting DNA in the wrong place. And even when the molecules do their jobs correctly, cells can make mistakes as they repair the loose ends of DNA left behind.

A number of scientists have invented new versions of CRISPR that overcome some of these shortcomings. At Harvard, for example, Dr. Liu and his colleagues have used CRISPR to make a nick in one of DNAs two strands, rather than breaking them entirely. This process, known as base editing, lets them precisely change a single genetic letter of DNA with much less risk of genetic damage.

Dr. Liu has co-founded a company called Beam Therapeutics to create base-editing drugs. Later this year, the company will test its first drug on people with sickle cell anemia.

Dr. Liu and his colleagues have also attached CRISPR molecules to a protein that viruses use to insert their genes into their hosts DNA. This new method, called prime editing, could enable CRISPR to alter longer stretches of genetic material.

Prime editors are kind of like DNA word processors, Dr. Liu said. They actually perform a search and replace function on DNA.

Rodolphe Barrangou, a CRISPR expert at North Carolina State University and a founder of Intellia Therapeutics, predicted that prime editing would eventually become a part of the standard CRISPR toolbox. But for now, he said, the technique was still too complex to become widely used. Its not quite ready for prime time, pun intended, he said.

Advances like prime editing didnt yet exist in 2018, when Dr. He set out to edit human embryos in Shenzen. He used the standard CRISPR-Cas9 system that Dr. Doudna and others had developed years before.

Dr. He hoped to endow babies with resistance to H.I.V. by snipping a piece of a gene called CCR5 from the DNA of embryos. People who naturally carry the same mutation rarely get infected by H.I.V.

In November 2018, Dr. He announced that a pair of twin girlshad been born with his gene edits. The announcement took many scientists like Dr. Doudna by surprise, and they roundly condemned him for putting the health of the babies in jeopardy with untested procedures.

Dr. Baylis of Dalhousie University criticized Dr. He for the way he reportedly presented the procedure to the parents, downplaying the radical experiment they were about to undertake. You could not get an informed consent, unless you were saying, This is pie in the sky. Nobodys ever done it, she said.

In the nearly four years since Dr. Hes announcement, scientists have continued to use CRISPR on human embryos. But they have studied embryos only when theyre tiny clumps of cells to find clues about the earliest stages of development. These studies could potentially lead to new treatments for infertility.

Bieke Bekaert, a graduate student in reproductive biology at Ghent University in Belgium, said that CRISPR remains challenging to use in human embryos. Breaking DNA in these cells can lead to drastic rearrangements in the chromosomes. Its more difficult than we thought, said Ms. Bekaert, the lead author of a recent review of the subject. We dont really know what is happening.

Still, Ms. Bekaert held out hope that prime editing and other improvements on CRISPR could allow scientists to make reliably precise changes to human embryos. Five years is way too early, but I think in my lifetime it may happen, she said.

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CRISPR, 10 Years On: Learning to Rewrite the Code of Life

CRISPR is the basis of a revolutionary gene editing system. One day, it could make it possible to do everything from resurrect extinct species to develop cures for chronic disease.

Its built on a natural adaptation found in the DNA of bacteria and single-celled organisms.

CRISPR stands for Clustered Regularly Interspaced Short Palindromic Repeats.

Theyre really just bits of genetic code with a specific, recognizable format. They contain a sequence that shows up over and over again, though its often reversed each time.

Thats what makes it "Palindromic: palindromes are words that can be read the same backwards as forwards. Palindromes are common in DNA. Some serve as backups for damage to our genetic code, while others are common in cancer mutations.

With CRISPR, a group of enzymes recognize certain repeats, and break the DNA there to insert important information in the middle. These insertions are called spacers, and they contain the genetic code of different viruses that have invaded in the past.

Such previous invasions served a very important evolutionary purpose: immunizing against foreign threats.

Researchers first discovered CRISPR in E. Coli in the 1980s. When E. coli survives viral attacks, it incorporates some of the virus DNA into its own genetic code.

E. Coli isnt unique in using this strategy. Between the 1980s and 2000, scientists found that lots of bacteria and single-celled organisms incorporate viral DNA this way.

Cells use these sequences as templates for transcribing complementary strands of RNA.

When viruses matching the template sequence enter the cell, the complementary RNA binds to them, and directs a series of CRISPR-associated enzymes, or Cas enzymes, to attackcutting invader DNA at the binding site. That neutralizes the viral threat.

The CRISPR-Cas system is incredibly effective. Its also easy to manipulate, letting us alter a cells genetic code however we want.

In 2012, French microbiologist Emmanuelle Charpentier and American biochemist Jennifer Doudna discovered that Cas enzymesspecifically Cas9can be reprogrammed to cut nearly any part of the genome, using RNA sequences made in a lab. Those guide RNA molecules tell Cas9 where to cut DNA in a cell.

For their discovery, Charpentier and Doudna won the Nobel Prize in Chemistry in 2020.

And the use of CRISPR has taken off in science since their breakthrough.

But scientists are still far from realizing CRISPRs potential.

Cas9 is great at suppressing or knocking out unwanted genes. But for most medical purposes, its not enough to cut unwanted DNA out. Scientists need to control how the DNA repairs itself.

Left to their own devices, cells tend to repair broken DNA using a method that introduces lots of random errors. Researchers can provide cells with templates to guide the repair process, but theyre still working on making that more reliable.

Researchers have found lots of applications for CRISPR in animals, like making disease-resistant chickens and pigs, and mosquitos that cant bite or lay eggs. But theyve got many projects underway, like making disease-resistant cropsincluding wine grapes. More ambitiously, theyre working to genetically alter pigs so their organs could be transplanted into humans. And bring extinct species such as the passenger pigeon back to life, by tweaking the genomes of similar birds.

When it comes to the human genome, though, scientists have been more hesitant. Editing our own DNA could easily end up causing more problems than it solves.

While Cas9 reliably cuts DNA where we want it to, recent experiments have shown it can also affect genes far off-target. And even if we could get it to work reliably, many experts have flagged ethical concerns about using the technology for eugenics and designer babies. If parents can one day pay scientists to edit their babies DNA, making them stronger and smarter, CRISPR could make the world even more unequal and prejudiced.

In 2018, Chinese researcher HEH JEE'-an-qway claimed hed used CRISPR to make HIV-resistant children. Whether or not he succeeded, his work violated Chinas National Health Commission rules, and he was sentenced to three years in prison.

Using CRISPR on babies is widely illegal. But there are some cases where using CRISPR on humans may be worth the risk.

In 2020, American researchers began the first clinical trials injecting CRISPR directly into living humans, aiming to repair a genetic mutation that causes blindness.

Many researchers hope CRISPR-based therapies could eventually cure hereditary diseases; theyve already seen promising results in various animal studies. Though given the risks of editing the human genome, were still a ways off from widespread use of CRISPR in medicine.

CRISPR has given science a tool to reliably tinker with the code of life. But the question remains: can we do so safely and ethically, while avoiding the unintended consequences of such power?

