Conditional CRISPR-mediated deletion of Lyn kinase enhances differentiation and function of iPSC-derived megakaryocytes

Abstract

Background:

Thrombocytopenia leading to life-threatening excessive bleeding can be treated by platelet transfusion. Currently, such treatments are totally dependent on donor-derived platelets. To support future applications in the use of in vitro-derived platelets, we sought to identify genes whose manipulation might improve the efficiency of megakaryocyte production and resulting hemostatic effectiveness. Disruption of Lyn kinase has previously been shown to improve cell survival, megakaryocyte ploidy and TPO-mediated activation in mice, but its role in human megakaryocytes and platelets has not been examined.

Methods:

To analyze the role of Lyn at defined differentiation stages during human megakaryocyte differentiation, conditional Lyn-deficient cells were generated using CRISPR/Cas9 technology in iPS cells. The efficiency of Lyn-deficient megakaryocytes to differentiate and become activated in response to a range of platelet agonists was analyzed in iPSC-derived megakaryocytes.

Results:

Temporally controlled deletion of Lyn improved the in vitro differentiation of hematopoietic progenitor cells into mature megakaryocytes, as measured by the rate and extent of appearance of CD41⁺CD42⁺ cells. Lyn-deficient megakaryocytes also demonstrated improved hemostatic effectiveness, as reported by their ability to mediate clot formation in rotational thromboelastometry. Finally, Lyn-deficient megakaryocytes produced increased numbers of platelet-like particles (PLP) in vitro.

Conclusions:

Conditional deletion of Lyn kinase increases the hemostatic effectiveness of megakaryocytes and their progeny as well as improving their yield. Adoption of this system during generation of in vitro-derived platelets may contribute to both their efficiency of production and their ability to support hemostasis.

Introduction

Bone marrow megakaryocytes (MKs) release ~2×10⁶ platelets per second, maintaining a platelet count of around 150,000 to 450,000 platelets per microliter of circulating blood. Platelet levels less than this are classified as thrombocytopenia, which can be caused by defective MK/platelet production, platelet depletion by platelet-specific auto- or allo-antibodies, blood loss due to trauma or surgery, or as a result of chemo- or radiation therapy. A platelet count of <30,000 per microliter of blood puts a patient at an increased risk of bleeding, and platelets are often transfused, either prophylactically or therapeutically, to reduce the risk of spontaneous bleeding. Maintaining adequate supplies of platelets is often a challenge due to (1) their short five-day shelf life; (2) requirement of room-temperature storage to maintain their activity; (3) variable functionality between donors; and (4) their short circulating half-life following transfusion. These properties make it difficult to maintain platelet products at the required level [1]. Repeated transfusions also increase the likelihood of developing an alloimmune response against ABO blood group antigens, Class I Human Leukocyte Antigens (HLA), or Human Platelet-specific Alloantigens (HPA), and the resulting anti-platelet antibodies can lead to development of post-transfusion purpura (PTP) or refractoriness to donor platelets [2]. Donor-derived blood products are also subject to bacterial and viral contamination; as a result, sepsis caused by contaminated platelet product represents the most frequent infectious complication from blood products today [3]. Finally, the >10⁶ units of platelets administered in the US each year still entirely relies on voluntary donations, the large majority of which are antigen mismatched. To address these issues, a number of laboratories have been working to develop processes to produce functional platelets in vitro that have well-defined antigenic composition to reduce the risk of alloimmune rejection and bacterial contamination [4–9].

Human induced pluripotent stem cells (iPSCs) are increasingly being developed as a safe, reliable starting material from which to produce large numbers of MKs and platelets [7, 10–12]. Improvement in the number of platelets generated per MK, combined with adoption of novel bioreactor technology, has enabled the generation of sufficient quantities of platelets to begin phase I safety trials in humans [6]. Strategically modifying the iPSC genome has the additional potential to improve both the yield and function of in vitro generated platelets and platelet-like-particles, however culturing constitutive gene deleted iPSCs before undertaking their differentiation into blood cells has the potential to skew lineage selection, block differentiation altogether, and cause cellular transformation. To study the effect of gene disruption at the desired phase of blood cell differentiation, we adopted an approach that permits precise temporal control of gene expression and deletion.

Lyn is a member of the Src family of tyrosine kinases (SFKs). The roles of Lyn in MK and platelet function have to date only been studied in mice expressing or lacking this enzyme. Disruption of Lyn in mice has been shown to lead to mild thrombocytosis and megakaryopoiesis [13]. Additionally, Lyn-deficient (KO) MKs show higher ploidy[14] for reasons that are not entirely understood. Finally, overexpression of a kinase-inactive form of Lyn in a B cell line expressing Mpl (BaF3/Mpl) improved cell proliferation [14].

