Prospective isolation of mouse and human hematopoietic stem cells using Plexin domain containing 2

doi:10.21203/rs.3.rs-1080470/v1

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Prospective isolation of mouse and human hematopoietic stem cells using Plexin domain containing 2

https://doi.org/10.21203/rs.3.rs-1080470/v1

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Numerous strategies exist to isolate hematopoietic stem cells (HSCs) using complex combinations of markers and flow cytometry. However, robust identification of HSCs using imaging techniques is substantially more challenging which has prompted the recent development of HSC reporter mice. To date, none of the molecules used in these reporters have been useful for human HSC identification. Here we report that PLXDC2 is a useful marker for both mouse and human HSCs. Using a green fluorescent protein (GFP) knock-in at the Plxdc2 locus in mice (hereafter denoted as Plxdc2-GFP), we showed that Plxdc2-GFP is highly expressed in HSCs with 1 in 2.8 Plxdc2-GFP⁺CD150⁺ cells giving long-term multi-lineage reconstitution in transplantation. Moreover, we developed a novel human PLXDC2 antibody and showed that human PLXDC2⁺ HSCs have stronger long-term multilineage reconstitution ability compared with PLXDC2^- HSCs in a xenograft model. Thus, our study identifies PLXDC2 as a highly relevant molecule in HSC identification, potentially allowing greater purity and live in vivo tracking of these cells.

Biological sciences/Immunology/Haematopoiesis

Biological sciences/Stem cells/Haematopoietic stem cells

A key aspect of stem cell biology is the definition of the different phenotypic and functional compartments, especially the most immature compartment, which replenishes all cells within a given tissue. In this regard, hematopoietic stem cells (HSCs) are the most comprehensively characterized tissue stem cell. Over the past three decades, the identification of HSCs has been improved by using both multicolor fluorescence-activated cell sorting analysis (FACS) and the in vivo bone marrow (BM) repopulation assay^{1,2,3,4 5,6,7,8,9,10,11,12}. c-Kit⁺, Sca-1⁺, lineage marker⁻ (KSL) cells represented the first major step in HSC identification, but have been further enriched by additional negative (CD34², Flk2¹³, CD48 and CD41¹⁴) and positive (CD150¹⁴, CD105¹⁵, CD201¹⁶ and Esam1¹⁷,¹⁸) markers. Most of the FACS-based HSC purification methods use the CD150⁺CD34^low/−c-Kit⁺Sca-1⁺lineage marker-negative (Lin–) BM population to purify adult mouse HSCs, which contain cells with long-term multilineage reconstitution and self-renewal ability^19,20,21. However, the CD150⁺CD34^low/−c-Kit⁺Sca-1⁺Lin^– BM cells still contain non-long term HSCs (LT-HSC). Additionally, identification of negatively stained cells is straightforward with FACS analysis but is substantially more complicated with non-quantitative immunofluorescence on fixed and embedded tissue sections despite the urgent need to understand HSC behavior in the context of the in vivo HSC-microenvironment (niche). Several HSC-specific reporters such as Fgd5-mCherry or ZsGreen, Hoxb5-mCherry, and α-catulin-GFP already exist as powerful tools for visualizing HSCs throughout the BM^22,23,24. However, these HSC visualization technologies are not accessible to all researchers because they require mice. Therefore, simple methods using a few antibodies for positive markers are preferable to meet the above requirements.

Human transplantation and xenograft repopulation assays show that human HSCs are CD34⁺, but the majority of CD34⁺ cells are lineage-restricted progenitors and HSCs consist of a minor component of the population. The standard purification strategy to date for human HSCs use the cell surface marker profile Lin⁻CD34⁺CD38⁻CD90⁺CD45RA^{− 11,25,26,27,28} and the most recent purification method for human HSCs uses CD49f in addition to these markers to arrive at a population that is ~ 10% functional HSCs when human cord blood is used as the starting material¹¹. While this is a major milestone, the enriched CD34⁺CD38⁻CD90⁺CD45RA⁻CD49f⁺ population remains more heterogeneous than mouse HSCs, therefore requiring new markers to further purify functional human HSCs.

