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      BIG-TREE: Base-Edited Isogenic hPSC Line Generation Using a Transient Reporter for Editing Enrichment

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          Summary

          Current CRISPR-targeted single-nucleotide modifications and subsequent isogenic cell line generation in human pluripotent stem cells (hPSCs) require the introduction of deleterious double-stranded DNA breaks followed by inefficient homology-directed repair (HDR). Here, we utilize Cas9 deaminase base-editing technologies to co-target genomic loci and an episomal reporter to enable single-nucleotide genomic changes in hPSCs without HDR. Together, this method entitled base-edited isogenic hPSC line generation using a transient reporter for editing enrichment (BIG-TREE) allows for single-nucleotide editing efficiencies of >80% across multiple hPSC lines. In addition, we show that BIG-TREE allows for efficient generation of loss-of-function hPSC lines via introduction of premature stop codons. Finally, we use BIG-TREE to achieve efficient multiplex editing of hPSCs at several independent loci. This easily adoptable method will allow for the precise and efficient base editing of hPSCs for use in developmental biology, disease modeling, drug screening, and cell-based therapies.

          Highlights

          • Generation of hPSC-MSCs by stepwise and chemically defined protocol

          • Ascorbate promotes the specification and chondrogenesis of hPSC-MSCs

          • Ascorbate promotes the specification of hPS-MSCs and promotes osteochondrogenesis

          • hPSC-MSCs are able to fully repair the cartilage defects

          Abstract

          In this study, Brafman and colleagues develop a method entitled base-edited isogenic hPSC line generation using a transient reporter for editing enrichment (BIG-TREE) that allows for the highly efficient single-nucleotide modification of hPSCs without the need for introduction of deleterious double-stranded DNA breaks.

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          Most cited references9

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          Repair of double-strand breaks induced by CRISPR–Cas9 leads to large deletions and complex rearrangements

          CRISPR-Cas9 is poised to become the gene editing tool of choice in clinical contexts. Thus far, exploration of Cas9-induced genetic alterations has been limited to the immediate vicinity of the target site and distal off-target sequences, leading to the conclusion that CRISPR-Cas9 was reasonably specific. Here we report significant on-target mutagenesis, such as large deletions and more complex genomic rearrangements at the targeted sites in mouse embryonic stem cells, mouse hematopoietic progenitors and a human differentiated cell line. Using long-read sequencing and long-range PCR genotyping, we show that DNA breaks introduced by single-guide RNA/Cas9 frequently resolved into deletions extending over many kilobases. Furthermore, lesions distal to the cut site and crossover events were identified. The observed genomic damage in mitotically active cells caused by CRISPR-Cas9 editing may have pathogenic consequences.
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            Continuous evolution of base editors with expanded target compatibility and improved activity

