CRISPR-Cas9 RAG2 Correction via Coding Sequence Replacement to Preserve Endogenous Gene Regulation and Locus Structure

doi:10.21203/rs.3.rs-2565742/v1

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CRISPR-Cas9 RAG2 Correction via Coding Sequence Replacement to Preserve Endogenous Gene Regulation and Locus Structure

https://doi.org/10.21203/rs.3.rs-2565742/v1

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published 27 Oct, 2023

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RAG2-SCID is a primary immunodeficiency caused by mutations in Recombination-activating gene 2 (RAG2), a gene intimately involved in the process of lymphocyte maturation and function. ex-vivo manipulation of a patient’s own hematopoietic stem and progenitor cells (HSPCs) using CRISPR-Cas9/rAAV6 gene editing could provide a therapeutic alternative to the only current treatment, allogeneic hematopoietic stem cell transplantation (HSCT). Here we show a first-of-its-kind RAG2 correction strategy that replaces the entire endogenous coding sequence (CDS) to preserve the critical endogenous spatiotemporal gene regulation and locus architecture. Expression of the corrective transgene led to successful development into CD3⁺TCRαβ⁺ and CD3⁺TCRγδ⁺ T cells and promoted the establishment of highly diverse TRB and TRG repertoires in an in-vitro T-cell differentiation platform. We believe that a CDS replacement technique to correct tightly regulated genes, like RAG2, while maintaining critical regulatory elements and conserving the locus structure could bring safer gene therapy techniques closer to the clinic.

Biological sciences/Stem cells/Haematopoietic stem cells

Biological sciences/Immunology/Lymphocytes

Severe combined immunodeficiency (SCID) is a group of multiple rare monogenic disorders characterized by defects in both cellular and humoral adaptive immunity. Patients are born healthy and due to being extremely susceptible to infections, they present with reccurent infections early in life which if left untreated can be fatal^1–3. The Recombination-activating genes 1 and 2 (RAG1 and RAG2, respectively) are tightly linked and have convergent transcriptional orientations on chromosome 11 separated by ~ 12 kb. The RAG genes encode proteins that, when complexed together, commence the lymphoid-specific variable (V), diversity (D), and joining (J) gene [V(D)J] recombination process by catalyzing DNA double-strand breaks (DSBs) at the recombination signal sequences (RSSs) which flank the V, D, and J gene segments⁴. V(D)J recombination is a critical step in the maturation of T and B cells as it is responsible for the generation of a diverse repertoire of T- and B-cell receptors (TCR and BCR, respectively). Thus, patients with disease-causing variants in the RAG genes typically present with the complete absence or significant reduction of T and B cells and the T-B-NK + immune phenotype^{2, 5–9}. V(D)J recombination has three main mechanisms of regulation: 1) lineage specificity, namely BCR and TCR gene rearrangement occurs in B cells and T cells, respectively; 2) immunoglobulin heavy-chain rearrangement occurs before immunoglobulin light chain; and 3) allelic exclusion, namely once a T or B cell rearranges its receptor locus, only one functional allele will be expressed in that cell. In addition to these general mechanisms, the transcription of the RAG1 and RAG2 genes is regulated by numerous highly-conserved, lineage-specific, cis-acting sequences surrounding their respective CDSs which control the spatial genomic organization inside the locus^10–15. The RAG genes are expressed exclusively in the G₀/G₁ stage and display a tightly-linked genomic organization with regulated expression during specific phases of T and B lymphocyte development. More specifically, two limited waves of RAG1 and RAG2 expression are necessary for Ig heavy and light chain rearrangements after which their expression is promptly terminated¹⁶. This process is orchestrated by a plethora of transcriptions factors and machinery that complex with cis-regulatory elements and promoter regions in the RAG1/2 locus. Together, they creating a chromatin hub super enhancer for the expression of the RAG genes^12,14,17,18. This formation, as well as the chromatin structure and 3D architecture, are crucial to ensuring that RAG1/2 are only expressed during the requisite developmental window¹⁹. Overexpression or expression of the RAG genes outside of this precise window, can result in genomic instability and lymphocyte malignancy through the formation of translocations and/or deletions in cancer-causing genes^{11,12, 19–23}. Unsuccessful termination of RAG1 and RAG2 expression is also associated with atypical thymus development, an aberrant lymphatic system, and immunodeficiency²⁴.

Currently, the only definitive curable treatment for SCID patients is allogeneic hematopoietic stem cell transplantation (HSCT) from a human leukocyte antigen (HLA)-matched donor²⁵. However, finding a HLA-matched donor is rare, and the alternative treatment, haploidentical HSCT, reduces the survival rate from > 80% with an HLA-matched donor to 60–70%^26,27. Although successful HSCT promotes lymphoid lineage development resulting in a long-term patient survival rate, it is accompanied by a high risk of graft-versus-host disease^28,29.

An ideal alternative to searching for an HLA-matched donor is to genetically edit the patient’s own CD34⁺ HSPCs ex-vivo to be subsequently returned to the patient as an autologous HSCT. CD34⁺ HSPCs’ marked ability to reconstitute the immune system from a small number of cells, make these cells an attractive platform for gene-therapy applications^30,31. To that end, transgene delivery via lentiviral (LV) or gammaretroviral (γRV) vectors for ex-vivo editing of patients’ CD34⁺ HSPCs was previously reported in gene-therapy clinical trials^32–36. However, genome-editing-based treatments using γRV of Chronic Granulomatous Disease (CGD)³⁷, Wiskott-Aldrich Syndrome (WAS)^38,39, SCID-X1⁴⁰, and Adenosine Deaminase (ADA)-SCID⁴¹ resulted in the activation of proto-oncogenes leading, in some patients, to a leukemic transformation^42,43. Although steps have been taken to improve the safety of these viral vectors, transgene integration into tumor-suppressor loci has been observed and incomplete phenotypic correction, toxicity, dysregulated hematopoiesis, and insertional mutagenesis related to the semi-random integration and constitutive expression of the transgene into the genome remain major safety concerns^44,45. Despite these concerns, which are particularly severe for highly controlled and regulated genes such as the RAG1/2, LV-based gene therapy for RAG1 is currently undergoing clinical trials⁴⁶.