Our previous screening on HSC-specific genes identified several cell surface genes²⁹. We noted plexin domain containing 2 (Plxdc2) among them. PLXDC2 is a single-pass type I membrane protein which encodes a 350-amino acid long plasma membrane protein. PLXDC2 plays an important role in cell proliferation and differentiation during nervous system development^30,31. PLXDC2 is also known as one of the membrane receptors of pigment epithelial-derived factor (PEDF), an endogenous anti-angiogenesis factor ³². Moreover, PLXDC2 is an activating ligand for ADGRD1 displayed on cumulus cells³³. In addition, PLXDC2 has a regulatory function and a significant impact on modulation of the host immune response against Helicobacter pylori³⁴. However, the function of PLXDC2 in the hematopoietic system remains unknown.

In the present study, we identified PLXDC2 as an HSC purification marker using a Plxdc2-GFP knock-in mouse line ³¹ and found that the combination of two positive markers, Plxdc2-GFP and CD150, provided sufficient purification of functional HSCs without any additional markers. Moreover, PLXDC2 was also highly expressed in human HSCs derived from both cord blood and BM. We showed that the anti-PLXDC2 antibody, clone 4G3 which we previously developed³⁵, was useful for further purification of human functional HSCs from the conventional human HSC fraction.

Plxdc2 is highly expressed in mouse HSCs

Our previous gene expression analysis between CD34⁻ KSL cells as HSCs and other hematopoietic cells identified HSC-specific genes²⁹. In the present study, we studied Plxdc2 as a candidate surface marker for purification of HSCs among them. Plxdc2 is a single pass type I membrane protein which is expressed in tumor vessels but not in most normal tissues. Quantitative reverse transcription PCR revealed that Plxdc2 was highly expressed in CD34⁻KSL cells and CD4⁺CD8⁺ double positive T cells in mouse hematopoietic cells (Fig. 1A). We further confirmed that Plxdc2 was highly expressed in long-term HSCs (LT-HSCs) compared with short term HSCs (ST-HSCs), multipotent progenitors (MPPs) and progenitors (Fig. 1B). Moreover, public gene expression data showed that Plxdc2 was highly expressed in HSCs and CAR cells in mouse BM (Figures S1A-D). Whole immunostaining of BM slice³⁶ showed that Plxdc2-GFP marked HSCs defined by CD31, Emcn and CD150 expression as well as adherent cells (Figure S1C and Movie S1).

To assess Plxdc2 expression in hematopoiesis in detail, we analyzed Plxdc2-GFP knock-in mice³⁷. Both Plxdc2^GFP/+ and Plxdc2^GFP/GFP mice were born and survived into adulthood with expected Mendelian frequencies. Plxdc2^GFP/+ and Plxdc2^GFP/GFP mice showed normal hematopoiesis (Tables S1 and S2). Colony-forming assay of whole BM cells revealed no significant difference between Plxdc2^+/+ and Plxdc2^GFP/GFP littermates in the frequency of lineage-restricted hematopoietic progenitors (Figure S2A). Plxdc2^GFP/+ and Plxdc2^GFP/GFP mice showed normal HSC frequency and normal HSC function upon transplantation into lethally irradiated mice, indicating that Plxdc2 is dispensable for HSC function (Figures S2B and S2C).

Mature hematopoietic cells in the BM from Plxdc2^GFP/+ mice did not express Plxdc2-driven GFP (hereafter referred to as “Plxdc2-GFP”), whereas most of the T cells except CD4⁻CD8⁻c-Kit⁻CD25⁻ cells in the thymus expressed Plxdc2-GFP (Figures S3A and S3B). Overall, just 0.19% of total BM cells in Plxdc2^GFP/+ mice were Plxdc2-GFP⁺ (Fig. 1C). Neither common lymphoid progenitors (CLPs) nor myeloid progenitors (MPs) expressed Plxdc2-GFP (Figure S3C). Plxdc2-GFP expression was highly enriched in KSL cells (9.23%), CD150⁺ KSL cells (43%) and CD34⁻CD150⁺ KSL cells (72.7%) suggesting that it was highly enriched in LT-HSCs (Fig. 1D). Of note, Plxdc2-GFP expression in KSL cells was negatively correlated with CD34 expression and most of the Plxdc2-GFP cells were CD150⁺ (Fig. 1E), further supporting the notion that Plxdc2-GFP⁺ cells are highly enriched in HSCs.