            Base editors use DNA-modifying enzymes targeted with a catalytically impaired CRISPR protein to precisely install point mutations. Here we develop phage-assisted continuous evolution of base editors (BE-PACE) to improve their editing efficiency and target sequence compatibility. We used BE-PACE to evolve new cytosine base editors (CBEs) that overcome target sequence context constraints of canonical CBEs. One evolved CBE, evoAPOBEC1-BE4max, is up to 26-fold more efficient at editing GC, a disfavored context for wild-type APOBEC1 deaminase, while maintaining efficient editing in all other sequence contexts tested. Another evolved deaminase, evoFERNY, is 29% smaller than APOBEC1 and edits efficiently in all tested sequence contexts. We also evolved a CBE based on CDA1 deaminase with much higher editing efficiency at difficult target sites. Finally, we used data from evolved CBEs to illuminate the relationship between deaminase activity, base editing efficiency, editing window width, and byproduct formation. These findings establish a system for rapid evolution of base editors and inform their use and improvement. Genome editing has revolutionized the life sciences and entered clinical trials to treat genetic diseases. 1 The use of programmable nucleases to generate double-stranded DNA breaks (DSBs) followed by homology-directed repair can introduce a wide variety of modifications but is inefficient in non-dividing cells, and is typically accompanied by an excess of unwanted insertions and deletions (indels), translocations, or other chromosomal rearrangements. 2 Base editing directly modifies target DNA bases in living cells and has become widely used to correct or install point mutations in organisms ranging from bacteria to human embryos. 3 Base editors use a catalytically impaired Cas9 to open a single-stranded DNA loop at a specified genomic site (Fig. 1a). Bases within the editing window (typically ~5 nt wide) in this region are modified by a tethered base-modification enzyme that only accepts single-stranded DNA. Two classes of base editors have been developed to date: cytosine base editors (CBEs) convert C•G to T•A, and adenine base editors (ABEs) convert A•T to G•C. 3 CBEs such as BE3, 4 BE4, 5 and BE4max (a state-of-the-art CBE) 6 use the cytidine deaminase APOBEC1 to modify target Cs. 4,7 In cells that express them robustly, CBEs can edit some sites with high (≥50%) efficiency, 6,8 but perform poorly at others. One factor that limits the most commonly used CBEs is the native sequence context preference of APOBEC1, which deaminates GC motifs poorly. 4,9 While a GC target positioned in the center of the editing window may be edited efficiently by APOBEC1-based CBEs, 9 other GCs are not. 4,10 CBEs incorporating different cytidine deaminases can edit GC targets more efficiently; for example, a CBE based on the CDA1 deaminase 7 edited GC 3 in the HEK3 site (numbering shown in Fig. 1a) more efficiently than the corresponding APOBEC1-based CBE (20% vs. 2%). 5 Non-APOBEC1 CBE alternatives, however, showed lower average performance than APOBEC1-CBE across a variety of targets in human cells. 5 Cytosine base editing would therefore benefit from new CBE variants that edit with high efficiency regardless of target sequence context. Such CBEs would be especially useful for applications that involve challenging target sites or cell types, multiplexed base editing, or large-scale screening. 11 Moreover, a general platform to tailor base editor properties would enable the development of editors ideally suited for specific applications, which differ widely in their requirements for efficiency, sequence compatibility, and tolerance for unwanted editing. Given the complexity of CBEs, we envisioned harnessing phage-assisted continuous evolution (PACE) 12,13 to generate editors with improved target sequence context compatibility and higher activity. PACE performs dozens of generations of mutation, selection, and replication per day and has facilitated dramatic alterations of protein function. 13–24 Here we describe a PACE system for evolving base editors (BE-PACE), and its application to generate CBEs with high editing activity on both GC and non-GC targets. Results Development of a genetic circuit that responds to base editing During PACE, an activity of interest is coupled to the propagation of M13 bacteriophage that encode a biomolecule with that activity (Fig. 1b). To achieve this coupling, the desired activity—here, cytosine base editing—is linked to the expression of gene III, which is required to produce infectious progeny phage, using a genetic circuit encoded in E. coli host cells. In BE-PACE, this circuit must be activated by a single-base change, respond to small numbers of editing events, and turn on rapidly enough to support phage propagation under continuous dilution. To meet these requirements, we designed a gene circuit in which cytosine deamination occurs on the transcription template strand to revert an inactivating mutation in a protein (Fig. 2a). This design allows a transcription-level response independent of DNA replication and repair, since E. coli RNA polymerase accepts uracil-containing templates. 