For these reasons, transgene delivery via a targeted HDR-mediated genome-editing approach such as the combination of Clustered regularly interspaced short palindromic repeats (CRISPR) and CRISPR-associated nuclease 9 (Cas9) and recombinant adeno-associated virus serotype 6 (rAAV6) could prove particularly beneficial for treating RAG-SCIDs^47–49. Although rAAV6 provides a number of benefits over LV and γRV vectors, we have shown that rAAV6 vectors trigger a toxic DNA damage response (DDR) proportional to the multiplicity of infection (MOI), or the amount of virus used⁵⁰. Thus, for a CRISPR-Cas9/rAAV6 genome-editing strategy to be therapeutically relevant, a delicate balance between maintaining high-quality HDR while reducing viral load as much as possible is required. Therefore, we tested a number of different donor constructs in order to find the optimal rAAV6 donor design for efficient HDR.

Additionally, a highly-specific CRISPR-Cas9-based approach is particularly beneficial for disorders where the affected gene is highly-regulated, such as RAG-SCIDs^11,12,51. CRISPR-Cas9/rAAV6 genome editing provides the potential for specific gene integration that maintains the strict endogenous spatiotemporal gene regulation and expression of the transgene. Modulation of transgene expression levels by including mRNA stability and/or nuclear export cis-acting elements such as polyadenylation (polyA or pA) signals or Woodchuck Hepatitis Virus Posttranscriptional Regulatory Element (WPRE) segments has been widely reported⁵². Polyadenylation at the end of a RNA transcript affects mRNA stability, nuclear export, and translation, thus, playing a crucial role in gene expression. In-vitro studies using rAAV6 vectors have highlighted the effects of different synthetic polyA signals, such as bovine growth hormone (BGH) polyA signal sequences, in boosting transgene expression^53,54. Similarly, cis-acting post-transcriptional regulatory elements (PREs), such as WPRE, are required for the nuclear export of intronless RNA, and the addition of such PREs has been reported to enhance transgene expression in-vitro^53,55. Through the use of such sequences to regulate our RAG2 transgene, we aim to establish transgene expression that closely resembles that of the the endogenous gene.

We previously reported on a proof-of-concept CRISPR-Cas9/rAAV6 RAG2 gene correction approach⁵⁶ which successfully led to the development of T cells with diverse TCR repertoires from RAG2-SCID patient-derived CD34⁺ HSPCs. Here, we describe a CRISPR-Cas9/rAAV6-mediated genome-editing approach that replaces the entire RAG2 CDS with a corrective transgene in CD34⁺ HSPCs, to maintain the endogenous spacial regulatory elements of the RAG2 locus. We achieved this by constructing three RAG2 CDS replacement correction donors where each maintains the 5’ endogenous promoter and regulatory elements^{10,15, 57–59}. However, in addition to the regulatory elements upstream to the RAG2 gene, at least one silencer sequence has been discovered between the RAG1 and RAG2 genes that is counteracted by an anti-silencing element 5’ of the RAG2 promoter⁶⁰. Additionally, there are evolutionarily conserved genes within the RAG1/2 locus' intronic sequences that are believed to play a role in transcriptional regulation^11,12,61,62. Thus, to most closely mimic the endogenous expression of RAG2, we designed one donor to preserve the 3’ UTR and downstream region in addition to the 5' UTR and promoter region. The two additional correction donors were engineered in an effort to increase the expression levels of the transgene by maintaining the RAG2 5’ UTR endogenous region, while introducing synthetic 3’ UTRs (WPRE-BGHpA and BGHpA sequences).

Developing a proof-of-concept gene-correction therapy typically requires large amounts of patient samples. However, since untreated SCID patients rarely survive past infanthood, neither invasive bone-marrow procedures nor drawing large volumes of peripheral blood (PB) are viable options for procuring such samples. Due to this challenge, we used enrichment of HD-derived CD34⁺ HSPCs with engineered genotypes after multiplex HDR (knock-in/knock-out [KI-KO]) to simulate single-allelic gene-correction therapies for RAG2-SCID. Since RAG2-SCID is an autosomal recessive disorder, correction of only one mutated allele is sufficient to cure the patient. In this strategy, we mimicked monoallelic correction in SCID-patient cells by knock-in (KI) of a diverged codon-optimized RAG2 (dcoRAG2) cDNA cassette in place of the endogenous RAG2 CDS in one allele (thereby preserving regulatory non-coding elements) and by knock-out (KO) of the second allele with a green fluorescent protein (GFP) gene-disrupting cassette. Via cell sorting, we were then able to enrich for cells with the desired KI-KO engineered genotype to model and track their progression into T-cell development. Thus, in this study, we show the first-of-its-kind RAG2 gene correction simulation in HD-derived CD34⁺ HSPCs by replacement of the entire endogenous CDS.