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What Is CRISPR, and Why Is It So Important? - Scientific American

Global CRISPR Technology Market Is Projected To Grow At A 22% Rate Through The Forecast Period  EIN News

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Global CRISPR Technology Market Is Projected To Grow At A 22% Rate Through The Forecast Period - EIN News

CNN

Two women have won the Nobel prize in chemistry for the development of a revolutionary gene editing tool thats been described as rewriting the code of life.

The technique discovered by Emmanuelle Charpentier, the director at the Max Planck Institute for Infection Biology, and Jennifer A. Doudna, a biochemist at the University of California Berkeley, is known as CRISPR/Cas9.

It hit the headlines in 2018 when a Chinese scientist used the technology to create the first gene-edited babies, shocking the world and sparking a highly charged ethical debate about its use.

What is CRISPR (pronounced crisper) and why has it been controversial?

DNA is like the instruction manual for life on our planet, and CRISPR/Cas9 can target sites in genetic material.

This allows scientists to change it by knocking out a particular gene or inserting new genetic material at a predetermined site in our DNA.

Cas9, a type of modified protein, acts like a pair of scissors that can snip parts of DNA strands. CRISPR stands for clustered regularly interspaced short palindromic repeats a repeated DNA sequence in genomes.

Doudna and Charpentier showed that CRISPR works like a pair of scissors that can be targeted to cut specific DNA sequences, said Andrew Holland, an assistant professor in the Department of Molecular Biology and Genetics at Johns Hopkins School of Medicine. After cutting, the repair of the DNA code enables it to be altered. This has allowed scientists to change the DNA code in a targeted way to help understand and treat genetic disease, he told CNN via email.

The technology has worked in pretty much every organism that it has been used on, including plants, microbes and humans.

What the system does is that it can recognize (a) certain specific gene in the genome of ourselves and correct mutations, do some copy pasting, do some editing like we edit a text. The system can edit the genome and change the properties of the genes, Charpentier said in 2016 when she was interviewed by CNN.

It is already having a major impact on biomedical research, clinical medicine and agriculture. For example, its been used to grow rice that accumulates lower levels of potentially toxic heavy metals and create livestock with more desirable traits.

It was used for the first time in humans in 2016 and a trial is underway in the United States to use the experimental technology to treat a dozen patients with sickle cell disease, a group of inherited blood disorders.

Related technologies may be able to potentially correct up to 89% of genetic defects, scientists have said.

Its not an exaggeration to say that the technology that arose from Doudna and Charpentiers discoveries has revolutionized the field, Jessica Downs, the deputy head of the Division of Cancer Biology at the Institute of Cancer Research in the UK, told the Science Media Centre in London.

We adopted the technology in our lab to investigate molecular changes that lead to cancer development. Its been transformative in terms of what we can achieve, but there is also great potential for using this technology in the clinic. And on a more personal level, its inspiring and uplifting to see two women honored for their work in this way.

While it has immense potential to transform our lives, the technology has raised many ethical questions.

Chinese scientist He Jiankui was jailed for three years in 2019 after announcing that twin girls had been born with modified DNA to make them resistant to HIV, which he had managed using the gene-editing tool CRISPR/Cas9 before birth.

An associate professor at the Southern University of Science and Technology in Shenzhen at the time, he said that he was proud of the achievement. But he was condemned by many of his peers, with the experiment labeled monstrous, unethical and a huge blow to the reputation of Chinese biomedical research.

Claes Gustafsson, secretary of the Nobel committee in chemistry and a professor of biochemistry and biophysics at Stockholm University, said that with every really powerful technology, in life sciences or elsewhere, theres a possibility of misuse.

Clearly this Chinese researcher was way out of line in applying it in this particular way, he told CNN.

Everyone has agreed that it cannot be used for germline engineering. You cant make heritable changes to human DNA. That is far too uncertain at this point, added Gustafsson. There are specific genetic diseases you can think of curing for the individual but not in a heritable way.

Scientists have called for a moratorium on human germline editing, while efforts are being made to better regulate use of the technology. An international commission said in September it was too early for gene-edited human embryos to be used to create a pregnancy.

Doudna has expressed deep concern about Hes work, telling CNN it was not medically necessary and there was no way to defend using an experimental technology when there were established ways of avoiding HIV transmission.

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What is CRISPR and why is it controversial? | CNN

CRISPR, in full clustered regularly interspaced short palindromic repeats, short palindromic repeating sequences of DNA, found in most bacterial genomes, that are interrupted by so-called spacer elements, or spacerssequences of genetic code derived from the genomes of previously encountered bacterial pathogens. CRISPR elements are found naturally in many bacteria and archaea, where they provide a sort of genetic memory, enabling the cells to efficiently detect and destroy pathogens, particularly viruses known as bacteriophages.

As a naturally occurring adaptive defense system, CRISPR functions by destroying nucleic acids from pathogens that invade the cell. The effectiveness and efficiency of CRISPR immunity is directly linked to the presence of spacer elements. Spacer elements essentially are recognition sequences that match sequences inpathogen genomes; as spacers from newly encountered pathogens are added to the bacterial genome, the cell gains the ability to recognize those pathogens on repeat encounters. Most new spacer elements are inserted only at one end of the CRISPR region; hence, across the length of the CRISPR region exists a record of pathogens that have been encountered by the cell and its ancestors over time. Less often, spacers are added in other places in a process called ectopic integration.

The CRISPR system works by producing small guide RNA sequences that correspond to specific DNA targets. Guide RNAs, generated via transcription of the CRISPR region,include hairpin formations, derived from the palindromic repeats, that are linked to sequencesderivedfrom the spacer elements. When guide RNAs bind to their DNA targets, an RNA-DNA heteroduplex is formed. The heteroduplex binds to a nuclease called CRISPR-associated (Cas), which catalyzes the cleavage of double-stranded DNA at a position near the junction of the target-specific sequence and the palindromic repeat in the guide RNA. In this way, the nuclease destroys invading pathogenic genomes.

CRISPR interacts with multiple Cas proteins as part of the defense response, and thus there are distinct CRISPR-Cas defense systems. The three major systems are type I, type II, and type III. The type I system is defined by the presence of Cas3 protein. Cas3 forms part of the so-called CRISPR-associated complex for antiviral defense (or Cascade) -like complex, which binds a guide RNA and identifies the target sequence for destruction. The type II system is based on the presence of several proteins, namely Cas1, Cas2, Cas9, and, in some cases, Cas4. The Cas9 protein is considered a signature element of the type II system, owing to its essential role in facilitating cellular adaptation to new pathogens and to its participation in RNA processing and cleaving of target DNA. The signature protein of the type III system is Cas10. The type III system differs from its two counterparts in that, in addition to targeting DNA, it identifies RNA targets.

