Gene Correction of Wiskott–Aldrich syndrome iPS Cells Rescues Proplatelet Defects and Improves Platelet Size

Abstract

Wiskott–Aldrich syndrome (WAS) is a severe X-linked disorder caused by loss-of-function mutations in the WAS gene, responsible for encoding WAS protein (WASP), a key regulator of the actin cytoskeleton in all hematopoietic cells, except red blood cells. The mechanism underlying microthrombocytopenia, a distinctive feature of WAS and a major contributor to mortality, remains not fully elucidated. In this study, using different gene-editing strategies, we corrected mutations in patient-derived WAS-induced pluripotent stem cell (iPSC) lines, generating isogeneic WAS-iPSC lines. These included lines with direct mutation-specific correction and lines incorporating a WASP transgene cassette regulated by the MND or WAS1.6 kb promoter integrated at the safe harbor AAV1 site. Our results demonstrated that direct mutation correction successfully restored WASP levels to the equivalent of the wild-type in iPSC-derived megakaryocytes (MKs). In contrast, the AAV1-targeted strategy using the MND and WAS1.6 promoters yielded a lower level of WASP. Notably, only the mutation-specific correction lines exhibited improvements in proplatelet structures and generated larger-sized platelets. Our findings underscore the crucial roles of WASP during human thrombopoiesis and suggest that therapeutic approaches, such as direct gene correction, which can achieve physiologic levels of WASP in MKs, hold promise for ameliorating platelet defects in individuals with WAS.

 

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Keywords

thrombocytopenia - Wiskott–Aldrich syndrome - gene targeting

Introduction

Wiskott–Aldrich syndrome (WAS) is a severe, X-linked primary immunodeficiency disorder originally described as a clinical triad of bleeding diathesis associated with microthrombocytopenia, eczema, and recurrent infections.[1] Patients with WAS have an increased risk of developing severe autoimmune disorders and lymphoid malignancies. WAS is caused by mutations in the gene encoding WAS protein (WASP), a scaffolding protein involved in mediating signals from membrane receptors to the actin cytoskeleton.[2] WASP is expressed exclusively in non-erythroid cells of the hematopoietic system, while other members of the WASP family, including N-WASP, WHAMM, WAVE, and WASH are more ubiquitously expressed. Consistent with the crucial role of actin polymerization in regulating cellular processes required for immune cell functions including podosome formation, cell migration, phagocytosis, and immunological synapse formation, cells of both adaptive and innate immunity have been reported to be defective in patients with WAS.[3] On the other hand, the mechanisms of microthrombocytopenia, the most unique and consistent finding and a leading cause of death in patients with WAS are still debatable.

WAS can be successfully treated with allogeneic hematopoietic stem cell transplantation (HSCT).[4] However, this curative treatment is only available for a limited number of patients. Gene therapy has emerged as an attractive treatment strategy for WAS patients without HLA-matched hematopoietic stem and progenitor cells (HSPCs) donors. Transplantation of genetically modified autologous HSPCs provides an opportunity to avoid toxicity from a full-conditioning regimen and reduces the incidence of graft-versus-host from allogeneic HSCT. An early study with a gamma retrovirus vector, despite the development of leukemia in several patients through insertional mutagenesis, proved that it could produce long-term improvements in immune cell function.[5] More recently, a study using a self-inactivating lentiviral vector containing a WASP 1.6-kb promoter has demonstrated a successful reduction of the autoimmune symptoms of WAS patients without causing significant side effects after a median follow-up of 8 years supporting the potential of this strategy.[6] [7] Nevertheless, platelet defects persist in a majority of patients. A better understanding of the roles of WASP defects in thrombocytopenia is crucial for advancing and refining this promising therapy.