25 To amplify editing signal, we chose T7 RNA polymerase (T7 RNAP) as the target gene for editing and placed expression of gene III and a luciferase reporter under control of a T7 promoter. We inactivated T7 RNAP by fusing a proteolytic degradation tag (degron) to its C-terminus, 26 which targets the enzyme for degradation and disrupts its folding and catalytic mechanism (Supplementary Fig. 1). 27 The degron is linked to T7 RNAP through a TGG tryptophan codon (template strand CCA) such that deamination of either or both template strand cytidines results in a W884STOP codon substitution in mRNA that removes the degron. This design allows the bases upstream of the target to be freely varied, changing the sequence context for deamination (Fig. 2a). We implemented this circuit using two plasmids, one encoding T7 RNAP and the other encoding the gIII-luciferase operon and a guide RNA targeting the T7 RNAP C-terminus. We began with a GTCCA editing target to match the native TC context preference of APOBEC1 and to minimize selection stringency. We tested the circuit using plasmid-encoded BE2 in a luciferase reporter assay (Fig. 2b). Activation was dependent on all components of the system, including fusion of APOBEC1 to dCas9. The circuit’s signal amplification, and thus selection stringency during PACE, can be tuned by adjusting the expression of T7 RNAP. We assayed a series of ribosome binding site and promoter strengths and chose a combination that provided strong circuit activation (>500-fold) (Supplementary Fig. 2). Despite the strong activation observed with plasmid-encoded BE2, phage encoding BE2 (Supplementary Discussion 1) did not measurably activate the circuit within 3 h (Supplementary Fig. 3). A discrete overnight phage propagation assay (Fig. 2c) showed that although the circuit was competent to propagate phage and was dependent on base editing activity, phage encoding BE2 did not enrich strongly enough to support PACE, which typically requires ≥10-fold enrichment in this assay. Split base editor phage support an optimized PACE selection To achieve robust, base editing-dependent phage propagation, we increased the copy number of the gIII-containing plasmid. We also cloned BE2 into an evolved phage backbone (“generation 2”) containing 41 mutations that previously emerged from extensive PACE 14 and observed that reporter gene expression increased (Supplementary Fig. 4) and BE2 phage propagation improved by as much as 100-fold (Supplementary Table 1). However, these phage still enriched 1,000-fold selectivity for base editor phage (Supplementary Table 1). Importantly, the BE-PACE circuit can be used to assess deaminase kinetics in cells in a CBE context based on expression of the luciferase reporter. Because the circuit responds to deaminated cytosine directly via transcription, the activation rate of the circuit as measured by the rate of change of luminescence over time should reflect the rate of cytosine deamination. We tested BE-PACE using the low-stringency selection circuit (TCC1, Supplementary Table 2) in a competitive continuous propagation experiment (Fig. 3c). A small amount of APOBEC1–intein phage was seeded along with a large excess of phage encoding red fluorescent protein (RFP). The phage titer initially plunged as the non-replicating RFP phage were diluted out of the lagoon, but by 45 h, the titer of APOBEC1–intein phage recovered and no trace of RFP phage was detected by PCR. These results validate CBE activity-dependent phage propagation during BE-PACE. BE-PACE generates improved deaminases To address the sequence context limitations of APOBEC1—which strongly favors TC and disfavors GC targets 4,9,31 —we constructed a low-stringency circuit (GCC1, Supplementary Table 2) with an AGC 4 C 5A target that requires editing of a GC to support maximal phage propagation. Initial activity of APOBEC1–intein phage as measured by luciferase assay on this GC target was negligible. Therefore, we sought to first increase overall APOBEC1 activity through PACE on a series of TC 4 C 5 circuits with increasing stringency (Fig. 4a, Supplementary Discussion 2, and Supplementary Fig. 5). After successive PACE on the TCC1 and TCC2 circuits, we isolated mutant phage that showed weak but measurable activity on the GCC1 circuit. Several mutations, including predominant A165S and F205S substitutions, were similar to mutations that arose during the PACE of APOBEC1 to maximize its soluble expression. 23 Further PACE on either GCC1 or TCC3 led to top-performing phage clones that showed up to 28-fold improvements in apparent activity when tested on the GCC1 circuit by luciferase assay (Supplementary Fig. 6). The GCC1 and TCC3 populations independently evolved H122L and D124N. To evaluate the contributions of these and other mutations to GCC editing activity we cloned evolved deaminases into a standardized phage backbone (“generation 3”) isolated after 210 h of PACE and tested them on circuits containing all four ANCCA targets in the luciferase assay (Fig. 4b). The results for wild-type APOBEC1 recapitulated its known sequence context preferences. 