Lyn appears to exert somewhat complex, time-dependent functions during the platelet activation process. Thus, while early activation responses to the glycoprotein VI (GPVI) agonist collagen-related peptide (CRP) are delayed due to disruption in early immunoreceptor tyrosine-based activation motif (ITAM)-mediated signaling, the late signaling responses such as the degree of maximal platelet aggregation and secretion are potentiated due to lack of negative feedback regulation by PECAM-1/SHP-2 complexes [15, 16]. In contrast, Lyn appears to only minimally affect responses to thrombin-mediated signaling stimulated by G protein-coupled receptor (GPCR) PAR1/PAR4 agonists.

Despite the above studies in mice and the use of moderately specific pharmacological inhibitors, there are no studies that specifically target Lyn in human MKs or platelets. The purpose of the present investigation therefore was to employ a conditional deletion strategy to examine the hypothesis that temporally controlled disruption of this SFK in hematopoietic progenitor cells (HPCs) might enhance MK maturation, improve activation responses to selected agonists, and improve platelet formation from iPSC-derived MKs. Adoption of this approach to conditionally delete genes known to regulate platelet production and function has the potential to facilitate the eventual development of in vitro derived platelets for transfusion therapy.

Materials and Methods

Reagents

FITC-conjugated mouse anti-human CD42a, PAC-1, and PE-conjugated CD41a, and APC-conjugated CD235a were from BD Biosciences (San Jose, CA). Antibodies against Akt-pTyr308 and ERK-pThr202/Tyr204 were from Cell Signaling Technology (Danvers, MD). Antibodies against Lyn was from Santa Cruz Biotechnology, Inc. (Dallas, TX). Antibody against human GPVI was from Blood Research Institute Hybridoma Core Laboratory. Horseradish peroxidase-conjugated secondary antibodies were from Jackson Immuno Research (West Grove, PA). Human fibrinogen was from Enzyme Research Laboratories (South Bend, IN). CRP was prepared by the Blood Research Institute Protein Core Laboratory. Recombinant human SCF and TPO were from R&D systems (Minneapolis, MN). mTeSR1 media and EasySep FITC positive selection kit were from STEMCELL Technologies (Vancouver, BC, Canada). All other reagents were from Sigma-Aldrich (St Louis, MO).

Construction of donor plasmids and sgRNAs

For FRT knock-in, a donor plasmid containing LoxP flanked puromycin resistance gene (pL552) was obtained from Addgene (Addgene plasmid #68407, Watertown, MA) [17]. DNA fragments with a 1 kb Lyn homologous recombination (HR) arm with FRT sites, as described in Fig. 2, were synthesized by gBlock (IDT, Coralville, IA, USA). DNA fragments were then cloned into the pL552 donor plasmid using multiple restriction enzyme sites. For sgRNA construction, sgRNAs designed to surround exon 4 of Lyn were cloned into pX459 V2.0 (Addgene plasmid #62988) [18]. Cas9 cleavage sites used in this study were sgRNA1: 5’- AGACCCTGAGCCCAC-TAGGC AGG and sgRNA2: 5’- TCGGTCTGTGAGCACACTTC AGG.

Figure 2. Generation of inducible Lyn knock-out iPSCs.

(A) Schematic of CRISPR/Cas9-mediated FRT knock-in to the Lyn locus. Key features of the donor plasmid include Lyn exon 4 flanked by FRT sites and a puromycin-resistance cassette flanked by LoxP sites for later removal by Cas9-Cre. PCR genotyping of FRT knock-in human iPSC clones show that the expected PCR product for the targeted Lyn locus of 1950 bp (red arrow). Clones containing the properly targeted allele were checked for homozygosity by PCR and were either heterozygous (814 bp PCR product – blue arrow) or homozygous (3876 bp product – not shown). (B) Clones containing the FRT/Lyn exon4/puromycin knock-in in both alleles of the Lyn locus underwent a second round of CRISPR/Cas9-mediated gene editing to insert a Flpe-ER^T2 cassette containing the neomycin resistance gene into the AAVS1 locus. The expected PCR product for a properly targeted AAVS1 locus is 641 bp (red arrow). (C) Genotyping strategy for Cre recombinase-mediated puromycin resistance cassette removal from the Lyn locus. Clones lacking the puromycin cassette (963 bp product – blue arrow) were further examined by PCR for homozygosity (no PCR product) or heterozygosity (1950 bp band – red arrow).

Human iPSC culture

The OT1–1 human iPSC line was generated as previously described [19]. Human iPSCs were maintained on Matrigel (Corning, Corning, NY) in Stemflex media (Thermo Fisher Scientific, Waltham, MA). Medium was changed daily, and cells were passaged using ReLeSR (STEMCELL Technologies). Cells were cultured at 37°C with 4% O₂/5% CO_2.

