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. Author manuscript; available in PMC: 2015 Dec 8.
Published in final edited form as: Dev Cell. 2014 Dec 8;31(5):640–653. doi: 10.1016/j.devcel.2014.11.007

Interferon gamma Signaling Positively Regulates Hematopoietic Stem Cell Emergence

Suphansa Sawamiphak 1, Zacharias Kontarakis 1, Didier YR Stainier 1
PMCID: PMC4371141  NIHMSID: NIHMS645431  PMID: 25490269

Summary

Vertebrate hematopoietic stem cells (HSCs) emerge in the aorta-gonad-mesonephros (AGM) region from “hemogenic” endothelium. Here we show that the pro-inflammatory cytokine Ifn-γ and its receptor Crfb17 positively regulate HSC development in zebrafish. This regulation does not appear to modulate the proliferation or survival of HSCs or endothelial cells, but rather the endothelial to HSC transition. Notch signaling and blood flow positively regulate the expression of ifng and crfb17 in the AGM. Notably, Ifn-γ overexpression partially rescues the HSC loss observed in the absence of blood flow or Notch signaling. Importantly, Ifn-γ signaling acts cell-autonomously to control the endothelial to HSC transition. Ifn-γ activates Stat3, an atypical transducer of Ifn-γ signaling, in the AGM, and Stat3 inhibition decreases HSC formation. Together, our findings uncover a developmental role for an inflammatory cytokine and place its action downstream of Notch signaling and blood flow to control Stat3 activation and HSC emergence.

Introduction

Hematopoietic stem cells (HSCs) are multipotent cells with the self-renewal potential to replenish all blood cell types throughout life. Their embryonic endothelial cell (EC) origin was proposed decades ago (Murray, 1932; Sabin, 2002); however, in amniotes, the process of embryonic endothelial to HSC conversion has only recently been revealed by live-imaging of the mouse aorta (Boisset et al., 2010). Importantly, the detailed progression of this specific cell transition, in which ECs in the ventral wall of the dorsal aorta initiate the hematopoietic program and enter circulation, has also been documented in zebrafish (Bertrand et al., 2010; Kissa and Herbomel, 2010; Lam et al., 2010). These findings not only confirmed the anatomical and functional equivalence of the zebrafish and mammalian aorta-gonad-mesonephros (AGM) region (Taoudi and Medvinsky, 2007), but also suggested that similar molecular pathways might be operating amongst different vertebrates to regulate this endothelial to HSC conversion (Clements and Traver, 2013).

A large number of studies in HSC biology have focused on the identification and study of transcription factors, cytokines, and other signaling molecules that are capable of promoting HSC survival, proliferation or self-renewal (Sorrentino, 2004; Walasek et al., 2012). The major proinflammatory cytokine IFN-γ have been widely viewed as negative regulators of HSC self-renewal. This notion might not be entirely unexpected particularly under certain stress conditions such as chronic infection, when robust cell proliferation and differentiation to restore blood and other immune cell types could drive dormant HSCs to exhaustion (Cheng et al., 2000; Sato et al., 2009). More intriguingly, both pro- and anti-proliferative effects of IFN-γ on HSCs have been reported (Yang et al., 2005; Baldridge et al., 2010; King et al., 2011; de Bruin et al., 2013). These seemingly opposite findings on the relationship between this inflammatory cytokine and HSC responses led us to speculate that its mode of action might be broader than anticipated, and that environmental cues might influence its biological effects. Importantly, involvement of IFN-γ in developmental hematopoiesis has not yet been investigated.

Here, we show that Ifn-γ signaling plays an unexpected role in HSC development. Embryonic endothelial to HSC conversion is positively regulated by Ifn-γ signaling, albeit HSCs do not appear to need Ifn-γ signaling for their survival or proliferation in their site of origin. We show that Notch signaling and blood flow implement at least part of their effect on HSC emergence by controlling the expression of Ifn-γ signaling components in the AGM region. Induction of Ifn-γ signaling in ECs is able to confer HSC fate in a cell-autonomous manner. Furthermore, we show that Ifn-γ signaling activates Stat3, which is required for HSC formation. Altogether, our work reveals a previously unappreciated role for an inflammatory cytokine in HSC development.

Results

Ifng1-2-induction of HSC generation

Initially, in order to investigate the role of Ifn-γ in macrophage development, we generated a transgenic line expressing a zebrafish Ifn-γ homolog, Ifng1-2, tagged with a V5 epitope under control of the heat-inducible hsp70l promoter. We included a cryaa-promoter-driven Cerulean expression cassette in the construct (Fig. 1a) to facilitate identification of the transgenic embryos by virtue of their fluorescent blue lens. V5-tagging does not significantly affect the biological activity of Ifng1-2 as shown by the ability of Ifng1-2-V5 to induce IFN-γ target gene expression (Fig. S1a). After heat-shock, strong expression of the V5 epitope was detected in numerous cells all over the trunk of Tg(hsp70l:ifng1-2-V5), abbreviated Tg(hsp:ifng1-2), embryos (Fig. 1 c). No Ifng1-2-V5 immunostaining was detected after heat-shocking embryos that do not carry the hsp70l:ifng1-2-V5 transgene (Fig. 1b). Ifng1-2-V5 positive cells were also found in the vicinity of the AGM, where the definitive HSCs emerge, as marked by the fluorescent axial vasculature in the kinase insert domain receptor (also known as Vegfr2) like (kdrl) reporter line Tg(kdrl:Hsa.HRAS-mCherry), abbreviated Tg(kdrl:HRAS-mCherry) fish (Chi et al., 2008).

Figure 1. Ifng1-2 and its receptor Crfb17 positively regulate HSC development.