Functional HSC activity is highly restricted to Plxdc2-GFP⁺CD150⁺ cells

To examine HSC potential of Plxdc2-GFP⁺ cells in the BM, 500 Plxdc2-GFP⁺ cells or 2x10⁵ Plxdc2-GFP⁻ cells were transplanted into lethally irradiated mice alongside 2x10⁵ cells of competitor cells (Fig. 2A). Long-term multilineage chimerism was observed in peripheral blood (PB) from Plxdc2-GFP⁺ cell-transplanted mice over 21 weeks, whereas only short-term lymphoid lineage chimerism was observed at low levels in PB from Plxdc2-GFP⁻ cell-transplanted mice (Fig. 2B). Limiting dilution analysis revealed that 1 in 3.53 Plxdc2-GFP⁺ KSL cells gave long-term multilineage reconstitution of irradiated primary recipients (Fig. 2D) and single-cell transplantations showed that 1 in 2.05 Plxdc2-GFP⁺CD150⁺KSL cells gave long-term multilineage reconstitution of irradiated primary recipients. To simply the purification method of HSCs to a two-color strategy, we assessed the combination of Plxdc2-GFP and CD150. Nearly all of the Plxdc2-GFP⁺CD150⁺ cells were also CD34⁻CD150⁺ KSL cells and Plxdc2-GFP⁺CD150⁻ cells were c-Kit⁻ cells (Fig. 2C). Limiting dilution analysis of the Plxdc2-GFP⁺CD150⁺ fraction of whole BM cells revealed that the HSC frequency of Plxdc2-GFP⁺CD150⁺ cells was 1 in 2.8 (Fig. 2D). Single cell transplantation studies revealed that 44.7% (17/38) of Plxdc2-GFP⁺CD150⁺ cells and 48.8% (21/43) of Plxdc2-GFP⁺CD150⁺KSL were functional HSCs capable of long-term multi-lineage reconstitution (Fig. 2D), indicating that Plxdc2-GFP⁺CD150⁺ fraction was roughly equivalent to Plxdc2-GFP⁺CD150⁺KSL fraction. Collectively, these data suggest that Plxdc2-GFP⁺CD150⁺ cells in the whole BM are functional HSCs and therefore the combination of Plxdc2-GFP and CD150 is sufficient to purify functional HSCs in the whole BM.

Plxdc2 stably marks mobilized HSCs.

To further extend analysis for clinical setting, we examined whether Plxdc2-GFP also marked mobilized HSCs in the spleen after cyclophosphamide/granulocyte-colony stimulating factor (G-CSF) treatment. To this end, we administered a single dose of cyclophosphamide on day 0 and constitutive administration of G-CSF for one week from day 1 and then analyzed Plxdc2-GFP expression in mobilized HSCs in the spleen (Fig. 3A). No Plxdc2-GFP⁺ cells were detected in spleen in the steady state condition, whereas 0.11% of cells were Plxdc2-GFP⁺ after administration of G-CSF (Fig. 3B). 8.15% of mobilized KSL cells in the spleen expressed Plxdc2-GFP and CD150. 500 Plxdc2-GFP⁺ cells isolated from mobilized spleen gave long-term multi-lineage chimerism in lethally irradiated mice, whereas 2x10⁵ cells of Plxdc2-GFP⁻ cells in mobilized spleen contained no HSCs (Figs. 3C and 3D). These data indicate that Plxdc2-GFP expression constantly marks HSCs in the mobilization condition.