Electroporation

Electroporation was performed using an Amaxa 4D-Nucleofector (Lonza, Morristown, NJ) using program CB-150. To improve survival, 10 μM Rho-associated protein kinase (ROCK) inhibitor Y27632 (StemRD Inc., Burlingame, CA) was added to the transfected cells. For FRT knock-in experiments, sgRNA expressing pX459 V2.0 plasmids and donor plasmid were used for nucleofection. Two days after electroporation, puromycin (0.5 μg/mL, Gibco, Gaithersburg, MD) was added to the medium for selection for 7–10 days. For Flpe-ERT2 knock-in and puromycin-resistant cassette removal, Cas9-2A-Cre, AAVS1-neo-CAG-FlpeERT2 (Addgene plasmid #68468 and #68460) [17], and sgRNA T2 (Addgene plasmid #41818) [20] were used for nucleofection. Two days after nucleofection, cells were treated with G418 (50 μg/mL, Corning) for 7–10 days and further treated with 100 μg/mL G418 for additional 10 days.

MK differentiation

Human iPSCs were differentiated into HPCs using the previously described protocol [19]. Cells were plated on Matrigel for differentiation and the medium and cytokine changes were followed as described. Cells were cultured at 37°C with 4% O₂/5% CO₂ for 9 days, and HPCs were collected. Cells were analyzed by flow cytometry to confirm the surface expression of CD41a and CD235a (BD Bioscience). The HPCs were further differentiated to MKs in serum-free differentiation medium, Iscove’s Modified Dulbecco’s Medium (Thermo Fisher Scientific) containing 25% Ham’s F-12 (Corning), 0.5% N2 (Thermo Fisher Scientific), 1% B27 without vitamin A (Thermo Fisher Scientific), 0.05% bovine serum albumin (BSA; Sigma), 2 mM l-glutamine, and penicillin /streptomycin supplemented with 50 ng/mL SCF and 50 ng/mL TPO at 37°C, 5% CO₂ for 6–7 days. Tamoxifen was added at day 0 of MK differentiation for 24 h to disrupt Lyn unless stated otherwise. HPC proliferation was analyzed by counting cell number from day 0–6 of MK differentiation.

Genomic DNA extraction and genomic PCR

Cells were lysed with lysis buffer [10 mM Tris-HCl (pH 7.5), 10 mM EDTA, 10 mM NaCl, 0.5% SDS, 10 μg/mL proteinase K] at 60°C. Genomic DNA was precipitated by adding ×3 volume precipitation buffer (150 mM NaCl in EtOH). Genomic PCR was done using Herculase II Fusion DNA polymerase (Agilent, Santa Clara, CA).

Western blotting

Cells were lysed by adding an equal volume of ice-cold 2×lysis buffer [300 mM NaCl, 20 mM Tris, 2 mM EGTA, 2 mM EDTA, and 2 % NP40 (pH 7.5)]. The samples were diluted with an equal volume of 2×sample buffer [4 % SDS, 10 % 2-mercaptoethanol, 20 % glycerol, and 50 mM Tris (pH 6.8)], separated by SDS-PAGE, and transferred to a poly (vinylidene difluoride) membrane. Western blotting was performed with the indicated antibodies. Bands were quantitated by densitometry using IMAGE J software (NIH, Bethesda, MD).

MK flow cytometry

MK surface receptor expression was measured at the indicated time points, following staining with FITC-CD42a, PE-CD41a, and APC-GPVI using a BD Accuri^™ C6 plus (BD Biosciences). The viable MK fraction was gated based on FSC vs SSC. Data were analyzed using FlowJo software (Tree Star, Ashland, OR). The extent of agonist-induced MK activation was assessed by staining agonist-stimulated MKs with an activation-specific marker (FITC-conjugated PAC-1, BD Biosciences). MKs were stimulated with CRP or thrombin for 20 min at 37°C, fixed with 5% neutral buffered formalin (NBF, Thermo Fisher Scientific) and analyzed using the BD Accuri^™ C6 plus.

Ploidy analysis

Cells at day 7 of MK culture were collected and stained with FITC-CD42. Cells were then fixed with 5% NBF and washed. Cells were resuspended in PI staining buffer containing 0.05 mg/mL propidium iodide (PI, Invitrogen, Waltham, MA), 0.1% Triton X, and 100 μg/mL RNAse and incubated for 15 min and analyzed using the BD FACSCelesta^™ Cell Analyzer (BD Biosciences).

Blood collection

Ethical approval was obtained from the institutional review board of the Medical College of Wisconsin. All study members provided informed, written consent to participate. Blood samples from healthy volunteers free from medication for two weeks was collected into 3.2% sodium citrate.