Figure 1

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a, Schematic drawing of construct for heat-shock-inducible expression of ifng1-2. The area imaged and analyzed in all experiments (red box) is shown in the embryo illustration. b–c, Immunofluorescent labeling of V5-tagged Ifng1-2 (blue) in the vicinity of the AGM of control animals lacking the hsp:ifng1-2-V5 transgene (b) compared to Tg(hsp:ifng1-2) animals (c). The AGM region is recognized by Tg(kdrl:HRAS-mCherry) expression (red) in the vasculature. d–f, Increased number of HSCs at the ventral wall of the dorsal aorta and in the cardinal vein lumen upon ifng1-2 overexpression. HSCs (white arrowheads) in control (d) and Tg(hsp:ifng1-2-V5) (e) embryos were fluorescently labeled by Tg(itga2b:EGFP) (green) and Tg(kdrl:HRAS-mCherry) (red) expression. f, Number of HSCs per 500 µm aortic length. Values represent means +/− SEM. n=21–22 embryos. * p ≤ 0.05. All embryos were heat-shocked at 24 and 48 hpf, and imaged at 52–54 hpf. Of note, the numbers of Tg(itga2b:EGFP)+;Tg(kdrl:HRAS-mCherry)− (only green) pronephric duct cells appear unaffected by Ifng1-2 overexpression. g–i, Impaired HSC development in crfb17−/− embryos. Control (crfb17+/+) (g) and crfb17−/− (h) siblings were imaged at 52–54 hpf and itga2b:EGFP+kdrl:HRAS-mCherry+ HSC (white arrowheads) numbers were analyzed prior to genotyping. i, HSCs per 500 µm aortic length. Values represent means +/− SEM. n=8–14 embryos ** p ≤ 0.01. j–l, Ifng1-2 knockdown recapitulates the HSC phenotype of crfb17 mutants. itga2b:EGFP+kdrl:HRAS-mCherry+ HSC (white arrowheads) in mock (1% phenol red in nuclease-free distilled water)-injected (j) or ifng1-2 MO-injected (k) embryos imaged at 52–54 hpf. Numbers of HSCs per 500 µm aortic length is shown in l. Values represent means +/− SEM. n=32–40 embryos. *** p ≤ 0.001. m–o, Reduction of runx1 expression upon Ifng1-2 knockdown is restored by ifng1-2 induction. Expression of the HSC marker runx1 in mock-injected (m), and ifng1-2 MO-injected embryos without (n) and with (o) the hsp70:ifng1-2-V5 transgene. p–q, ifng1-2 overexpresssion is unable to rescue Crfb17 knockdown. runx1 expression in crfb17 MO-injected embryos without (p) and with (q) the hsp70:ifng1-2-V5 transgene. All embryos were heat-shocked at 24 hpf and runx1 expression was assessed by in situ hybridization at 36 hpf. The number of embryos showing the representative phenotype per total number of embryos analyzed is indicated in the lower left corner. Red brackets identify the dorsal aorta (DA), blue brackets identify the cardinal vein (CV). All images are lateral views, dorsal up, anterior to the left.

Surprisingly, overexpression of ifng1-2-V5 caused an increase in the number of cells expressing Tg(-6.0itga2b:EGFP) (abbreviated Tg(itga2b:EGFP) (Traver et al., 2003). While itga2b (also known as cd41) expression is one of the earliest markers of HSC commitment (Mikkola et al., 2003; Bertrand et al., 2008), it is also expressed in pronephric cells (in zebrafish) and thrombocytes (in mouse and zebrafish) (Bertrand et al., 2008; Zhang et al., 2007). To further examine the possible involvement of Ifng1-2 in HSCs development, we crossed the Tg(hsp:ifng1-2) line to Tg(itga2b:EGFP);Tg(kdrl:HRAS-mCherry) animals in which HSCs can be distinguished from other Itga2b+ cell types by virtue of their co-expression of cytoplasmic EGFP and plasma membrane HRAS-mCherry, owing to their endothelial origin. When comparing HSCs that had emerged from the aortic floor or migrated ventrally into the subaortic space or cardinal vein during definitive hematopoiesis in control embryos (11.8±1.1 cells/500µm aortic length), we found that Ifng1-2-V5 overexpression resulted in an approximately 31% increase in HSC number (15.4±1.2 cells/500µm aortic length) (Fig. 1d–f).

Ifng1-2 and its receptor Crfb17 positively regulate HSC development

Studies in mammalian models have provided essential molecular details of IFN-γ signaling. A functional unit of IFN-γ receptor (IFNGR) is a tetramer consisting of two IFNGR1 and two IFNGR2 chains. IFN-γ ligands bind as dimers with high-affinity to IFNGR1 to initiate signal transduction (Pestka et al., 2004). In zebrafish, the transmembrane receptor Crfb17 possesses the conserved JAK1 and STAT1 binding motifs found in IFNGR1 and is responsible for Ifng1-2 signal transduction (Aggad et al., 2010). To test the requirement of Ifng1-2 signaling for HSC development, we generated a crfb17 mutant using transcription activator-like effector nucleases (TALEN) (Bedell et al., 2012) (Fig. S1b–c). In comparison to control (crfb17+/+) siblings, in which HSCs were readily observed at the ventral wall of the dorsal aorta (13.3±1.0 cells/500µm aortic length), the aortic floor of crfb17−/− embryos harbored substantially fewer HSCs (7.5±1.1 cells/500µm aortic length) (Fig. 1g–i).

In agreement with this hematopoietic deficiency in crfb17−/− embryos, morpholino (MO)-mediated Ifng1-2 knockdown resulted in a decrease in the number of HSCs found within the AGM region (3.9±0.4 cells/500µm aortic length) when compared to mock-injected control siblings (10.2±0.7 cells/500µm aortic length) (Fig. 1j–l). Furthermore, we found that the dorsal aorta expression of the HSC marker runx1 was diminished in ifng1-2 morphants (Fig. 1m–n). The specificity of the ifng1-2 MO was addressed by testing the ability of ifng1-2-V5 overexpression to rescue runx1 expression in these morphants (Fig. 1o).