The high sequence specificity of the CRISPR system has drawn significant interest in the field of gene editing. The functional precision of CRISPR allows researchers to remove and insert DNA in desired locations within a genome, making it possible to correct genetic defects in animals and to modify DNA sequences in embryonicstem cells. These types of sequence corrections and alterations are possible because RNA-DNA heteroduplexes are stable and because designing an RNA sequence that binds specifically to a unique target DNA sequence is based simply on the Watson-Crickbase-pairingrules (adenine binds to thymine [or uracil in RNA], and cytosine binds to guanine).

The possibility of using CRISPR as a gene-editing technology was recognized in 2012 by American scientistJennifer Doudna, French scientistEmmanuelle Charpentier, and colleagues. These researchers discovered that guide RNAs produced by CRISPR bind to nucleases, which then target particular DNA sequences, and that such RNAs could be modified to bind to a desired sequence. The researchers found that the type II CRISPR-Cas9 system was especially versatile for correcting or altering desired target sequences. Doudna and Charpentier shared the 2020 Nobel Prize in Chemistry for their work.

In 2015 American scientistFeng Zhang and colleagues developed a new version of CRISPR technology using the microbial nuclease CRISPR from Prevotella and Francisella 1 (Cpf1) in place of Cas9. Unlike Cas9, Cpf1 requires only a single CRISPR guide RNA for specificity and introduces staggered (rather than blunt) cuts in double-stranded DNA, which in certain instances can give greater control over the modification of target DNA sequences. Zhang and colleagues subsequently developed multiple other CRISPR gene-editing tools, including CRISPR-Cas13 systems, which target RNA.

CRISPR gene-editing technology has a wide array of research and medical applications. For example, in the laboratory, CRISPR systems can be used to modify genes in bacteria and in animal and plant models, enabling researchers to gain new understanding of the effects of genetic modification. Although preexisting genetic engineering technologies have allowed researchers to investigate various types of genetic modifications and alterations for decades, CRISPR is less costly, more efficient, and more reliable.

In addition, different CRISPR-based therapies are being explored in clinical trials for the treatment of certain human diseases. Some examples include novel treatments for diabetes; for sickle cell disease; for cancers of blood-forming tissues, such as multiple myeloma, leukemia,and lymphoma; for chronic infectious diseases, such as AIDS; and for a form of inherited impairment in vision known as Leber congenital amaurosis. Investigations of CRISPR-based therapies in humans are helping to shed light on how DNA alterations induced by CRISPR enzymes affect cells, on how the human immune system responds to CRISPR-derived interventions, and on risks associated with unwanted off-target alterations in DNA.

The ability to easily and accurately edit genes using CRISPR technology has raised significant ethical issues. In particular, CRISPR can be used to modify DNA sequences in embryonicstem cells, such as in germ-line (spermandegg) genome modification in humans. Critics point out that this ability, applied to embryos in the womb, may be used to improve traits such as intelligence, appearance, and athletic ability, potentially introducing permanent changes in human DNA. The generation of such designer babies sparked debates about the morality of tampering with human development and the ethics of who would have access to the technology. The worlds first gene-edited human babies were born in late 2018 in China; the infants, twin girls, carried an edited gene that reduced the risk of HIV infection.

Following the birth of gene-edited babies, some medical and bioethics researchers, including Charpentier, called for a moratorium on editing human genes in eggs, sperm, or embryos. They contend that because there remain many unknowns about the technology, scientists may unintentionally introduce as many genetic errors as they attempt to fix. Nonetheless, critics argue that CRISPR technology is a remarkable achievement with significant potential to improve human health, although under rigorously controlled conditions.

Excerpt from:
CRISPR | Description, Technology, Uses, & Ethical Concerns

Figure 4.. Terminal differentiation of T EFF cells is dependent upon Pofut1 and associated with

(A) Differentially expressed genes in sgPofut1- compared to sgNTC-transduced P14 cells at day 7.5 post-infection (p.i.). Upregulated (orange) or downregulated (blue) transcripts [false discovery rate (FDR) < 0.05] are highlighted. Selective MP- and TE-associated genes are labelled. (B) Enrichment plots of cell cycle-related signatures. NES, normalized enrichment score. (C) Flow cytometry (left) and quantification (right) of BrdU incorporation. (D) UMAP plot of published scRNA-seq dataset of P14 cells at day 8 p.i. (Chen et al., 2019). Each dot corresponds to an individual cell. The number and frequency of cells in each of the color-coded clusters (clusters 13) are indi cated. (E) Violin plots of Klrg1, Cxcr3 or Il7r expression in clusters 13 from (D). (F) Gating str ategy (left) and quantification (right) of the proportions of TE (KLRG1hiCXCR3loCD127lo), MP (KLRG1loCXCR3hiCD127hi) and TINT (CXCR3hiCD127lo) cells among WT P14 cells. (G) PCA plot of TE, MP and TINT cells [gating strategy in (F)] at day 7.5 p.i., with the percentage of variance shown. (H) Quantification of the relative frequency of BrdU+ cells in MP and TINT cells compared to TE cells. (I) Diagram of the in vivo differentiation assay (left), flow cytometry of KLRG1 versus CXCR3 expression (middle), and quantification of TINT, TE and CXCR3hiCD127hi cells (right). Only representative plots of KLRG1 versus CXCR3 are shown (TE population is largely defined by KLRG1hiCXCR3lo cells, which constitute ~ 95% of TE cells). (J) Quantification of TE, MP and TINT cells in the indicated P14 cells. (K) UMAP plot of Pofut1-dependent signature [downregulated genes as identified in (A)] in published scRNA-seq dataset from (D) (Chen et al., 2019). (L) UMAP plot of scRNA-seq data from sgNTC- (in black, left) and sgPofut1- (in red, right) transduced P14 cells (from dual-color transfer system) at day 7 p.i. Gray shadow indicates location of all cells; the number of analyzed cells in each group is indicated. (M) UMAP plot of the expression of Klrg1 (left), Cxcr3 (middle) and Il7r (right) in scRNA-seq data described in (L). (N) Flow cytometry of KLRG1 versus CXCR3 expression (left) and quantification (right) of TE cells in the in vivo differentiation assay similar as (I), except for the use of both wild-type and Pofut1-null TINT groups as the pretransfer cells. Data are from one (A, B, D, E, G, and KM), representative of two (C, H, and N), or compiled from at least two (I, J, and N) independent experiments, with 4 (A, C, G, H, and I), 17 (F), 11 (J), or 3 (L and N) biological replicates per group. *P < 0.05, **P < 0.01, and ***P < 0.001; NS, not significant; two-tailed paired Students t-test (C), two-tailed unpaired Students t-test (I, J, and N), or one-way analysis of variance (ANOVA) (F and H). Data are presented as mean s.e.m. See also Figures S4S6 and Tables S3S6.

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In vivo CRISPR screening reveals nutrient signaling processes ... - PubMed

CRISPR is a technology that can be used to edit genes and, as such, will likely change the world.

The essence of CRISPR is simple: its a way of finding a specific bit of DNA inside a cell. After that,the next step in CRISPR gene editing is usually to alter that piece of DNA. However, CRISPR has also been adapted to do other things too, such as turning genes on or off without altering their sequence.

