Murine hematopoietic stem cell activity is derived from pre-circulation embryos but not yolk sacs

Ganuza, Miguel; Chabot, Ashley; Tang, Xing; Bi, Wenjian; Natarajan, Sivaraman; Carter, Robert; Gawad, Charles; Kang, Guolian; Cheng, Yong; McKinney-Freeman, Shannon

doi:10.1038/s41467-018-07769-8

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Published: 20 December 2018

Murine hematopoietic stem cell activity is derived from pre-circulation embryos but not yolk sacs

Nature Communications volume 9, Article number: 5405 (2018) Cite this article

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Abstract

The embryonic site of definitive hematopoietic stem cell (dHSC) origination has been debated for decades. Although an intra-embryonic origin is well supported, the yolk sac (YS) contribution to adult hematopoiesis remains controversial. The same developmental origin makes it difficult to identify specific markers that discern between an intraembryonic versus YS-origin using a lineage trace approach. Additionally, the highly migratory nature of blood cells and the inability of pre-circulatory embryonic cells (i.e., 5–7 somite pairs (sp)) to robustly engraft in transplantation, even after culture, has precluded scientists from properly answering these questions. Here we report robust, multi-lineage and serially transplantable dHSC activity from cultured 2–7sp murine embryonic explants (Em-Ex). dHSC are undetectable in 2–7sp YS explants. Additionally, the engraftment from Em-Ex is confined to an emerging CD31⁺CD45⁺c-Kit⁺CD41⁻ population. In sum, our work supports a model in which the embryo, not the YS, is the major source of lifelong definitive hematopoiesis.

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Introduction

The embryonic origin of cells that sustain lifelong mammalian hematopoiesis and blood production has long been debated. Resolving this debate is complicated by the emergence of sequential waves of blood cells at distinct sites within the embryo:¹ blood-islands composed of primitive nucleated erythrocytes first appear at E7-E7.5 in the YS. Definitive erythroid-myeloid precursors also emerge from the YS at E8.5. Finally, around E10.5-E11.5, the first definitive HSC (dHSC) capable of reconstituting the hematopoietic system of adult recipients using existing assays are detected and presumably these precursors support lifelong blood production^2,3. The site of origin of these dHSC has been contentious^{2,3,4,5,6,7,8,9,10,11,12,13,14,15,16}. An intra-embryonic origin, concentrated around the para-aortic splanchnopleura (PSp)-derived aorta-gonad-mesonephros region (AGM), is currently the favored model. In contrast, the contribution of YS to the dHSC compartment is controversial¹. Early work implicated the YS blood islands as a source of both primitive-erythroblasts and dHSC;^1,4,5,6,8,15 however later work challenged this hypothesis. In particular, Dieterlen-Lievre and colleagues demonstrated an intra-embryonic origin for definitive hematopoiesis in vertebrates using quail-chick chimeras^7,16. Recent work has formally demonstrated in chicken the presence of bona fide dHSC originating from the embryo aortas but not from the YS, allantois or head¹⁷. An intra-embryonic origin for dHSC in mammals was later supported by studies showing that the first dHSC capable of reconstituting adult recipients are detected in the PSp/AGM region^2,3. Despite these findings, the potential contribution of YS to lifelong hematopoiesis has not been completely excluded^13,14,18,19.

YS-derived and AGM-derived hematopoietic progenitors both arise from hemogenic endothelial (HE) precursors that are mesodermal in origin^{14,20,21,22,23,24,25}. Very few markers have been identified that could potentially distinguish between AGM and YS hematopoietic precursors. The highly migratory nature of blood cells in circulating embryos and the inability of cells isolated from pre-circulation embryos to robustly engraft in transplantation assays, even after ex vivo culture, has precluded definitively addressing if the YS hemogenic endothelium (YS-HE) contributes to lifelong hematopoiesis and the adult dHSC pool^12,26. PSp tissue from pre-circulation embryos generated long-term multi-lineage engraftment while YS did not, but reconstitution was extremely low (1–5%) in these experiments, raising concerns that lower activity present in the YS would have been very difficult to detect¹². Furthermore, PSp-derived reconstitution was only observed in severely immunocompromised recipient mice (i.e., Rag2γc^−/−)¹². Indeed, it has recently been suggested that the YS may be a major embryonic source of dHSC¹⁴. Lineage tracing studies exploiting the high expression of LYVE1 (lymphatic vessel endothelial hyaluronan receptor-1) in the YS and vitelline-endothelium concluded that >40% of adult blood may ultimately derive from these sites in mice¹⁴.

Here, we present a platform that supports the ex vivo development of robust dHSC activity from pre-circulation embryos, allowing us to rigorously interrogate the dHSC-forming potential of both the early embryo and YS. We find that cultured pre-circulatory Em-Ex, but not YS explants (YS-Ex), yield robust dHSC activity. Importantly, this activity in cultured Em-Ex was restricted to an emerging CD31⁺CD45⁺c-Kit⁺CD41⁻ population that also develops in cultured YS-Ex. Additionally, in pre-circulation embryos, we identify LYVE1⁺CD31⁺ aortic endothelial cells, confirming that LYVE1 expression is found outside the YS and vitelline HE at this early stage of development¹⁴. We further demonstrate that pre-circulatory Em-Ex-derived LYVE1⁺ precursors yield robust dHSC activity, indicating that LYVE1 constitutes an early marker of intraembryonic definitive hematopoiesis. In sum, our work strongly supports a model in which the YS is not a major source of lifelong definitive hematopoiesis.

Results

Robust dHSC activity generation from pre-circulatory embryos

Blood circulation is established around 5–7sp (≈E8.5). The first heartbeat is detected at 5sp, a consistent blood flow begins around 7sp and fully functional circulation is established after E10, as evidenced by the uneven distribution of erythroblasts within the embryo vasculature prior to 35sp^27,28. Although dHSC activity has previously been observed from cultured pre-circulation embryos, this activity was low, and only observed after transplantation into Rag2γc^−/− immunocompromised recipients¹². Here, we sought to determine if it might be possible to coax robust dHSC activity from E8-E8.5 concepti.