Different HDR Strategies: Cut-Site Insertion vs. Coding Sequence (CDS) Replacement and Adjusting Homology Arm Length

RAG2-SCID is caused by mutations scattered throughout the CDS of the RAG2 gene⁴. Therefore, a universal correction technique that would suit all RAG2-SCID patients requires the delivery of an intact copy of the complete RAG2 CDS. While KI of an intact CDS at the endogenous locus would achieve this, this strategy could interfere with the 3D chromatin architecture and critical endogenous gene regulation by moving regulatory elements further downstream from the transgene. This could potentially disrupt spatial cross-talk between functional elements upstream and downstream of the RAG2 gene, such as promoter and/or enhancer sequences^12,14,17,18 (Figure S1A-D). Hence, we hypothesized that a donor DNA with a left homology arm (LHA) upstream adjacent to the cut site, and a right homology arm (RHA) distanced from the cut site, downstream of the RAG2 stop codon would ensure preserving RAG2 regulatory elements by replacing the entire RAG2 CDS. To examine the feasibility of our hypothesis, we produced two rAAV6 vectors that would integrate a GFP expression cassette under the regulation of a spleen focus-forming virus (SFFV) promoter and BGHpA sequence after delivery of a chemically modified RAG2 sgRNA/Cas9 ribonucleoprotein (RNP) complex via electroporation into CD34⁺ HSPCs. In previous studies, we demonstrated this sgRNA’s high on-target editing efficiency and accuracy^50,63,64. The first donor⁵⁶, herein CSI_GFP-BGHpA_400x400, uses 400bp homology arms immediately flanking the Cas9-induced cut site for donor insertion (hereafter termed a cut-site-insertion [CSI] vector), while the second donor, herein CDSR_GFP-BGHpA_400x400, uses a 400bp LHA spanning the immediate sequence upstream to the Cas9 cut site and a 400bp RHA spanning the immediate sequence downstream to the RAG2 stop codon, to replace the entire RAG2 CDS with the DNA donor (herein a CDS-replacement [CDSR] vector) (Fig. 1A and Table S1). Two days post-editing, via flow cytometry, we found that the HDR efficiency of the CSI_GFP-BGHpA_400x400 vector was significantly higher than that of the CDSR_GFP-BGHpA_400x400 vector (21.8% and 9.1%, respectively) (Fig. 1B and S1E). Attempting to improve the HDR efficiency of the CDSR technique, we designed an additional two rAAV6 donors with RHAs extended from 400bp to 800bp and 1,600bp spanning the immediate region downstream to the RAG2 stop codon (herein CDSR_GFP-BGHpA_400x800 and CDSR_GFP-BGHpA_400x1600, respectively [Figure 1A and Table S1]). While elongation to 800bp produced significantly higher HDR efficiency than the CDSR_GFP-BGHpA_400x400 donor (14.8%), only after elongation to 1,600bp, did we observe HDR efficiency comparable to that of the CSI_GFP-BGHpA_400x400 donor (25.2%). (Fig. 1B and Figure S1E). Using a uniform pair of primers and probe for droplet digital PCR (ddPCR), we confirmed that the HDR efficiencies as determined by flow cytometry were accurate and locus-specific (Fig. 1C). Interestingly, via flow cytometry, we observed a significantly higher mean fluorescence intensity (MFI) of GFP-expressing cells after integration of the CDSR vectors compared to the CSI vectors (Fig. 1D-E), highlighting that different integration strategies have unique effects on the genomic locus and impact subsequent transgene expression.

Synthetic polyA Sequences and/or Cis-acting PREs Affect Transgene Expression

To modulate transgene expression further, we aimed to test the impact of synthetic 3’ regulatory elements on transgene expression. Thus, we designed two CDSR vectors (herein CDSR_GFP-WPRE-BGHpA_400x1600 and CDSR_GFP-NoBGHpA_400x1600 [Figure 2A and Table S1]) with a homology arm pattern similar to that of the CDSR_GFP-BGHpA_400x1600 donor. However, whereas the CDSR_GFP-BGHpA_400x1600 vector contained a BGHpA sequence alone and the CDSR_GFP-WPRE-BGHpA_400x1600 contained a WPRE-BGHpA sequence, the CDSR_GFP-NoBGHpA_400x1600 vector lacked both regulatory elements, thus allowing GFP expression to be controlled by the endogenous RAG2 3’-UTR. While the CDSR_GFP-BGHpA_400x1600 and CDSR_GFP-WPRE-BGHpA_400x1600 donors produced comparable HDR efficiencies, the CDSR_GFP-NoBGHpA_400x1600 donor induced lower HDR as observed by flow cytometry and confirmed by ddPCR (Fig. 2B-C and S2). Interestingly, the three donors produced significantly different MFI levels, with CDSR_GFP-BGHpA_400x1600 (2.8x10⁶) being the highest followed by CDSR_GFP-WPRE-BGHpA_400x1600 and CDSR_GFP-NoBGHpA_400x1600 (1.6x10⁶ and 0.3x10⁶, respectively) (Fig. 2D-E), highlighting the strength of the synthetic 3’-UTRs in boosting expression and the significance of preserving the endogenous RAG2 regulatory elements.

KI-KO Genotype Engineering in HD-derived HSPCs Using Two-part Enrichment Strategy

Since RAG2 gene regulation is critical, we aimed to fine-tune our previously published RAG2-correction strategy⁵⁶ by using the CDSR method. We hypothesized that replacing the entire CDS would allow for transgene expression to be driven by the RAG2 endogenous promoter and 3’-UTR, thus enabling the transgenic dcoRAG2 cDNA expression patterns to most similarly resemble that of endogenous RAG2. Additionally, by replacing the entire CDS (~ 1.5kb), as opposed to pushing the sequence ~ 4kb downstream in the case of HDR via insertion, we are able to more closely maintain the proximity of the RAG genes, conserving the ability to form the chromatin hub super enhancer necessary for proper expression. The dcoRAG2 cDNA produces a protein identical to WT RAG2, while the introduction of wobble changes leads to reduced similarity to the genomic sequence precluding the Cas9 from re-cutting the inserted sequence or from the inserted sequence serving as a homology arm causing premature cessation of HDR. We constructed a CDSR correction donor (herein CDSR_Corr_Endo3’UTR [Figure 3A and Table S1]) with a 400x800bp homology arm pattern for KI of the dcoRAG2 cDNA. To track the expression of dcoRAG2 cDNA and enrich for cells with successful integration, the dcoRAG2 stop codon was eliminated and replaced with a T2A self-cleaving peptide sequence followed by a truncated nerve growth factor receptor (tNGFR) reporter gene, producing in-frame transcription of the two sequences (dcoRAG2 cDNA and tNGFR). Following translation of the fusion protein, the T2A self-cleaves producing two proteins (RAG2 and tNGFR) at a 1:1 ratio in the cell. The use of tNGFR is particularly advantageous since it enables tracking and enrichment of corrected cells and has been approved for clinical applications⁶⁵. Additionally, we constructed two donors with synthetic 3’-UTRs following the tNGFR, one with WPRE-BGHpA sequences and one with only the BGHpA sequence each with a 400x800bp homology arm pattern (herein CDSR_Corr_WPRE-BGHpA and CDSR_Corr_BGHpA, respectively [Figure 3A and Table S1]). While we observed highest HDR for CDSR donors with a RHA of 1,600bp (Fig. 1B-C), we could not design correction donors with a 400x1,600bp homology arm pattern due to the limited carrying capacity (~ 4.8kb) of rAAV6 vectors⁶⁶. For comparative purposes, we utilized our previously published CSI KI donor which contained dcoRAG2 cDNA followed by the RAG2 endogenous 3’-UTR sequence along with a tNGFR reporter gene cassette under the regulation of a constitutive phosphoglycerokinase (PGK) promoter and BGHpA sequence between 400bp homology arms (herein CSI_Corr)⁵⁶ (Figure S3A and Table S1). We tested these four correction donors individually and observed highly effective locus-specific HDR for them all, determined by ddPCR (Figure S3B).