PLXDC2 is heterogeneously expressed on human HSCs

In order to determine whether PLXDC2 is relevant for human HSCs, we extended our analysis by first assessing its expression using newly established rabbit monoclonal antibody for human PLXDC2³⁵. PLXDC2 expression was highest on the conventional human HSCs (CD34⁺CD38⁻CD90⁺CD45RA⁻CD90⁺CD49f⁺ cells) among hematopoietic stem and progenitor cells (HSPCs) in the cord blood (Figs. 4A and S4A). Interestingly, this expression pattern varied in the BM and G-CSF-mobilized PB HSPCs. MEPs showed highest expression of PLXDC2 in the BM HSPCs, whereas GMPs did in the G-CSF-mobilized PB HSPCs (Figures S4B). Unlike in mouse, PLDXC2 expression was not entirely restricted to the HSC compartment, showing some expression in progenitor cells and CD14⁺ monocytes (Figures S4B, S4C and S4D). Flow cytometric analysis of the human peripheral blood mononuclear cells (PBMCs) showed that PLXDC2 was detected on CD41⁺ cells (12.6%), CD56⁺ cells (2.8%), CD19⁺ cells (13.9%), CD3⁺ cells (3.05%), CD33⁺CD66b⁻ cells (15.5%) and CD14⁺CD15⁺ cells (49.4%) (Figure S4E). In order to test whether PLXDC2 expression tracked with HSC function, we separated human HSCs derived from cord blood into PLXDC2^+/high (referred to here as PLXDC2⁺) and PLXDC2^−/low (referred to here as PLXDC2⁻) and performed functional assays (Fig. 4B). We first evaluated their hematopoietic colony-forming capacity in serial replating assays. PLXDC2⁻ cells formed 1.6-fold more colonies than PLXDC2⁺ cells in the first round. In contrast, PLXDC2⁺ cells formed 1.6-fold more colonies than PLXDC2⁻ cells in the second round of plating (Fig. 4C), suggesting that PLXDC2⁺ cells were more immature than PLXDC2⁻ cells. We next evaluated their capacity for long-term multilineage reconstitution by transplanting them into NOD-scid-IL2Rgc^−/− (NSG) recipients (200 cells of either PLXDC2⁺ HSCs or PLXDC2⁻ HSCs per mice). PLXDC2⁺ cells gave higher human CD45⁺ chimerism in peripheral blood than PLXDC2⁻ cells during the observation period (Fig. 4D). Although levels of chimerism remained high in the peripheral blood of mice transplanted with PLXDC2⁺ HSCs, engraftment of PLXDC2⁻ HSCs peaked at 2 months and then declined. The chimerism pattern of PLXDC2⁻ HSCs indicated that PLXDC2⁻ HSCs were mainly short-term HSCs. Moreover, PLXDC2⁺ cells gave much higher mean chimerism in hematopoietic organs than PLXDC2⁻ cells (Fig. 4E). PLXDC2⁺ cells gave more B cell output (Chimerism of CD19⁺ cells) than PLXDC2⁻ cells (Fig. 4F) whereas erythrocyte output was similar between PLXDC2⁺ and PLXDC2⁻ cells (Fig. 4G). Limiting dilution assays demonstrated that 1 in 49.9 PLXDC2⁺ cells could give rise to long-term multilineage engraftment compared with 1 in 249.1 PLXDC2⁻ cells (P = 0.00348, Fig. 4H). Our HSC frequency (2%) was much less than that shown by the Dick group (10%)¹¹. This discrepancy may be due to differences in experimental conditions such as an irradiation dose and an injection method.

Next, we checked the surface expression of PLXDC2 on HSPCs within HSC culture by flow cytometric analysis to examine whether PLXDC2 is a useful marker within the HSC culture or not. PLXDC2 expression was positively correlated with CD90 and CD201 in 6-day culture (Figure S5A). Furthermore, we compared expression of CD112, recently reported latency marker, between PLXDC2 + and PLXDC2- HSCs. Both HSCs expressed CD112 to the same extent (Figure S6A and S6B). Together, although PLXDC2 was not a specific marker for HSCs in human, PLXDC2 was a useful marker to further purify human HSCs and for the HSC culture.

PLXDC2⁺ HSCs express more HSC signature genes than PLXDC2⁻ HSCs

To resolve PLXDC2 mRNA expression in human HSPCs, we analyzed public single-cell RNASeq data for fresh cord blood CD34 + cells. PLXDC2 was highly expressed in Cluster 1, which were defined as HSPCs by MECOM, MLLT3 and HLF, indicating that PLXDC2 was enriched in human HSPCs (Fig. 5A). To examine transcriptomic differences between PLXDC2⁺ and PLXDC2⁻ HSCs derived from the cord blood, we performed RNA-Seq analysis. Although transcriptomes of PLXDC2⁺ HSCs and PLXDC2⁻ HSCs were highly similar, THBS1, SORBS2, SGSH and SAMHD1 were identified as highly differentially expressed genes (DEGs) in PLXDC2⁺ HSCs (FDR < 0.01), whereas CPA3, ABCA1, CDK6 and IRF8 were highly DEGs in PLXDC2⁻ HSCs (FDR < 0.01) (Figs. 5C and 5D). Interestingly, CDK6 expression has been shown to mark active HSCs in mouse and human studies^38,39,40 and IRF8 is a differential marker for myeloid cells^41,42. The gene expression data therefore suggest that PLXDC2⁻ HSCs express a more differentiated gene signature than PLXDC2⁺ HSCs. Interestingly, although functions of THBS1, SORBS2, SGSH and SAMHD1 in HSCs were not well understood, but their functions were related with important HSC signatures such as active TGFβ signaling, adhesion character, activation of lysosomal function and resistance to viral infection (Table S3). Next, we investigated the biological pathways differentially represented between PLXDC2⁺ HSCs and PLXDC2⁻HSCs using gene set enrichment analysis (GSEA) (Fig. 5E, FDR < 0.1). Again, PLXDC2⁺ HSCs showed an enrichment for the HSC signature. Conversely, we found downregulation of essential biosynthetic processes such as mRNA processing, oxidative phosphorylation and MYC targets in PLXDC2⁺ HSCs compared with PLXDC2⁻ HSCs. Together, these data reinforced the functional data above, indicating that the PLXDC2⁺ fraction contains more cells expressing HSC-related genes than the PLXDC2⁻ fraction.