We collected CD45.2⁺ embryos at E8.0-E8.5 and staged them by number of somite pairs. Concepti were organized into two groups: 2–3sp (pre-circulation) and 5–7sp (peri-circulation). Embryos were separated from the YS and cultured at the air-liquid interface in the presence of SCF, FLT3L, and IL-3, as previously described^29,30,31. We hypothesized that an extended culture period might be required for maturation of dHSC from these early embryos (Fig. 1a). Hence, embryos were cultured for 9–10 days. Cultured Em-Ex were then recovered, dissociated and transplanted into lethally irradiated CD45.1⁺CD45.2⁺ recipients along with 2 × 10⁵ CD45.1⁺ whole bone marrow (WBM) cells for radio-protection (Fig. 1a). Remarkably, both 2–3sp and 5–7sp cultured embryos generated robust dHSC activity. 5–7sp embryos engrafted 9/11 (82%) recipients while 2–3sp embryos engrafted 5/7 (71%) recipients (Fig. 1b). Globally, >15% of recipient peripheral blood (PB) was CD45.2⁺ in 7/18 total recipients at 16 weeks post-transplant. Four recipients displayed >50% CD45.2⁺ PB reconstitution (Fig. 1b). Importantly, most engrafted recipients (11/14) displayed multi-lineage engraftment at 16 weeks post-transplant (Fig. 1b). To further test if Em-Ex developed robust dHSC activity, we performed secondary limiting dilution transplants. 0.8 × 10⁶ (n = 9), 1 × 10⁶ (n = 6), 2 × 10⁶ (n = 8) or 5 × 10⁶ (n = 8) WBM cells from three independent primary recipients of 5–7sp Em-Ex were transplanted into lethally irradiated CD45.2⁺CD45.1⁺ recipients (Fig. 2a). At the three highest doses of WBM, 100% of secondary recipients displayed significant Em-Ex-derived CD45.2⁺ chimerism (1–92%) at 16 weeks post-transplant (Fig. 2a). CD45.2⁺ reconstitution was >25% in 11/31 secondary recipients and >90% in two recipients. Multiple independent secondary recipients also yielded robust long-term Em-Ex-derived CD45.2⁺ chimerism after transplantation into lethally irradiated CD45.1⁺CD45.2⁺ tertiary recipients, although interestingly this reconstitution displayed dramatic lineage skewing (Fig. 2b). The robust hematopoietic reconstitution of primary, secondary, and tertiary recipients by cultured 5–7sp Em-Ex is indicative of self-renewing dHSC and long-term repopulating cells, confirming that our explant culture system supports the emergence and maturation of dHSC from embryos isolated at the onset of blood circulation.

**Fig. 1: Pre-circulation embryos but not YS yield robust dHSC activity.**

**Fig. 2: Pre-circulation Em-Ex display serially transplantable multi-lineage potential.**

Pre and peri-circulation YS do not produce dHSC ex vivo

Dissected CD45.2⁺ YS from pre-circulation and peri-circulation concepti (2–3sp and 5–7sp) were cultured as explants (YS-Ex) in parallel and under the same conditions as Em-Ex. None of the cultured pre-circulation or peri-circulation CD45.2⁺ YS-Ex yielded detectable dHSC activity when transplanted into lethally irradiated CD45.1⁺CD45.2⁺ recipients along with 2 × 10⁵ WBM CD45.1⁺ cells (0/5 and 0/11 recipients, respectively, Fig. 1b). It is possible that YS hemogenic precursors undergo distinct temporal maturation compared to intra-embryonic hemogenic endothelium. Indeed, YS-Ex displayed substantially greater CFU-GEMM (i.e., colony forming unit granulocyte, erythrocyte, monocyte, megakaryocytic) potential at day 5 versus day 10 of ex vivo culture (Supplementary Fig. 1A). Thus, 5–7sp YS-Ex were cultured for five or seven days prior to dissociation and transplanted into lethally irradiated recipients. These cultures also failed to produce detectable dHSC activity (0/8 for five-day YS-Ex cultures and 0/3 recipients for seven-day YS-Ex, Fig. 1c).

These data indicate that the only precursors capable of maturing into functional dHSC under these conditions are found in the embryo at this early stage of development. Additionally, we tested the ability of YS isolated from later developmental time-points (e.g., E9.5 (19–22 sp, n = 3), E10.5 (32–36 sp, n = 3) and E11.5 (46–47sp, n = 7)) to engraft lethally irradiated recipients after being cultured ex vivo for five days prior to transplantation. 100 and 71% of CD45.1⁺CD45.2⁺ recipients transplanted with CD45.2⁺ E10.5 and E11.5 cultured YS-explants showed robust CD45.2⁺ YS-derived engraftment, respectively (Fig. 1c). In contrast, CD45.1⁺CD45.2⁺ recipients transplanted with CD45.2⁺ E9.5 YS-derived explants displayed no evidence of CD45.2⁺ engraftment (Fig. 1c). To test if E9.5 hematopoietic precursors are capable of maturing into dHSC under our ex vivo culture conditions, embryonic caudal halves and YS isolated from CD45.2⁺ E9.5 concepti were cultured for five, seven or 10 days and then transplanted into lethally irradiated CD45.1⁺CD45.2⁺ recipients (Fig. 1d). 100% of recipients transplanted with embryonic caudal halves cultured for five or seven days (3/3 and 6/6, respectively, Fig. 1d) displayed robust CD45.2⁺ dHSC activity, confirming that post-circulation E9.5 dHSC can mature under these culture conditions. However, all E9.5 YS-derived explants failed to generate detectable dHSC in these experiments (Fig. 1d).

In sum, these findings support a model where the precursors in pre-circulation and peri-circulation concepti capable of maturing into functional dHSC under these conditions are located in the embryo.

SCF and IL-3 induce CD31⁺CD45⁺c-Kit⁺CD41⁻ dHSC emergence

We next sought to phenotypically characterize functional dHSC that emerge during the ex vivo culture of peri-circulation Em-Ex. We took advantage of cell surface markers to isolate distinct populations from CD45.2⁺ 5–7sp Em-Ex for transplantation into lethally irradiated CD45.1⁺CD45.2⁺ recipients (Fig. 3a). Previous work showed that dHSC activity in E10.5-E11 Em-Ex cultured under similar conditions is contained in the CD31⁺CD45⁺ fraction^29,31. This population was apparent in the Em-Ex cultures (Fig. 3b). CD41 and c-Kit are also expressed by nascent hematopoietic populations in early embryos^{30,31,32,33,34,35,36,37,38}. Distinct populations expressing both CD41 and c-Kit were clear within CD31⁺CD45⁺ Em-Ex cells (Fig. 3b, Fig. 4b and Supplementary Fig. 1B). We therefore isolated CD31⁺CD45⁺c-Kit⁺CD41⁺, CD31⁺CD45⁺c-Kit⁺CD41⁻and CD31⁺CD45⁺c-Kit⁻ cells by FACS from cultured 5–7sp Em-Ex and transplanted them into lethally irradiated CD45.1⁺CD45.2⁺ recipients (n = 5, Fig. 3a–c). All dHSC activity was restricted to CD31⁺CD45⁺c-Kit⁺CD41-cells (Fig. 3c). Interestingly, this same population was also apparent in 5–7sp YS cultures (Fig. 4b and Supplementary Fig. 1B).