Several lines of evidence suggest that thrombocytopenia may be influenced by both reduced platelet production and accelerated platelet consumption. For reduced platelet production, there are conflicting data on the role of WASP in proplatelet formation. The mouse model of WAS exhibited normal-size platelets, premature proplatelet formation, and fragmentation which were observed in the bone marrow (BM).[8] [9] However, deletion of genes known to interact with WASP such as ARP2/3 [10] and profilin-1 [9] resulted in microthrombocytopenia with premature platelet release in the BM murine models. In humans, an early report showed that HSPCs from WAS patients displayed reduced proplatelet formation.[11] Using the WAS patient-derived induced pluripotent stem cell (iPSC) model, we showed that WAS iPSC-derived megakaryocytes (MKs) exhibited qualitative proplatelet formation defects and produced small platelets in vitro.[12] [13] In contrast, there was also a report showing that HSPCs from WAS patients generated normal proplatelets.[14] In addition, a recent study has shown that proplatelet formation relies on NWASP rather than WASP.[15] For accelerated platelet consumption, it was revealed that platelets from WAS patients showed an increase in ex vivo phagocytosis.[16] The mouse model of WAS demonstrated that increased uptake of WAS^−/− platelets by splenic red pulp could also contribute to thrombocytopenia.[17]

An important issue that needs to be considered in iPSC disease modeling is the variability in the differentiation potential of each iPSC line due to genetic background and residual epigenetic memory. In this study, we used different genome editing strategies to create isogenic WAS-iPSC lines with different levels of WASP expression. As a result, we demonstrated a WASP level-dependent effect on MK properties. We also created new WAS-iPSC lines from a wild-type (WT) iPSC line using Clustered Regularly Interspaced Short Palindromic Repeats/CRISPR-associated protein 9-mediated gene editing and showed that it recapitulated defects in proplatelet structures. In summary, our findings support the application of a genome editing strategy for the treatment of WAS and suggest that the WAS-iPSCs differentiation system could serve as a viable tool in refining the next generation of therapies.

 

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Materials and Methods

WAS Gene Correction Design, AAVS1 Gene Targeting Design, Induced Pluripotent Stem Cell Gene Correction, Western Blot - [Supplementary Material S1] (available in the online version).

Induced Pluripotent Stem Cell Generation and Culture

WT-iPSCs, WASX503R-iPSCs, and WASQ19X-iPSCs were previously generated using temperature-sensitive sendai virus vector (TS7) encoding Oct-3/4, SOX2, Klf4, and c-Myc (a gift from DNAVEC, Japan).

WT-iPSCs were generated by using episomal vectors, pCXLE-hOCT3/4-shp53-F (Addgene plasmid #27077), pCXLE-hSK (Addgene plasmid # 27078), and pCXLE-hUL (Addgene plasmid #27080; a gift from Shinya Yamanaka).[18] The reprogramming procedure was performed as previously described.

All iPSC lines were cultured in feeder-free condition using dishes precoated with Corning™ Matrigel™ Human embryonic stem cell (hESC)-Qualified Matrix (Corning) and mTeSR1 media (Stemcell Technologies).

 

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Hematopoietic Cell Differentiation

For hematopoietic progenitor cell differentiation, small clump iPSCs dissociated with collagenase IV/trypsin, knockout (CTK) solution were cultured on mitotically inactivated C3H/10T1/2 cells (ATCC) in hematopoietic differentiation media, Iscove's Modified Dulbecco's Medium (IMDM) with 10 μg/ml human insulin, 5.5 μg/ml human transferrin, 5 ng/ml sodium selenite, 2 mM L-glutamine, 0.45 mM α-monothioglycerol, 50 μg/ml ascorbic acid, 15% FBS, supplemented with 20 ng/ml human vascular endothelial growth factor (VEGF; R&D Systems, Minneapolis, MN). At day 14 of differentiation, hematopoietic progenitor cells produced within the sac were collected by gently crushed with a pipette.

For MK differentiation, hematopoietic progenitor cells were reseeded onto freshly irradiated OP9 feeder cells and cultured in hematopoietic cell differentiation media supplemented with 50 ng/ml human thrombopoietin (R&D Systems), 10 ng/ml human stem cell factor (R&D Systems), and 25 ng/ml heparin (Sigma–Aldrich, St. Louis, MO).

 

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Time-lapse Proplatelet Formation

MKs on day 6 of differentiation were stained with 1 μM SiR-tubulin (Spirochrome, Cytoskeleton) and transferred onto matrigel-coated 96-well CellCarrier microplates (PerkinElmer). The image was captured every 2 hours for a total of 14 hours with Z-stack at 0, 1, 2, 3, and 4 μm using 63× magnification (Opera Phenix, PerkinElmer). For image analysis, five images (Z-stack) of each position were combined using maximum intensity projection. The proplatelet tip diameter and tubulin thickness were measured using Zen software (Blue edition; Carl Zeiss).