Rotational thromboelastometry (ROTEM) analysis

Whole blood was collected from healthy adult volunteers into 3.2% sodium citrate. Thrombocytopenic blood was prepared as previously described [21]. Hemostasis in human whole blood at the described platelet count was measured by ROTEM delta (Instrumentation Laboratory, Werfen, Bedford, MA). Mature MKs purified by its CD42a expression were added to the ROTEM reaction at the cell number described.

Mature MK purification

iPSC-derived MKs cultured in TPO for 6–7 days were collected and resuspended in a buffer (PBS, 0.5% BSA, 5 mM EDTA). Cells were stained with FITC-conjugated CD42a antibody and CD42-positive cells were purified using EasySep FITC positive selection kit II (Stemcell Technologies).

MK spreading

Eight-chamber glass tissue-culture slides (Corning) were coated with 1% bovine serum albumin (BSA), 50 μg/mL cross-linked CRP or 30 μg/mL fibrinogen at 4 °C overnight. Wells were blocked with 1% BSA. Cultured MKs were allowed to spread for the indicated times at 37 °C. Adherent MKs were fixed with 5 % neutral buffered formalin (NBF) and permeabilized with 0.1 % Triton-X100. The fixed MKs were stained with phalloidin-TRITC (10 μg/mL, Sigma) and DAPI. Images were taken with an inverted microscope (Nikon, Melville, NY) with a ×100 or ×60 oil immersion lens. MK spreading area was analyzed using NIS-Elements software (Nikon) and results reported as the mean area of spread MKs (μm²/MK). Proplatelet forming MKs were identified as cells displaying at least one filamentous extension. In Fig. 5c, proplatelet forming MKs on to different substrates were calculated as the percentage over the total MK population adhering to the substrate in each field.

Figure 5. Lyn-KO MKs exhibit enhanced proplatelet formation on substrates for both GPVI and integrin αIIbβ3.

Wild-type (WT) and Lyn-deficient (Lyn-KO) MKs at day 7 of differentiation were plated on fibrinogen- or CRP-coated cover slips at 37°C for 3 hours. Adherent cells were fixed and stained for immunofluorescence analysis with TRITC-phalloidin (white) and DAPI (blue). (A) Images of spread MKs that are representative of five independent experiments are shown. Bar represent 10 μm. Non-spreading cells are shown in green arrows, spreading MKs are shown in orange arrows, and proplatelet forming MKs are shown in red arrows. (B) The average spreading area of individual MKs was quantified from 100–120 MKs on (i) fibrinogen- and (ii) CRP-coated surfaces. Spreading area is reported as the mean ± SEM. (C) Percentage of proplatelet forming MKs on (i) fibrinogen (30 μg/mL) and (ii) CRP (50 μg/mL) were analyzed and reported as the mean ± SEM. (D) (i) Representative FSC vs SSC and CD41 vs CD42 from human platelet-rich plasma (PRP) and platelet-like particle (PLP) from MK culture media. WT and Lyn-KO MKs at day 4 of differentiation were analyzed for PLP count for 1–4 days. PLPs were defined by FSC/SSC and CD41/CD42 expression. (ii) relative difference in number of PLP obtained from Lyn-KO MKs compared to WT MKs and (iii) number of CD41⁺CD42⁺ PLP in culture are shown as the mean ± SEM (n=7). Statistical analysis was performed using the unpaired t test. *P<0.05, **P<0.01, ****P<0.0001 of Lyn-KO, ns= not significant as compared to WT.

In vitro release of platelet-like particles

MKs from day 4 MK culture were seeded on 48-well plates (10⁴ cells /200 μL per well. The cultured cells were collected at the indicated time points from day 1 to day 4. Cells were stained with FITC-CD42 and PE-CD41 for 20 min at RT and analyzed with the BD FACSCelesta^™ Cell Analyzer. Cells were gated based on human platelet-rich plasma (PRP) FSC/SSC and CD41⁺CD42⁺ platelet number was counted. CountBright absolute counting beads (Invitrogen) was added to the samples to obtain the absolute number of CD41⁺CD42⁺ cells per microliter.

Statistics

All data expressed as the mean ± standard error of the mean (SEM). Statistical analyses were performed by the unpaired t-test using Prism 8.4.3 software (GraphPad, San Diego, CA).

Results

One or more Src family kinases function as negative regulators of MK differentiation

To better understand the role of SFKs in human MK differentiation, we treated iPSC-derived HPCs with the SFK inhibitor PP2 during the course of their differentiation into mature MKs. As shown in Figure 1a, the CD41⁺CD42⁺ mature MK population increased over time. Interestingly, PP2-treated cells consistently produced more mature MKs in culture, suggesting that one or more SFKs act as a negative regulators of MK differentiation. Inhibition of global SFK activity also resulted in increased responsiveness to TPO-stimulation, as assessed by ERK and Akt phosphorylation in PP2-treated cells (Fig. 1b). To identify the specific SFK responsible for negatively regulating MK differentiation and activation, we first examined the potential role of Lyn, which is not only one of the most abundant SFKs in MKs and platelets [22, 23], but also functions as a negative regulator of GPVI-mediated platelet activation [16, 22].