**Fig. 3: HSC potential is restricted to CD31⁺CD45⁺c-Kit⁺CD41⁻ embryo explant cells and requires SCF and IL-3.**

**Fig. 4: scRNAseq analysis identifies a rare HSC-like population specific to embryo explant cultures.**

We next tested whether the emergence of the CD31⁺CD45⁺c-Kit⁺CD41⁻ engrafting population was selectively dependent on any of the three growth factors added to these cultures (i.e., SCF, IL-3, FLT3L). Embryos and YS isolated from 5–7sp concepti were cultured in the presence of different combinations of growth factors and assessed by flow cytometry for the emergence of CD31⁺CD45⁺ and CD31⁺CD45⁺c-Kit⁺CD41⁻ cells (Fig. 3d, e). CD45.2⁺ Em-Ex cultures were also transplanted into lethally irradiated CD45.1⁺CD45.2⁺ recipients to test for dHSC activity (Fig. 3d, f). Removal of FLT3L had no effect on the frequency of CD31⁺CD45⁺ or CD31⁺CD45⁺c-Kit⁺CD41⁻ cells in YS-Ex or Em-Ex cultures (Fig. 3e). Moreover, removal of FLT3L did not affect the emergence of dHSC in Em-Ex cultures: the contribution of CD45.2⁺ cells to the PB of transplant recipients was 18% in the presence of all three factors and 21% in the absence of FLT3L (Fig. 3e, f and Supplementary Fig. 1B). In contrast, removal of IL-3 or SCF reduced the frequency of CD31⁺CD45⁺ or CD31⁺CD45⁺c-Kit⁺CD41⁻ cells, respectively, in YS-Ex and Em-Ex cultures, which correlated with a loss of dHSC activity from CD45.2⁺ Em-Ex cultures (Fig. 3e, f and Supplementary Fig. 1B). The average CD45.2⁺ hematopoietic reconstitution in these recipients dropped to about 5 and 10% in the absence of IL-3 or SCF, respectively (Fig. 3f). Em-Ex cultured in the absence of both FLT3L and IL-3 also displayed reduced hematopoietic engraftment potential (p < 0.1, exact Wilcoxon rank sum test (eWrs-test)) (Fig. 3f). Em-Ex cultured in the absence of all three cytokines displayed dramatic reductions in the frequency of CD31⁺CD45⁺ and CD31⁺CD45⁺c-Kit⁺CD41⁻ cells and lacked detectable dHSC activity (Fig. 3e, f and Supplementary Fig. 1B). In sum, administration of both SCF and IL-3 is required for the efficient development of CD31⁺CD45⁺c-Kit⁺CD41⁻ cells from pre-circulation concepti during YS-Ex and Em-Ex culture and dHSC potential in Em-Ex cultures. These results are consistent with the observation that SCF and IL-3 can both promote the in vitro derivation of bona fide HSC from other cell sources^{29,30,31,39,40,41}.

An HSC-like population is generated by embryo explants

YS and embryos are separated and cultured before the onset of the circulation in our experimental platform. Thus, this represents an opportunity to selectively study embryo versus YS-derived hematopoiesis under controlled conditions that allow for the maturation of dHSC from early embryonic precursors. Indeed, novel cell surface markers of dHSC-derived from early embryos could be useful reporters in efforts to isolate dHSC from pluripotent stem cells or other sources. Although CD31⁺CD45⁺c-Kit⁺CD41⁻cells develop in both YS-Ex and Em-Ex cultures, cultured YS-Ex-CD31⁺CD45⁺c-Kit⁺CD41⁻ cells do not harbor dHSC activity (Figs. 1b, 3b, 3f, Supplementary Fig. 1B). Therefore, we decided to interrogate the gene expression profiles of 5–7sp YS-Ex-CD31⁺CD45⁺c-Kit⁺CD41⁻ cells and Em-Ex-CD31⁺CD45⁺c-Kit⁺CD41⁻ cells on day 10 post-culture for molecularly distinct cell populations that might explain the absence of dHSC activity in YS explants by single cell RNA sequencing (scRNAseq) (Fig. 4).

First, we determined the frequency of the engrafting population within cultured Em-Ex. Limiting dilution transplantation revealed 0.83 repopulating units (RUs) per Em-Ex (95% C.I. = (0.48, 1.42); results fit the LDA model: p-value = 0.717, t-test) (Fig. 4a). Next, 49 concepti were dissected and cultured as separate YS-Ex and Em-Ex for scRNAseq. In this experiment we are interrogating approximately 40.67 RUs in the Em-Ex sample (i.e.,: 49Em-Ex*0.83RUs).

To identify Em-Ex-derived populations of putative biological interest, the resulting scRNAseq data was projected as tSNE1 Vs tSNE2. A population specific to the Em-Ex sample was apparent (X-cells, Fig. 4c). It was comprised of 623 cells (17.8% of the Em-Ex population). To interrogate the developmental state of this population, we compared its expression profile to Zhou and colleagues scRNAseq dataset of embryonic hematopoiesis (Fig. 4d)⁴². Em-Ex cells were most similar transcriptionally to E11-T2-pre-HSC (CD31⁺CD45⁺cKit⁺CD41^low, 60% of the X-cells were assigned to this group) or E12.5 Fetal Liver (FL) HSC (Lineage⁻Sca-1⁺Mac-1^lowCD201⁺, 37% of the X-cells were assigned to this group) and thus ‘assigned’ to these populations computationally (Fig. 4d). Interestingly, Em-Ex-derived X-cells were highly enriched for cells assigned to FL HSC. Indeed, 70 and 6% of Em-Ex-derived X-cells were assigned to E12.5 FL HSC and E14.5 FL HSC (CD45⁺CD150⁺CD48⁻CD201⁺), respectively (Fig. 4d). These data suggest that X-cells are enriched for developmentally more mature hematopoietic progenitors⁴³. Interestingly, the vast majority of profiled YS-Ex cells (92%) were assigned to E11.0-T2-pre-HSC, suggesting developmental immaturity relative to Em-Ex cells (Fig. 4d). A comparison of E11.0-T2-pre-HSC and E12.5-FL HSC revealed 353 downregulated genes and 80 upregulated genes (FDR<0.05 and Log₂ fold change <−2 or >2, respectively; eWrs-test and Benjamini and Hochberg p-value adjustment) (Fig. 4e). Remarkably, compared to other analyzed cells in our scRNAseq dataset, X-cells showed lower expression for the downregulated genes and higher expression for the upregulated genes (Fig. 4f). Upregulated genes were enriched in proliferation related functional terms (i.e., E2F cell cycle targets and DNA-repair genes (FDR q-values = 3.98 × 10⁻³ and 1.58 × 10⁻², respectively; Fisher’s exact test and Benjamini and Hochberg p-value adjustment). This is consistent with increasing proliferation as HSC transition from the AGM to the FL¹. Genes defining epithelial-to-mesenchymal downregulated in the E11.0-pre-HSC to E12.5-HSC transition were also downregulated in the X-cells (FDR q-value = 2.85 × 10⁻⁶; Fisher’s exact test and Benjamini and Hochberg p-value adjustment). Of note, multiple cell surface proteins appear selectively upregulated in Em-Ex X-cells (e.g., Cxcr4, Mpl, Il6ra, and Tnfrsf1, Supplementary Fig. 2). Further work will be required to determine if these cell surface markers hold functional utility for further enrichment of transplantable dHSC from Em-Ex. Indeed, identifying a specific marker for embryonic populations with bona fide dHSC potential could be very useful for in vitro screens.