There were ways to edit the genomes of some plants and animals before the CRISPR method was unveiled in 2012 but it took years and cost hundreds of thousands of dollars. CRISPR has made it cheap and easy.

CRISPR is already widely used for scientific research, and in the not too distant future many ofthe plantsandanimalsinour farms, gardens or homes may have been altered with CRISPR. In fact, some people already are eating CRISPRed food.

CRISPR technology also has the potential to transform medicine, enabling us to not onlytreatbut alsopreventmany diseases. We may even decide to use it tochange the genomesofour children. An attempt to do this in Chinahasbeen condemned as premature and unethical, but some think it could benefit children in the future.

CRISPR is being used for all kinds of other purposes too, from fingerprinting cells andlogging what happensinside them todirecting evolutionand creatinggene drives.

The key to CRISPR is the many flavours of Cas proteins found in bacteria, where they help defend against viruses. The Cas9 protein is the most widely used by scientists. This protein can easily be programmed to find and bind to almost any desired target sequence, simply by giving it a piece of RNA to guide it in its search.

When the CRISPR Cas9 protein is added to a cell along with a piece of guide RNA, the Cas9 protein hooks up with the guide RNA and then moves along the strands of DNA until it finds and binds to a 20-DNA-letter long sequence that matches part of the guide RNA sequence. Thats impressive, given thatthe DNA packed into each of our cellshas six billion letters and is two metres long.

What happens next can vary. The standard Cas9 protein cuts the DNA at the target. When the cut is repaired, mutations are introduced that usually disable a gene. This is by far the most common use of CRISPR. Its called genome editing or gene editing but usually the results arenot as preciseas that term implies.

CRISPR can also be used tomake precise changessuch as replacing faulty genes true genome editing but this is far more difficult.

Customised Cas proteins have been created that do not cut DNA or alter it in any way,but merely turn genes on or off: CRISPRa and CRISPRi respectively. Yet others, called base editors,change one letter of the DNA code to another.

So why do we call it CRISPR? Cas proteins are used by bacteria to destroy viral DNA. They add bits of viral DNA to their own genome to guide the Cas proteins, and the odd patterns of these bits of DNA are what gave CRISPR its name: clustered regularly interspaced short palindromic repeats. Michael Le Page

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What is CRISPR? | New Scientist

Many bacterial clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR-associated (Cas) systems employ the dual RNA-guided DNA endonuclease Cas9 to defend against invading phages and conjugative plasmids by introducing site-specific double-stranded breaks in target DNA. Target recognition strictly requires the presence of a short protospacer adjacent motif (PAM) flanking the target site, and subsequent R-loop formation and strand scission are driven by complementary base pairing between the guide RNA and target DNA, Cas9-DNA interactions, and associated conformational changes. The use of CRISPR-Cas9 as an RNA-programmable DNA targeting and editing platform is simplified by a synthetic single-guide RNA (sgRNA) mimicking the natural dual trans-activating CRISPR RNA (tracrRNA)-CRISPR RNA (crRNA) structure. This review aims to provide an in-depth mechanistic and structural understanding of Cas9-mediated RNA-guided DNA targeting and cleavage. Molecular insights from biochemical and structural studies provide a framework for rational engineering aimed at altering catalytic function, guide RNA specificity, and PAM requirements and reducing off-target activity for the development of Cas9-based therapies against genetic diseases.

Keywords: CRISPR; Cas9; genome engineering; mechanism; off-target; structure.

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CRISPR-Cas9 Structures and Mechanisms - PubMed

In July, an HIV-positive man became the first volunteer in a clinical trial aimed at using Crispr gene editing to snip the AIDS-causing virus out of his cells. For an hour, he was hooked up to an IV bag that pumped the experimental treatment directly into his bloodstream. The one-time infusion is designed to carry the gene-editing tools to the mans infected cells to clear the virus.

Later this month, the volunteer will stop taking the antiretroviral drugs hes been on to keep the virus at undetectable levels. Then, investigators will wait 12 weeks to see if the virus rebounds. If not, theyll consider the experiment a success. What were trying to do is return the cell to a near-normal state, says Daniel Dornbusch, CEO of Excision BioTherapeutics, the San Francisco-based biotech company thats running the trial.

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Crispr isbeing used in several other studiesto treat a handful of conditions that arise from genetic mutations. In those cases, scientists are using Crispr to edit peoples own cells. But for the HIV trial, Excision researchers are turning the gene-editing tool against thevirus. The Crispr infusion contains gene-editing molecules that target two regions in the HIV genome important for viral replication. The virus can only reproduce if its fully intact, so Crispr disrupts that process by cutting out chunks of the genome.

This is an excerpt. Read the full article here

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A CRISPR cure for HIV? Gene-editing technology may be able stop viral replication in its tracks and wipe out infections - Genetic Literacy Project

Studies in non-human primates demonstrated nearly 100% gene editing and knockout of endogenous RHO gene and more than 30% replacement protein levels using a dual vector AAV approach

Treated eyes showed morphological and functional photoreceptor preservation

EDIT-103 advancing towards IND-enabling studies

CAMBRIDGE, Mass., Oct. 13, 2022 (GLOBE NEWSWIRE) -- Editas Medicine, Inc. (Nasdaq: EDIT), a leading genome editing company, today announced ex vivo and in vivo preclinical data supporting its experimental medicine EDIT-103 for the treatment of rhodopsin-associated autosomal dominant retinitis pigmentosa (RHO-adRP). The Company reported these data in an oral presentation today at the European Society of Gene and Cell Therapy 29th Annual Meeting in Edinburgh, Scotland, UK.

EDIT-103 is a mutation-independent CRISPR/Cas9-based, dual AAV5 vectors knockout and replace (KO&R) therapy to treat RHO-adRP. This approach has the potential to treat any of over 150 dominant gain-of-function rhodopsin mutations that cause RHO-adRP with a one-time subretinal administration.

These promising preclinical data demonstrate the potential of EDIT-103 to efficiently remove the defective RHO gene responsible for RHO-adRP while replacing it with an RHO gene capable of producing sufficient levels of RHO to preserve photoreceptor structure and functions. The program is progressing towards the clinic, said Mark S. Shearman, Ph.D., Executive Vice President and Chief Scientific Officer, Editas Medicine. EDIT-103 uses a dual AAV gene editing approach, and also provides initial proof of concept for the treatment of other autosomal dominant disease indications where a gain of negative function needs to be corrected.

Key findings include:

Full details of the Editas Medicine presentations can be accessed in the Posters & Presentations section on the Companys website.

About EDIT-103EDIT-103 is a CRISPR/Cas9-based experimental medicine in preclinical development for the treatment of rhodopsin-associated autosomal dominant retinitis pigmentosa (RHO-adRP), a progressive form of retinal degeneration. EDIT-103 is administered via subretinal injection and uses two adeno-associated virus (AAV) vectors to knockout and replace mutations in the rhodopsin gene to preserve photoreceptor function. This approach can potentially address more than 150 gene mutations that cause RHO-adRP.