We aimed to utilize the KI-KO strategy to engineer genotypes via multiplex HDR in HD-derived CD34⁺ HSPCs to simulate the therapeutic outcome of RAG2-SCID single-allelic correction following a gene-editing-based treatment. This strategy has two main advantages over more extensive editing methodologies: 1) In contrast to the use of induced pluripotent stem cells (iPSCs)^67–70, HD-derived CD34⁺ HSPCs are biologically authentic since they are the same cells used in HSCT⁷¹; and 2) Lengthy culturing protocols are insufficient, since CD34⁺ HSPCs lose their regenerative ability as well as their engraftment potential after elongated culturing⁷². Thus, we chose to apply our KI-KO strategy in HD-derived CD34⁺ HSPCs by utilizing multiplex HDR to obtain a cell population with one allele targeted with one of the four aforementioned correction donors and the other allele with a KO template (Fig. 1A [CDSR_GFP-BGHpA_400x800 was paired with the three CDSR correction donors and CSI_GFP-BGHpA_400x400 was paired with the CSI_Corr]).

For the CSI_Corr donor, enrichment of KI-KO CD34⁺ HSPCs was achieved by sorting for biallelic double-positive tNGFR⁺/GFP⁺ expression two days post-electroporation (herein day 0) and immediately seeding the cells into the in-vitro T-cell differentiation (IVTD) system (Figure S3C-E). However, with CDSR correction donors, since tNGFR expression is under the regulation of the RAG2 promoter (and the RAG2 expression window occurs later in the T-cell developmental process), there is no expression of tNGFR on day 0. Therefore, a novel enrichment strategy was required to isolate KI-KO cells. On day 0, the CDSR donors were sorted only for KO GFP expression, and the GFP⁺ cells were immediately seeded into the IVTD system. On day 14 of IVTD, when RAG2 is highly expressed (thus, tNGFR is expressed [Figure S3F]) tNGFR⁺ cells were sorted for and seeded back into the IVTD system (Fig. 3B and S3G-H). Additionally, all samples were sorted for CD7 expression to enrich for cells that have begun to differentiate, namely, only cells that were CD7⁺ were subjected to days 14–28 of IVTD (Figure S3G). ddPCR was performed on gDNA from the KI-KO populations to confirm that the two-step enrichment method indeed led to enrichment of a cell population with ~ 100% edited alleles. Indeed, in all four correction donor multiplex HDR combinations, ~ 100% of targeted alleles were found to be positive (Fig. 3C). For confirmation that the integration of the donors occurred as expected, we conducted an 'in-out' PCR with one primer located on the tNGFR sequence and one primer downstream to the RHA (Figure S3I). Indeed, the observed amplified bands were consistent with the expected integration patterns and corroborated effective HDR (Figure S3J).

Additionally, since the CDSR donors produce a fusion protein separated by a self-cleaving T2A sequence resulting in a 1:1 ratio between transgenic RAG2 and tNGFR, tNGFR MFI measurement was used as a proxy measurement for transgenic dcoRAG2 expression levels. As expected, we observed that the MFI for CDSR_Corr_WPRE-BGHpA was 2x that of CDSR_Corr_Endo3’UTR and 1.4x greater than CDSR_Corr_BGHpA on day 28 of IVTD with a similar trend on day 14 (Fig. 3D) indicating higher expression of the dcoRAG2.

KI-KO HSPCs Produce CD3⁺TCRγδ⁺ and CD3⁺TCRαβ⁺ T Cells in the IVTD System

With a robust method to isolate cells with the KI-KO genotype, we aimed to validate that specifically the expression of dcoRAG2 enabled the KI-KO cells to differentiate into CD3⁺ T cells with diverse TCR repertoires, to present a proof-of-concept for gene correction. Quantitative real-time PCR (qRT-PCR) using transcript-specific primer pairs (Figure S4A) revealed that the expression of endogenous RAG2 was ostensibly eliminated in the KI-KO populations while robust dcoRAG2 cDNA expression was found exclusively in the KI-KO engineered cells (Fig. 4A and S4B). Additionally, when we directly compared the total RAG2 mRNA levels between all groups, we found that expression of the dcoRAG2 transgenes does not exceed that of the Mock samples indicating that the transcription is still tightly controlled and that the gene is not being overexpressed (Fig. 4B).

Importantly, the expression of the dcoRAG2 cDNA indeed facilitated T-cell development highlighted by the successful differentiation of RAG2 KI-KO cells into CD7⁺, CD5⁺, and CD1a⁺ pre-T cells on day 14 and CD3⁺ T cells by day 28 (Fig. 4C-D and S4C-D). Additionally, robust TCRγδ expression in the CD3⁺ population was observed by flow cytometry on day 28 with the CD3⁺TCRγδ⁻ cells presumed to be CD3⁺TCRαβ⁺ T cells (Fig. 4E). Lastly, PCR amplification using primers flanking the V-J regions of the TRG locus highlighted the successful recombination of KI-KO cells comparable to that of the Mock cells on day 28 (Figure S4E).