Intraembryonic LYVE1⁺ blood precursors yield dHSC activity

Our studies suggest that the pre-circulation and peri-circulation YS is not a major source of dHSC. However, it was recently reported that >40% of adult blood may emerge from the early YS and vitelline endothelium¹⁴. Lee and colleagues used Lyve1-eGFP-hCre knock-in mice (Lyve1^Cre) (Fig. 5a) to claim an extraembryonic origin for a significant fraction of adult blood, but not primitive erythrocytes or the definitive hematopoietic precursors of the embryo, placenta or umbilical vessels. LYVE1 expression was reportedly highest in the endothelium of the YS and vitelline vessels (VV) in the mid-gestation conceptus¹⁴.

**Fig. 5: LYVE1 is expressed by E9.5-E10.5 putative hematopoietic progenitors in embryonic and YS tissues.**

We used Lyve1^Cre, Rosa26^+/mTmG and Rosa26^+/Confetti mice to further examine the contribution of LYVE1⁺ cells to lifelong hematopoiesis. The unrecombined Rosa26^mTmG allele ubiquitously labels all cells with the tdTOMATO protein (Fig. 5a). CRE recombinase (CRE) drives the excision of the tdTomato cassette and stably labels CRE⁺ cells and their progeny with GFP (Fig. 5a)⁴⁴. The unrecombined Rosa26^Confetti multicolored cassette does not drive any fluorescent expression. Here, CRE induces recombination of the Rosa26^Confetti allele, which results in cells and their progeny stably labeled with one of four fluorescent proteins (GFP, CFP, RFP, or YFP, Fig. 5a)⁴⁵. We observed ≈20 and ≈5% labeled cells in the adult blood of Lyve1^+/CreRosa26^+/mTmG and Lyve1^+/CreRosa26^+/Confetti mice, respectively (Fig. 5b), confirming that cells expressing CRE via the Lyve1^Cre allele at some point during their development contribute to adult hematopoiesis, albeit in our hands less than the 40% previously reported¹⁴. Similar to previous reports, we observed that the endogenous eGFP signal from the Lyve1^Cre knock-in allele (Fig. 5a) was faint and did not hinder detection of the eGFP signal from the Rosa26^mTmG reporter allele^14,46.

To further interrogate LYVE1 as a marker of early hematopoietic progenitors, we next assessed LYVE1 expression in post-circulation concepti. We observed both LYVE1^low and LYVE1^high cells in E9.5 (19–22sp) and E10.5 (32–36sp) YS and embryos (Fig. 5c–f). Most embryo LYVE1⁺ cells were LYVE1^low (Fig. 5c–f). Indeed, at E9.5 and E10.5 the absolute numbers of embryo- and YS-LYVE1^lowVE-Cadherin⁺CD31⁺CD41⁺c-Kit⁺ cells were not significantly different (Fig. 5d, f). Thus, LYVE1 expressing cells are readily detectable in post-circulation embryos and YS. Further, examination of public gene expression databases revealed Lyve1 mRNA in dHSC precursors in E11-E11.5AGM (Supplementary Fig. 3)^42,43. Altogether, these data confirm that LYVE1 is expressed by putative hematopoietic progenitors in both YS and embryonic tissues post-circulation¹⁴.

Next, we examined LYVE1 expression in peri-circulation (5–7sp concepti) mouse concepti using flow cytometry and confocal microscopy. We again observed distinct LYVE1^high and LYVE1^low populations in both YS and embryos (Fig. 6a). Although both populations were more abundant in YS relative to embryos, especially LYVE1^high cells, LYVE1 expressing cells were clearly detectable by flow cytometry in 5–7sp embryos (% of YS-LYVE1^high = 2.86; % of Em-LYVE1^high = 0.11; % of YS-LYVE1^low = 3.11; % of Em-LYVE1^low = 0.56) (Fig. 6a, b). Most LYVE1^high and LYVE1^low cells in both 5–7sp YS and embryos also expressed CD41 (≈70 and ≈85%, respectively, Fig. 6a, c). Moreover, LYVE1⁺CD31⁺ cells were apparent in the endothelium of the paired aorta of E8.5 embryos (Fig. 6d). Altogether, these data demonstrate that LYVE1 expression is not restricted to YS in peri-circulation embryos.

**Fig. 6: LYVE1 is expressed by both yolk sac and embryo cells at E8.5.**

To gain further insight into the fidelity of LYVE1 as a specific marker of YS and vitelline hemogenic endothelium, YS and embryos were collected at the onset of the heartbeat from 5–7sp- Lyve1^+/CreRosa26^+/mTmG concepti (Fig. 7a). Here, GFP⁺ cells were readily detected in YS (Fig. 7b, c), with an average frequency of 3.9% (Fig. 7c). Labeled cells were much less frequent in 5–7sp embryos (average labeling of 0.067%, Fig. 7b, c). We previously observed that the vast majority of LYVE1⁺ cells in Lyve1^+/+Rosa26^+/+ concepti are CD41⁺ (Fig. 6c). Similarly, most GFP⁺ cells in 5–7sp Lyve1^+/CreRosa26^+/mTmG concepti were CD41⁺ (Fig. 7b, d). Additionally, the %CD41⁺ cells was about 20 times larger in the YS than in the embryo (Fig. 7e). As CD41 labels hematopoietic precursors at this early developmental stage, we focused on the recombination efficiency in the CD41⁺ compartment and found it to be about 5% in the YS and 1.1% in the embryo (Fig. 7f)^{30,32,33,34,36,37}. Importantly, GFP⁺ cells in E8.5 embryos were confined to the endothelium of the paired aortas (Fig. 7g).