MO-mediated knockdown of Crfb17 similarly reduced runx1 expression (Fig. 1m, p). Importantly, this effect could not be rescued by ifng1-2-V5 overexpression (Fig. 1q), indicating that Crfb17 is required to mediate Ifng1-2 signaling during HSC development.

Ifn-γ signaling does not appear to modulate the proliferation or survival of HSCs or ECs in the AGM region

The alteration of HSC number observed upon Ifn-γ signaling modulation could be the result of changes in cell proliferation and/or cell death. To test these possibilities, we first analyzed the cell-cycling activity of HSCs in the AGM region after ifng1-2-V5 overexpression. PCNA (produced during G1, abundant in S, and declining in G2/M phase), immunofluorescence showed no significant changes in the percentage of itga2b:EGFP+kdrl:HRAS-mCherry+ cells that exited the quiescent state of the cell-cycle in Tg(ifng1-2-V5) (53.66±4.03%) as compared to control (55.35±6.12%) animals (Fig. 2a–c). Live-imaging of zebrafish embryos (Bertrand et al., 2010; Kissa and Herbomel, 2010) has previously revealed that the generation of HSCs from specialized aortic ECs occurs via transdifferentiation. Nevertheless, to test a potential effect of Ifn-γ signaling on EC division, we assessed EC cell-cycling rate upon ifng1-2-V5 overexpression. Interestingly, we detected fewer PCNA+Kdrl+ cells in Tg(hsp:ifng1-2) embryos (21.44±1.67 cells/500 µm aortic length) as compared to controls (26.54±1.50 cells/500 µm aortic length) (Fig. S2a–c). Therefore, the increase of HSCs observed within the AGM area caused by elevated levels of Ifng1-2-V5, does not appear to be caused by a change in HSC or EC proliferation in this region.

Figure 2. Cell proliferation and survival in the axial vessels do not appear to be affected by gain-or loss-of-function of Ifng1-2 signaling.

Figure 2

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a–c, ifng1-2 overexpression increases HSC number without affecting cell division. HSCs (itga2b:EGFP+kdrl:HRAS-mCherry+, green+ red+) that exit the G0 phase, labeled by PCNA immunostaining (blue), in control embryos not harboring the Tg(hsp:ifng1-2-V5) transgene (a), and Tg(hsp:ifng1-2-V5) embryos (b) are indicated by white circles. PCNA− HSCs are marked with white arrowheads. c, 32 Percentage of PCNA+ HSCs per total HSCs in the dorsal aorta. Values represent means +/− SEM. n=9 embryos per group. n.s. not significant (p > 0.05). d–m, Ifng1-2 and Crfb17 knockdown has no effect on apoptosis. Apoptotic cells (green, white arrowheads) visualized by Tg(Tbp:GAL4 UAS:secA5-YFP) expression (d–i) and TUNEL assay (j–m) in the axial vessels of MOCK-(d, g, j), crfb17 MO-injected (e, l), and ifng1-2 MO-injected (h, k) embryos at 52–54 hpf. Numbers of Tbp:GAL4;UAS:secA5-YFP+ cells (f, i) and TUNEL+kdrl:HRAS-mCherry+ cells per 500 µm dorsal aorta length (m) are shown as means +/− SEM. n=11–25 embryos. n.s. p > 0.05. All images are lateral views, dorsal up and anterior to the left.

We next tested whether the reduction in HSCs caused by Ifng1-2 or Crfb17 knockdown could be attributed to increased cell death. To visualize apoptotic cells in live embryos, we used a transgenic line expressing YFP tagged with the phosphatidylserine-binding protein Annexin V (secA5-YFP) under the control of a ubiquitous TATA-box binding protein (TBP) promoter (van Ham et al., 2010). Using this tool, we found that the numbers of apoptotic cells within the axial vessels along the AGM region were comparable in ifng1-2 morphants (8.3±0.6 cells/500µm aortic length), crfb17 morphants (6.0±1.1 cells/500µm aortic length) and control mock-injected siblings (6.6±0.7 and 7.7±0.7 cells/500µm aortic length, respectively) (Fig. 2d–i). Similarly, terminal deoxynucleotidyl transferase (TUNEL)-mediated labeling showed that Ifng1-2 or Crfb17 knockdown did not significantly alter EC apoptosis (6.5±1.0 cells and 7.3±1.5/500µm aortic length, respectively) compared to control animals (5.6±1.1 cells/500µm aortic length) (Fig. 2j–m). Similarly, we did not observe any significant difference in the apoptotic rate of HSCs (Kdrl+Itga2b+) upon Ifng1-2 knockdown (Fig. S2d–f). Together, these results indicate that increased cell death is not a cause of HSC decrease following the disruption of Ifng1-2/Crfb17 signaling.

Ifn-γ positively regulates HSC emergence

Since we did not find evidence for the involvement of Ifng1-2 signaling in the proliferation or survival of HSCs or ECs in the AGM region, we directed our attention for the cellular mechanisms underlying the change in HSC number to the process of EC-to-HSC transition. We performed time-lapse imaging of Tg(hsp:ifng1-2) embryos and control siblings to follow HSC emergence from the dorsal aorta. Starting from 8 h after heat-induction of ifng1-2-V5 expression, the emergence of HSCs, evident by the initiation of itga2b:EGFP expression in kdrl:HRAS-mCherry+ ECs at the aortic floor, was more frequent in Tg(hsp70l:ifng1-2-V5) embryos over the entire 14.6 h observation time (movie S1–2, Fig. 3a–f).

Figure 3. Ifng1-2 signaling positively regulates HSC emergence.