AboutEditas MedicineAs a leading genome editing company, Editas Medicine is focused on translating the power and potential of the CRISPR/Cas9 and CRISPR/Cas12a genome editing systems into a robust pipeline of treatments for people living with serious diseases around the world. Editas Medicine aims to discover, develop, manufacture, and commercialize transformative, durable, precision genomic medicines for a broad class of diseases. Editas Medicine is the exclusive licensee of Harvard and Broad Institutes Cas9 patent estates and Broad Institutes Cas12a patent estate for human medicines. For the latest information and scientific presentations, please visit http://www.editasmedicine.com.

Forward-Looking StatementsThis press release contains forward-looking statements and information within the meaning of The Private Securities Litigation Reform Act of 1995. The words "anticipate," "believe," "continue," "could," "estimate," "expect," "intend," "may," "plan," "potential," "predict," "project," "target," "should," "would," and similar expressions are intended to identify forward-looking statements, although not all forward-looking statements contain these identifying words. The Company may not actually achieve the plans, intentions, or expectations disclosed in these forward-looking statements, and you should not place undue reliance on these forward-looking statements. Actual results or events could differ materially from the plans, intentions and expectations disclosed in these forward-looking statements as a result of various factors, including: uncertainties inherent in the initiation and completion of preclinical studies and clinical trials and clinical development of the Companys product candidates; availability and timing of results from preclinical studies and clinical trials; whether interim results from a clinical trial will be predictive of the final results of the trial or the results of future trials; expectations for regulatory approvals to conduct trials or to market products and availability of funding sufficient for the Companys foreseeable and unforeseeable operating expenses and capital expenditure requirements. These and other risks are described in greater detail under the caption Risk Factors included in the Companys most recent Annual Report on Form 10-K, which is on file with theSecurities and Exchange Commission, as updated by the Companys subsequent filings with theSecurities and Exchange Commission, and in other filings that the Company may make with theSecurities and Exchange Commissionin the future. Any forward-looking statements contained in this press release speak only as of the date hereof, and the Company expressly disclaims any obligation to update any forward-looking statements, whether because of new information, future events or otherwise.

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Editas Medicine Presents Preclinical Data on EDIT-103 for Rhodopsin-associated Autosomal Dominant Retinitis Pigmentosa at the European Society of Gene...

Gene editing has long been primarily used for research, treatment, and disease prevention. Currently, this technology is increasingly being applied to modify agricultural products to create more perfect species. More and more genetically edited foods are appearing on the market, including high-nutrient tomatoes and zero-trans-fat soybean oil.

Some argue that gene-edited foods are safer than genetically modified (GM) foods (pdf). The U.S. Department of Agriculture (USDA) specified in 2018 that most genetically edited foods do not need to be regulated. However, are these foods, which will increasingly appear on the table, really risk-free?

In September 2021, the first gene-edited foodSicilian Rouge tomatoesmade with CRISPR-Cas9 technology were officially on sale.

This gene-edited tomato contains high levels of gamma-aminobutyric acid (GABA), which helps lower blood pressure and aids relaxation.

Japanese researchers remove a gene from the genome of the common tomato. After the gene is removed, the activity of an enzyme in tomatoes increases, promoting the production of GABA. The GABA content in this tomato is four to five times higher than that of a regular tomato.

Warren H. J. Kuo, an emeritus professor of the Department of Agronomy at National Taiwan University, explains that both gene editing and transgenic organisms are genetic modification, also known as genetic engineering.

The earliest technique was genetic modification, that is, transgenicin which a plant or animal is being inserted a gene from another species, such as a specific bacterial gene. The purpose of artificially modifying plants and animals is to improve their resistance against diseases and droughts, promote growth rates, increase yields, or improve nutrient content. However, the finished product will exhibit the foreign species genes.

Kuo says that transgenic modification is genetic modification 1.0, while gene editing is genetic modification 2.0. Gene editing is directly modifies the genes of the organism itself, so most of them do not exhibit foreign genes. However, the most common gene editing technique, CRISPR-Cas9, introduces foreign genes as the editing tool, and then removes the transplanted foreign genes.

While gene-edited tomatoes were on the market, Japan also approved two types of fish genetically edited with CRISPRtiger pufferfish and red seabream. These fish are genetically edited to accelerate muscle growth. Among them, the gene-edited tiger pufferfish weighs nearly twice that of the ordinary species.

Back in 2019, the United States had used another earlier gene-editing technique to create soybean oil with zero trans fat and introduced it into the market.

Gene-edited foods which have also been approved for sale worldwide by now include soybeans, corn, mushrooms, canola, and rice.

The number of genetically edited foods on the market is likely to increase. Patent applications relating to CRISPR-edited commercial agricultural products have skyrocketed since the 2014/2015 period.

Proponents ofgenetic modification believe this is a method to perfect agricultural produce and solve problems such as pests, droughts, and nutritional deficiencies. But the technology is still a double-edged sword.

Genetic engineering indeed has its benefits in the short term, but it may bring long-term pitfalls, said Joe Wang, molecular biologist. Wang is currently a columnist with The Epoch Times.

Hornless cattle were once the celebrity of the animal kingdom, appearing in news stories one after another.

Many breeds of dairy cattle have horns, but they are dehorned to prevent them from harming humans and other animals, and to save more feeding trough space. To solve the problem of horns, the gene editing company Recombinetics successfully produced hornless cattle with gene-editing techniques many years ago.

The company simply added a few letters of DNA to the genome of ordinary cattle and their offspring didnt grow horns, either.

However, a few years later, an accident happened.

The FDA found that a modified genetic sequence of a bull contained a stretch of bacterial DNA including a gene conferring antibiotic resistance, which has been one of the global health crises in recent years. Scientists arent clear whether this gene in gene-edited cattle will pose a greater risk than expected or not, and the FDA has stressed that its hazard-free. However, John Heritage, a retired microbiologist from Leeds University, told MIT Technology Review that the antibiotic resistance gene could be absorbed by gut bacteria in cattle and could create unpredictable opportunities for its spread.

In fact, this is one of the currently perceived risks of genetically edited foods.

The problem with unexpected accidents in the genetic modification process occurs in GM foods because transgenic techniques cannot control where the foreign gene is embedded in the chromosome.

Kuo used the example of a study that compared the protein of transgenic soybeans and non-transgenic soybeans. These transgenic soybeans were initially embedded with one foreign gene, and should have had only one protein that didnt exist before. However, the comparison showed that there was a difference of about 40 proteins between the two: Half of the proteins were originally present, but disappeared after transgenic modification; the other half were not present but were added after the transgenic modification.

In contrast, emerging gene editing techniques allow for more precise modification of specific genes (pdf). Its like a tailor modifying a section of a zipper by cutting off a specific segment and replacing it with a new one. However, there may be mistakes and unexpected changes in the process of cutting and repairing, and another similar section of the zipper may also be cut off.