**Fig. 7: Blood precursors emerge from intra-embryonic LYVE1⁺ cells at the onset of the heartbeat.**

As our explant culture system allows us to visualize the emergence of phenotypic hematopoietic precursors from pre-circulation and peri-circulation concepti, we next examined GFP labeling in cultured YS-Ex and Em-Ex-derived hematopoietic precursors (CD31⁺CD45⁺c-Kit⁺CD41⁻). We observed that, on average, 61% of YS-Ex-derived and 47% of Em-Ex-derived CD31⁺CD45⁺c-Kit⁺CD41⁻cells were GFP⁺ (Fig. 7h, i). Lethally irradiated CD45.1⁺CD45.2⁺ recipients were next transplanted with cells isolated from Lyve1^+/CreRosa26^mTmG 5–7sp Em-Ex and 2 × 10⁵ CD45.1⁺ WBM cells (Fig. 7j-l). As expected from our previous experiments (Fig. 1b), 5–7sp Lyve1^+/CreRosa26^mTmG cultured Em-Ex harbored dHSC activity and reconstituted 15–80% of recipient PB (Fig. 7k). Importantly, 3/4 recipients of E8.5 Lyve1^+/CreRosa26^mTmG-derived Em-Ex displayed GFP⁺ engraftment, demonstrating that Lyve1^Cre labels the progeny of intra-embryonic HE precursors (Fig. 7l). Indeed, in one recipient, 88% of Em-Ex-derived PB was GFP⁺ (Fig. 7l). Differences in the %GFP⁺ recipient PB reflects the small number of Em-Ex-derived RUs transplanted per recipient (3.32 RUs = 4EE*0.83RUs per explant, Fig. 4a) and low recombination efficiency of the Rosa26^mTmG allele (about 20%, Fig. 5b). We also tested cells isolated from 19–22sp Lyve1^+/CreRosa26^mTmG Em-Ex and YS-Ex for dHSC activity via transplantation. Here, Em-Ex-derived PB reconstitution was high (averaging 51%, n = 10). Although the %GFP⁺ PB varied widely (Fig. 7l), 8/10 recipients displayed GFP + PB reconstitution. As expected (Fig. 1c, d), Lyve1^+/CreRosa26^mTmG YS-Ex did not engraft recipient mice (Fig. 7k, l). In sum, our data reveal that the embryo harbors LYVE1⁺ blood precursors capable of giving rise to dHSC.

Discussion

Identifying the original site of dHSC specification during embryogenesis has been a major challenge due to the migratory nature of blood precursors in the circulating embryo and the absence of assays to rigorously test hematopoietic potential of pre-circulatory tissues. Here, we report the robust generation of dHSC activity from pre-circulation and peri-circulation murine embryos. dHSC activity could only be detected in cultured embryo explants and not cultured YS explants, indicating that the YS is unable to autonomously generate dHSC activity, at least under the conditions tested here. This implies that either the YS does not contain dHSC precursors or that it lacks the essential support environment for maturation of such precursors. Our findings are consistent with those of Cumano and colleagues, but in those studies, the authors were only able to detect very low levels of HSC activity from pre-circulation intra-embryonic tissues (<5%), and only after transplantation into immunocompromised Rag2γc^−/− recipients¹². Poor reconstitution activity left open the possibility that YS harbored dHSC activity that fell below their detection threshold. Indeed, very recent studies suggest an extraembryonic (i.e., YS and/or vitelline hemogenic endothelium) origin for a large fraction of lifelong hematopoiesis^13,14. However, the high levels of dHSC activity detected in our culture system (up to >75% reconstitution of PB after transplantation into immunologically competent mice) allows us to rigorously compare YS and embryos and demonstrate that pre-circulation YS lacks significant dHSC potential (Fig. 1b). None-the-less, our data support a model in which dHSC originate from intra-embryonic precursors in pre-circulation and peri-circulation concepti.

CD31⁺CD45⁺c-Kit⁺CD41⁻cells harbored all Em-Ex-derived dHSC activity in our experimental system. The development of this population required exogenous administration of SCF and IL-3 (Fig. 3). This agrees with previous studies where SCF is also required for the maturation of E9.5 pro-HSCs and E10.5 type I and type II pre-HSCs. Scf is expressed in the E9.5 dorsal aorta and upregulated at E10.5 when HSPC specification accelerates^29,30,31. Scf^−/− mice display reduced dHSC activity and peri-natal death³⁹. IL-3 induces maturation of dHSC between E10.5 and E11.5 in the AGM^40,41. In contrast, exogenous administration of FLT3L is dispensable for dHSC maturation ex vivo³¹. Our finding that 5–7sp Em-Ex also require SCF and IL-3 to effect dHSC maturation ex vivo suggests that even very early dHSC precursors require these same signals.

Interestingly, CD31⁺CD45⁺c-Kit⁺CD41⁻ cells also emerged in cultures of early YS (Fig. 4b), where they clearly lack detectable dHSC activity (Fig. 1b–d). The ability of our culture conditions to yield bona fide dHSC from pre-circulation concepti provides a unique opportunity to study YS-specific and embryo-specific hematopoiesis and the critical molecular features that allow Em-Ex to render dHSC activity. This could be useful for efforts to derive bona fide dHSC from alternative sources ex vivo and provide specific reporters for in vitro screening. Remarkably, single cell global gene expression profiling identified an Em-Ex-derived population (X-cells, Fig. 4c) that does not emerge from YS cultures and displays enrichment for genes expressed by dHSC found in the FL (Fig. 4d). Further analysis identified multiple cell surface molecules selectively expressed by this unique population (e.g., Cxcr4, c-Mpl, Il6ra, and Tnfrsf1, Supplementary Fig. 2). These cell surface markers harbor putative utility to both refine the Em-Ex population harboring dHSC activity and explore for in vivo surrogates that may represent critical transitions in the maturation of dHSC in vivo. Exploring the biological significance of these and other markers will be the subject of future investigations.

An alternative approach to defining the embryonic origins of lifelong hematopoiesis is the use of YS versus embryo-specific lineage markers. This approach has proved challenging as YS and embryo-derived hematopoietic precursors exhibit similar developmental programs¹. Both are mesodermal in origin and emerge from endothelial precursors. Previously, temporally controlled Runx1⁻based lineage tracing suggested YS-derived contribution to adult hematopoiesis¹³. However, the duration of the labeling pulse was unclear and only a low percentage of adult blood cells were labeled^1,13. Additionally, RUNX1⁺ cells were apparent outside the YS, raising doubts that all Runx1-induced Cre recombinase activity in early embryos was YS-restricted^1,13. Further, this study employed a Cre recombinase knock-in mouse model that was hemizygous for Runx1. Runx1^−/+ mice are severely perturbed in both the timing of hematopoietic specification and the cell surface phenotype of emerging hematopoietic progenitors⁴⁷. Thus, observations made in a hemizygous Runx1 mouse model are unlikely to reflect normal steady-state hematopoietic ontogeny.