Figure 3

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a–f, Higher frequency of ECs at the aortic floor that turn on HSC marker expression upon ifng1-2 overexpression. Still images captured at different time points during time-lapse imaging of control (lacking the hsp:ifng1-2-V5 transgene) (a–c) and tg(hsp:ifng1-2-V5) (d–f) embryos starting at 33 hpf, 8h after heat-shock (see also movies S1 and S2). White arrowheads point to HSCs, labeled by Tg(itga2b:EGFP) (green) and Tg(kdrl:HRAS-mCherry) (red) expression, just arisen from the aortic wall at each indicated time point. g–l, Ifng1-2 knockdown reduces the emergence of HSCs and consequently the development of myeloid and lymphoid cells. runx1 expression was assessed during initiation of definitive HSC development at 26 hpf in mock-injected (g) and ifng1-2 MO-injected (h) embryos. Myeloid marker mpx1 expression in 48 hpf mock-injected (i) and ifng1-2 MO-injected (j) embryos. Lymphoid marker rag1 expression in the developing thymus (black circle) of mock-injected (k) and ifng1-2 MO-injected (l) 4 dpf larvae. m–p, Elevated expression of runx1 upon ifng1-2 overexpression. runx1 expression in 36–38 hpf wild-type (WT) embryos heat-shocked (h.s.) at 10–12 hpf (m) or 20–22 hpf (o) and Tg(hsp:ifng1-2) embryos heat-shocked at 10–12 hpf (n) or 20–22 hpf (p). The number of embryos showing the representative phenotype per total number of embryos analyzed is indicated in the lower left corner. Red brackets identify the dorsal aorta (DA), blue brackets identify the cardinal vein (CV). All images are lateral views, dorsal up and anterior to the left.

Since Runx1 is a critical regulator of the endothelial-to-HSC transition (Chen et al., 2009), we checked whether runx1 expression during the earliest establishment of HSC fate (i.e., just after the onset of blood circulation) was affected by Ifng1-2 signaling. MO-mediated knockdown of Ifng1-2 resulted in a reduction of runx1 expression in 26 hpf (hours post fertilization) embryos (Fig. 3g–h). Consequently, the development of hematopoietic progenitor cells including myeloid lineage cells, marked by mpx1 expression, in the caudal hematopoietic tissue at 48 hpf (Fig. 3i–j) and lymphoid lineage cells, marked by rag1 expression, in the developing thymus at 4 dpf (days post fertilization) (Fig. 3 k–l) was severely compromised in ifng1-2 morphants. Conversely, heat-induction of ifng1-2 expression was able to elevate runx1 transcripts within the AGM region when heat-shock was performed at 20–22 hpf, but not at 10–12 hpf (Fig. 3m–p, S3). Together, our findings suggest that Ifng1-2 signaling functions as a positive regulator of HSC specification.

Ifn-γ signaling acts downstream of Notch signaling in HSC development

Notch signaling is known to play crucial roles during the establishment of HSC fate. Mice and zebrafish lacking components of Notch signaling including Notch1, Jagged1, Rbpj, and Mind bomb (a ubiquitin ligase essential for activation of Notch signaling) fail to execute definitive hematopoiesis at the AGM region (Kumano et al., 2003; Burns et al., 2005; Robert-Moreno et al., 2005; Robert-Moreno et al., 2008). Notch was proposed to act upstream of Runx1 to specify HSCs since Runx1 induction in a Notch signaling deficient background can rescue the loss of HSCs (Burns et al., 2005; Nakagawa et al., 2006). These studies and the developmental hematopoiesis role of Ifng1-2 signaling uncovered here prompted us to investigate the connection between Notch and Ifng1-2 signaling in HSC specification.

To investigate possible interaction between Ifng1-2 and Notch signaling, we first tested whether blockage of Notch signaling would exacerbate the HSC defect caused by Crfb17 deficiency. Wild-type embryos treated with N-[N-(3,5-difluorophenacetyl)-l-alanyl]-S-phenylglycine t-butyl ester (DAPT), which blocks Notch intracellular domain (NICD) release from the plasma membrane, developed fewer HSCs (8.05±0.49 cells/500 µm aortic length) than DMSO-treated siblings (12.63±0.77 cells/500 µm aortic length) (Fig. 4a–b, e), in agreement with previous observations (Kim et al., 2012). By contrast, HSC numbers in crfb17−/− embryos treated with DAPT (7.16±0.62 /500µm aortic length) were not significantly different from those treated with DMSO (7.01±0.34 cells/500µm aortic length) (Fig. 4c–e), indicating that Notch and Ifng1-2 function in the same pathway. Furthermore, the potency of NICD overexpression in Tg(-1.5hsp70l:Gal4; 5xUAS-E1b:6xMYC-notch1a), abbreviated Tg(hsp:GAL4; UAS:NICD), to induce HSCs in mock-injected control embryos (from 9.86±0.96 to 16.03±2.27 cells/500µm aortic length) was markedly reduced when Ifng1-2 was knocked down (4.87±1.33 cells/500µm aortic length) (Fig. 4f–i). More importantly, we found that the HSC defect caused by DAPT-mediated Notch inhibition could be rescued by ifng1-2-V5 overexpression. HSC numbers in embryos with reduced Notch but enhanced Ifng1-2 signaling (12.41±0.89 cells/500µm aortic length) were comparable to those seen in control siblings (11.76±1.06 cells/500µm aortic length) (Fig. 4j–m). Correspondingly, ifng1-2-V5 overexpression restored the loss of HSCs caused by MO-mediated knockdown of mind bomb (mib) (Fig. S4 a–d).

Figure 4. Ifng1-2 functions downstream of Notch signaling during HSC development.