 

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Immunofluorescence Staining

For proplatelet formation, MKs on day 6 of differentiation were collected and transferred onto matrigel-coated cover slides and cultured for 24 hours. For proplatelet formation on the BMEC-1 feeder, 1 × 10⁵ BMEC-1 cells were cultured on a 5-μm transwell for 3 days (100% confluence) and treated with 10 ng/ml interleukin (IL)-1β for 18 hours. MKs were then transferred onto the BMEC-1 feeder and cultured for 24 hours. For platelet production, MKs on day 6 of differentiation, were collected and cultured in feeder-free condition for 6 days. Platelets produced in supernatant were collected by centrifugation at 500 × g for 15 minutes at 25 °C in the presence of 1 μM prostaglandin E1. The platelet pellet was resuspended and smeared onto cover slides. Then prepared slides were fixed with 4% paraformaldehyde for 15 minutes and incubated with blocking solution supplemented with 0.3% of Triton X-100 for 60 minutes at room temperature. Slides were stained with anti-α-tubulin (Abcam) and Alexa Fluor™ 647 Phalloidin or Alexa Fluor™ 488 Phalloidin. Nuclei were counter-stained with 4′,6-diamidino-2-phenylindole (molecular probe). All fluorescence images were obtained by using the Axio observer fluorescence microscope, Zeiss LSM800 confocal microscope, and Zeiss LSM980 confocal microscope (Carl Zeiss, Jena, Germany). For platelet size measurement, fluorescence images were obtained using the Axio observer fluorescence microscope (Carl Zeiss). The shortest diameter of tubulin-stained discoid-shaped platelets was measured using Zen software (Blue edition; Carl Zeiss).

For podosome formation, MKs at day 6 of differentiation were cultured on matrigel-coated 96-well CellCarrier microplates (PerkinElmer). After 72 hours, cells were stained with SPY555-FastAct (Spirochrome) for 1 hour. Then cells were fixed and counter-stained with DAPI. The image was captured using 63× magnification (Opera Phenix, PerkinElmer). A total of 100 to 150 cells were counted for each replicate (n = 3).

 

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Statistical Analysis

Statistical analyses were conducted using an unpaired t-test for platelet size and one-way analysis of variance (ANOVA) for tubulin thickness, proplatelet tip diameter, and percentage of podosome-forming cells. All data are presented as mean ± standard deviation.

 

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Results

CRISPR-mediated Wiskott–Aldrich syndrome Knockout Induced Pluripotent Stem Cells Recapitulate Proplatelet Formation and Platelet Size Defects of Induced Pluripotent Stem Cells derived from Wiskott–Aldrich syndrome Patients

To address the effect of genetic background on MK abnormality we previously observed in our two WAS-iPSC lines (WASQ19X and WASX503R),[12] we designed a Cas9-guide RNA (gRNA) plasmid to introduce double-strand breaks (DSBs) at the end of the WAS first exon and used it to generate isogenic WAS knockout (KO) iPSCs from control WT-iPSCs ([Fig. 1A]). WT-iPSCs were transfected with the Cas9-gRNA plasmid. Green fluorescent protein (GFP)-positive iPSC clones were expanded and screened for mutations in WAS exon 1 using the T7 assay. Mutations were subsequently confirmed by DNA sequencing. The WT-iPSC clones with frameshift mutations that created a premature stop codon were selected for further experiments ([Fig. 1B]). When iPSCs were differentiated into MKs and platelets using the ES-SAC method,[19] the isogenic WAS-KO iPSCs still retained the capability to generate MKs comparable to the parent iPSC line ([Fig. 1C, D]). Western blot analysis showed the absence of WASP expression in MKs derived from WAS-KO iPSCs ([Fig. 1E]). Remarkably, MKs from WAS-KO iPSCs demonstrated qualitative defects in proplatelet formation ([Fig. 1F]) and generated smaller platelets than MKs from the WT-iPSCs ([Fig. 1G, H]). Thus, abnormalities we observed from iPSCs derived from WAS patients could be recreated from WT cells with different genetic backgrounds.