Our study shows that PLXDC2 is a useful marker for identifying both mouse and human HSCs. Because of the limited availability of human HSCs in the BM or fetal liver and their different purification methods from those of mouse HSCs, features of human HSCs are not well understood. In order to make robust comparisons across species, it is therefore essential to identify common markers. To date, however, only a few common markers, such as EPCR and ESAM, have been reported^{17,43.44,45,46,47}. Interestingly, these markers, including PLXDC2, are all endothelial markers, suggesting that HSCs retain some aspects of an endothelial program, which is perhaps reflective of their origin from endothelial cells during development and may be a useful feature that allows their discrimination from other hematopoietic cells. Therefore, developing common purification methods between mouse and human HSCs using these endothelial markers is useful for furthering our understanding of human HSCs.

PLXDC2 in combination with CD150 is enough to highly purify mouse functional HSCs compared with the conventional multi-antibody combination. 1 in 2.8 Plxdc2-GFP⁺CD150⁺ cells in whole BM were functional HSCs (Fig. 2D). This HSC frequency is equivalent to the frequency reported using α-catulin in combination with c-Kit (1 in 3.1 cells)²⁴ and collectively represent the best two-color strategies to date. That said, PLXDC2 is a more attractive marker for purification of HSCs than α-catulin because PLXDC2 is a membrane protein, whereas α-catulin is an intracellular protein. In this study, we developed a flow cytometry approach using a recently available anti-human PLXDC2 antibody. Unfortunately, this antibody does not recognize mouse Plxdc2 despite homology between mouse and human PLXDC2 being over 90%, suggesting that another antibody in the future may be able to universally detect both mouse and human PLXDC2. One possible reason for this discrepancy is that PLXDC2 has been reported to be a substrate of γ-secretase and β-site amyloid precursor protein (APP) cleaving enzyme 1 (BACE-1), which cleaves the extracellular domains of membrane proteins^48,49. This suggests that mouse Plxdc2 on HSCs may be cleaved by these enzymes and that using inhibitors of these enzymes before purification might be useful in future studies. Nonetheless, developing a flow cytometric analysis-available antibody against mouse PLXDC2 will allow us to highly purify mouse functional HSCs in combination with CD150 and to easily detect mouse functional HSCs without the need for a reporter mouse. Our two-color strategy is also useful for imaging studies. It is hoped that the new antibody will improve the sensitivity and enable HSC detection.

Our study also shows that PLXDC2 further purifies human HSCs from the conventional human CD34⁺CD38⁻CD90⁺CD45RA⁻CD49f⁺ fraction, which are most purified faction for human functional HSCs to date. The finding that further enrichment of functional human HSCs could be achieved by using an anti-PLXDC2 antibody will aid researchers looking to further purify functional human HSCs for cellular and molecular studies. Moreover, PLXDC2⁺ HSCs showed a negative correlation with gene signatures for mRNA processing, upregulation of Myc-regulated genes and oxidative phosphorylation, indicating that PLXDC2⁺ HSCs are likely to be quiescent (Fig. 5C). In addition, the mature cell output ratio between B cells and myeloid cells in PLXDC2⁺ HSCs was B lymphoid dominant, consistent with that of conventional CD34⁺CD38⁻CD90⁺CD45RA⁻CD49f⁺ HSCs. In contrast, repopulation in PLXDC2⁻ HSCs was balanced, which could be explained by higher expression of IRF8 in PLXDC2⁻ HSCs compared to PLXDC2⁺ HSCs.