Figure 4

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a–e, Notch inhibition has no additional effect on HSC impairment in crfb17 mutants. HSCs, identified by Tg(itga2b:EGFP) (green) and Tg(kdrl:HRAS-mCherry) (red) expression (white arrowheads), in DMSO-treated (a) or DAPT-treated (b) wild-type (WT) and DMSO-treated (c) or DAPT-treated (d) crfb17−/− embryos. Siblings were imaged at 52–54 hpf and HSC numbers were analyzed prior to genotyping. e, itga2b:EGFP+kdrl:HRAS-mCherry+ cells per 500 µm aortic length. Values represent means +/− SEM. n=12–13 embryos. f–i, Reduced efficiency of Notch signaling to induce HSCs in the absence of Ifng1-2. itga2b:EGFP+ cells (green) in the ventral wall of the dorsal aorta of mock-injected control (f) and Tg(hsp:GAL4;UAS:NICD-myc) embryos (g), and ifng1-2 MOinjected Tg(hsp:GAL4;UAS:NICD-myc) embryos (h). Elevated NICD was 34 visualized by Tg(Tp1:H2B-mCherry) expression (red). White arrowheads point to itga2b:EGFP+Tp1:H2B-mCherry+ cells. i, itga2b:EGFP+Tp1:H2B-mCherry+ cells per 500 µM aortic length. Values represent means +/− SEM. n=8–24 embryos. * p ≤ 0.05, ** p ≤ 0.01, *** p ≤ 0.001. j–m, ifng1-2 overexpression rescues the reduction of HSCs caused by Notch inhibition. itga2b:EGFP+ (green) kdrl:HRAS-mCherry+ (red) HSCs (white arrowheads) in control animals treated with DMSO (j) and DAPT (k) and DAPT-treated Tg(hsp:ifng1-2) animals (l). HSC numbers per 500 µm aortic length are shown as means +/− SEM. n=22–33 embryos. n.s. p > 0.05, * p ≤ 0.05, ** p ≤ 0.01, *** p ≤ 0.001. All heat-shock treatments were done at 24 and 48 hpf and all embryos were imaged at 52–54 hpf. n–q, Inhibition of Notch signaling down-regulates crfb17 and ifng1-2 expression. crfb17 (n, o) and ifng1-2 (p, q) expression in 36 hpf embryos treated with DMSO (n, p) or DAPT (o, q) starting at 24 hpf. The number of embryos showing the representative phenotype per total number of embryos analyzed is indicated in the lower left corner. Red brackets identify the dorsal aorta (DA), blue brackets identify the cardinal vein (CV). All images are lateral views, dorsal up and anterior to the left.

Many genes with pivotal roles in diverse developmental processes have been identified as Notch transcriptional targets (Hamidi et al., 2011). Thus, we examined whether Notch signaling could also modulate ifng1-2 or crfb17 expression. Inhibition of Notch signaling with 12 h-DAPT treatment resulted in a reduction of ifng1-2 and crfb17 transcripts, typically present in the axial vessels of DMSO-treated control embryos (Fig. 4n–q, S4e). Expression of other homologs of IFN-γ and IFNGR1, ifng1-1 and crfb13, respectively (Aggad et al., 2010), appeared unaffected by Notch inhibition (Fig. S4f). Recent findings indicate that Notch signaling functions downstream of retinoic acid signaling to mediate hemogenic EC specification through cell-cycle control. Retinoic acid-deficient mice show a reduction in the proportion of ECs in the G0 phase and a concomitant increase in the S/G2/M fraction (Marcelo et al., 2013). Therefore, we tested whether ifn-γ signaling-deficient animals also exhibited this phenotype. Indeed, MO-mediated Ifng1-2 or Crfb17 knockdown resulted in an increase in the number of Pcna+ ECs (Fig. S4g–j), suggesting that Ifng signaling might be required to restrain ECs in a quiescent state. Collectively, these findings suggest a signaling network in which Ifng1-2 signaling acts downstream of Notch signaling during HSC specification.

Blood flow-regulation of HSC development is mediated, in part, by Ifn-γ signaling

Another key regulator of HSC development is blood flow (North et al., 2009). To investigate the relationship between blood flow and Ifng1-2 signaling, we checked whether Ifng1-2 induction could rescue HSC loss in the absence of blood flow. Embryos injected with tnnt2 (previously known as silent heart) MO (Sehnert et al., 2002) lack a heartbeat and consequently show disturbed vascular development and a reduction of HSCs in the AGM (2.80±0.61 cells/500µm aortic length) as compared to mock-injected controls (10.02±0.84 cells/500µm aortic length). Overexpression of ifng1-2-V5 partially restored HSCs in tnnt2 morphants (5.67±0.38 cells/500µm aortic length) (Fig. 5a–d). Correspondingly, the decrease of runx1 expression in tnnt2 morphants could be rescued by ifng-1-2-V5 overexpression (Fig. 5e–g). The moderate efficiency of ifng1-2-V5 overexpression to restore HSCs in the absence of circulation led us to test whether flow affects the expression of Ifn-γ signaling components in hematopoietic tissues. Indeed, we found that the expression of both ifng1-2 and crfb17 was nearly absent from the axial vessels of tnnt2 morphants (Fig. 5h–k). This reduction in ifng1-2 and crfb17 mRNA expression in flow-deficient embryos was further confirmed by qPCR (Fig. 5l). The role of Ifn-γ/Crfb17 in HSC formation and the requirement of blood flow for their expression in the AGM suggests that Ifn-γ signaling is a part of the blood flow-regulated mechanism underlying HSC development.

Figure 5. Reduced Ifng1-2 signaling is partially responsible for the HSC developmental defects caused by the lack of blood flow.