Fig. 1 WAS isogenic model. (A) Schematic diagram of WASP gene knockout (KO) using CRISPR/Cas9. (B) DNA sequencing analysis of the WASP gene of CRISPR/Cas9-transfected iPSC clones. (C) Flow cytometry analysis of CD34 expression in WT-iPSCs and WAS mutated WT-iPSCs derived from sac. (D) Flow cytometry analysis of CD41 and CD42b expression in WT-iPSC and WAS mutated WT-iPSC derived megakaryocytes. (E) Western blot analysis revealed the absence of WASP expression in WAS-mutated WT iPSC-derived MKs. (F) Immunofluorescence staining of proplatelet formation showed defects in proplatelet structures of WAS-KO iPSC-derived MKs. (G) Immunofluorescence staining of platelets generated from the WT and WAS-KO iPSCs using an anti-α-tubulin antibody (green). (H) Box plot showing platelet diameter of tubulin-stained discoid-shaped platelets generated from the WT and WAS-KO iPSCs. Data are presented as mean ± SEM, n = 300 (****p < 0.0001). iPSC, induced pluripotent stem cell; MK, megakaryocyte; WAS, Wiskott–Aldrich syndrome; WASP, WAS protein; WT, wild-type.

 

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Zinc Finger Nucleases-mediated Gene Correction of WAS Mutation Restores Wiskott–Aldrich syndrome Protein Levels in Megakaryocytes and Increases Platelet Size

From the WASX503R-iPS cell line derived from fibroblasts of a WAS patient we previously characterized,[12] we designed zinc finger nucleases (ZFNs)[20] that generated a DSB 28 bp after the mutation site within exon 12 of the WAS gene ([Fig. 2A]). WASX503R-iPSCs were nucleofected with ZFNs and a donor vector plasmid containing WT homology arms and a hygromycin selection cassette flanked by Cre–lox sequences. After selection with hygromycin, 8 of 11 clones were confirmed by PCR-Sanger sequencing that the mutation site was corrected ([Fig. 2B]).

Fig. 2 WASX503R gene correction. (A) Schematic diagram of WASX503R gene correction using ZFN. (B) DNA sequencing showed single base correction at c.1507 position of the WASP gene in cWASX503R iPSCs. (C) Western blot analysis revealed WASP expression in the WT-iPSC and ccWASX503R-derived MKs. (D) Electron microscopic examination of platelets generated from WT-iPSC, WASX503R-iPSC, and ccWASX503R-iPSC-derived megakaryocytes. Bottom panel, immunofluorescence staining of platelets generated from WASX503R and ccWASX503R iPSCs using an anti-α-tubulin antibody (red). (E) Box plot showing platelet diameter of tubulin-stained discoid-shaped platelets generated from WT, WASX503R, cWASX503R, and ccWASX503R iPSCs. Data are presented as mean ± SEM, n = 120 (****p < 0.0001). iPSC, induced pluripotent stem cell; MK, megakaryocyte; WASP, Wiskott–Aldrich syndrome protein; WT, wild-type; ZFN, Zinc finger nuclease.

Two of these corrected clones (cWASX503R-iPSCs) were treated with Cre recombinases to remove the hygromycin cassette, resulting in the generation of isogenic Cre-treated corrected (ccWASX503R-iPSCs) lines. All isogenic WASX503R-iPSC lines retained the ability to differentiate into hematopoietic cells. Although all isogenic WASX503R-iPSC lines generated an indistinct number of CD34+ hematopoietic stem/progenitor cells, WASP expression was observed exclusively in MKs derived from ccWAS-iPSCs, and not in those derived from WAS-iPSC ([Fig. 2C]) and cWAS-iPSC lines ([Supplementary Fig. S1] [available in the online version]). This result suggested that the unremoved selection cassettes in the cWAS line interfered with WASP translation.

While the number of platelet-like particles generated in our in vitro differentiation system was not remarkably different, the size of ccWASX503R-derived platelets was larger than those produced from the parent line, as shown by electron microscopic analysis and immunofluorescence imaging ([Fig. 2D]). To eliminate the possibility of cell debris in our co-culture system, we specifically quantified α-tubulin-stained particles and found a significant increase in mean platelet size ([Fig. 2E]). In contrast, corrected WAS-iPSC lines retaining the hygromycin selection cassette (cWAS-iPSC), and lacking expression of the WASP, did not exhibit a correction in size.

