Bnip3lb-driven mitophagy maintains fate of the embryonic hematopoietic stem cell pool

Domain: chemistry
Published: 2026-02-24
Editorial outcome: accept

Abstract
Embryonic hematopoietic stem and progenitor cells (HSPCs) have the clinically valuable ability to undergo substantial proliferative expansion while maintaining multipotency, which remains difficult to replicate in culture. Here, we show that newly specified HSPCs achieve this unique state by precise spatio-temporal regulation of reactive oxygen species (ROS) via Bnip3lb-associated developmentally-programmed mitophagy, a distinct autophagic regulatory mechanism from that of adult HSPCs. While ROS drives HSPC specification in the dorsal aorta, scRNAseq and live-imaging of mitophagy-reporter zebrafish indicate that mitophagy initiates during endothelial-to-hematopoietic transition and colonization of secondary niches. Knockdown of bnip3lb reduces mitophagy and HSPC numbers in the caudal hematopoietic tissue by promoting myeloid-biased differentiation and apoptosis, which can be rescued by antioxidant exposure. Conversely, chemical or genetic induction of mitophagy enhances embryonic HSPC and lymphoid progenitor numbers. Significantly, compound-mediated mitophagy activation improves ex vivo function of HSPCs derived from human-induced pluripotent stem cells, enhancing serial-replating hematopoietic colony forming potential.

Introduction
Compared to adult HSCs, embryonic HSCs possess distinctive functional characteristics, including naïve immunity, which allows less rigorous matching for use in therapeutic transplantation and greater ability to proliferatively expand while maintaining full multipotent erythro-myeloid and lymphoid potential1. This notable difference implies activation of specialized mechanisms for establishing these attributes unique to early development. Here, we show that spatio-temporally programmed mitophagy initiated by Bnip3lb, a pathway not utilized by adult HSCs2, is the key to the precise regulation of metabolic state that grants embryonic HSCs their unusual maintenance properties.

In both the embryo and adult, Reactive Oxygen Species (ROS), byproducts of mitochondrial oxidative phosphorylation (OX/PHOS), act as potent regulators of HSC fate. Normally contained within the mitochondrial outer membrane, overproduction or loss of membrane integrity allows for ROS leakage into the cytoplasm, causing oxidative stress, which can impact or damage cellular components3. Despite this potentially negative attribute, ROS is an essential signaling molecule that influences the balance between hematopoietic proliferation and differentiation3–6. In the adult bone marrow (BM), quiescent HSCs maintain low ROS and glycolytic metabolism4,7,8, then transition to OX/PHOS metabolism, and subsequent ROS production, when cycling and/or driving maturation9. Although this switch is necessary for the production of lineage-restricted blood progenitors, excess or unregulated ROS can lead to exhaustion of the HSC pool via unrestricted differentiation or apoptosis10–19. Conversely, if ROS production is completely inhibited, HSCs are prevented from cycling and producing downstream progenitors, resulting in an inability to fight infection or maintain hematopoietic homeostasis7,15,20,21, particularly in the context of stem cell transplantation. Thus, metabolic state and ROS levels must be exquisitely balanced to support hematopoietic demand for the organism’s lifespan11,22.