Kuo says that this process may have unforeseen side effects; for example, if during this, new allergy-causing proteins or new toxins are produced.

The genetic engineering procedure, and this includes gene editing, has the potential to damage DNA, said molecular geneticist Dr. Michael Antoniou, head of the Gene Expression and Therapy Group at Kings College London, in an interview in April 2022. If you alter gene function, you automatically alter the biochemistry of the plant included within that altered biochemistry can be the production of novel toxins and allergens that is my main concern.

Another major concern with GM foods is herbicide residue.

Most crops, whether genetically edited or genetically modified, have herbicide-resistant genes incorporated into them. This is done so that when herbicides are applied to crops for weed control, the crops themselves wont be harmed.

When planting herbicide-resistant crops, farmers can use herbicides rather liberally. But, long term, the weeds the farmers are targeting become increasingly herbicide-resistant as well, resulting in a cycle of increased herbicide use and resistance.

Since the introduction of herbicide-resistant GM crops in 1996, herbicides have experienced a significant growth in application every year. The herbicides residue in the crops grown are increasing as well.

One of the most widely used herbicides is glyphosate under the trade name Roundup. The International Agency for Research on Cancer (IARC) classifies glyphosate as a Group 2A carcinogen that is probably carcinogenic to humans.

Massachusetts Institute of Technology (MIT) researcher Stephanie Sene and scientific consultant Anthony Samsel said in their study that 80 percent of GM crops, especially corn, soybeans, canola, cotton, sugar beets, and alfalfa, are specifically introduced with glyphosate resistance genes.

In addition to carcinogenic concerns, glyphosate may have more harmful effects. They have collected and reviewed 286 studies and indicated that glyphosate inhibits the activity of an enzyme in the mitochondria of liver cellscytochrome P450which has the ability to detoxify and decompose foreign toxic substances. Moreover, glyphosate also has adverse effects on the gut microbiota.

These effects are not immediately apparent, but in the long run may contribute to inflammatory bowel disease, obesity, depression, attention deficit hyperactivity disorder (ADHD), autism, Alzheimers, Parkinsons, amyotrophic lateral sclerosis (ALS), multiple sclerosis, cancer, infertility, and developmental abnormalities.

An animal study published in Environmental Health shows that long-term exposure to ultra-low doses of glyphosate still causes liver and kidney diseases in rats.

The debate over whether GM food is safe or not has not yet settled. Many advocates of transgenic modification and gene editing believe that people have been eating GM crops for 20-plus years and still there is no evidence that they have caused problems to human health. Other argue they contribute to long term harm that is still being measured.

Kuo said that GM food is not a highly toxic drug causing immediate problems. Health problems can be the result of something cumulative, and hard to relate back to a single food cause. Whether GM foods are the culprit of such health problems has not been proven, nor ruled out.

At present, various countries have adopted an early warning principle for GM foods, stipulating that merchants label their products. It is the consumers decision to purchase them or not.

Will gene-edited food require specific labeling? Some argue that because these foods do not exhibit foreign genes, there should not be such regulation. Kuo believes this is a misleading argument, given that the tool used to edit the original genes were in fact foreign genes, and the method carries the risk that these foreign genes may not be completely removed.

Currently, the regulations for gene-edited foods in various countries are much looser than those for GM foods.

The USDA has consistently stated that gene-edited agricultural products are not regulated. Plant technologists are usually given the green light within months after submitting inquiries to the agency, allowing them to grow gene-edited foods without oversight.

In addition to the United States, Brazil and Australia and other countries have also adopted similar regulatory approaches. European regulations are still more stringent.

Antoniou argues that since these GM agricultural products are not monitored, the unexpected genes that they carry are released into the environment and will cause harm to it. They may also cause harm to the public due to the scientific communitys insufficient understanding of their risks.

Wang said that scientists who support gene editing believe that what they are doing now will also happen in nature, albeit at a slower pace. They simply speed it up. However, humans are not gods and cannot control everything. When humans do such things, the odds of mistakes and danger are definitely higher than what happens naturally, Wang said.

We humans have violated the laws of nature for a long time, Kuo said.

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Camille Su is a health reporter covering disease, nutrition, and investigative topics. Have a tip? kuanmi.su@epochtimes.com

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More Foods Will Be Gene-Edited Than You Think - The Epoch Times

There are over 6 billion letters, or nucleotides, of DNA in the genome. These contain all the information needed to create an individual organism. Certain sequences of DNA, called genes, contain instructions for making proteins that determine everything about how we look and how we function. We expect there to be some differences in those sequences that lead to differences in individual humans, but sometimes these instructions have significant mutations, or changes, that can lead to serious diseases such as cancer.

Imagine having to figure out which changes in which sequences in that long string of 6 billion letters are important for targeting treatments for diseases. And, once you identify some of those important genes, how do you fix those mutations?

A game-changing discovery in 2012 of a system called CRISPR has triggered a revolution in biomedical breakthroughs over the last decade. Scientists can use it to target, edit, modify and regulate genes and put any enzyme or protein they want at any location in the genome. This allows them to find new treatment targets and understand how different genes affect cells in a way that was previously impossible.

But how do we apply CRISPR to understanding cancer? We spoke with Traver Hart, Ph.D., associate professor in Bioinformatics and Computational Biology, to learn more about CRISPR and how it could be used to advance cancer treatment.

What is CRISPR?

CRISPR stands for Clustered Regularly Interspaced Short Palindromic Repeats. Thats a mouthful, so scientists refer to it as simply CRISPR. These are repeated sequences in the genetic code that were first found in bacteria and were later found to be part of a novel bacterial adaptive immune system against phages, which are viruses that attack bacteria.

This system combines the CRISPR DNA sequences and a set of Cas (CRISPR associated) proteins to identify and destroy invading viral DNA. It also embeds a sample of that viral DNA between these CRISPR sequences so that it can easily recognize and attack the same virus in the future. Thanks to this unexpected discovery in E. coli bacteria, scientists can now harness this method and use it in a similar way within human cells.

How does CRISPR work?

The main part of the CRISPR system is the Cas endonuclease, the Cas protein that cuts DNA strands. These Cas proteins can be programmed to find a 17- to 24-letter sequence by attaching a guide RNA that uniquely matches the specific DNA target. Its similar to a key matching a lock. Researchers have a large library of guide RNAs available that can match certain parts of different genes in the human genome.

Once CRISPR is added to a cell, it searches for and binds to that matching target sequence in the DNA, and the attached Cas protein gets activated to do what scientists have asked it to do. Some Cas proteins such as Cas9 can cut or break the DNA. This is the original protein that was found in bacteria. Others have been engineered to turn a gene on or off without having to cut it. This allows researchers to find out more about what happens if cells make too much (upregulation) or too little (downregulation) of a certain protein and how that can affect the outcome of a cell.

How do we use CRISPR to study cancer in human cells?

For the past several decades, studies have been done in yeast cells and other model organisms where scientists can efficiently edit the genome. The discovery of CRISPR has been instrumental in changing that.