 

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Gene Correction Strategies Effects on Wiskott–Aldrich syndrome Protein Level and Platelet Size

The mutations associated with WAS are heterogeneous. Aside from individually correcting each mutation, employing a universal correction strategy, such as gene insertion at the safe harbor site of the genome, may provide benefits comparable to those of a viral vector. Simultaneously, it has the potential to mitigate possible side effects associated with random integration. To evaluate the effect of different gene correction strategies on in vitro platelet formation, the WASQ19X iPSC was gene-edited to create multiple isogenic lines ([Fig. 3A, B]). We first employed a strategy resembling WASX503R correction to correct the mutation at the target sequence by using TALEN2[21] [22] and the donor vector containing the loxP-flanked phosphoglycerate kinase (PGK)-hygromycin cassette designed to integrate into intron 1 of the WAS gene. Although we could generate many iPSC clones with the correct integration site, after the successful removal of the selection cassette, WASP expression remained absent from the corrected WAS iPSC-derived MKs. This result suggested that the 32-bp loxP residual nucleotides could interfere with WASP splicing. We subsequently employed a piggyBac donor vector for gene correction to avoid interference by the insertion of exogenous sequences. Puromycin-resistant clones from three experiments were selected; however, a corrected WASQ19X-iPSC clone could not be obtained. We next evaluated gene correction of the WASQ19X using CRISPR/Cas9 and single-strand oligonucleotides (ssODNs) as a template donor. We designed 90-bp donor oligonucleotides with a silent mutation at the gRNA-binding site to prevent recutting. Cas9 and gRNA-expressing plasmids, containing GFP as a selectable marker, were co-nucleofected with ssODNs. Three corrected WASQ19X iPSC clones (cWASQ19X-D1, cWASQ19X-D3, and cWASQ19X-F1) were selected from 40 GFP-positive sorted iPSCs ([Fig. 3C]).

Fig. 3 WASQ19X gene correction. (A) Schematic diagram of endogenous gene correction at WASQ19X using TALEN with a donor vector and CRISPR/Cas9 with ssODNs. (B) Schematic diagram of WASP gene knock-in at the AAVS1 site using TALEN with a donor vector. (C) DNA sequencing analysis showed mutation correction at c.55 positions of the WASP gene with a silent mutation at positions −1 and −4 of the mutation site. (D) mRNA expression of the WASP gene under the control of the MND promoter and WAS1.6 promoter in the AAVS1 locus compared with normal expression levels using droplet digital PCR. (E) Western blot analysis of WASP expression in iPSCs and iPSC-derived megakaryocytes. The expression of WASP was detected in AAVMNDPW(Q19X) iPSCs and AAVW1.6PW(Q19X) iPSCs but not in WASQ19X iPSCs. In megakaryocytes, WASP was detected in WT and cWASQ19X-derived MKs and rarely detected in AAVW1.6PW(Q19X). (F) Box plot showing platelet diameter of tubulin-stained discoid-shaped platelets generated from WASQ19X, cWASQ19X, AAVW1.6W(Q19X), and AAVMNDW(Q19X) MKs. Data are presented as mean ± SEM, n = 300 (****p < 0.0001). (G) Immunofluorescence staining of platelets generated from WASQ19X, cWASQ19X, AAVW1.6W(Q19X), and AAVMNDW(Q19X)-derived MKs using an anti-α-tubulin antibody (green). (h) Western blot analysis of N-WASP expression in iPSC-derived MKs. iPSC, induced pluripotent stem cell; MK, megakaryocyte; WASP, Wiskott–Aldrich syndrome protein; WT, wild-type.

