Highly Efficient Genome Editing of Murine and Human Hematopoietic Progenitor Cells by CRISPR/Cas9

Abstract

Our understanding of the mechanisms that regulate hematopoietic stem/progenitor cells (HSPCs) has been advanced by the ability to genetically manipulate mice; however, germline modification is time consuming and expensive. Here, we describe fast, efficient, and cost-effective methods to directly modify the genomes of mouse and human HSPCs using the CRISPR/Cas9 system. Using plasmid and virus-free delivery of guide RNAs alone into Cas9-expressing HSPCs or Cas9-guide RNA ribonucleoprotein (RNP) complexes into wild-type cells, we have achieved extremely efficient gene disruption in primary HSPCs from mouse (>60%) and human (∼75%). These techniques enabled rapid evaluation of the functional effects of gene loss of Eed, Suz12, and DNMT3A. We also achieved homology-directed repair in primary human HSPCs (>20%). These methods will significantly expand applications for CRISPR/Cas9 technologies for studying normal and malignant hematopoiesis.

Keywords: CRISPR/Cas9; HSC; gene therapy; genome editing; hematopoietic stem cells; homology-directed repair; human CD34; progenitor; sgRNA; transplantation.

Figure 1. Gene editing in murine HSPCs

(A) A representative flow cytometry histogram showing efficient ablation of GFP by electroporating GFP-sg1 into Cas9-expressing HSPCs. Black histogram represents GFP⁻ HSPCs, and green and red histograms represents Rosa26 (R26) and GFP disrupted HSPCs, respectively (n=3). (B) Survival of HSPCs was determined by trypan blue staining of cells cultured without electroporation, cells mock electroporated without sgRNA, and cells electroporated with R26 or GFP sgRNA. 1 μg of sgRNA was used to electroporate 10⁵ cells (n=3). (C) Deletion efficiencies of GFP exhibiting sgRNA dose-dependent response. A plateau in gene editing efficiency was reached by 1 μg of sgRNA per 10⁵ cells (n=3). (D) A brief culture of murine HSPCs for 1 to 3 hours increased gene-editing frequency, while overnight (O/N) culture did not further increase gene editing (n=3). (E) After electroporating c-kit⁺ HSPCs with GFP-sg1, HSCs were sorted clonally into methylcellulose media. Most (40 out of 48) HSC colonies exhibited loss of GFP expression, as shown by the representative flow cytometric histograms for 3 HSC-derived colonies from one donor mouse (n=3 independent experiments). (F) A representative histogram demonstrating efficient ablation of GFP expression by electroporating Cas9/GFP-sg1 RNP into GFP expressing HSPCs (n=3). (G) Quantification of results in (F). Even as little as 200 ng of GFP-sg1 efficiently ablated GFP upon delivery with Cas9 protein (1 μg). (H) T7E1 assays performed with GFP amplicon derived from R26- or GFP-disrupted HSPCs. PCR amplicons were either treated (+) or untreated (−) with T7E1. Arrowheads indicate the bands with expected size assuming small indels, based on the Cas9 cleavage site. 1 μg of sgRNA was used to electroporate 10⁵ Cas9-expressing cells unless otherwise noted. All data represent mean ± standard deviation; *, p<0.05; **, p<0.01; and ***, p<0.001 by Student’s t-test. See also Figure S1.

Figure 2. Editing endogenous genes in murine HSPCs

(A) Gene editing of Eed or Suz12 using Cas9-sgRNA RNP increased the ability of murine HSPCs to serially replate in culture. 1 μg Cas9 protein and 1 μg of sgRNA were used (n=4). (B) T7E1 assays performed with Eed (upper panel) or Suz12 (lower panel) amplicon derived from Rosa26- (R26), Eed-, or Suz12- disrupted HSPCs 48 hours after electroporation. The numbers below the gel image represents the cleavage efficiency determined by densitometric analysis (n=3). (C, D) T7E1 assays performed with Eed (C) or Suz12 (D) amplicons derived from Rosa26- (R26), Eed-, or Suz12-disrupted colonies, with (+) or without (−) the nuclease (n=4). Arrowheads indicate the bands with expected size based on the Cas9 cleavage site, while asterisks indicate non-specific bands. (E, F) Sequencing results of representative 4 clones each (out of 12) after electroporating with Eed (E) or Suz12 (F) sgRNA. All colonies analyzed (Eed: 12/12, Suz12: 12/12) acquired indels. Red line represents the position of the PAM sequence. All data represent mean ± standard deviation; ***, p<0.001 by Student’s t-test. See also Figure S2.