It has been reported that the timing of exit from quiescence in human HSC subsets is controlled by CDK6 expression levels³⁸. Since PLXDC2⁻ HSCs show higher CDK6 expression than PLXDC2⁺ HSCs, this would further suggest that PLXDC2⁺ HSCs are more quiescent than PLXDC2⁻ HSCs. THBS1, SORBS2, SGSH and SAMHD1 were highly differentially expressed in PLXDC2⁺ HSCs as compared with PLXDC2⁻ HSCs. THBS1 is known to activate latent TGFβ⁵⁰ and active TGFβ is essential for maintenance of HSCs⁵¹. SORBS2 (Sorbin and SH3 Domain Containing 2), also known as ArgBP2, is an adapter protein that can localize to both focal adhesions and actomyosin filaments⁵² and suppresses ovarian cancer metastasis⁵³. Defect of focal adhesion alters maintenance of HSCs^54,55. These features might be involved in adhesion in niche cells and hibernation of HSCs. SGSH encodes N-sulfoglucosamine sulfohydrolase; one of several enzymes involved in the lysosomal degradation of heparan sulfate. SGSH mutations causes the Sanfilippo syndrome, or mucopolysaccharidosis III, which is a lysosomal storage disease due to impaired degradation of heparan sulfate. HSC daughter cells with low levels of lysosomes after asymmetric cell division are induced to differentiate, indicating that lysosomes are important for maintenance of stemness⁵⁶. Therefore, high expression of SGSH in PLXDC2⁺ HSCs indicates that lysosome in PLXDC2⁺ HSCs is more active than that in PLXDC2⁻ HSCs. SAMHD1 (The sterile alpha motif (SAM) domain and HD domain-containing protein 1 ) is a potent restriction factor for human immunodeficiency virus^57,58. SAMHD1 is also known as an interferon stimulated gene and exclusively expressed in HSCs⁵⁹. Collectively, high expression of these 4 genes is likely to contribute to maintain the stem cell phenotype in PLXDC2⁺ HSCs.

Taken together, our data show that Plxdc2 is a potent marker for enrichment of both mouse and human HSCs. Until we have common markers across species, we will be limited in our ability to make comparisons between mouse and human HSCs. Therefore, finding of PLXDC2 as a common marker between mouse and human has a great impact in this regard.

Acknowledgements

We thank Shiori Shikata and Akiho Tsuchiya for expert technical assistance. We thank the IMSUT FACS Core Laboratory and the IMSUT Animal Research Center. This work was supported by a Grant-in-Aid for Challenging Exploratory Research (17K19645, Y.T.), KAKENHI grant number (20K07639, Y.K.), a grant from The SENSHIN Medial Research Foundation (Y.T.), a grant from Leukemia Research Fund (S.G.), Tokyo Metropolitan Small and Medium Enterprise Support Center (T.A. and Y.T.) and MRC-AMED joint award (MR/V005502/1) (DG.K. and Y.T). We finally thank Shin-ichi Nishikawa for supervision of starting this study.

Author contributions

Conceptualization and Methodology, Y.T., T.A. and Y.K.; Investigation, Y.T., Y.K., T.K., T.Y., JL.B., and JW.B.; Visualization, C.S., T.S., M.N., Y.K., T.M., T.A.; Writing – Original Draft, Y.T.; Writing – Review & Editing, Y.T., Y.K., DG.K., S.Y., and T.S.; Funding Acquisition, Y.T., T.A., DG.K. and Y.K.; Resources, Y.T., Y.K., S.G., I.L., C.S., T.S., M.N., Y.K., T.M., C.J.O., T.A. and T.M.J.; Supervision, S.N. S.K. T.A. and T.K.

Competing interests

The authors declare no competing interests.

Quantitative PCR

Total RNA was harvested from FACS-purified CD34^-/low KSL cells, CD34⁺ KSL progenitor cells, Gr-1⁺ neutrophils, Mac-1⁺ monocytes/macrophages, B220⁺ B cells, thymic CD4⁺ CD8⁺ T cells, TER119⁺ erythroblasts, or CD41⁺ megakaryocytes. Total RNA was then subjected to one round of in vitro transcription as described in the previous paragraph. Quantitative PCR was performed using a QuantiTect SYBR Green PCR kit (QIAGEN) according to the manufacturer’s protocol. Each sample was normalized to hypoxanthine phosphoribosyl transferase.

Hematopoietic progenitor assays

Colony-forming activity (CFU-C) assays were performed using MethoCult M3434 (Stem Cell Technologies). A total of 2x10⁴ BM mononuclear cells (MNCs) were plated on 35-mm culture dishes and then incubated at 37°C in humidified chambers containing 5% CO₂. Colonies were counted using a dissecting microscope after 10 to 14 days of culture.