For the universal correction strategy, the WAS1.6-WASP ORF and MND-WASP ORF expression cassettes[23] were targeted to the AAVS1 locus using TALEN ([Fig. 3B]). When the selected clones of the AAVW1.6PW(Q19X) iPSCs and AAVMNDPW(Q19X) iPSCs were differentiated into MKs, transgene WASP mRNA expression was detected at 25% of the endogenous WASQ19X mRNA in AAVW1.6PW(Q19X) iPSC-derived MKs and 4% in AAVMNDPW(Q19X) iPSC-derived MKs as analyzed by digital PCR ([Fig. 3D]). As shown in the western blot, CRISPR-corrected WASQ19X iPSC-derived MKs expressed WASP at a level equal to the WT-iPSC-derived MKs. In contrast, lower levels of the WASP were detected in MKs differentiated from both universal correction lines compared with lines generated from CRISPR-mediated Homology directed repair (HDR) ([Fig. 3E]). Therefore, various strategies could result in a significant difference in the levels of WASP observed in MKs derived from isogenic iPSC lines.

We analyzed the in vitro platelet production capabilities of these isogenic corrected WAS-iPSC lines. Remarkably, corresponding with the level of WASP expression, only the WAS-iPSC line with correction at the mutation site produced larger platelets than the uncorrected line ([Fig. 3F, G]). It should be noted that we observed a stable level of NWAS in all our isogenic lines ([Fig. 3H]). This observation suggested that NWAS did not contribute to platelet defects observed in our system.

 

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Wiskott–Aldrich syndrome Protein Level-dependent Effect on Proplatelet Branching and Tubulin Thickness

To study the effect of WASP on proplatelet formation, iPSC-derived MKs were cultured on matrigel-coated coverslip for 24 hours before being fixed and analyzed by immunofluorescent staining for cytoskeletal protein tubulin. When compared with the WT MKs, WASX503R and WASQ19X exhibited thinner and less branching proplatelet processes and small proplatelet tips. All gene-corrected WAS MKs, except the AAVMNDPW(Q19X), showed marked improvements in proplatelet morphology ([Fig. 4A]).

Fig. 4 Proplatelet defects in WAS-derived MKs. (A) Immunofluorescence staining of proplatelet formation. MKs were cultured on a matrigel-coated coverslip for 24 hours. Cells were fixed and stained with anti-α-tubulin antibody (green) and phalloidin (red). WT, ccWASX503R, and cWASQ19X-derived proplatelets exhibited thicker proplatelet shafts and larger proplatelet tips compared with WASX503R, WASQ19X, AAVW1.6PW(Q19X), and AAVMNDW(Q19X). (B) Time-lapse fluorescence image of proplatelet formation in WASX503R and ccWASX503R-iPSC-derived MKs stained with SiR-tubulin. At the early stage of proplatelet formation, the extension of pseudopodia was observed in both conditions, but with a difference in its size and microtubule concentration. WASX503R showed a rapid proplatelet process. The microtubule bundles in ccWASX503R-derived proplatelet shafts are thickened and form larger proplatelet tips compared with those observed in WASX503R. Box plot showing proplatelet tip diameter (C) and tubulin thickness of proplatelet (D) of WT, WAS-KO, WASX503R, ccWASX503R, WASQ19X, cWASQ19X, AAVW1.6PW(Q19X), and AAVMNDW(Q19X)-iPSC-derived proplatelets. Data are presented as mean ± SEM, n = 63 and n = 59, respectively (****p < 0.0001, **p < 0.005, *p < 0.05). iPSC, induced pluripotent stem cell; KO, knockout; MK, megakaryocyte; WAS, Wiskott–Aldrich syndrome; WASP, WAS protein; WT, wild-type.

We then employed time-lapse analysis to study the effect of WAS gene correction on microtubule dynamic during proplatelet processing ([Fig. 4B]). WASX503R and ccWASX503R-derived MKs were treated with SiR-tubulin, a fluorescence-tagged tubulin, and visualized every 2 hours for 14 hours. WASX503R MKs extended multiple thin proplatelet processes around the cell margin without microtubule-forming cortical bundles. In contrast, in the ccWASX503R MKs, there was evident microtubule condensation at the protrusion sites, resembling the pattern previously described by Thon et al.[24] Notably, compared with the ccWASX503R, the branching and elongation process of WASX503R-derived proplatelets was more rapid, and their structures were less complex.