Expression of dcoRAG2 cDNA Induces Normal TCR Repertoire

Deep-sequencing analysis of TRB and TRG recombination on day 28 revealed diverse V(D)J rearrangement repertoires in the RAG2 KI-KO populations following expression of the dcoRAG2 cDNA (Fig. 5A) with no significant differences in either TRB or TRG clonotypes between the Mock and RAG2 KI-KO populations as calculated by Shannon’s H and Simpson’s 1-D diversity indices (Fig. 5B-C). Lastly, the complementarity determining region 3 (CDR3) lengths frequency distribution was comparable in all RAG2 KI-KO and Mock populations for both the TRB and TRG sequencing (Figure S5). CDR3 is the region of the TCR responsible for recognizing processed antigen peptides and its sequence and length varies from one clone to another. Thus, sequencing the CDR3 regions of a cell population is used as a measurement of TCR diversity⁷³. Together, these data indicate that KI and expression of the dcoRAG2 cDNA promotes successful V(D)J recombination, subsequent differentiation into CD3⁺TCRαβ⁺ and CD3⁺TCRγδ⁺ T cells, and the development of highly diverse TRB and TRG repertoires.

When it comes to highly-regulated genes such as RAG2, a correction strategy that replaces the entire endogenous CDS is particularly advantageous, since it maintains the gene’s endogenous regulatory elements and safeguards the locus architecture, compared to insertion of the corrective donor at the Cas9-induced cut site which distorts the locus by approximately 4 kb (see Figure S1A-D)⁵³. Previous works to correct the RAG genes lacked the ability to maintain endogenous regulatory and spatiotemporal elements since the integration of the transgene was either semi-random or via insertion of thousands of bp to the Cas9-induced cut site, thus distancing the RAG genes one from the other and potentially altering the genomic locus and hindering optimal gene expression. In our work, we show that the CDS replacement method is able to induce efficient expression of the dcoRAG2 cDNA without exceeding naturally-occuring RAG2 expression levels found in Mock samples (25–50% expression of RAG2). Taken together with the efficient T-cell differentiation induced by the corrective transgenes, we describe a novel approach to simulating genome-editing correction for the treatment of autosomal recessive inborn errors of immunity. While our approach specifically addresses RAG2, the methodology can be applied to any tightly controlled gene.

Interestingly, when we designed a CDS replacement rAAV6 donor by distancing the RHA from the cut site (CDSR_GFP-BGHpA_400x400), HDR frequencies were significantly reduced compared to the insertion donor with equivalent homology arm lengths (CSI_GFP-BGHpA_400x400). Recently, Cromer et al. showed that elongating the length of the homology arms can lead to higher rates of HDR⁵³. Thus, to account for the reduced efficiencies with the CDS replacement donor, we elongated the RHA to 800 bp which induced higher HDR frequencies, however, only once we elongated the RHA to 1,600 bp were we able to completely abrogate the low HDR efficiency observed with 400 bp. We hypothesize that by increasing the length of the distal homology arm, we enhanced the possibility for recognition and subsequent incorporation of the exogenous donor template by the cellular HDR mechanism. Additionally, we observed a significant change in expression of the reporter gene between integration and CDS replacement donors, attributable to substantial and unique conformational changes in the edited locus which affect expression. Lastly, we found that we were able to further modulate the reporter gene expression via the addition or removal of synthetic polyA sequences. Synthetic 3’ UTRs are important and easily implementable tools, some of which have been already implemented in gene-therapy applications^46,74. Although these strategies proved to be effective for improving HDR and/or reporter-gene expression in the RAG2 locus, it is well documented that every genomic locus has its own unique characteristics and genomic topology. Thus, we surmise that the exact length and positioning of homology arms to induce efficient HDR in both an integration and CDS replacement manner is target- and locus-specific.

To establish that the expression of our transgene alone is capable of differentiating CD34⁺ HSPCs into CD3-expressing T cells with diverse TRB and TRG repertoires, we utilized a KI-KO approach for each of our CDS replacement correction donors in HD-derived CD34⁺ HSPCs. Since large amounts of SCID-patient samples are scarce, many studies have used iPSCs to develop gene-correction models^67–70. However, in contrast to iPSCs, HD-derived CD34⁺ HSPCs provide a biologically authentic platform since they are the same cells used in HSCT⁷¹. The use of the KI-KO strategy outlined in Iancu et al. to mimic single-allelic correction of SCID-patient cells provides that all T-cell differentiative progress is due solely to the expression of the transgene and not due to the presence of endogenous RAG2⁵⁶. KI of the transgene in one allele and reliance on Cas9-induced insertions or deletions (INDELs) for KO of the second allele would not be sufficient since the Cas9 does not induce KO INDELs in 100% of the relevant alleles (50% of the total alleles), and there would be no way to isolate the KI-KO cells in this case. Thus, using multiplex HDR with two distinct reporter genes to indicate the KI-KO genotype and FACS enrichment of the double-positive population is the most effective method for achieving ~ 100% KI-KO enrichment in HD-derived CD34⁺ HSPCs. Since our correction donors rely on the endogenous RAG2 promoter for expression which only begins seven days into IVTD (see Figure S3F), we developed a two-step enrichment process over the course of T-cell differentiation which provided a robust platform for testing our dcoRAG2 cDNA correction donors. Additionally, due to the rapid expansion of cells over the first 14 days of IVTD, our two-step enrichment strategy enabled us to both use fewer cells and to reduce the MOI from 25,000 viral genomes (VG)/cell (used in Figs. 1 and 2) to 12,500 VG/cell while still maintaining requisite HDR. This is particularly beneficial since our group previously showed, in Allen et al., that rAAV6 vectors can trigger a potentially toxic DDR after entering cells, proportional to the MOI used⁵⁰. Thus, reduction of the rAAV6 toxicity is required in order for a rAAV6-based method to be implemented as a clinical therapy for the purpose of gene correction. Follow-up studies to test different methods of improving HDR to allow for further reduction of the MOI, including inhibiting the NHEJ repair pathway via molecules such as i53 and/or DNA-PK inhibitors^75–77, are currently underway.