It is possible that the YS harbors dHSC precursors that fail to ‘mature’ in our system due to the absence of key embryo-derived non-cell-autonomous cues. In this scenario, YS-precursors may need to migrate to a maturation niche in the embryo, as proposed by Matsuoka and colleagues¹⁸. Indeed, Runx1⁺Gata-1⁻ (hemogenic angioblasts) have been reportedly observed migrating from the E7.5 extraembryonic YS into the embryo proper in cultured embryo explants, where they were proposed to receive a currently undefined signal that induces competence to contribute to lifelong hematopoiesis¹⁹. However, here again the duration of the 4-OHT labeling pulse was unclear, raising the possibility that rare Runx1 + intraembryonic precursors could have been labeled between E8 and E8.5. Further, it has not been explicitly demonstrated that Runx1⁺Gata-1⁻ cells seen migrating in ex vivo cultures from the extra-embryonic to the intra-embryonic region are capable of contributing to lifelong hematopoiesis in vivo. In our explant system, pre-circulation and peri-circulation YS explants failed to generate any dHSC activity.

Although, Lyve1⁻Cre lineage tracing was recently employed to discern blood derived from YS versus embryonic endothelial precursors¹⁴, we observed two distinct LYVE1⁺ populations (LYVE1^high and LYVE1^low) in both YS and embryos isolated from E8.5, E9.5 and E10.5 concepti (Figs. 5–6). Although the frequency and absolute numbers of LYVE1^high cells is higher in the YS, the absolute numbers of LYVE1^low cells between YS-derived and embryo-derived blood precursors is not significantly different (Figs. 5–6). Moreover, we observed LYVE1⁺ endothelial cells in the endothelium of the paired embryo aortas prior to the onset of circulation (Fig. 6d and Fig. 7g). Furthermore, Lyve1^+/Cre Rosa26^+/mTmG embryos isolated at the onset of the first heartbeat (5–7sp) and cultured as explants gave rise to GFP⁺CD31⁺CD45⁺c-Kit⁺CD41⁻ cells, which is the transplantable population that emerges in this system (Fig. 3c and Fig. 7h, i). On average, about 50% of emerging CD31⁺CD45⁺c-Kit⁺CD41⁻ cells were GFP⁺ in these cultures, with one Em-Ex yielding 100% GFP⁺CD31⁺CD45⁺c-Kit⁺CD41⁻ cells (Fig. 3c and Fig. 7h, i). Most importantly, transplantation of 5–7sp Lyve1^+/Cre Rosa26^+/mTmG Em-Ex revealed that LYVE1⁺ embryo-derived progeny consistently contributed to the engraftment of lethally irradiated recipients (Fig. 7j–l). Thus, embryo-derived hematopoietic precursors must also express LYVE1. Altogether, these data confirm LYVE1 as an early marker of definitive hematopoiesis, reveal that LYVE1 expression is not restricted to YS hematopoietic precursors and suggest that most LYVE1⁺ contribution to adult hematopoiesis is embryonic in origin.

In summary, here we report the generation of robust dHSC activity from pre-circulation and peri-circulation embryo explants during ex vivo culture. Cultured YS explants failed to yield dHSC activity under these same conditions, suggesting that the YS contains few precursors capable of maturing into dHSC. We also present evidence that both YS and embryo-derived hemogenic precursors express LYVE1 during ontogeny. Our study supports a model in which most of lifelong hematopoiesis derives from hematopoietic precursors that develop de novo in the embryo proper. Future studies will be required to better define the pre-circulation and peri-circulation embryonic precursors that ultimately mature into dHSC. These studies would benefit any effort to coax bona fide dHSC from pluripotent stem cells.

Methods

Mice

C57BL/6 J, C57BL/6.SJL-PtprcaPep3b/BoyJ, ROSA26^+/mTmG (Gt(ROSA)26Sor^{tm4(ACTB-tdTomato,-EGFP)Luo}/J), ROSA26^+/Confetti (Gt(ROSA)26Sor^{tm1(CAG-Brainbow2.1)Cle/J}) and Lyve1-eGFP-hCre knock-in mice (Lyve1^Cre) mice were acquired from The Jackson Laboratory (Bar Harbor, Maine) and housed in a pathogen-free facility. Both females and males were used. All animal experiments were carried out according to procedures approved by the St. Jude Children’s Research Hospital Institutional Animal Care and Use Committee and comply with all relevant ethical regulations regarding animal research.

Genotyping

Polymerase chain reactions (PCR) were performed using Go Taq DNA Polymerase (Promega, Madison WI) as indicated by the manufacturer. PCR conditions: (95 °C, 2’);((94 °C, 30”; 56 °C, 30”; 72 °C, 30”]×35); (72 °C, 10’).

Primers: Cre F1 (5’ CTGTTACGTATAGCCGAAAT 3’), Cre

R1 (5’ CTACACCAGAGACGGAAATC T 3’). CRE Positive PCR band: 203 bp. Detection of the GFP allele was used for mTmG and Confetti allele genotyping: GFP Fw (5’ CAGATGAAGCAGCACGACTTCT 3’), GFP Rv (5’ AACTCCAGCAGGACCATGTGAT 3’). GFP PCR band: 400 bp.

Transplants

For all transplants, 8–12 weeks old CD45.2⁺/CD45.1⁺ C57BL/6 J recipients were treated with 11 Gy of ionizing radiation in split doses of 5.5 Gy prior to transplant. Males and females were used as recipients. In each experiment, sexes of the recipient mice were evenly distributed among experimental groups. All cells were transplanted by tail vein injection. E8.5, E9.5, E10.5, and E11.5 CD45.2⁺ Em-Ex-derived or YS-derived cells were transplanted along with 2 × 10⁵ CD45.1⁺ C57BL/6.SJL WBM cells into recipients. For secondary and tertiary transplantation, 5 × 10⁶, 2 × 10⁶, 1 × 10⁶, or 0.8 × 10⁶ WBM cells were transplanted from primary or secondary recipients. Engraftment was defined as >0.5% CD45.2⁺ cells in each lineage (T cells, B cells, and myeloid cells) and >1% CD45.2⁺ total PB.