To quantify the change in proplatelet morphology among different corrected iPSC lines, we performed a time-lapse analysis on MKs generated from other WAS iPSC lines and measured the proplatelet tip diameter ([Fig. 4C]) and tubulin thickness of the proplatelet shaft ([Fig. 4D]). Our results showed that MKs from the direct gene-corrected lines (ccWASX503R and cWASQ19X-D3) and the universal correction line, AAVW1.6PW(Q19X) significantly increased both proplatelet tip size and proplatelet shaft thickness compared with the parent lines. Nevertheless, compared with the direct gene-corrected cWASQ19X-D3, the AAVW1.6PW(Q19X) produced significantly smaller proplatelet tips and thinner proplatelet shafts. The line corrected with an MND promoter cassette slightly increased proplatelet tip diameters, but the tubulin thickness did not significantly increase. Altogether, our data suggested that the level of WASP expression correlated with proplatelet phenotypes.

 

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Wiskott–Aldrich syndrome Protein Level-dependent Effect on Podosome Formation of Megakaryocytes

WASP is a key podosome formation regulator essential for cell migration and invasiveness. Immune cells and MKs from WAS^−/− mice failed to assemble podosomes.[8] Similar to previous reports, MKs generated from the uncorrected WASX503R, WASQ19X, and WAS-KO iPSC lines could not form podosomes when attached to the matrigel-coated surface ([Fig. 5A]). In contrast, MKs derived from the direct gene-corrected lines, ccWAS503R and cWASQ19X, could form podosomes. MKs derived from the AAVW1.6PW(Q19X) were able to form podosomes, but the number of podosome-positive MKs was significantly lower than those derived from the cWASQ19X ([Fig. 5B]). Thus, in our study, the WASP level correlated with the ability of MKs to form podosomes.

Fig. 5 Podosome formation. (A) Immunofluorescence staining of podosomes. MKs were cultured on a matrigel-coated coverslip for 72 hours. Cells were stained with SPY555-FastAct and fixed. Podosome-forming MKs were detected in WT, ccWASX503R, cWASQ19X, and AAVW1.6PW(Q19X). (B) The bar graph showed the percentage of podosome-positive cells in each condition (n = 3; ****p < 0.0001). (C) Fluorescence image with an orthogonal view of MKs cultured on interleukin (IL)-1β-treated BMEC1 feeder layer (8-μm cell culture insert) in the presence of SDF1α in the bottom well. Twenty-four hours after culture, cells were fixed and stained with CD42b (purple), anti-α-tubulin antibody (green), and phalloidin (red). In WASQ19X condition, only planar contact between MK and BMEC1 was observed without podosome formation. For corrected WASQ19X-D3, white arrowheads showing podosome forming MKs which protruded their cytoplasm and nucleus toward BMEC1 feeder cells. Crossed gray lines indicate the selected orthogonal planes. MK, megakaryocyte; WT, wild-type.

A recent study has suggested that MKs could use podosome-like structures to protrude the proplatelet process through sinusoidal endothelium before shedding platelets.[25] To test whether WAS mutation correction could affect endothelial invasion, we then co-cultured the WASQ19X and cWASQ19X derived-MKs on the BMEC-1 cell line and performed immunofluorescent staining. The confocal image revealed podosome-like structures that protruded through the BMEC layer from the corrected WASQ19X-derived MKs but not the uncorrected WASQ19X ([Fig. 5C]). This result suggested that defects in podosome formation could contribute to thrombocytopenia in WAS.

 

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Discussion

In our previous study, we described proplatelet defects and the generation of small-size platelets in vitro from WAS-iPSCs. To ascertain the influence of heterogeneity among iPSC lines and genetic background, in this study, we generated multiple isogenic iPSC lines. Remarkably, our results consistently demonstrated level-dependent impacts of WASP on the regulation of proplatelet structures and platelet size in vitro.

Poor proplatelet structures could lead to premature proplatelet rupture in BM similar to what was observed in mutant mice.[8] [9] [10] [11] Although the low levels of WASP driven by the MND or WAS1.6 promoter integrated at the AAVS1 site were sufficient to restore MK actin polymerization and podosome formation, the resulting impact on proplatelet morphology and platelet size was significantly less pronounced compared with gene-corrected WAS-iPSC lines. These iPSC lines successfully restored WASP levels to those observed in the WT MKs. We demonstrated that the corrected WAS MKs could form podosome-like structures while invading through endothelial cells. Nevertheless, it remained unclear how much quantitative or qualitative defects of MK podosome formation could contribute to residual defects in thrombopoiesis following gene therapy. Therefore, achieving a physiologic level of MK function may necessitate a higher level of WASP than other immune cells. This observation mirrors the clinical presentation of X-linked thrombocytopenia, where a low WASP level leads primarily to thrombocytopenia without notable defects in immune cells. The N-WASP level remained unchanged in our system, making it unlikely to be the causative factor for the observed proplatelet phenomena. However, the role of N-WASP in the overall proplatelet process should be further investigated.