Utilizing the KI-KO strategy, we were able to track the different correction donors individually and pinpoint the effect that their expression patterns had on T-cell development. We observed higher dcoRAG2 mRNA levels in the CDS replacement correction donors than in the insertion correction samples, highlighting the effect of different gene-editing and transgene-integration strategies on the resulting gene expression. Notably, even between CDS replacement donors, we observed a significantly higher level of dcoRAG2 mRNA when the transgene expression was under the 3' regulation of the WPRE-BGHpA sequence than the expression level under the endogenous 3' UTR. Additionally, the RAG2 mRNA levels produced by the CDS replacement donors were observed to be 25–50% that of the Mock samples indicating that overexpression of RAG2 transgene is not occurring. We expect that the difference between mRNA levels in Mock and KI-KO samples is in part a function of only one active allele in the correction simulation samples compared to two active alleles in the Mock samples. Since parents of RAG2-SCID patients (carriers of RAG mutations ["naturally-occuring" KI-KO]) present clinically as normal and together with the fact that our KI-KO samples are able to differentiate effectively, we conclude that the mRNA levels observed are sufficient. Lastly, since the dcoRAG2 and tNGFR are expressed from the CDS replacement donors as a fusion protein, when the T2A peptide that separates the two proteins self-cleaves, the result is a 1:1 ratio of RAG2:tNGFR. Thus, the MFI of tNGFR can serve as a proxy measurement for the level of translation of RAG2 in the cell. Indeed, higher MFI was observed for the CDS replacement donor under the 3' regulation of the WPRE-BGHpA sequence, consistent with the difference identified on the mRNA level. Together, these data indicate that the synthetic 3’ UTRs have a significant effect on the mRNA stability and/or nuclear export efficiency leading to greater translation into RAG2. Lastly, all three CDS replacement RAG2 correction donors promoted successful V(D)J recombination and subsequent differentiation into CD3⁺TCRαβ⁺ and CD3⁺TCRγδ⁺ T cells and developed highly diverse TRB and TRG repertoires. Although the CDS replacement correction donors with synthetic 3' UTRs may be efficient for demonstrating robust transgene expression and successful T-cell development in our IVTD system, they retain the risk of leading to aberrant expression pattern in patient cells; and more extensive studies are ongoing in our lab to assess this possibility.

In summary, the aforementioned results describe a novel genome-editing strategy that replaces the entire RAG2 CDS (1,584 bp [in contrast to the CSI_Corr donor that leads to an introduction of an additional 3,815 bp at the Cas9 cut site]), thus treating any and all possible mutations in the gene while introducing a transgene that maintains endogenous regulatory and spatiotemporal elements. We believe that the CRISPR-Cas9/rAAV6-based CDS replacement strategy for a proof-of-concept gene therapy, which is site-specific and in a clinical setting would use a patient’s own cells, can alleviate a number of current barriers to treatment for not only RAG2-SCID but other monogenic diseases of the blood and immune system as well. Firstly, autologous gene therapy would eliminate the need for finding an HLA-matched donor and would reduce the risk of graft rejection. Secondly, using a CRISPR-Cas9/rAAV-based approach avoids the risk of dysregulated hematopoiesis, incomplete phenotypic correction, and insertional mutagenesis associated with the semi-random integration of a γRV- or LV-based approach. Thirdly, since the 3D genomic architecture of the RAG1/2 locus is critical for proper gene expression^12,14,17,18 we were able to avoid inserting kilobases of new DNA between the two genes by using a CDS replacement strategy. This allows for transgene integration while maintaining important regulatory and spatiotemporal elements which can be crucial, especially in genes like RAG2 that are expressed in a very limited and tightly controlled window. We show that expression patterns of the corrective transgene are both sufficient to induce quality T-cell differentiation while not exceeding endogenously occurring levels in the Mock samples. For these reasons, we believe that adoption of such a strategy can assist in reducing the risks associated with gene therapy and lead to safer and more accurate applications.

Cells and cell-culture conditions

Cord blood (CB)-derived CD34⁺ HSPC were obtained from Sheba Medical Center CB bank under Institutional Review Board-approved protocols. CD34⁺ HSPCs were isolated via magnetic bead-separation (Miltenyi Biotec). CD34⁺ HSPCs were cultured in SFEM II enriched with 100 ng/ml Flt3-Ligand, 100 ng/ml TPO, 100 ng/ml SCF, 0.035 mg/µl UM171, 0.75 mg/µl SR1 (Stemcell Technologies), 20 unit/ml penicillin, and 20 mg/ml streptomycin (Biological Industries, Beit Haemek, Israel) at 37°C, 5% CO₂, and 5% O₂.

CRISPR-Cas9 preparation and nucleofection

CRISPR-Cas9 RNP complex preparation and nucleofection were conducted in accordance with the extensive protocol published in Shapiro et al.⁷⁸. A single guide RNA (sgRNA, Alt-R^®sgRNA, IDT) with end chemical modifications was complexed with Cas9 at a molar ratio of 1:2.5 (Cas9:sgRNA) for 10-20 min at 25°C to form the RNP complex. The RAG2 sgRNA variable region sequence is 5'-UGAGAAGCCUGGCUGAAUUA-3'. RNP complexes were added to CD34⁺ HSPCs reconstituted in P3 Primary Cell electroporation solution, according to the manufacturer’s protocol (Lonza) at a final molar concentration of 4 μM. The cell solution was electroporated in the Lonza 4D-Nucleofector using the DZ-100 program.

rAAV6 DNA donor design and vector production

All rAAV6 vector plasmids were designed and cloned into the pAAV-MCS plasmid containing AAV2-specific inverted terminal repeats (ITRs). The pDGM6 plasmid, containing the AAV6 cap genes, AAV2 rep genes, and adenovirus helper genes, was a gift from David Russell (University of Washington). The final rAAV6 vectors were produced by The University of North Carolina (UNC) Vector Core in large-scale rAAV6 batches (UNC Vector Core).