Figure 5

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a–d, ifng1-2 overexpression partly restores HSCs in embryos with no circulation. itga2b:EGFP+ (green) kdrl:HRAS-mCherry+ (red) HSCs (white arrowheads) in mock-injected controls (a), and tnnt2 MO-injected embryos without (b) and with 35 (c) hsp:ifng1-2 transgene. All embryos were heat-shocked at 24 and 48 hpf and imaged at 52–54 hpf. d, HSC numbers per 500 µm aortic length are shown as means +/− SEM. n=12–20 embryos. ** p ≤ 0.01, *** p ≤ 0.001. e–g, Loss of runx1 expression from the lack of blood flow is rescued by ifng1-2 overexpression. runx1 expression in mock-injected controls (e), and tnnt2 MOinjected embryos without (f) and with (g) the hsp:ifng1-2 transgene. All embryos were heat-shocked at 24 hpf and harvested at 36 hpf. h–k, crfb17 and ifng1-2 expression depends on blood flow. crfb17 (h, i) and ifng1-2 (j, k) expression in mock-injected (h, j) and tnnt2 MO-injected (i, k) 36 hpf embryos. The number of embryos showing the representative phenotype per total number of embryos analyzed is indicated in the lower left corner. Red brackets identify the dorsal aorta (DA), blue brackets identify the cardinal vein (CV). All images are lateral views, dorsal up and anterior to the left. qPCR assay of ifng1-2 and crfb17 expression at 36 hpf in tnnt2 MO-injected embryos compared to mock-injected siblings. n=3, 30 embryos per sample. * p ≤ 0.05

Cell-autonomous action of Ifn-γ signaling in HSC emergence

Notch was reported to function cell-autonomously in HSC specification (Hadland et al., 2004). Our finding that Ifn-γ might act downstream of Notch signaling during this process prompted us to check whether ECs must experience Ifn-γ signaling to become HSCs. Therefore, we generated a nls-Cerulean:14xUAS:crfb17 construct (Fig. 6a) utilizing a bidirectional 14xUAS promoter to drive the expression of crfb17 and nuclear-localized Cerulean in a cell-type specific manner. We injected the nls-Cerulean:14xUAS:crfb17 construct or a control construct nls-Cerulean:14xUAS into Tg(cdh5:GAL4FF) embryos to induce crfb17 expression in ECs. We observed higher percentages of Crfb17 overexpressing cells activating the HSC marker itga2b:EGFP (86.70±4.62) when compared to Cerulean overexpression alone (55.10±8.12) (Fig. 6b–d), suggesting that Crfb17 functions cell-autonomously in the aortic endothelium.

Figure 6. Crfb17 signaling acts cell-autonomously to regulate HSC emergence.

Figure 6

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a–d, crfb17 overexpression in endothelial cells induces HSC emergence. a, Schematic drawing of the bidirectional 14xUAS-driven nls-Cerulean and crfb17 expression construct. Tg(cdh5:GAL4FF) embryos were injected with the control construct nls-cerulean:14xUAS (b) or the nls-Cerulean:14xUAS:crfb17 construct (c). nls-Cerulean+ (red) cells that turned on the HSC marker itga2b:EGFP+ (green) are indicated by white asterisks. White 36 arrowheads point to itga2b:EGFP−nls-Cerulean+ cells. d, percentages of itga2b:EGFP+nls-Cerulean+ per total nls-Cerulean+ cells are shown as means +/− SEM. n=13–16 embryos. ** p ≤ 0.01. e–h, Crfb17 is required cell-autonomously in the HSC lineage. e, Schematic drawing of the transplantation assay. 52–54 hpf embryos with transplanted cells from control (crfb17+/+) (f) or crfb17−/− (g) embryos carrying the Tg(itga2b:EGFP) (green) and Tg(kdrl:HRAS-mCherry) (red) transgenes. White arrowheads point to green+red+ HSCs. h, Numbers of itga2b:EGFP+kdrl:HRAS-mCherry+ cells per 100 µm aortic length are shown as means +/− SEM. n=16–20 embryos. * p ≤ 0.05.

To further test this model, we carried out cell transplantations and found that crfb17−/− cells integrated into the dorsal aorta of wild-type (crfb17+/+) animals but showed reduced contribution to the host definitive HSCs (1.23±0.28 crfb17−/− as compared to 2.29±0.42 crfb17+/+ HSCs/100 µm) (Fig. 6e–h). Together these data indicate that signaling through Crfb17 acts cell-autonomously in dorsal aorta ECs during their differentiation to HSCs.

Stat3 activated by Ifn-γ signaling positively regulates HSC development

Most biological responses of IFN-γ signaling are thought to be mediated through STAT1 (Platanias, 2005). However, in certain settings, activation of STAT3 by IFN-γ has been reported (Caldenhoven et al., 1999; Qing and Stark, 2004). Interestingly, targeted deletion of STAT3 in hematopoietic tissues cause a reduction in the number of long-term (LT)-HSCs in the bone marrow (Mantel et al., 2012). However, mutation of the STAT1/3 binding site in Gp130, a receptor that mediates IL-6 signal transduction, does not affect HSC repopulation and self-renewal (Wang et al., 2012). Therefore, the ligand/receptor pair that activates STAT3 in HSC development remains unidentified.

To examine the possible involvement of Stat3 in the Ifng1-2/Crfb17-regulation of HSCs, we first checked whether Ifng1-2 induction could activate Stat3. Elevated levels of tyrosine-phosphorylated (i.e., activated) Stat3 were observed in the AGM following ifng1-2-V5 overexpression (Fig. 7a–d). Moreover, induction of Notch signaling by NICD overexpression similarly increased activated Stat3, whereas Notch inhibition by DAPT had the opposite effect (Fig. 7c–d). Total Stat3 levels were altered correspondingly (Fig. 7c, e).

Figure 7. STAT3 is activated upon ifng1-2 induction and positively regulates HSC development.