Figure 1. Inhibiting Src family kinase activity improves the rate and extent of megakaryocyte differentiation.

Human OT1–1 iPS cells were differentiated into hematopoietic progenitor cells (HPCs) and then further differentiated into megakaryocytes (MK) in culture media supplemented with TPO and SCF. (A) HPCs were cultured in the presence or absence of 10 μM PP2 at Day 0 of MK differentiation, and the cells were analyzed for CD41 and CD42 expression by flow cytometry for six days (left). The percentage of mature CD41⁺CD42⁺ MKs in culture is reported as the mean ± SEM (n=5–7) (right). Statistical analysis was performed using the unpaired t test. *P<0.05, **P<0.01 of PP2 treated cells compared to control cells. (B) MKs at Day 7 were collected and stimulated with 100 ng/mL TPO for the indicated time. Lysates were analyzed for phospho-ERK (Thr202/Tyr204), phospho-Akt (Thr308), and actin. Blot shown is representative of three independent experiments.

Generation of a Lyn-inducible KO (iKO) iPS cell line

Previous studies on Lyn-KO mice have shown that disruption of Lyn leads to increased numbers of myeloid progenitors, splenic myeloid cells, peripheral blood monocytes, and immature MKs [13, 24, 25]. To avoid potential undesirable effects on differentiation, we adopted a strategy originally developed by Chen et al. [17] that allows inducible disruption of protein expression in a temporally controlled manner. To disrupt Lyn protein expression, we decided to delete exon 4 in a temporally controlled manner based on the Lyn conditional KO mice in the International Mouse Phenotyping Consortium (IMPC) data base (www.mousephenotype.org). As depicted in Figure 2a, we employed CRISPR/Cas9 gene editing technology to insert a cassette containing exon 4 of Lyn flanked by short flippase recognition target (FRT) sequences to induce flippase (Flp)-dependent deletion and a loxP-flanked puromycin-resistance gene into iPS cells. Gene edited cells were genotyped to confirm homozygous FRT knock-in in the Lyn allele (Fig. 2a). Of twenty clones obtained, six were found to contain homozygous insertion of the targeted FRT-containing cassette on both Lyn alleles. These iPSC clones were then subjected to a second round of CRISPR/Cas9 gene editing to insert between exons 1 and 2 of the AAVS1 locus, a safe harbor for hosting DNA transgenes that is on human chromosome 19, an expression cassette encoding neomycin resistance, and Flp recombinase fused to a tamoxifen-responsive form of the human estrogen receptor ligand-binding domain (Flpe-ER^T2) (Fig. 2b). Transfection of correctly targeted clones with a plasmid expressing Cre-Cas9 resulted in removal of the puromycin-resistance cassette from the Lyn locus (Fig. 2c). Thus, addition of tamoxifen at the desired time to the final Lyn-iKO clones should in theory activate the Flpe-ERT2, resulting in removal of Lyn exon 4 surrounded by FRT sites and disrupt Lyn protein expression.

Tamoxifen treatment efficiently disrupts Lyn protein expression in Lyn iKO cells

To analyze the efficiency of gene disruption in the established Lyn iKO lines, cells were treated with 4-hydroxytamoxifen (4-OHT). A pair of genotyping primers were used to validate deletion of exon 4 from the Lyn locus (Fig. 3a). As shown in Figure 3b, exon 4 was lost from genomic DNA within 24 hours of 4-OHT treatment, whereas Lyn protein largely disappeared by 48 hours, with complete loss by 72 hours post 4-OHT treatment. Disruption of Lyn DNA/protein was dose-dependent (Fig. 3c), requiring only 100 nM 4-OHT for deletion of exon 4 and loss of Lyn protein expression. These results demonstrate that Lyn expression can be efficiently disrupted in a temporally controllable manner within 72 hours after addition of 4-OHT.

Figure 3. Tamoxifen-inducible disruption of Lyn.

(A) Schematic of the PCR primer set for genotyping the gene-edited Lyn locus. (B) Time-dependent deletion of Lyn DNA (left) and protein (right) after addition of 2.5 μM tamoxifen (4-OHT) to the culture media. Depletion of Lyn protein and the FRT-flanked exon 4 was quantified. Data are represented as mean ± SEM (n=3–5). (C) Dose-dependent deletion of Lyn DNA and protein after 4-OHT treatment at the indicated concentrations 72 hours after treatment. Red arrow depicts the intact PCR product, while the blue arrow indicates the size of the exon 4-deleted PCR product.