Genome targeting and quantification

After electroporation with the RAG2 RNP complex, CD34⁺ HSPCs were seeded at a density of 0.4x10⁶ cells/ml for 24 hrs. Following the incubation period, the cell density was adjusted to 0.25x10⁶ cells/ml for an additional 24 hrs. In HDR experiments, the cells were transduced with the rAAV6 donor at MOIs of either 12,500 or 25,000 VG/cell within 5 min of electroporation. Cells were cultured at 37°C, 5% CO₂, and 5% O₂ in StemSpan™ SFEM II enriched medium as noted above. Flow cytometry analyses were performed on the BD LSRFortessa™ (BD Biosciences), Aria III cell sorter (BD Biosciences), or the Accuri C6 flow cytometer (BD Biosciences)

IVTD system and immunostaining

CD34⁺ HSPCs were cultured in the StemSpan™ T-Cell Generation Kit (STEMCELL Technologies, Inc.). For the first 14 days, cells were cultured in StemSpan™ SFEM II medium containing Lymphoid Progenitor Expansion Supplement in plates pre-coated with the Lymphoid Differentiation Coating Material. Cells were then harvested and re-seeded for an additional 14 days in StemSpan™ SFEM II medium contanining the T-Cell Progenitor Maturation Supplement on pre-coated plates. Flow cytometry analysis was conducted on days 14 and 28 of IVTD using the LSR Fortessa™ (BD Biosciences). On day 14, cells were stained with PE/Cy7-anti-CD7 (clone: CD7-6B7, BioLegend), BV421-anti-CD5 (clone: UCHT2, BioLegend), PE-anti-CD1a (clone: BL6, Beckman Coulter), and APC-anti-NGFR (clone: ME20.4, BioLegend) antibodies. On day 28, cells were stained with PE/Cy7-anti-CD4 (clone: RPA-T4, BioLegend), APC-r700-anti-CD8a (clone: RPA-T8, BD Horizon™) BV421-anti-CD3 (clone: UCHT1, BioLegend), and APC-anti-NGFR (clone: ME20.4, BioLegend) antibodies. BD Horizon™ Fixable Viability Stain 510 was performed on all collected cells at both time points. Gating strategies were based on fluorescence minus one (FMO) plus isotype control samples using the following isotypes: PE/Cy7 Mouse IgG2a κ, (BioLegend), BV421 Mouse IgG1 κ (BioLegend), PE Mouse IgG1 κ (BioLegend), PE/Cy7 Mouse IgG1 κ (BioLegend), APC-r700 Mouse IgG1 κ (BD Biosciences), and APC Mouse IgG1 κ (BioLegend).

Digital Droplet PCR™ (ddPCR™)

Genomic integration quantification for HDR experiments was performed by Digital Droplet PCR™ (ddPCR™, Bio-Rad). DNA was extracted from cell populations using GeneJET Genomic DNA Purification Kit (Thermo Fisher Scientific). Each ddPCR reaction contained a HEX reference assay detecting the CCRL2 gene to quantify the chromosome 3 copy number input⁷⁹. FAM assays for either KO (disruption) or KI (correction) donors were designed to detect the locus-specific donor integration. The ddPCR reaction was carried with the following reagents: 10 μl of ddPCR Supermix for Probes No dUTP (Bio-Rad), 1 μl each of FAM and HEX PrimeTime^® Standard qPCR Assay (IDT), 1 μl restriction enzyme mix [5 μl EcoRI-HF^® (NEB), 2 μl nuclease-free water, 1 μl CutSmart Buffer 10X (NEB)], genomic template DNA, and supplemented to a total of 20 μl with nuclease-free water. Droplet samples were prepared according to the manufacturer’s protocol (Bio-Rad) and 40 µl of the droplet output was transferred to a 96-well plate and amplified in a Bio-Rad PCR thermocycler (Bio-Rad) with at the following PCR conditions: 1 cycle at 95°C for 10 min, then 40 cycles of 95°C for 30 sec and 55°C for 3 min, followed by 1 cycle at 98°C for 10 min with a ramp rate of 2.2°C/sec. Following the PCR, the plate was read in the QX200 Droplet Reader (Bio-Rad) and the data was analyzed using the QuantaSoft analysis software (Bio-Rad). Primer and probe sequences are listed in Table 1.

mRNA quantification

RNA was extracted using Direct-zol™ RNA Miniprep Plus (Zymo Research) from differentiated T cells obtained on day 28 of IVTD. cDNA preparation was executed from RNA, using Oligo d(T)23 VN- S1327S (NEB), dNTPs 10 mM (Sigma-Aldrich), and M-MuLV Reverse Transcriptase (NEB). qRT-PCR reactions were conducted using the TaqMan^® Fast Advanced Master Mix (Thermo Fisher Scientific) in the StepOnePlus™ Real-Time PCR System (Thermo Fisher Scientific). PCR conditions were as follows: uracil-N-glycosylase gene (UNG) incubation at 50°C for 2 min, polymerase activation at 95°C for 20 sec, and 40 cycles at 95°C for 1 sec and at 60°C for 20 sec. Primer and probe sequences are presented in Table 1.