Figure 7

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a–b, ifng1-2 overexpression stimulates Stat3 activation in the AGM region. a, Immunofluorescent staining of phosphorylated Stat3 (pStat3, green) in embryos without (a) and with (b) the hsp:ifng1-2 transgene. All embryos were heat-shocked at 24 and 48 hpf and harvested at 52–54 hpf. The AGM region is recognizable by the vascular expression of kdrl:HRAS-mCherry (red). c–e, Ifng1-2 and Notch signaling regulate Stat3 activation. c, Western blot analysis of pStat3, total Stat3, and Actin expression in the dissected yolk-spanning region of the trunk. Dissected area is outlined by dashed lines in the embryo illustration. 36 hpf embryos were heat-shocked for 5 h and 8 h for ifng1-2 and NICD overexpression, respectively. DMSO and DAPT treatments were done from 24–48 hpf. d, Relative expression of pStat3 normalized to Actin. e, Relative expression of total Stat3 normalized to Actin. 37 Values represent means +/− SEM. n=3–4 independent experiments. n.s. p > 0.05, * p ≤ 0.05, ** p ≤ 0.01. f–i, Stat3 inhibition causes a reduction in HSC numbers which cannot be restored by ifng1-2 overexpression. itga2b:EGFP+ (green) kdrl:HRAS-mCherry+ (red) HSCs (white arrowheads) in control embryos (not carrying the hsp:ifng1-2-V5 transgene) treated with DMSO (f) or Stat3 inhibitor (inh) (g) and Tg(hsp:ifng1-2-V5) embryos treated with Stat3 inh (h). All embryos were heat-shocked at 24 and 48 hpf and imaged at 52–54 hpf. DMSO and Stat3 inh treatments started at 24 hpf. i, Numbers of itga2b:EGFP+kdrl:HRAS-mCherry+ cells per 500 µm aortic length are shown as means +/− SEM. n=19–24 embryos. n.s. p > 0.05, ** p ≤ 0.01.

We next treated embryos with the Stat3 inhibitor S3I-201, which blocks Stat3 phosphorylation by targeting its Src homology (SH2) domain, and found a decrease in the number of HSCs emerging from the AGM. Moreover, overexpression of ifng1-2-V5 was unable to rescue this defect (Fig. 7f–i). Taken together, these findings suggest that Stat3 acts as a mediator of Ifn-γ signaling to control HSC formation. A similar mechanism could be present in mammals, thereby explaining the reduction of LT-HSCs in STAT3 mutants.

Discussion

Ifn-γ in HSC development

We report here a role for the potent inflammatory cytokine Ifn-γ as a positive regulator of developmental hematopoiesis. During the early emergence of definitive HSCs at the dorsal aorta, Ifn-γ signals through the Crfb17 receptor to stimulate the emergence of HSCs. Furthermore, we find that Notch signaling and blood flow, known regulators of HSC development, are required for the expression of ifng1-2 and crfb17 in this hematopoietic tissue.

Extensive studies on the immune and inflammatory functions of IFN-γ have contributed to the general view that it mediates the differentiation of hematopoietic cells from HSCs that reside in the bone marrow at the expense of their self-renewal potential (Yang et al., 2005; Baldridge et al., 2010; MacNamara et al. 2011; de Bruin et al., 2013). Within the bone marrow, a composite of several cell types maintain a specialized microenvironment that regulates HSC homeostasis. In addition to the endothelial and mesenchymal stromal cells that are in close contact with HSCs, sympathetic nerves and osteolineage cells also provide molecular cues that affect the maintenance, differentiation and mobilization of HSCs (Morrison and Scadden, 2014). A clear example of the niche influence on IFN-γ function is the recent finding that IFN-γ secreted from cytotoxic T-cells acts on bone marrow mesenchymal stromal cells to induce IL-6 production, which then mediates myeloid differentiation from hematopoietic progenitor cells (Schurch et al., 2014). Therefore, HSCs are exposed to distinct signals at their site of origin in the AGM in comparison to their site of maintenance in the bone marrow, and these signals could affect the outcome of IFN-γ signaling.

Moreover, while previous studies have focused on the role of IFN-γ in quiescent adult HSCs, we now provide evidence for a cell-autonomous action of Crfb17 in hemogenic ECs. Under inflammatory stress, IFN-γ was reported to activate STAT1 and stimulate dormant LT-HSCs to enter the cell-cycle (Baldridge et al., 2010). Furthermore, the interferon regulatory factor (IRF)7, a transcriptional mediator of Ifng-Stat1 signaling (Farlik et al., 2012), negatively affects HSC development when its post-transcriptional inhibitor, miR-142a-3p, is knocked down (Lu et al., 2013). Our work, on the other hand, identifies Stat3 as a mediator of Ifn-γ signaling during the EC-to-HSC transition. Thus, the dual roles of Ifn-γ signaling in HSC specification and differentiation appear to be cell-type dependent and these distinct outcomes likely involve different effector molecules.

Notch and Ifn-γ signaling during HSC specification

We identified in our studies a Notch-Ifng1-2 signaling network in the regulation of HSC emergence. However, the relationship between Notch and Ifn-γ might be more complicated than the mere transcriptional control of Ifn-γ signaling components by Notch signaling. The involvement of other mechanisms in the reduction of ifng1-2 and crfb17 expression in the axial vessels following Notch inhibition reported here cannot be ruled out. One possibility is that this reduction might be a secondary effect of the loss of arterial identity, typically observed in Notch loss-of-function manipulations (Duarte et al., 2004). In support of this hypothesis, we found that the decrease of ifng1-2 and crfb17 expression following DAPT treatment was more noticeable in the dorsal aorta as compared to the cardinal vein (Fig. 4n–q). Therefore, additional and yet unidentified artery-specific or Notch-dependent proteins might modulate the transcription and/or post-transcriptional control of Ifn-γ signaling components.