31 Results for the PACE-evolved deaminases show that the D124N and especially H122L mutations dramatically improve activity on non-TC targets, reducing the TCC/GCC activity ratio from 27-fold in wild-type APOBEC1 to between 1.9 and 0.9 in evolved clones. Ancestral sequence reconstruction of deaminases 6 could provide promising starting points for BE-PACE. We chose five ancestral sequences from nodes within our APOBEC phylogeny 6 (Supplementary Fig. 7) and constructed corresponding deaminase–intein phage. We excluded from these sequences the N- and C-termini of rat APOBEC1, which have low sequence similarity to other APOBEC deaminases and are implicated in functions irrelevant to base editing. 32,33 After subjecting a mixture of all five genotypes to BE-PACE on the TCC1 circuit, (Fig. 4a and Supplementary Fig. 5) we obtained improved phage clones from the ancestral node 656 sequence, which we named “FERNY” for its five N-terminal amino acids (Supplementary Fig. 7). We further evolved FERNY–intein phage on the TCC3 circuit (Fig. 4a), resulting in H102P and D104N mutations at positions corresponding to H122 and D124 in APOBEC1. These mutations substantially improved apparent activity on the GCC target (Fig. 4b), further implicating these positions as sequence compatibility determinants. 34 Wild-type CDA1–intein phage showed much higher starting activity on both TCC and GCC targets in the luciferase assay than APOBEC1–intein phage (Fig. 4b), in agreement with yeast mutational assays. 35 We sought to improve CDA1 activity further through three stages of BE-PACE with increasing stringency (Fig. 4a and Supplementary Fig. 5), resulting in conserved mutations including A123V. All tested variants exhibited increased apparent activity by luciferase assay (Fig. 4b). Evolved deaminases improve base editing in mammalian cells To determine whether the apparent activity improvements in the luciferase assay translated into improved mammalian cell base editing, we subcloned a panel of evolved deaminase variants into the BE4max architecture 6 and transfected them into HEK293T cells along with guide RNAs targeting five genomic sites previously shown to undergo efficient editing. Under optimal plasmid dosing and conditions (Fig. 4b, Supplementary Figs. 8–10, and Supplementary Table 3), we observed that editing efficiency at the center of the activity window reaches a maximum of ~60–80% (“plateau levels”), likely limited by CBE-independent factors such as transfection efficiency or cellular DNA repair processes (Discussion). Notably, editing at positions away from the center of the activity window was improved for all evolved BEs, and editing values at these positions correlated with luciferase assay activity (Supplementary Fig. 11). Among APOBEC1 CBEs, evolved mutations including H122L and D124N resulted in a striking improvement in base editing of GC targets. For example, editing of GC 3 at the HEK3 site rose from very low (2.3±0.42%) to plateau levels (58±3.7%), a 25-fold improvement; editing of GC 3 at HEK4 increased from 5.0±0.41% to 64±5.2%; and editing of GC 8 at HEK4 increased from 12±2.1% to 58±5.6%. The unevolved FERNY-CBE exhibited higher activity on GC targets than APOBEC1-CBE (e.g. 20% at HEK3 GC 3, Fig. 4b), and the H102P D104N double mutant CBE achieved plateau levels at GC 3 (70±4.8%, Fig. 4b). In the BE4max context, the wild-type CDA1-CBE showed a much wider editing window than was reported for Target-AID 7 (which places CDA1 C-terminal to Cas9), resulting in plateau levels of editing (63±9.2%) across protospacer positions 3–8 (Fig. 4b). Evolved CDA1 CBEs showed further broadening of the editing window to include positions such as HEK3 AC 9, EMX1 GC 10, HEK2 GC 11, and RNF2 TC 12. Based on these results, we selected one high-performing evolved variant of each deaminase to characterize in depth: evoAPOBEC1 (clone 330–1), evoFERNY (164–1), and evoCDA1 (184–1). We created fully codon-optimized 6 BE4max variants and tested their editing activity using optimal dosing levels (Fig. 5a) and in a dose titration experiment (Supplementary Fig. 12). We also characterized a panel of 24 CBE variants to dissect the roles of evolved mutations, confirming the role of H122 and D124 mutations in GC activity of the evolved APOBEC-family CBEs, and finding additive improvements from the three mutations in evoCDA1-BE4max (Supplementary Discussion 3 and Supplementary Figs. 13–16). Interestingly, the critical H122L D124N mutations evolved during APOBEC1 PACE are present in 40% editing at off-target sites for which BE4max editing was undetectable (Fig. 5c and Supplementary Fig. 20). In agreement with a previous report, 9 we found that A3A-BE4max also exhibits high off-target editing activity (Supplementary Fig. 20). These observations support the expectation that higher activity deaminases lead to higher off-target editing. We examined indel formation by CBEs at all tested sites. We observed divergent behavior between the well-edited sites in HEK293T cells and the poorly edited sites in other cell types. At many, but not all, well-edited sites, evolved CBEs generated higher levels of