Long-term competitive reconstitution assay

Test cells from Plxdc2^+/+, Plxdc2^+/GFP, and Plxdc2^GFP/GFP mice (CD45.2) were mixed with 2×10⁵ CD45.1/CD45.2 adult BM MNCs and injected into adult recipient mice (CD45.1) irradiated at a dose of 9.5 Gy using a Cesium137 GammaCell40 Exactor Irradiator (MDS Nordia). Peripheral blood cells were stained with

FITC-conjugated anti-CD45.1 antibodies, biotinylated anti-CD45.2 antibodies followed by APC-Cy7 streptavidin, PE-conjugated anti Mac-1 and Gr-1 antibodies, APC-conjugated anti-CD4 and -CD8 antibodies, and PE-Cy7–conjugated anti-B220 antibodies. The test cell-derived chimerism was evaluated on a FACSCanto (BD Bioscience) system.

Blood count

Peripheral blood was collected and analyzed on an automated blood cell counter, KX-21 (Sysmex), according to the manufacturer’s instructions.

Human cord blood processing

All experiments using human cord blood (CB) cells were approved by the Ethics Committee at the Institute of Medical Science, the University of Tokyo (approval number: 27-34-1225). Human CB cells were obtained from the Japanese Red Cross Kanto-Koshinetsu Cord Blood Bank (Tokyo, Japan) following an institutional review board-approved protocol. Informed consent was obtained in accordance with the Declaration of Helsinki. Various cord blood samples were pooled and an equal volume of PBS/2% FBS was added prior to layering on Lymphoprep^TM(STEM CELL Tecnologies) in 50mL conical tubes. Tubes were subjected to 30min centrifugation at 800xg followed by careful removal of mononuclear layer and washed with PBS/2% FBS. CD34⁺ cells were enriched by Direct CD34⁺ Progenitor Isolation Kit (Miltenyi Biotec) according to manufacture protocol. CD34⁺ cells were stored in liquid nitrogen.

Cell preparation for cell sorting

CD34⁺ cells were stained with CD45RA (FITC, clone HI100), CD90 (APC/, clone 5E10), CD49f (PE-DazzleTM594, clone GoH3), CD34 (APC-Cy7, clone 581), CD38 (PE-Cy7, clone HIT2), Lineage marker cocktail (BV510, CD3/14/16/19/20//56, clone OKT3/M5E2/3G8/HIB19/2H7/HCD56) (BioLegend) and PLXDC2 (PE/APC, rabbit monoclonal clone 4G3) (Kyokuto Pharmaceutical Industrial Co. Ltd. and Abwiz Bio Inc.) and incubated for 30min at 4℃. Cells were washed with PBS/2% FBS and stained with streptavidin conjugated BV605 (BioLegend) if CD90 biotin-conjugated antibody was used. Cells were subsequently washed and resuspended in PBS/2% FBS prior to sort. Cells were sorted on FACS AriaII (Becton Dickinson) with a 100-micron nozzle operating mode and collected in 1.5mL microtubes.

Xenotransplant assays

NOD.Cg-Prkdc^scidIl2rg^tm1Wjl/SzJ (NSG) (Jackson Laboratory) were purchased. Animal experiments were performed in accordance with institutional guidelines approved by University of Tokyo. Various cell doses of PLXDC2⁺ or PLXDC2^- human HSCs were transplanted into 2.5Gy irradiated NSG mice by tail vein injection.

Assessment of human cell engraftment

Bone marrow, thymus, peripheral blood and spleen from NSG mice were analyzed for human cell engraftment with CD45 (BV421, clone). Cells were stained in PBS/2% FBS and analyzed by flow cytometry (FACSAriaII, Beckton Dickinson). Lineage committed cells in bone marrow, peripheral blood and spleen were analyzed with CD33 (PE, clone), CD19(PE-Cy7, clone HIB19), CD3(FITC, clone OKT3). Thymocytes were analyzed with CD4 (FITC, clone ) and CD8 (PE, clone ). Stem progenitor cells in bone marrow were analyzed with CD34, CD38, CD90, CD45RA, CD49f and PLXDC2. All antibodies used in this study were obtained from BioLegend unless otherwise indicated.