Lyn deficiency promotes MK maturation and proliferation

Previous studies have shown that Lyn-KO mice have a mild increase in immature Mpl⁺CD41⁺ MKs in their bone marrow and spleen [13]. To see whether MK differentiation and/or maturation are similarly affected by Lyn disruption in human cells, we differentiated Lyn iKO iPSCs to HPCs and treated them with 4-OHT at the start of MK differentiation. As shown in Figure 4a, addition of 4-OHT to HPCs resulted in an increase in mature CD41⁺CD42⁺ MKs starting from day 2 of differentiation, compared to untreated cells, indicating that disrupting Lyn expression can enhance the efficiency of MK differentiation. There were no changes in the expression levels of cell surface CD41, CD42, or GPVI between wild-type (WT) and Lyn-KO mature MKs (Supplemental Fig. 1). Lyn-KO cells started to proliferate earlier compared to WT cells, but overall cell number was comparable (Fig. 4b). This is consistent with the previous report that kinase inactive Lyn promotes cell proliferation [14]. Lyn-KO HPC and immature MKs proliferate better than WT cells, but mature MKs do not proliferate, resulting in overall comparable cell number in WT and Lyn-KO cells. The increase in MK maturation occurred when 4-OHT was added to the cells up to day 3 of MK differentiation, but adding 4-OHT after day 4 did not lead to further improvements in MK differentiation rate (Supplemental Fig. 2a,b), suggesting that disruption of Lyn promoted early MK differentiation. 4-OHT itself did not have any effect on MK differentiation in WT cells (no genetic modification) as shown in Supplemental Figure 2c. Addition of 4-OHT resulted in loss of Lyn protein expression beginning on 48 hours of 4-OHT treatment and was virtually complete by 72 hs regardless of the MK differentiation state (Fig. 4c). Disruption of Lyn did not have any effect on the expressions of other Src family kinases (Supplemental Fig. 2d). Finally, consistent with observations in Lyn-KO mouse MKs, we saw a significant increase in the ploidy of Lyn-KO MKs (Fig. 4d). These results demonstrate that Lyn-deficiency enhances MK differentiation and maturation.

Figure 4. Tamoxifen-inducible disruption of Lyn improves megakaryocyte differentiation.

Inducible Lyn knock-out (iKO) iPS cells were differentiated into hematopoietic progenitor cells (HPCs) and then further differentiated into megakaryocytes (MK) in culture supplemented with TPO and SCF. Tamoxifen (4-OHT) 2.5 μM was added to disrupt Lyn expression (Lyn-KO) at the start of MK differentiation. (A) HPC differentiation was analyzed by flow cytometry for six days after the start of MK differentiation (left). The percentage of mature CD41⁺CD42⁺ MKs in culture is reported as the mean ± SEM (n=5–7) (right). (B) Cell proliferation during MK differentiation. Total cell number during differentiation is reported as the mean ± SEM (n=6–8). (C) MKs were collected 0, 24, 48, 72 hours after 4-OHT addition. Lysates were analyzed for Lyn and actin. Blot shown is representative of three independent experiments. (D) Ploidy distribution of WT and Lyn-KO MKs at day 7 of differentiation is reported as the mean ± SEM (n=5). Statistical analyses were performed using an unpaired t test. *P<0.05, **P<0.01, ***P<0.001 of Lyn-KO cells compared to WT cells.

Lyn-KO MKs exhibit enhanced formation of proplatelets and platelet-like particles

To investigate their relative ability to produce platelets, Lyn-KO MKs were plated on coverslips that had been coated with immobilized CRP or fibrinogen and proplatelet formation was assessed three hours post-plating (Fig. 5a). As shown in Figure 5b and 5c, although Lyn-KO MKs spread to a similar extent as WT MKs on fibrinogen, the percentage of proplatelet forming MKs was increased. The same was true for MKs plated on CRP-coated surfaces. It has been previously reported that the collagen—GPVI interaction has an inhibitory effect on MK proplatelet formation, and this could be reversed by PP2 treatment [26]. Consistent with this, we also saw an increase in proplatelet forming MKs on collagen (Supplemental Fig. 3). Furthermore, PLP release into MK culture media was analyzed to measure platelet productivity in Lyn-KO MKs. Lyn-KO MKs consistently produced more PLP in the culture compared to WT MKs (Fig. 5d). These results demonstrate that Lyn disruption promotes MK differentiation and platelet production.