indels compared to starting CBEs (Supplementary Fig. 21). BE4max generated 2.6±0.52% indels across all five sites, while evoAPOBEC1-BE4max and evoFERNY-BE4max generated between 2.5 and 15% indels. While CDA1-BE4max generated 0.8 to 8.7% indels, evoCDA1-BE4max generated 6.8 to 20% indels, and A3A-BE4max showed similar levels to evoCDA1-BE4max (Supplementary Fig. 21). By contrast, at the TMC1, APOE, and WFS1 disease-relevant sites, indels were much lower ( 1010 pfu/mL) in a clear-bottom black-walled 96-well assay plate (Costar). A biological replicate constituted a host cell culture grown independently from a colony or −80 °C stock mixed with a single clonal phage stock for each phage genotype tested. For plasmid-expressed base editor assays, cultures were induced with arabinose (10 mM unless otherwise indicated) and grown for a further 3h, then transferred to assay plates (150–200 μL per well). Absorbance at 600 nm (OD600) and luminescence (integration time 500 ms, no attentuation) were monitored using an Infinite M1000 Pro microplate reader (Tecan) with temperature set to 37 °C. For kinetic assays, readings were made every 3.5 minutes during the monitoring period and the plate was shaken for 20 s between reads (double orbital, 168 r.p.m.). OD600-normalized luminescence values were obtained by dividing raw luminescence by background-subtracted 600 nm absorbance. The background value was set to the 600 nm absorbance of wells containing DRM only. For phage-based CBE assays, the slope of OD600-normalized luminescence vs. time (s) was calculated by least squares linear regression over a span of at least eight time points from between 2.25 and 3 h post-infection, with the range of time points chosen such that regression gave a Pearson correlation coefficient R2 of >0.9 for the majority of conditions having slopes of at least 0.5 luminescence units OD600 −1 s−1. A single time range was used to calculate slopes across all conditions within a single assay. For Fig. 3b, data for each of the four sets of host cell biological replicates were collected on the same day. For Fig. 4b (top panel), data for each of the three sets of host cell biological replicates were collected on different days. Plaquing. Phage were plaqued on S206020 host cells containing plasmid pJC175e (activity-independent propagation), 13 plasmids pJC175e + pDB01615 (for negative selection against T7 RNAP activity; this plaquing strain was used routinely for plaquing samples from PACE experiments), plasmid pT7-AP 13 (to check for the presence of T7 RNAP recombinants), or no plasmid (to check for the presence of gene III recombinant phage). To prepare a cell stock for plaquing, overnight culture of host cells (fresh or stored at 4 °C for up to ~1 week) was diluted 50-fold in DRM containing appropriate antibiotic(s) and grown for 3–5 h at 37 °C, then stored at 4 °C for up to ~2 weeks. Serial dilutions of phage (10-fold) were made in LB medium or water. To prepare plates, molten 2xYT medium agar (1.5% agar, 55 °C) was mixed with Bluo-gal (10% w/v in DMSO) to a final concentration of 0.04% Bluo-gal. The molten agar mixture was pipetted into quadrants of quarted Petri dishes (1.5 mL per quadrant) or wells of a 12-well plate (~1.2 mL per well) and allowed to set. To prepare top agar, DRM (30 mL) was warmed by microwave heating for 15 s and 15 mL of molten (55 °C) 2xYT medium agar (1.5%) was added to give 45 mL top agar (0.5% agar final). Top agar was maintained tightly capped at 55 °C for up to 1 week. To plaque, cell stock (75–100 μL) and phage (10 μL) were mixed in 2 mL library tubes (VWR), and 55° C top agar added (400 or 900 μL for 12-well plate or Petri dish, respectively), then the mixture was immediately pipetted (without mixing) onto the solid agar medium in one well of a 12-well plate or one quadrant of a quartered Petri dish. Top agar was allowed to set undisturbed (10 min at room temperature), then plates or dishes were incubated (without inverting) at 37 °C overnight inside an unsealed plastic bag (to prevent desiccation). Phage propagation assays (Fig. 2c and Supplementary Table 1). Host cells in DRM were prepared as described above for luciferase assays and grown for ~1.5 h following dilution. Cells were diluted 2-fold with previously titered phage stocks to a final concentration of 106 plaque forming units per mL and a volume of 1 mL and grown overnight in a 96-well deep well plate (Eppendorf) fitted with porous sealing film. The cultures were centrifuged (3,600 g, 10 minutes) to remove cells and the supernatants titered as previously described. 