TRB and TRG V(D)J assessment

Genomic DNA was extracted using the GeneJET Genomic DNA Purification Kit (Thermo Fisher Scientific) from differentiated T cells on day 28 of IVTD. For TRG assessment via PCR amplification, 12 possible CDR3 clones were amplified using combinations of 4 primers for the Vγ regions and 3 primers for the Jγ regions (primer sequences are presented in Table 2). The PCR products were run on a 2% agarose gel. For deep sequencing of the TRB and TRG repertoires, the same genomic DNA was amplified using a multiplex master mix from either the LymphoTrack^® TRB assay and/or LymphoTrack^® TRG assay kits (Invivoscribe, Inc.). The amplicons were purified and sequenced using the MiSeq V2 (500 cycles) kit, with 250-bp paired-end reads (Illumina). The resulting FASTQ files were analyzed by the LymphoTrack Software (Invivoscribe, Inc.) and by the IMGT^® Software (The International ImMunoGeneTics Information System^®, HighV-QUEST, http://www.imgt.org). The analysis of the incidence and clonality of TRB and TRG rearrangement sequences were performed for visual representation by the TreeMap Software (Macrofocus GmbH). Unique CDR3 sequence and length were determined from the total productive sequences. Lastly, the Shannon’s H and Simpson’s 1-D diversity indices were calculated using the PAST Software⁸⁰.

Data sharing statement

The TRB and TRG sequencing data were deposited to the Sequence Read Archive (SRA), under accession number: PRJNA926613.

Acknowledgements

We would like to thank the members of the Somech and the Hendel Labs for reading the manuscript and providing practical advice. Additionally, we would like to thank D. Russell for providing the pDGM6 plasmid. This study was supported in part by research funding from the European Research Council (ERC) under the European Union Horizon 2020 research and innovation program (Grant No. 755758) as well as the Israel Science Foundation (ISF) - Israel Precision Medicine Partnership (IPMP) (Grant No. 3115/19) and Israel Science Foundation (ISF) - Individual Research Grants (Grant No. 2031/19).

Authorship Contributions

D.A., O.K., and B.I. designed and conducted the experiments; D.A. evaluated, and analyzed the data and performed the bioinformatics analyses, with the help and guidance of O.I. and Y.N.L.; K.B., and A.N. provided cord blood samples; K.B., Y.N.L., A.N., and R.S. critically reviewed the experiments and provided important advice; A.H. supervised and conceived the research; D.A. and A.H. wrote the manuscript, with contributions and input from all authors.

Conflict of Interest Disclosures

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest. Figures 1A, 2A, 3A, S1A-D, S3A, S3G, S3I, S4A, and Table S1 were created with BioRender.com.

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RAG2 KO )DNA(	Forward	CTCTCCCAGAGCAACAAAGAC
	Reverse	TTTCCATGCCTTGCAAAATGG
	Probe	56-FAM/CCTACAGGT/ZEN/GGGGTCTTTCATTCC/3IABkFQ/
RAG2 KI Correction )DNA(	Forward	TCTCCCAGAGCAACAAAG
	Reverse	GAAGTTCATGAGGCTGAAG
	Probe	56-FAM/CATAGCCTT/ZEN/AATCCAGCCCG/3IABkFQ/
CCRL2 )DNA(	Forward	GCTGTATGAATCCAGGTCC
	Reverse	CCTCCTGGCTGAGAAAAAG
	Probe	5HEX/TGTTTCCTC/ZEN/CAGGATAAGGCAGCTGT/3IABkFQ
Endogenous RAG2 )RNA(	Forward	CCAAGTGCTGACAATTAATACCTG
	Reverse	GACATGGTTATGCTTTACATCCAG
	Probe	56-FAM/CATCAGTGA/ZEN/GAAGCCTGGCTGAATTAAGG/3IABkFQ
dcoRAG2 cDNA )RNA(	Forward	CAAGTGCTGACAATTAATACCT
	Reverse	GAAGTTCATGAGGCTGAAG
	Probe	56-FAM/CATAGCCTT/ZEN/AATCCAGCCCG/3IABkFQ
Comparison RAG2 )RNA(	Forward	CCAAGTGCTGACAATTAATACCTG
	Reverse	GACATAGTTTCTGATGGTACGTAGA
	Probe	56-FAM/TCACGCCTC/ZEN/TCTGAATCTTTGCCG/3IABkFQ
GAPDH )RNA(	Forward	ACATCGCTCAGACACCATG
	Reverse	TGTAGTTGAGGTCAATGAAGGG
	Probe	56-FAM/AAGGTCGGA/ZEN/GTCAACGGATTTGGTC/3IABkFQ

Table 1 – ddPCR and qRT-PCR assay sequences

Jγ_{_1/2}	TACCTGTGACAACCAGTGTTG
Jγ_{_P}	ACTTACCTGTAATGATAAGCTTT
Jγ_{_P1/2}	TTACCAGGCGAAGTTACTATG
Vγ_{_9_2}	ACCTGGTGAAGTCATACAGTTC
Vγ_{_11}	CTTCCACTTCCACTTTGAAA
Vγ_{_f1}	ACTGGTACCTACACCAGGAGG
Vγ_{_10-2}	AGCATGGGTAAGACAAGCAA

Table 2 – TRG PCR amplification primers for V(D)J recombination assessment

There is NO Competing Interest.

Cord blood (CB)-derived CD34+ HSPC were obtained from Sheba Medical Center CB bank under Institutional Review Board-approved protocols. Donations of cord blood to public cord blood bank are recruited among labors in the obstetric delivery department. Informed consent is signed that cord blood specimens that are not suitable for banking will be used for research. No compensation is granted upon donation.

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published 27 Oct, 2023

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CRISPR-Cas9 RAG2 Correction via Coding Sequence Replacement to Preserve Endogenous Gene Regulation and Locus Structure

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Abstract

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Introduction

Results

Discussion

Methods

Cells and cell-culture conditions

CRISPR-Cas9 preparation and nucleofection

rAAV6 DNA donor design and vector production

Genome targeting and quantification

IVTD system and immunostaining

Digital Droplet PCR™ (ddPCR™)

mRNA quantification

TRB and TRG V(D)J assessment

Data sharing statement

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References

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Supplementary Files

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