We can edit the genome directly in human cells with unprecedented ease thanks to CRISPR.

Once CRISPR cuts the target DNA, it gets repaired or replaced with a different sequence. Scientists use this method to knock out human genes in cancer cells and identify which of those genes are essential for the growth of tumor cells without harming normal cells. This allows us to nominate gene candidates for drug targets that can be very highly tumor-specific. My lab is trying to find better ways to kill tumor cells by disabling multiple genes at a time using a different Cas protein called Cas12a. This gives us more insight into how different genes and proteins work together in tumor cells to promote cancer progression.

A recent study by Yohei Yoshihama, Ph.D., and Ronald Depinho, M.D., used CRISPR to screen cancer cells and identify a protein called JMJD1C as a candidate target in castration-resistant prostate cancer. Another study by Chao Wang, Ph.D. and Junjie Chen, Ph.D., used CRISPR to screen human cancer cells growing in mouse models and discovered a protein named KIRREL, which was shown to be important for tumor suppression.

Can CRISPR fix genes in people?

While the idea of being able to fix bad genes to cure diseases is a worthy pursuit, science isnt at the point to be able to safely and effectively do so yet. Researchers are looking at how to use CRISPR to correct the genetic defects that cause beta-thalassemia and sickle cell anemia, diseases that affect the amount of hemoglobin in the body and cause patients to require constant blood transfusions. If approved, this type of therapy, called exa-cell, would become the first CRISPR-based medical treatment, which is incredibly exciting.

Whats next for CRISPR?

The possibilities are endless for the information that can be gained from using CRISPR systems and, just 10 years in, scientists have only scratched the surface. Newer Cas proteins and other enzymes are being studied, and there are still questions about how to make CRISPR more specific so that it doesnt accidentally have unintended targets.

Here at MD Anderson, our use of CRISPR continues to lead to a better understanding of how cancer cells function and helps uncover many ways to target individual treatments specific to certain tumors that will, hopefully, one day, achieve our goal to end cancer.

Request an appointment at MD Anderson online or by calling 1-877-632-6789.

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What is CRISPR? - MD Anderson Cancer Center

1. CRISPR infusion eliminates swelling in those with rare genetic disease  Science
2. Intellia Investors Are Dumping Stock on News of Crispr Gene-Editing Success  Barron's
3. Intellia's gene editing therapies both post early successes as evidence grows for CRISPR potential  FierceBiotech
4. Intellia says CRISPR treatment safely corrects DNA of six patients with rare disease  STAT
5. Intellia offers first look at CRISPR drug for rare swelling disorder  BioPharma Dive
6. View Full Coverage on Google News

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CRISPR infusion eliminates swelling in those with rare genetic disease - Science

The move comes at a crucial time for the nearly nine-year-old company, which is on the cusp of asking regulators to approve its first therapy for two genetic blood conditions. Crispr is a pioneer in developing therapies based on the experimental CRISPR gene editing technology, which can make precise changes to DNA. The company, which is valued at more than $5.5 billion, also has earlier-stage programs in treatments for cancer, cardiovascular diseases, diabetes, and neurological conditions.

As were growing and getting more programs into the clinic and ultimately to commercialization, we need more space, said Kulkarni, who has led the company since 2017.

Crisprs new building sits near the southwestern edge of Bostons Seaport, a neighborhood that is a burgeoning new hub for biotech companies looking for an alternative to the increasingly expensive and crowded Kendall Square.

Traditionally, there was a dogma that if youre a biotech company you needed to be in Cambridge, Kulkarni said. But the growing number of biotech firms based there, along with big pharma firms such as AstraZeneca and Bayer opening labs in Cambridge, has made affordable lab space harder to come by.

Only 2.2 percent of the roughly 13 million square feet of lab space in Cambridge was vacant in the second quarter of this year, compared with 13 percent of the nearly 3.8 million square feet of lab space in Boston, according to a market report from the real estate firm Newmark. The average asking price for renting lab space in Boston was about $9 cheaper per square foot as well $105 in Boston versus $114 in Cambridge.

Several drug companies have announced plans this year for new labs in the Seaport, including Boston-based Vertex Pharmaceuticals, the French firm Servier Pharmaceuticals, and the Indianapolis-based Eli Lilly & Co.

The Seaport is really emerging as a vibrant area, Kulkarni said.

A recent report from the Massachusetts Biotechnology Council showed that a slight majority of venture capital money raised by biotech startups in the first half of this year went to firms based outside of Cambridge a reversal from the year prior. Nearly 30 percent of those funds more than $1.5 billion went to startups in Boston.

The number of biotech firms planting roots south of the Charles River is likely to continue. According to Newmark, theres 4.3 million square feet of lab space under construction in Boston largely in the Seaport compared to 2 million square feet under construction in Cambridge.

Kulkarni would not disclose specifics on how much the new building cost, but he said it is more cost-efficient to be in the Seaport. The company spent $241.4 million on research and development expenses in the first half of this year, a 58 percent increase from the first half of 2021, according to the firms quarterly financial statements. Some of these expenses were related to the new building.

The building includes a publicly accessible ground floor with a caf, meeting space, and a thruway to West Second Street. The building was built by Breakthrough Properties, through a joint venture with Tishman Speyer and Bellco Capital.

The firms corporate headquarters will remain in Switzerland, where it was founded. We continue to do business development deals out of there. Its where we started and weve maintained our roots there, Kulkarni said. The firm also has employees at a manufacturing facility in Framingham.

Crisprs most advanced program is a one-time treatment for the genetic blood conditions sickle cell disease and beta-thalassemia. The experimental approach is boosting levels of the crucial blood protein hemoglobin in these patients, and the results from clinical trials look promising so far. Thanks to the genetic fix, people with sickle cell disease are experiencing fewer pain crises and people with beta-thalassemia are requiring fewer blood transfusions to stay healthy.

Crispr plans to submit the therapy, which it is developing with Boston-based Vertex Pharmaceuticals, to European regulators by the end of the year, and potentially to the FDA around the same time or soon after. If approved, it will likely become the first commercial CRISPR therapy.

If someone had said 10 years ago that this new technology is going to be an approved medicine in 10 years, people would have laughed you out of the room. But today its almost a reality, Kulkarni said. And it could be a curative medicine for patients.

Ryan Cross can be reached at ryan.cross@globe.com. Follow him on Twitter @RLCscienceboss.

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Crispr Therapeutics becomes the latest biotech to open in the Seaport - The Boston Globe

BOSTON--(BUSINESS WIRE)--Carver Biosciences, Inc., a biotech company focused on the development of CRISPR/Cas13 antivirals, today announced the closure of a seed round of funding, led by Khosla Ventures.

Carver is focused on using the bacterially derived RNA-directed RNase Cas13 to target respiratory viruses.

The company was founded in 2021 by Dr. Cameron Myhrvold, Assistant Professor at Princeton University. Dr. Myhrvold has worked on developing Cas13-based technologies for studying viral host RNAs since 2016, the year the protein was first discovered. Dr. Walter Strapps joined as co-founder and CEO in February of 2022. Dr. Myhrvold will serve as the chair of Carvers Scientific Advisory Board.