Statics

Statistical analysis was performed by unpaired Student’s t test or two- or one-way ANOVA without correction for multiple comparison (Fisher LSD test). All data are presented as mean ± SEM. Significance levels were set at p* < 0.05, p** < 0.01 and p*** < 0.001. For statistical analysis GraphPad Prism was used.

RNASeq

We performed RNA-Seq as described previously⁶¹ with minor modification. In brief, using 100 sorted cells, the first strand of cDNA was synthesized by using PrimeScript RT reagent kit (TaKaRa Bio Inc.) and not-so random primers. Following the synthesis of the first strand, the second strand was synthesized by using Klenow Fragment (30, _50, exo-; New England Biolabs Inc.) and complement chains of not-so random primers. Using purified double-strand cDNA, the library for RNA-Seq was prepared and amplified using Nextera XT DNA sample Prep kit (Illumina Inc.). These prepared libraries were sequenced on Next-Seq system (Illumina Inc.), according to the manufacturer’s instruction. In addition, each obtained read was mapped to the reference sequence ‘‘GRCh38’’ using CLC genomic workbench v11.0.0 (QIAGEN), and expression levels were normalized and subjected to the statistical analyses based on EdgeR. Transcriptome data were subjected to GSEA using GSEA v3.0.0 software, available from the Broad Institute⁶². All Gene sets were obtained from the database of Broad Institute unless otherwise stated. Principle component analysis, hierarchal clustering analysis and visualization of gene expression or protein level were performed by Multi expression Viewer.

Single-cell RNA-seq

Analysis was performed using Seurat v.4.0 (ref. 47) in R. HLF, MECOM, MLLT3 and PLXDC2 expression in fresh cord blood data was obtained from the Gene Expression

Omnibus (GEO; GSE153370).

BM slice preparation, immunostaining and optical clearing

Methods for 3D imaging of BM were adapted from previously published protocols³⁶. Femurs were isolated from Plxdc2^GFP/+ mice, cleaned and immersed in PBS/2% paraformaldehyde for 6 h at 4 °C, followed by a dehydration step in 30% sucrose for 72 h at 4 °C. Femurs were then embedded in cryopreserving medium (optimal cutting temperature (OCT)) and snap frozen in liquid nitrogen. Bone specimens were iteratively sectioned using a cryostat until the BM cavity was fully exposed along the longitudinal axis. The OCT block containing the bone was then reversed and the procedure was repeated on the opposite face until a thick bone slice with bilaterally and evenly exposed BM content was obtained. Once BM slices were generated, the remaining OCT medium was removed by incubation and washing of the bone slices in PBS 3 times for 5 min. For immunostaining slices were incubated in blocking solution (0.2% Triton X-100, 5% skim milk, 1% bovine serum albumin (BSA), 10% donkey serum in PBS) overnight at 4 °C.

Primary antibody incubations were performed in blocking solution for 3 days at 4 °C, followed by overnight washing in PBS. Secondary antibody stainings were performed for another 3 days at 4 °C in blocking solution but in the absence of BSA to avoid cross-absorption. Immunostained thick femoral slices were successively washed in PBS overnight and incubated in RapiClear 1.52, for a minimum of 6 h, which typically increased imaging depth to 150 μm from the tissue surface without significant loss of signal intensity. The clearing protocol employed is fast, compatible with all fluorescent probes and proteins tested, and preserves the integrity of subcellular structures.

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Plxdc2-GFP abd CD150 mark hematopoietic stem cells
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Prospective isolation of mouse and human hematopoietic stem cells using Plexin domain containing 2

Status:

Version 1

Abstract

Figures

Introduction

Results

PLXDC2 is heterogeneously expressed on human HSCs

PLXDC2⁺ HSCs express more HSC signature genes than PLXDC2⁻ HSCs

Discussion

Declarations

EXPERIMENT AL MODEL AND SUBJECT DET AILS

References

Additional Declarations

Supplementary Files

Status:

Version 1

Prospective isolation of mouse and human hematopoietic stem cells using Plexin domain containing 2

Status:

Version 1

Abstract

Figures

Introduction

Results

PLXDC2 is heterogeneously expressed on human HSCs

PLXDC2+ HSCs express more HSC signature genes than PLXDC2− HSCs

Discussion

Declarations

EXPERIMENT AL MODEL AND SUBJECT DET AILS

References

Additional Declarations

Supplementary Files

Status:

Version 1

PLXDC2⁺ HSCs express more HSC signature genes than PLXDC2⁻ HSCs