PB Analysis

PB was collected from the retro-orbital plexus in heparinized capillary tubes (Fisherbrand, Pittsburgh, PA) and lysed in red blood cell lysis buffer (Sigma-Aldrich, St. Louis, MO). Cells were stained with the following antibodies: CD45.1-APC (A20) (Biolegend, San Diego, CA) or CD45.1-FITC (A20) (BD Biosciences, San Diego, CA), B220-PECy7 and CD8-PECy7 (53–6.7) (Tonbo Biosciences, San Diego, CA), CD45.2-V500 (104), B220- PerCPCy5.5 (RA3-6B2), Gr1-PerCPCy5.5 (RB6-8C5), Cd11b-PerCPCy5.5 (M1/70) and CD4-PECy7 (RM4-5) (BD Biosciences, San Diego, CA). All antibodies were used at 1:200 dilution. 4’,6-diamidino-2-phenylindole (DAPI) staining was used to gate live events. Analysis was performed on a LSR Fortessa and a BD FACSAria III SORP (Special Order Research Product, which contains the following LASERs: 405 nm, 445 nm, 488 nm, 562 nm, and 640 nm) (both BD Biosciences, San Diego, CA) and the data analyzed with FlowJo version 9.4.11 (Tree Star, Ashland, OR). To discern the four Confetti colors: the filter arrangements for each LASER were: CFP, 445 nm LASER—470/24 band-pass filter (BP); GFP, 488 nm LASER—515/20 BP and 505 long-pass filter (LP); YFP, 488 nm LASER—545/10 BP and 525 LP; RFP, 562 nm LASER—610/20 BP and 600 LP⁴⁸.

Explants cultures and flow cytometry analysis

Explant culture conditions were adapted from the protocol developed by Medvinsky and colleagues²⁹. Embryo and YS-explants were cultured at the air-liquid interface on 0.65 μm DV Durapore Membrane Filters (Merck Millipore, Cork, Ireland) in Iscove’s Modified Dulbecco’s Medium-Glutamax (IMDM-Glutamax, Thermo Fisher Scientific, Waltham, MA), 20% FCS (Lot #535905; Cat #FB-02; Omega Scientific, Tarzana, CA), 0.1 mM 2-Mercaptoethanol (Thermo Fisher Scientific, Waltham, MA), and 100 units/ml of Penicillin/Streptomycin (Thermo Fisher Scientific, Waltham, MA) supplemented with recombinant murine SCF, recombinant murine IL-3, and recombinant murine FLT3 Ligand (Peprotech, Rocky Hill, NJ, all at 100 ng/ml) (or with other cytokine combinations indicated along the text) for 9–10 days to allow engraftment from E8.5 tissues or for other time periods as detailed in each figure. Membrane filters were sterilized and rinsed three times in boiling water (five minutes/wash). Up to eight explants were carefully placed on each membrane filter. Filters were then placed on in-house-made ring stands (3.175 mm high; outside diameter: 32.3 mm; inside diameter: 23 mm; Teflon) positioned in the well of a non-tissue-culture-treated 6-well-plate (Corning, NY, USA) containing 2.25 mL of the above described culture media. Media was not refreshed during the culture. Explants were recovered from filters using a sterilized scalpel.

Freshly isolated embryonic tissues or cultured explants were dissociated with collagenase (0.0012 g/ml, Sigma-Aldrich, St. Louis, MO) in Phosphate-Buffered Saline (PBS, Thermo Fisher Scientific, Waltham, MA) supplemented with 10% FCS (Omega Scientific, Tarzana, CA). For flow cytometry cell sorting or analysis, dissociated cells were stained with one or more of these antibodies: CD45.2-V500 (104) (BD Biosciences, San Diego, CA), CD41-PerCP-eFluor710 (eBioMWReg30) (eBioscience, San Diego, CA), CD31-FITC (MEC13.3) (BD Biosciences, San Diego, CA), c-Kit-APC-eFluor780 (2B8) (eBioscience, San Diego, CA), Lyve1-PE (#223322) R&D Byosystems, Minneapolis, MN). All antibodies were used at 1:200 dilution. 4’,6-diamidino-2-phenylindole (DAPI) staining was used to gate live events. Analysis was performed on a LSR Fortessa and cell sorting on a BD FACSAria III SORP (Special Order Research Product) (both BD Biosciences, San Diego, CA). Cells were collected employing the BD FACSDiva Software (version 8.0.1) (BD Biosciences, San Diego, CA). Data was analyzed with FlowJo version 9.4.11 (Tree Star, Ashland, OR).

Statistics and reproducibility

Summary statistics, including mean and standard deviation, were reported for analyses. Two sample t or exact Wilcoxon rank sum test were used to test for a difference between two groups depending on the normality of the data, which was tested by the Shapiro-Wilk test. False discovery rate (FDR) method developed by Benjamini and Hochberg⁴⁹ was used to correct for multiple comparisons at a level of 0.05. Otherwise, p-values < 0.05 were considered statistically significant. Analyses were conducted in R-3.3.1.

For limiting dilution analysis, parameters were estimated using a generalized linear model with a complementary log-log link. The generalized Pearson Chi-square was used to assess the goodness-of-fit to the model. The analyses were done using program L-Calc and ELDA online software⁵⁰.

All in vitro experiments were reliably reproduced at least 3 times with the exception of Fig. 3Eii (the 5 days culture time point was n = 2) and Fig. 3Ei (within the 5 days culture time point the SCF/IL3 and the SCF conditions were n = 2). Sample size and number of experiment replicates are detailed in each Figure Legend.

Single cell RNA-seq

Sorted YS-Ex-CD31⁺CD45⁺c-Kit⁺CD41⁻ and Em-Ex-CD31⁺CD45⁺c-Kit⁺CD41⁻ were resuspended in PBS containing 0.04%BSA at a concentration of 500–600 cells/ul. In total, 8600 cells per sample were loaded into each well of a 10X Chromium single cell capture chip with a recovery of 3000–3500 cells per sample. The captured cells then underwent lysis, reverse transcription, cDNA amplification, and library preparation with indexing per the manufacturer’s protocol (10X Genomics). The libraries were sequenced together on an Illumina Nextseq500 (Illumina Inc., San Diego, CA) using a high output 150 cycle kit with read lengths recommended by 10X Genomics.