Adult HSCs regulate ROS levels using a number of mechanisms, including maintaining quiescence, localizing within hypoxic niches, utilizing glycolytic metabolism15,16,23–27, and employing high levels of mitophagy9,10,28–33. The latter, a specialized form of cellular autophagy, is initiated when proteins on the outer membrane of a mitochondrion are tagged to mark the organelle for destruction or via context-driven transcriptional regulation of mitophagy receptor expression. Autophagy machinery constructs a membrane to surround the mitochondrion, which is then fused to a lysosome, allowing contents to be digested and recycled. Broadly, there are two major pathways controlling mitophagy: the PINK1-Parkin pathway and the receptor-mediated pathway, which can each be regulated both transcriptionally and post-transcriptionally to fine-tune ROS levels within the cell34. PINK1-Parkin-directed mitophagy occurs in tissues, including adult BM HSCs12,20,35, in response to damage and oxidative stress. Loss of membrane integrity leads to PINK1 accumulation on the mitochondrial outer surface36. PINK1 then phosphorylates Parkin, which draws in the autophagy machinery36–38. In contrast, receptor-mediated mitophagy occurs when autophagy machinery is activated by mitophagy receptors, such as FUNDC1, BNIP3, and BNIP3l (formerly NIX, which has two zebrafish homologs, Bnip3la and Bnip3lb), expressed on the surface of a mitochondrion39,40. Receptor-mediated mitophagy is responsive not only to oxidative stress41 but also to hypoxia42 and other developmental cues34; for example, BNIP3l/NIX-associated mitophagy is utilized to create space for hemoglobin in red blood cells2,43, or minimize interference to light in developing corneal lens fibers44–47. While damage-responsive mitophagy is well documented in adult HSCs, no role has been described for receptor-mediated mitophagy, although there is notable heterogeneity in autophagy pathways used by differentiating progenitors of various blood lineages48.

In contrast to detrimental roles generally ascribed to elevated ROS in the adult, our lab, and others, have shown that induction of ROS-mediated transcriptional activation is necessary for specification of HSPCs in the embryo. In this context, ROS elevation, coinciding with the onset of blood flow and initiation of embryonic OX/PHOS metabolism49, stimulates Hypoxia Inducible Factor 1a (HIF1α)-associated gene targets critical for hematovascular development50–52 and sterile inflammatory signaling via the NLRP3-inflammasome49. To explain this apparent shift in the regulatory impact of ROS on HSCs during early embryogenesis versus their mature state, it’s necessary to understand their developmental journey. After arising de novo from hemogenic endothelium (HE) in the ventral dorsal aorta (VDA), via an energetically demanding, metabolically active process termed endothelial-to-hematopoietic transition (EHT)53, HSCs immediately migrate to secondary niches: fetal liver (FL) sinusoids in mammals, or a vascular plexus in the tail called the caudal hematopoietic tissue (CHT) in zebrafish54. Following seeding, cells undergo proliferative expansion to produce sufficient HSCs to fill the terminal niche and aid in populating subsequent waves of myeloid, lymphoid, and erythroid lineages. Notably, during this expansion phase, FL HSCs utilize OX/PHOS metabolism more so than in the adult55. Following this window of exponential growth and differentiation, HSCs migrate to colonize the mammalian BM, or kidney marrow (KM) in fish, where they take on adult characteristics, such as quiescence56,57. The distinct environmental conditions of each niche, and unique activities undertaken by HSCs within them, presumably necessitate differential regulatory mechanisms to balance ROS, although little is known about the use of mitophagy prior to BM colonization.

The work described herein documents the shifting spatio-temporal impact of ROS in the vertebrate embryo and elucidates the key regulatory role of receptor-mediated mitophagy in establishing the lifelong HSPC pool. Through scRNAseq and in vivo fluorescent mitophagy reporter analysis, we show that mitophagy is initiated in nascent HSPCs during EHT, via developmentally programmed expression of the bnip3lb receptor. Gene knockdown studies confirm that embryonic induction of mitophagy is essential for maintenance of the self-renewing HSPC population within the CHT. Furthermore, antioxidant epistasis experiments indicate that mitophagy functions to protect the embryonic HSPC population by regulating ROS levels to balance maintenance of multipotency while allowing for proliferative expansion in the absence of apoptosis. Finally, we reveal that activation of mitophagy via genetic or chemical stimulation, such as via Nicotinamide Riboside (NR), can significantly enhance the numbers of HSPCs in vivo in the zebrafish embryo, as well as promote ex vivo maintenance of long-lived multipotent hematopoietic progenitors from human-induced pluripotent stem cells (iPSCs).