The effect of Notch inhibition by DAPT treatment on HSC numbers observed here seems moderate when compared to the reported nearly complete loss of runx1 expression in mind bomb mutants (Burns et al., 2005; Gering and Patient, 2005). One of the possible explanations is the developmental stage when the treatment was applied. To avoid gross developmental abnormalities including the loss of some posterior somites (Zhang et al., 2007), we chose to apply 12–24 h treatments starting at 24 hpf. This timing also matches the developmental stage when Ifn-γ signaling functions in HSC formation, since we observed elevated runx1 expression levels after induction of ifng1-2 expression by heat-shock at 20–22 hpf, but not at 10–12 hpf (Fig. 3m–p). Furthermore, the role for Notch signaling in HSC development likely extends beyond the EC-HSC transition. It has recently been reported that activation of Notch signaling in endothelial progenitor cells, initiated by their interaction with somitic tissues and before the formation of the dorsal aorta, is required for HSC formation (Kobayashi et al., 2014). Therefore, Ifn-γ signaling most likely implements only part of the broad range of Notch signaling functions during definitive hematopoiesis.

Conservation of IFN-γ signaling function in vertebrates

IFNGR1-deficient mice show reduced mitotic activity in the bone marrow-residing quiescent HSCs following bacterial infection. Nevertheless, the total number of HSCs, sorted by Hoechst 33342-effluxed (so-called side population, SP) Kit+Sca+Lin− (SPKSL), was not found to be significantly altered when compared to wild-type animals (Baldridge et al., 2010). However, analysis of the HSC pool with an additional marker, CD150, whose expression levels correlate with self-renewing potential (Kiel et al., 2005), revealed a reduction in the proportion of the CD150+KSL population in these mutant mice (Baldridge et al., 2010). Moreover, the numbers of KSL CD150+CD48+CD34+CD135− cells, which represent a more differentiated progenitor pool, were also lower in the absence of IFNGR1 (MacNamara et al., 2011). Accordingly, a significant reduction in the numbers of KSL CD34− cells, which represent a pool of LT-HSCs, was reported in the bone marrow of mice lacking STAT3 (Mantel et al., 2012). Together, these reports on the decrease of certain hematopoietic stem/progenitor cell pools observed in adult mouse mutants indicate that the role of Ifng1-2 signaling in HSC emergence, reported here in zebrafish, could represent a mechanism common to other vertebrate models, and that its requirement cannot be fully compensated for over the course of development.

Materials and Methods

Fish care and strains

Embryos and adult fish were raised under standard conditions following the institution’s ethical guidelines. Please refer to the supplemental information for the transgenic lines used in this study.

Generation of transgenic and mutant lines

Expression constructs, transgenic and mutant lines were generated as described in the supplemental information.

In situ hybridization and morpholinos

Whole mount in situ hybridization was performed as described previously (Thisse and Thisse, 2008). Primers used for probe generation are described in the supplemental information. Antisense morpholinos for Ifng1-2 (Sieger et al., 2009), Crfb17 (Aggad et al., 2010), Mib (mib-m2-MO) (Zhang et al., 2007), and Tnnt2 (Sehnert et al., 2002) knockdown were purchased from Gene Tools and injected into 1- or 2-cell stage embryos at a concentration of 0.12 mM, 0.3 mM, 1 mM, and 0.3 mM or approximately 2 ng, 5 ng, 17 ng, and 5 ng, respectively.

Immunofluorescence staining

Embryos were fixed in 4% PFA and permeabilized by incubation with 10 µg/ml proteinase K at room temperature for 30 min. Immunofluorescence staining was carried out as described (Sawamiphak et al., 2010). Mouse anti-Pcna (Santa Cruz) were used at a dilution of 1:50.

Immunoblotting

Immunoblotting was performed as described (Sawamiphak et al., 2010). Mouse anti-PhosphoStat3 (MBL), rabbit anti-Stat3 (Santa Cruz), and rabbit anti-actin (Cell Signaling Technology) were used at a dilution of 1:500, 1:500, and 1:1000, respectively.

Embryo heat-shock and chemical treatments

Embryos were heat-shocked once at 24 hpf or twice at 24 and 48 hpf by incubating the cell-culture plates containing the embryos at 37°C for 1 h. The plates were then transferred back to a 28°C incubator until imaging or harvesting time. To inhibit Notch signaling, embryos were treated with 100 µM DAPT (Sigma) in Danieau solution. Siblings treated with 1% DMSO in Danieau solution were used as controls.

Time-lapse microscopy

Embryos were embedded in 1.2% low-melting agarose in glass-bottom dishes (MatTek), with their lateral side fully exposed. The dishes were then filled with Danieau buffer containing 0.016% Tricaine to anesthetize the embryos. For confocal time-lapse imaging, Z-stacks of 4-or 4.5-µm intervals were acquired from the AGM area every 10 or 15 min for 14.5–15 h using a 20× water immersion objective. Individual images were rendered into time-lapse movies with Fiji software.

Quantitative analysis of HSCs, apoptosis, and proliferation

Quantitative analysis was performed using the Fiji software. All confocal z-stacks were first converted into “sum slices” projections. Aortic lengths were measured along the ventral wall of the dorsal aorta across the microscopic field, which covered approximately 85% of the yolk extension span. HSCs, apoptotic and proliferating cells were counted and normalized to aortic length or calculated as percentages of total numbers of HSCs or ECs. Equality of variance between the experimental and control sample sets were analyzed using the F-test. The two-tailed unpaired Student’s t test was used to calculate significance.

Quantitative PCR

Total RNA isolated from 20–30 embryos was reverse transcribed using Superscript II (Life Technologies) and gene expression was assayed relative to rps11 and/or gapdh using an Eco Real-Time PCR System (Illumina, San Diego, CA). All reactions were performed in technical triplicates and the results represent biological replicates including the standard error of the mean, unless otherwise stated. Primers are listed in the supplemental information.

Supplementary Material

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Acknowledgements

We are grateful to Herwig Baier for support and scientific resources, David Traver and Nancy Speck for discussions, and Alessandro Filosa for critically reading the manuscript. We are also thankful to Dominique Foerster and Anna Kramer for reagent sharing. S.S. is financed by a Human Frontier Science Program Fellowship, and Z.K. by an EMBO fellowship. This work was funded in part by support from the NIH, Packard Foundation, and Max Planck Society to D.Y.R.S.

Footnotes

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