14 Fold enrichment was calculated by dividing the titer of phage propagated on host cells by the titer of phage at the same input concentration shaken overnight in DRM without host cells. Phage-assisted continuous evolution (Fig. 4a and Supplementary Fig. 5). Unless otherwise noted, PACE apparatus, including lagoons, chemostats, pumps and media, were prepared and used as previously described 14 . S2060 host cells containing the appropriate plasmids were freshly transformed with MP6 or DP6 48 and plated on 2xYT medium with 1.5% agar supplemented with 0.5% (w/v) glucose and appropriate antibiotics. To verify the function of the mutagenesis plasmid, single colonies were resuspended in 50 μL DRM and serially diluted 10-fold; 1 μL of each dilution was plated on 2xYT medium with 1.5% agar containing either 0.5% (w/v) glucose or 10 mM arabinose and incubated overnight at 37 °C. The same colony stock dilutions (105-108 fold) were added to DRM containing antibiotics (5 mL) in 13 mL tubes and grown overnight at 37 °C with shaking. All colony stocks routinely showed robust growth on glucose-supplemented plates and zero growth (regardless of dilution level) on arabinose-supplemented plates, indicating uniform induction of mutagenesis. The entire overnight culture with the lowest visible cell density was used to inoculate a chemostat (80 mL), which was grown to OD600 ~0.6–1 then maintained under continuous dilution with fresh DRM at 1–1.5 volumes/h to keep cell density roughly constant. Lagoons were initially filled with DRM, then continuously diluted with chemostat culture for at least 2 h prior to seeding with phage. In the APOBEC1 stage 1 and 2 PACE experiments, stock solution of arabinose (1 M) was pumped directly into lagoons (10 mM final) as previously described, 14,23 with or without the appropriate concentration of aTc present in the stock solution to give the indicated final concentrations. Syringes containing aTc solution were covered in aluminum foil and work was conducted so as to minimize light exposure of tubing and lagoons. In all other PACE experiments, chemostat culture being pumped to lagoons was mixed using a Y junction with 1 M arabinose at a flow ratio of 100 volumes/h culture:1 volume/h arabinose, giving a ~10 mM final concentration of arabinose. Mixing with arabinose occurred before tubing was split to feed into each connected lagoon. In these experiments, aTc (800 ng/mL final) was added directly to the appropriate lagoons, either concurrently with phage seeding or at a later time-point, and was allowed to dilute over time. Lagoons were seeded at a starting titer of ~107 pfu/mL. Dilution rate was adjusted by modulating lagoon volume (5–20 mL) and/or culture inflow rate (10–20 mL/hr). Lagoons were sampled at indicated times (usually every 24 h) by removal of culture (500 μL) by syringe through the waste needle. Samples were centrifuged at 13,500 g for 2 minutes and the supernatant removed and stored at 4 °C. Titers were evaluated by plaquing on S2060 + pJC175e + pDB016 and recombinant phage titers assessed by plaquing on S2060 + AP-T7 A 13 . Phage genotypes were assessed from pool samples or single plaques by diagnostic PCR (0.5 μL phage + 14.5 μL PCR mixture) using primers BT-52F (5′-GTCGGCGCAACTATCGGTATCAAGCTG) and BT-52R2 (5′-AGTAAGCAGATAGCCGAACAAAGTTACCAGAAGGAAAC) and a two-step program (98° for 3 min, followed by 25 or 30 cycles of 98° for 10 s and 72 °C for 90 s, followed by 72° for 2 min). The resulting PCR products were subjected to 1% agarose gel electrophoresis (shown in Fig. 3c) and/or Sanger sequencing. Phage titers were determined by plaquing on pJC175e pDB16 S2060 host cells, which allow activity-independent propagation but negatively select against T7 RNAP recombinant phage. The presence of T7 RNAP or gene III recombinant phage was monitored by plaquing on S2060 cells containing pT7-AP and no plasmid. Details of PACE seed phage, aTc dosing, and workup are in Supplementary Fig. 5. HEK cell culture, transfections and genomic DNA extraction. See Figs. 4b, 5 and 6c, Supplementary Table 3 and Supplementary Figs. 8–14, 16 and 20. HEK293T culturing conditions, transfections for both single dose and titration experiments, and genomic DNA extraction were all conducted as previously described. 6 Briefly, HEK293T cells were seeded into 48-well Poly-D-Lysine-coated plates (Corning 354509) at 30,000 cells/well. 1 day after plating, cells were transfected by Lipofectamine 2000 (Thermo Fisher) with 750 ng of base editor plasmid, 250 ng of guide RNA plasmid, and 10 ng of fluorescent protein expression plasmid as a transfection control following the manufacturer’s directions. For titration experiments, base editor plasmid was replaced by equal mass of pUC to maintain the ratio of DNA to Lipofectamine 2000. Cells were cultured for 3 days before genomic DNA was extracted by replacement of culture media with 100 μL lysis buffer (10 mM Tris-HCl, pH 7.5, 0.05% SDS, 25 μg/mL proteinase K (NEB) and 37 °C incubation for 1 hour. Proteinase K was inactivated by 30-minute incubation at 80 °C. Replicates constituted independent transfections of separate splits of cells, performed on the same day or on different days. Nuclefection of baringo MEFs and ApoE4 astrocytes (Fig. 6a, 6b, Supplementary Table 3 and Supplementary Fig. 21). MEF cells were cultivated until confluent, then pooled. Replicates were performed on the same day using three separate nucleofections followed by cultivation in separate wells. Each nucleofection contained 400 ng base editor plasmid and 100 ng guide RNA plasmid. Transfection programs were optimized following manufacturer’s instructions (CZ-167, P4 Primary Cell 4D-Nucleofector® X Kit, Lonza). Cells were harvested