Safe harbor-targeted CRISPR-mediated HDR has been proposed as a cost-effective strategy for correcting various genetic diseases with heterogenous mutations while avoiding random integration of viral vectors.[26] Our data suggest that the genome editing approach, capable of restoring WASP levels in MKs to those of the WT, may offer a potential advantage over AAVS1 safe harbor gene targeting with the WAS1.6 promoter in the restoration of platelet defects. Improving the proplatelet formation defects with this strategy will likely require a stronger MK promoter, regulatory regions, or targeting different safe harbor sites. The screening of proplatelet formation in WAS-iPSCs can be employed for further optimizing the strategy. Nevertheless, whether improved proplatelet formation and platelet size in our in vitro system correlate with a better platelet count still needs to be evaluated clinically. Apart from WAS, our data suggest that the selection of promoters and regulatory regions for transgene expression needs to be tailored specifically for each disease. On the other hand, a direct gene editing approach requires that each mutation be specifically designed and optimized to increase editing efficiency, reduce off-target, and prevent insertion mutagenesis, which can be challenging to achieve and costly.

The limitation of the iPSC system in generating long-term repopulating HSCs prevents gene-corrected iPSC-derived HPC from being tested in patients.[27] Nevertheless, recently remarkable progress has been made in gene correction in adult HSPCs. A platform such as AAV6 has been demonstrated to produce a high rate of HDR in HSC/progenitors while maintaining the Hematopoietic stem cell (HSC) capability of serial transplantation in mouse models.[28] [29] [30] Approaches aiming for the universal correction of WASP expression using an AAV6 donor vector, containing a full-length WASP ORF for targeted integration at the first exon, have shown promising results in both in vitro experiments and animal models.[31] [32] This strategy is particularly relevant given the heterogeneous nature of mutations responsible for WAS. Nevertheless, genome injury signals from DSB during gene correction using the HDR-based strategy may reduce the long-term maintenance of corrected HSCs.[33] While it has been shown that CRISPR-mediated Non-homologous DNA end-joining (NHEJ) in HSCs can last more than 2 years in patients, it remains to be seen whether HSCs corrected with HDR can long-term self-renew in non-human primates and patients. Several new studies demonstrate effective strategies to overcome the genome toxicity and inflammatory signals caused by genome editing such as lipid nanoparticles,[34] small molecules,[35] and optimizing donor templates.[36] Alternatively, gene editing techniques without DSB such as base editing and prime editing,[37] theoretically cause less injury to the genome and may provide better long-term outcomes. However, it is important to note that certain toxicity issues have been reported as well.[38] Overall, although certain challenges persist, gene editing strategies hold promise for individuals with WAs. On the other hand, viral-mediated gene therapy has demonstrated efficacy lasting more than 5 years in patients and has proven effective in significantly reducing bleeding events. Further optimization of the vector to increase WASP expression in MKs has the potential to improve this already effective therapy.

 

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Conflict of Interest

None declared.

Acknowledgment

We thank Mahendra Rao, Jizhong Zou, and Sukhdeep Singh Dhadwar for the genome editing tool design.

Authors' Contribution

Conceptualization: N.I. and K.S. Designed the experiments: P.I. and N.I. Investigation: P.I., S.P., P.A., P.M., P.P., and N.S. Funding acquisition: K.S., N.I., and V.S. Writing original draft: N.I., P.I., and K.S. Review and editing: K.S., N.I., and V.S. Project administration: K.S. and N.I. Supervision: V.S.

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Publikationsverlauf

Eingereicht: 09. Juli 2024

Angenommen: 20. Dezember 2024

Accepted Manuscript online:
24. Dezember 2024

Artikel online veröffentlicht:
25. Februar 2025