Cas13 opens up a whole new world of possibilities for targeting RNA, much like Cas9 did for targeting DNA," said Dr. Myhrvold. Im particularly excited about using Cas13 to treat viral infections, as the vast majority of viruses dont have FDA-approved therapies or vaccines.

Dr. Strapps is an experienced biotech executive, most recently serving as Chief Scientific Officer at Gemini Therapeutics, a dry age related macular degeneration-focused biotechnology company. Prior to that Dr. Strapps has held various roles leading research efforts in oligonucleotide-based therapeutics, including at Sirna Therapeutics, Merck & Co., Inc and Intellia Therapeutics.

Im very excited by the potential of Cas13 to treat patients infected with respiratory viruses. Cas13 provides us with a programmable platform to treat various existing viruses and to rapidly respond to new viruses as they occur, said Dr. Strapps. Having helped build several therapeutic oligo platforms before, Im thrilled to bring that experience to a new CRISPR protein.

We invest in companies early that are bold and have the ability to make a large impact on society, said Alex Morgan, partner at Khosla Ventures. The major respiratory viruses with great impact on human health are RNA based and include respiratory syncytial virus, influenza virus, parainfluenza virus, metapneumovirus, rhinovirus, and something everyone can appreciate today, coronavirus. The ability to have a programmable therapy that can respond to newly emergent strains in a highly targeted way would revolutionize our approach to infectious disease, from the common cold to some of the major infectious causes of mortality. This team has a unique set of expertise that can hopefully unlock this potential.

Carver will use the seed funding to conduct initial proof of concept Cas13 experiments in cells and in models of disease at their laboratories in Boston and with CRO partners. Cas13 acts by using sequence specific targeting of mRNA to cleave and specifically degrade the targeted RNA. Similar to Cas9 which targets DNA, Cas13 is part of a bacterial immune system that can be used in mammalian cells.

About Carver Biosciences, Inc.

Carver is a Boston, MA-based gene therapy company developing CRISPR-based therapies for RNA viruses that infect humans. Carvers approach is a programmable, platform technology that could offer a generalizable solution for treating many viruses, as targeting is dictated by a guide RNA sequence. The companys underlying technology was developed by co-founder Dr. Cameron Myhrvold. For additional information, please visit http://www.carver.bio.

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Carver Biosciences Launches with Seed Financing to Advance CRISPR/Cas13-based Technologies - Business Wire

Newswise Montana BioAgriculture Inc. is to develop commercial product for wheat stem sawfly:it is now time to CRISPR edit and play with wheat stem sawfly gene and regulatory elements to develop a product for farmers said Dr. Hikmet Budak, Chief Science Officer.

Newswise Insects, diseases, and abiotic stressors cause losses of millions of tons of wheat and cost farmers $100s of millions each year. Solutions can come from understanding and working with interactions between naturally occurring fungi and plants in the common agroecology of US and Canadian wheat growing areas. MBA are working to develop biological solutions to cross border problems affecting North American farmers and global food security.

The wheat stem sawfly, WSS, is the number one insect pest of wheat in North America. There are no effective insecticides. The larvae which cause the damage live their life cycle protected inside the wheat stem. Solid stem wheat varieties can limit damage however lower yields than conventional varieties. WSS apparently first adapted from native grasses to infest spring wheat in North Central Montana spreading into North Dakota and into Canadian prairie provinces. WSS has now adapted to winter wheat, expanding east and south in the US. Our recent forum (International Virtual Wheat Stem Sawfly Forum:Farmer'sVoiceshttps://www.linkedin.com/feed/update/urn:li:activity:6906012389427290112/v) in June, 2022 declared that biopesticide and other products are necessary to control WSS.

Montana BioAgriculture (MT BioAg), has been working on WSS using interdisciplinary tools. It has received two large grants from the US Department of Agriculture to (i) develop a fungus which induces tolerance to drought in wheat and barley, and (ii) use genomics tools to fight against wheat stem sawfly, a major problem in great plains. MT BioAg licensed the fungus to develop as a commercial product.These awards is a major boost to Mt BioAgs work to develop integrated, biological solutions for farmers to reduce losses in grain yield from drought, insects and diseases. Noted by Cliff Bradley MT BioAg president. One of the awards from USDA to evaluate fungi to control wheat stem sawfly and Fusarium head blight (FHB), the most important insect and disease of wheat in Montana. MBA licensed use of fugal stains originally isolated from wheat in Montana by scientist with the USDA Agricultural Research Service. These fungi are natural pathogens of the WSS and are endophytes, that is the fungi colonize the stem without causing damage to the wheat. The fungi infect and kill the WSS larvae inside the stem. Dr. Hikmet Budak, Chief Science officer at MT BioAg said, these projects will utilize new genomics tools in combination with speed breeding. This has been further helping us to CRISPR edit gene(s) to control Wheat Stem Sawfly, FHB and drought stress. With these USDA grants and funds from the Montana Department of Commerce program, MT BioAg has been working on developing production and delivery systems needed to bring these naturally occurring fungi to commercial use. This could lead to higher incomes for grain growers, better nutrition for world populations and new wheat and barley varieties. The research and new findings especially small RNAs (microRNAs, LncRNA) targeting wheat stem sawfly gene and metabolites also offer immense potential for our breeding program to create new discoveries when it comes to advancing global food security." Montana BioAg. Inc., Chief Science Officer said.

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CRISPR Editing Wheat Stem Sawfly Genes and Small RNAs. - Newswise

NEW YORK GenScript and Ireland's Avectas said on Tuesday that they're partnering to develop a non-viral cell therapy manufacturing process.

Under the terms of the agreement, their research teams will apply Avectas' Solupore technology to permeabilize target cell membranesto deliver GenScript's GenCRISPR single guide RNA, CRISPR-Cas9 protein, and GenExact single-stranded DNA homology-directed repair templates.

"By combining Avectas' cell engineering technology and know-how with GenScript's expertise in synthetic long oligo production, the partnership aims to demonstrate a novel and efficient solution for cell therapy manufacturing and to improve editing efficiency and cell viability over traditional delivery methods," the firms said in a statement.

"We expect this method will provide our customers with more complete solutions for efficient gene editing using our GenCRISPR sgRNA and ss/dsDNA HDR templates," Ray Chen, president of GenScriptUSALife Science Group, added.

GenScript, a Piscataway, New Jersey-based synthetic DNAmaker, is listed on the Hong Kong Stock Exchange. In May 2021, the firm licensedthe ERS Genomics CRISPR-Cas9 patent portfolio, based on IP held by Emmanuelle Charpentier, one of the pioneers of CRISPR genome editing.

Avectas offers Solupore, a cell engineering technology that delivers payloads to cells by diffusion across a permeabilized membrane.

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GenScript, Avectas Partner on Cell Therapy Manufacturing - GenomeWeb