for genomic DNA extraction after ~96 h. ApoE4-expressing astrocytes were diluted to 200,000 astrocytes per 20-μL reaction and nucleofected with 750 ng base editor plasmid and 250 ng guide RNA plasmid (program EN-150, SF Cell Line 4D Nucleofector® X Kit, Lonza). Reactions were diluted to 100 μL with pre-warmed media, and half of the resulting solution was plated in 12-well dishes. After 72h, genomic DNA was extracted with 300 μL lysis buffer. Baringo PMEF generation. Baringo females at 3–4 weeks of age were treated with single intra-peritoneal injection of 5 U each of pregnant mare’s serum gonadotropin (Prospec) followed by human chorionic gonadotropin (Sigma) after 44–45 h and paired with baringo males. The following morning, females were examined for copulatory plugs to confirm matings and marked as 0.5 dpc. At day 13.5 females were sacrificed by CO2 inhalation followed by cervical dislocation. Embryos were harvested in PBS under aseptic conditions. To harvest primary embryonic fibroblasts, each embryo was eviscerated and head was removed. The remaining parts of each embryo were minced to prepare single-cell suspensions and treated with 0.25% Trypsin-EDTA (Gibco) at 37 °C for 10 minutes, followed by centrifugation for 10 minutes. Pellets were resuspended in growth media containing DMEM, 10% FBS, penicillin-streptomycin (100 U/mL) and plated on 15-cm tissue culture plates, then incubated at 37 °C until confluent. The baringo colony is maintained ad libitum and all animal procedures are approved by the Children’s Hospital IACUC in compliance with relevant ethical regulations. Library preparation and high-throughput sequencing. DNA sequencing libraries of edited sites were prepared as previously described. 4,6,10 PCR1 amplifications for new sites (TMC1, WFS1, APOE, off-target sites) were carried out by qPCR and amplification was stopped within the exponential phase to prevent overamplification and PCR bias, but were otherwise prepared for sequencing identically to previously tested sites. Primers used are shown in Supplementary Table 6. High-throughput sequencing data analysis. Editing was quantified using CRISPResso2, 58 available at http://crispresso.pinellolab.partners.org, using the guide and amplicon sequences given in Supplementary Table 6 and with the following options: base_editor_output, quantification_window_size 28, quantification_window_center −13, plot_window_size 24. Reads were not quality filtered. Reads containing both indels as well as base edits were counted toward base editing and towards indels. Read mapping for cytosines in and near the editing window were compiled from the amplicon nucleotide percentage summary output. Insertions and deletions were quantified using CRISPResso2 quantification of editing frequency output. The number of reads with substitutions only were subtracted from the number of modified reads and the result divided by the number of total mapped reads and multiplied by 100. For quantification of conversion to desired alleles (Figure 6a–c), the percentage counts of aligned reads around the target site that matched the sequences given in Supplementary Table 6, all of which contain the targeted coding mutation with no other non-silent mutations or indels, were summed for each replicate from the CRISPResso allele table. Ancestral sequence reconstruction (Supplementary Fig. 7). We performed ancestral sequence reconstruction using our previously reported alignment and phylogenetic tree of the APOBEC protein family. 6 Briefly, 468 APOBEC protein sequences where collected from the UniProt database aligned with MAFFT and a phylogenetic tree was built from the alignment using IQ-Tree 59 and the best fitting model (JTT + F + R5). Sequences at internal nodes were inferred using the FastML package 60 given the tree, alignment and substitution model. Structure-guided alignment and homology model (Supplementary Figs. 15 and 17). A BLAST 61 search was performed with the protein sequences P41238, H2P4E7, E1BTD6, and H2P4E9 (UNIPROT) against the Protein Data Bank archive (PDB). A match is considered a suitable template if it displays a minimum of 50% sequence identity with the source sequence and a minimum coverage of 70%. These criteria were chosen to limit the selected templates to close homologues whose alignment with the source is non-ambiguous. Ten structures resulted for structure-guided alignment. These templates where used to refine the alignment of 1,189 APOBEC homologs previously described 6 using the Expresso algorithm which is part of the T-Coffee software package. 62 The resulting alignment was manually searched for the presence of the LxN motif in the recognition loop. Homology models of APOBEC1, FERNY and CDA were generated by using the I-TASSER web server. 63,64 The quality of the models predicted by I-TASSER was assigned a TM-score of 0.62 +/− 0.14, 0.85 +/− 0.08 and 0.74 +/− 0.11 respectively. Typically, a TM-score value above 0.5 implies a correct topology for the model. Supplementary Material 1 2 3 4 5 6
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              Recent advances in the CRISPR genome editing tool set