Single-cell RNAseq analysis

Raw data processing: Cellranger 2.0.1⁵¹ was used to process the raw fastq files and to generate unique molecular identifiers (UMI) counts for each gene in each cell. The UMI count table was imported into R and analyzed with Seurat 2.1.0 package⁵². The raw counts were normalized using NormalizeData function with default parameters (normalization.method = LogNormalize). Cells expressing less than 1000 genes, more than 5000 genes or having over 5% mitochondria reads were removed from the analysis to exclude potential dead, broken or doublet cells. Bach effect is removed using R package scran v1.8.1⁵³ with k = 20, sigma = 0.1. To identify the X-cell population, R package Rtsne v0.13⁵⁴ was used to reduce data dimension to two for visualization with perplexity = 20, initial_dims = 100. X-cell population was defined by clusters generated with R package pdfCluster v1.0-2⁵⁵ and selected for further analysis. R package scmap v1.1.5⁵⁶ was used to map single cells to cell clusters from Zhou et al.⁴² with threshold = 0.7. In detail, R package scmap v1.1.5 computed the similarity between expression profile of a single cell and centroid of cell clusters in the reference data⁵⁶. Cells were assigned to reference cell clusters with the highest similarity. Multiple types of correlations (including Cosine similarity and Pearson and Spearman correlations) were calculated and at least one of them had to be larger than 0.7 (i.e., threshold = 0.7).

CFU assays

For analysis of CFU potential, 5–7sp embryos and YS were cultured as explants at the air-liquid interface for five or 10 days in the same conditions as indicated above (20% FCS, SCF, IL-3, FLT3L in IMDM). CD31⁺CD45⁺ and CD31⁻CD45⁻ cells were then isolated by FACS and plated in M3434 methylcellulose (STEMCELL Technologies, Vancouver, Canada). Approximately 60,000 sorted cells were dispensed to 3 mL of M3434 and equally distributed into two plates (#27150, STEMCELL Technologies, Vancouver, Canada). Colonies were analyzed 10 days after plating.

Confocal microscopy

E8.5 concepti were isolated and fixed in 4% paraformaldehyde (Electron Microscopy Sciences, Hatfield, PA) overnight at 4 °C. Concepti were then embedded in 4.5% low-melting-point agarose (Thermo Fisher Scientific, Waltham, MA) and sectioned (100 μm slices) using a vibratome (microtome with vibrating blade VT1200, Leica Byosystems Wetzlar, Germany). Sections were washed three times with PBS 0.3% Triton X-100 (Biorad, Hercules, CA) (30 min/wash) and permeabilized at 4 °C overnight in PBS 0.3% Triton X-100 (Biorad, Hercules, CA). Sections were then blocked in 0.3% Triton X-100 with 3% donkey serum (Sigma-Aldrich, St. Louis, MO) in PBS at 4 °C overnight. Sections were next incubated with previously conjugated anti-LYVE1 (Clone ALY7; eBioscience. San Diego, CA), anti-CD41 (Clone MWReg30; BD Bioscience, San Diego, CA), BV421⁻Rat Anti-Mouse CD31 (Clone MEC 13.3 (RUO); BD Biosciences, San Diego, CA) at 1:200 in 1% skimmed milk in 0.3% Triton X-100-PBS overnight. Anti-LYVE1 and anti-CD41 antibodies were conjugated following manufacture’s protocol to CFTM568 and CFTM488 fluorophores, respectively, using the Mix-n-StainTM labeling kits (Sigma-Aldrich, St. Louis, MO). Sections were then washed three times with 0.3% Triton X-100 in PBS (30 min/wash). Sections were mounted on Fisherbrand Superfrost Microscope Slides (Fisher Scientific, Pittsburgh, PA) and covered with 22 × 22 mm No.1.0 microscope cover glass (Infolab, Indianapolis, IN). Sections were imaged on a Zeiss LSM780 Confocal Microscope (Zeiss, Oberkochen, Germany) using a ×40, 1.1NA water immersion lens. Images were processed employing a ZEN 2012 software (Zeiss, Oberkochen, Germany).

Data availability

The authors declare that all data supporting the findings of this study are available within the article and its supplementary information files or from the corresponding author upon reasonable request. Single-cell RNAseq data have been deposited in NCBI’s Gene Expression Omnibus (GEO) database under accession code GSE110909. The source data underlying Figs. 1, 2, 3, 4A, 5, 6A-C and 7 and Supplementary Figs. 1A, and 3 are provided as a Source Data file.

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Acknowledgements

We thank W. Clements, H. Mikkola, the McKinney-Freeman laboratory and Department of Hematology at St. Jude Children’s Research Hospital (St. Jude) for critical discussions and reading of the manuscript; D. Ashmun, S. Schwemberger, and J. Laxton for FACS support; C. Davis-Goodrum, K. Millican, A. Reap, and C. Savage for help with timed pregnancies and transplants. V. Frohlich, S. King, J. Peters and A. Pitre for help with confocal imaging. This work was supported by the American Society of Hematology (S.M.-F.), the Hartwell Foundation (S.M.-F.), the NIDDK (K01DK080846 and R01DK104028, S.M.-F.), the American Lebanese Syrian Associated Charities (ALSAC) (S.M.-F. and St. Jude Cell and Tissue Imaging Center), and the NCI (P30 CA021765-35, SJCRH Cell and Tissue Imaging Center) and the Burroughs Wellcome Fund (C.G.).

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Authors and Affiliations

Department of Hematology, St. Jude Children’s Research Hospital, Memphis, TN, 38105, USA
Miguel Ganuza, Ashley Chabot, Xing Tang, Yong Cheng & Shannon McKinney-Freeman
Department of Biostatistics, St. Jude Children’s Research Hospital, Memphis, TN, 38105, USA
Wenjian Bi & Guolian Kang
Department of Oncology, St. Jude Children’s Research Hospital, Memphis, TN, 38105, USA
Sivaraman Natarajan, Robert Carter & Charles Gawad

Authors

Miguel Ganuza
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Ashley Chabot
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Xing Tang
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Wenjian Bi
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Sivaraman Natarajan
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Robert Carter
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Charles Gawad
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Guolian Kang
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Yong Cheng
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Shannon McKinney-Freeman
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Contributions

M.G. designed the study, performed and analyzed transplants, collected and analyzed data, and wrote the paper. A.C. analyzed transplanted mice and contributed to study design. X.T. and Y.C. performed analysis on single-cell RNA sequencing data. X.T. wrote relevant sections of the manuscript. S.N., R.C., and C.G. collected single-cell RNA sequencing data. G.K. and W.B. performed statistical analyses, S.M.-F. designed the study, analyzed data, and wrote the paper. All authors discussed the results and commented on the manuscript.

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Correspondence to Shannon McKinney-Freeman.

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Ganuza, M., Chabot, A., Tang, X. et al. Murine hematopoietic stem cell activity is derived from pre-circulation embryos but not yolk sacs. Nat Commun 9, 5405 (2018). https://doi.org/10.1038/s41467-018-07769-8

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Received: 08 March 2018
Accepted: 23 November 2018
Published: 20 December 2018
DOI: https://doi.org/10.1038/s41467-018-07769-8

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