Lyn-deficiency improves the hemostatic effectiveness of in vitro-derived MKs

Because MKs and platelets share many of the same cell surface receptors, signaling components, and cytoskeletal elements [27, 28], MKs have often been used as a reliable surrogate for evaluating the functionality of platelets that would be generated from them [29]. To determine whether Lyn deficiency might impart improved functionality, WT and Lyn-KO MKs were compared for their ability to mediate hemostasis, as measured in vitro using rotational thromboelastometry (ROTEM). Lyn-KO and WT MKs were added to thrombocytopenic blood prepared according to previously reported methods [21]. As shown in Fig. 6, addition of iPSC-derived MKs markedly enhanced maximum clot firmness (MCF) and clot firmness over time while decreasing the clot formation time (CFT) of thrombocytopenic (TCP) blood in a dose-dependent manner. Figure 7 shows that, compared to WT MKs, addition of Lyn-KO MKs further enhanced the MCF and increased the rate of clot formation. Taken together, these data demonstrate that Lyn functions as a negative regulator of hemostasis in human iPSC-derived MKs, and that cells lacking this abundant SFK exhibit improved hemostatic characteristics.

Figure 6. Human iPSC-derived megakaryocytes support ex vivo hemostasis.

(A) Rotational thromboelastometry (ROTEM) was performed using (i) whole blood (WB), (ii) thrombocytopenic blood (TCP) adjusted to 0.2×10⁵ platelets/μL, and (iii) TCP supplemented with 2×10⁵ human iPSC-derived MKs at day 7 of differentiation. The pink area shows the time to reach clot firmness of 20 mm. (B) Clot firmness over time, (C) clot formation time (CFT), and (D) maximum clot firmness (MCF) are reported as the mean ± SEM (n=3–6). Statistical analysis were performed using an unpaired t test. *P<0.05, **P<0.01, ***P<0.001 of MK-supplemented blood, as compared to TCP blood.

Figure 7. Lyn-deficiency improves the hemostatic effectiveness of in vitro-derived MKs.

(A) Rotational thromboelastometry (ROTEM) was performed using (i) whole blood (WB), (ii) thrombocytopenic blood (TCP) adjusted to 0.2×10⁵ platelets/μL, and (iii) TCP supplemented with 2×10⁵ wild-type (WT) or (iv) Lyn-deficient (KO) MKs at day 7 of differentiation. The pink area shows the time to reach clot firmness of 20 mm. (B) Clot firmness over time, (C) clot formation time (CFT), and (D) maximum clot firmness (MCF) are reported as the mean ± SEM (n=4–9). Statistical analysis were performed using an unpaired t test *P<0.05 of TCP supplemented with Lyn-KO MKs, as compared to WT.

Lyn-deficiency enhances MK activation in response to TPO and GPVI agonist

Previous studies have shown that MK differentiation is predominately controlled by thrombopoietin (TPO) and its downstream signaling pathways [30]. To examine whether enhanced TPO signaling might be contributing to the enhanced MK differentiation (Fig. 4) and hemostatic effectiveness (Fig. 7) in Lyn-KO MKs, we compared the activation of TPO signaling molecule in WT and Lyn-KO MKs. As shown in Figure 8a, Lyn-KO MKs exhibited significantly sustained TPO-induced ERK phosphorylation than did WT cells, providing a potential mechanistic explanation for the effects of Lyn deficiency on the processes described above. TPO stimulation leads to activation of the Jak/STAT pathway as well, but we did not see any difference in STAT3 phosphorylation in Lyn-KO MKs, suggesting that Lyn is not involved in activation of this pathway (Supplemental Fig. 4).

Figure 8. Lyn deficiency enhances TPO and GPVI agonist stimulation.

Wild-type (WT) and Lyn-deficient (Lyn-KO) MKs at day 7 of differentiation were stimulated with 100 ng/mL TPO for 10, 30 and 60 minutes. Lysates were analyzed for phospho-ERK (Thr202/Tyr204) and actin. (A) Blot shown is representative of six independent experiments. Quantification of band densities normalized to actin over all experiments is reported as the mean ± SEM. (B) PAC-1 binding as a reporter of integrin αIIbβ3 activation was measured by flow cytometry in MKs at day 7 of differentiation after 20 min of stimulation with thrombin (left) or CRP (right) at the indicated concentrations. Values represent the mean ± SEM of PAC-1 positive cells (%) (n=3–10). Statistical analyses were performed using the unpaired t test. *P<0.05 and **P<0.01 of Lyn-KO, as compared to WT MKs.

Finally, because Lyn has been shown to function as a negative regulator of collagen-induced activation of murine platelets [22], we analyzed whether it might have similar inhibitory properties in human cells. As shown in Figure 8b, whereas there was no difference in their responsiveness to thrombin, which operates via a PAR-1- and PAR-4-mediated G-protein-coupled signaling pathway, Lyn-KO MKs were significantly more responsive to CRP, which signals via the ITAM-coupled GPVI/FcRγ-chain complex as detected by PAC-1 binding after stimulation. These data indicate that Lyn, as in murine platelets, functions as a negative regulator of ITAM signaling in human cells of the MK lineage.