              Genome editing took a dramatic turn with the development of the clustered regularly interspaced short palindromic repeats (CRISPR)-CRISPR-associated proteins (Cas) system. The CRISPR-Cas system is functionally divided into classes 1 and 2 according to the composition of the effector genes. Class 2 consists of a single effector nuclease, and routine practice of genome editing has been achieved by the development of the Class 2 CRISPR-Cas system, which includes the type II, V, and VI CRISPR-Cas systems. Types II and V can be used for DNA editing, while type VI is employed for RNA editing. CRISPR techniques induce both qualitative and quantitative alterations in gene expression via the double-stranded breakage (DSB) repair pathway, base editing, transposase-dependent DNA integration, and gene regulation using the CRISPR-dCas or type VI CRISPR system. Despite significant technical improvements, technical challenges should be further addressed, including insufficient indel and HDR efficiency, off-target activity, the large size of Cas, PAM restrictions, and immune responses. If sophisticatedly refined, CRISPR technology will harness the process of DNA rewriting, which has potential applications in therapeutics, diagnostics, and biotechnology.
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                Author and article information

                Contributors
                Journal
                Stem Cell Reports
                Stem Cell Reports
                Stem Cell Reports
                Elsevier
                2213-6711
                30 January 2020
                11 February 2020
                30 January 2020
                : 14
                : 2
                : 184-191
                Affiliations
                [1 ]School of Biological and Health Systems Engineering, Arizona State University, 501 E. Tyler Mall, ECG 334A, Tempe, AZ 85287, USA
                [2 ]Molecular and Cellular Biology Graduate Program, Arizona State University, Tempe, AZ 85287, USA
                [3 ]Graduate Program in Clinical Translational Sciences, University of Arizona College of Medicine-Phoenix, Phoenix, AZ 85004, USA
                Author notes
                []Corresponding author xiaowang@ 123456asu.edu
                [∗∗ ]Corresponding author david.brafman@ 123456asu.edu
                [4]

                Co-first author

                Article
                S2213-6711(19)30452-7
                10.1016/j.stemcr.2019.12.013
                7013208
                32004495
                0bdf9c5b-6132-4028-879f-6066ef7ba10f
                © 2019 The Authors

                This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).

                History
                : 17 October 2019
                : 22 December 2019
                : 28 December 2019
                Categories
                Report

                crispr,genome modification,base editor,human pluripotent stem cells,multiplexing

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