Figure 3. Efficient CD45 knock-out in human hematopoietic cells

(A) Flow cytometry analysis of hCD45 expression in three AML cell lines and activated primary T cells 96 hours following electroporation with Cas9 only (blue) or Cas9/hCD45-sg1 RNP (red). (B) Effects of pre-culture before electroporation on gene disruption efficiency. hCD45 loss were examined in CD34⁺ cells cultured for 0, 24 and 48 hours in presence of cytokines before electroporation with Cas9/hCD45-sg1 RNP. hCD45 expression was evaluated 4 days after electroporation. Each experiment (n=8) was performed on CD34⁺ cells isolated from single donors. (C) Flow cytometry analysis of CD45 and CD34 expression in CD34⁺ cells 96 hours following electroporation with Cas9 only (left panel) or Cas9/hCD45-sg1 RNP (right panel). (D–E) Cell viability examined by trypan blue staining (D, n=8) or flow cytometry (E, n=6) of CD34⁺ cells electroporated either with Cas9 only (black) or with Cas9/hCD45-sg1 RNP (grey) relative to non-electroporated cells at the indicated time points. The cell counts (D) or viability (E) of Cas9 only and Cas9/hCD45-sg1 transfected cells were compared to the viable cell counts of non-electroporated cells. *, p<0.05; **, p<0.01; and ***, p<0.001 by non parametric t-test.

Figure 4. CRISPR-Cas9 mediated gene disruption and HDR in human HSPCs

(A) Table showing indel frequencies at targeted loci. When multiple sgRNAs were used in the same experiment, sg1 indicates alleles with disruption of sg1 only, sg2 indicates alleles with disruption of sg2 only and Δ indicates alleles with a deletion between sg1 and sg2. Tables containing raw allele counts for each indel/deletion are found in Table S2. (B) Indel frequencies at 3 predicted hCD45-sg1 off-target sites. Off-target sites (OT1-3) were predicted using CRISPRscan (Moreno-Mateos et al., 2015). (C) Plots showing percentages of human cells (left) in the bone marrow and the spleen of 16 NSG recipient mice (8 Cas9 only and 8 Cas9/hCD45-sg1 RNP) and the fraction of engrafted human cells that have lost hCD45 (right) in Cas9 only (black) and Cas9/hCD45-sg1 RNP (red). Human CD34⁺ cells from individual cord blood donors were electroporated with Cas9 only and Cas9/hCD45-sg1 RNP and transplanted into sublethally irradiated NSG mice. Engraftment was analyzed 8 weeks post-transplant. (D) Flow cytometry analysis of two engrafted NSG mice. Upper panels show the engraftment of normal human cells (CD45^posHLA-ABC^pos). Lower panels show the presence of hCD45 knock-out cells (highlighted in red) both in the bone marrow and the spleen. (E) Western blot analysis of DNMT3A expression in CD34⁺ cord blood cells 96 hours after electroporation with Cas9 only, Cas9/hCD45-sg1 RNP, Cas9/DNMT3A exon 7/8-sg (4 sgRNAs) RNP, or Cas9/DNMT3A exon 7-sg (2 sgRNAs) RNP. (F) Schematic representation of the CRISPR-mediated knock-in (top). Three single nucleotide changes, two of which (red) result in the formation of a BsiWI restriction site (italics), were introduced into hCD45 exon 25. The most common observed alleles from a representative sample (bottom), which include both precise (HDR; all three single nucleotide changes) and imprecise (HDR*; two out of three nucleotide changes) knock-in events, were assessed by high-throughput sequencing and their allele frequencies are displayed. The numbers in red represent frequencies of reads containing the BsiWI restriction site. (G) A gel image of BsiWI digested PCR amplicon prepared from CD34⁺ cord blood cells targeted with Cas9/hCD45-sg1 (1 μg each) RNP and different single-stranded DNA oligonucleotides (ssODN) containing BsiWI sites. Both symmetric (S) and asymmetric (A) homology arms were tested. BsiWI digests the 282 bp amplicon with HDR editing into 172 and 110 bp fragments. The numbers below the gel represent efficiency of restriction site knock-in (HDR+HDR*) as determined by high-throughput sequencing.