Discussion

Although the role of Lyn has previously been studied in mouse platelets and MKs, there are no reports on the function of this abundant SFK in human MKs. By taking advantage of CRISPR/Cas9 technology and in vitro iPSC→MK differentiation methods, we generated temporally controllable, inducible Lyn-KO MKs that allowed us to examine the role of Lyn in MK differentiation, activation and platelet formation. Importantly, addition of tamoxifen was able to efficiently disrupt Lyn expression at any stage of the iPSC→MK differentiation process, and we found that deleting Lyn expression at an early stage of differentiation significantly increased the rate of differentiation into mature MKs; an observation that might be a useful addition to currently developing forward programming in vitro platelet generation protocols [31].

In vitro platelet production is still a developing field, as is the ability to rapidly and faithfully predict the relative efficacy of genetically modified iPSC-derived MKs and their progeny to support hemostasis. As a surrogate for in vivo studies, we adopted and modified a strategy, first reported and validated by Lyde et al. [32], of evaluating the functionality of human iPSC-derived MKs using ROTEM. By comparing their ability to enhance the rate and extent of clot formation in thrombocytopenic human blood, we were able to show that Lyn normally functions as a net negative regulator of hemostasis, and that conditional deletion of this SFK results in improved hemostatic effectiveness of MKs, and presumably their daughter platelets. We expect that this strategy will be useful for evaluating the role of other presumed negative regulators of platelet function in human cells – a field previously and necessarily dominated by studies in mice, as using this methodology allows one to assess the overall hemostatic function of numerous genes of interest before resorting to time-consuming, cumbersome, and expensive animal models.

The function of MEK-ERK in TPO-induced MK differentiation is controversial. While phosphorylation and activation of ERK has been shown to be important for the differentiation of mouse MKs [33, 34], its role in the differentiation of human primary MKs remains less certain. Studies examining the differentiation of human cord blood-derived CD34⁺ cells found that MEK negatively regulates TPO-induced MK polyploidization and proplatelet formation [35, 36], while others observed that ERK activation actually promotes MK differentiation and polyploidization [37–40]. The reason for this discrepancy is unclear, but could be due to the use of different MEK inhibitors and/or the use of cells from different sources. In this study, we found that knocking-out Lyn in human iPSC-derived MKs resulted in sustained TPO-induced ERK phosphorylation, an event that correlated well with enhanced MK differentiation. Although it is possible that Lyn is affecting other pathways to regulate MK differentiation, our results are consistent with ERK functioning as a positive regulator of MK differentiation and activation.

The mechanism by which Lyn affects ERK activation under TPO stimulation is even less clear. Consistent with our findings, knocking out Lyn in mouse MKs has been shown to result in sustained ERK activation downstream of TPO stimulation [13]. One speculation for the enhanced ERK activation in Lyn-KO MKs is that Lyn might recruit, or be recruited by, known negative regulator of TPO signaling, like SHP-1/SHP-2 [41], although SHP-2 is also known as a positive regulator for Ras and its role could be agonist- and cell-type dependent. Along these lines, it has been reported using the c-Mpl–expressing Ba/F3 cell line that the TPO receptor becomes phosphorylated at tyrosine 591 upon TPO stimulation, which recruits SHP-1, Syk, and Btk and that point mutation of tyrosine 591 results in enhanced ERK phosphorylation [41]. Whether or not Lyn is a component of this signaling complex to negatively regulate TPO-induced MK activation and differentiation remains an intriguing avenue of future investigation.

Finally, although recent technological improvements in cell culture technology have enabled several labs to generate large numbers of MKs in vitro [6, 31], the yield and function of the resulting platelets and platelet-like particles leaves much room for improvement. In this study, we genetically modified the iPSC genome to inducibly disrupt Lyn expression with the goal of improving both the yield and the hemostatic effectiveness of the resulting MKs/platelets. We also observed a specific upregulation of GPVI-, but not thrombin-, mediated MK activation, suggesting that transfusion of Lyn-KO MKs/platelets should exhibit improved functionality without undesirable thrombus formation. By incorporating such functional improvements together with precise cell surface alloantigen matching [42, 43] and engineered loading of platelet alpha granules with proteins that promote hemostasis and wound healing [19], in vitro-derived platelet products have the potential to be used as novel therapeutics in a wide variety of applications where hemostatic effectiveness is urgently needed.

Essentials.

• The role of Lyn in the differentiation and function of human megakaryocytes (MK) is unknown.

• Inducible knockout cells were generated to study the function of Lyn in human iPSC-derived MKs.

• Lyn negatively regulates MK maturation, platelet-like particle formation and activation.

• Disrupting Lyn might improve yield and effectiveness of the in vitro produced MKs/platelets.