Results
ROS is essential for HSPC specification, but detrimental as HSPCs leave the VDA niche
We previously reported that elevation in ROS levels downstream of embryonic metabolic activation drives HSPC production in the VDA49,50. Given that sustained ROS stimulation negatively influences adult HSPC maintenance, we sought to understand at which developmental timepoint(s), and by what mechanism, the impact of ROS on embryonic HSPCs is regulated. To begin, we examined the effects of ROS manipulation on HSPCs in the specification niche (the VDA), compared to the expansion niche (the CHT), through targeted manipulation at discrete points in development (Fig. 1A). To elevate ROS levels in vivo, we microinjected morpholino oligonucleotides (MO) to knockdown expression of peroxiredoxin 1 (prdx1), an enzyme that acts to reduce hydrogen peroxide and other ROS50. Conversely, incubation with the antioxidant N-acetylcystine (NAC; 10 µM) was used to decrease ROS levels, as previously described50. In vivo alterations in ROS levels were measured via flow cytometry of disaggregated 72 hpf embryos using relative Mean Fluorescence Intensity (MFI) values of CellROX, a vital dye which fluoresces in the presence of ROS (Fig. 1B, left); use of the Tg(−6.0itga2b:EGFP)[referred to as CD41:EGFP] HSPC-reporter line58 allowed for quantification of ROS levels within CD41lo HSPCs, with statistically significant alterations across treatment groups noted (Fig. 1B, right), as expected. Consistent with our prior report50, increased ROS mediated by prdx1 knockdown promoted expression of the conserved HSPC markers runx1 and cmyb in the VDA at 36 hpf as determined by whole mount in situ hybridization (WISH); representative images of the beads-on-a-string expression pattern of the combined probes in the VDA and a graphic summary of the qualitative phenotypic distribution (high, medium, or low expression bins compared to matched controls) of total embryos scored per condition are shown (Fig. 1C, D, top row). Notably, by 72 hpf, after HSPCs migrated to the CHT, continued ROS elevation in prdx1 morphants substantially reduced runx1/cmyb expression (Fig. 1C, D, bottom row). Similar biphasic alterations in HSPC markers were observed when increasing ROS by H2O2 or 1% glucose treatment (Supplementary Fig. 1a, b). Specificity of impact on HSPCs in the CHT was confirmed by WISH for individual runx1 and cmyb probes (Supplementary Fig. 1c). In contrast, as opposed to negative effects on HSPC formation in the VDA (Fig. 1C, D, top row), exposure to NAC enhanced the appearance of HSPC markers in the CHT at 72 hpf (Fig. 1C, D, bottom row). This developmental flip was validated by in vivo imaging and further quantified by measuring the CD41:GFPlo HSPC population in the live cell population via flow cytometry58 (Fig. 1E, F; see Supplementary Fig. 1d for gating), confirming both a significant increase in the NAC-treated HSPC population and a significant decrease in the prdx1 morphant HSPC population at 72 hpf (Fig. 1F). Interestingly, the impact of ROS modulation at 48 hpf, a time point at which HSPCs continue to bud in the AGM, while simultaneously colonizing the CHT, is more ambiguous (Supplementary Fig. 1a, b, middle rows), reflecting a transitional phase for both HSPC localization and ROS regulation.

To assess whether ROS impacts the HSPC population at later stages of development, and/or address potential effects on HSPC-dependent lineage commitment, we analyzed embryos at 120 hpf. Knockdown of prdx1 significantly reduced CD41-labeled HSPCs in the CHT and KM, as confirmed by flow cytometry (Fig. 1G, H); morphants also had lower expression of rag1 in the thymus (Fig. 1I, J), which was confirmed by flow cytometry of Tg(rag2:GFP) embryos59 as a significant decrease in Rag2+ lymphocytes at 120 hpf (Fig. 1K and Supplementary Fig. 1e). Notably, prdx1 morphants showed only minor changes in expression of the neutrophil marker mpx at this time point by WISH or flow cytometry on Tg(mpx:GFP) embryos60, presumably reflective of primitive myeloid origin from earlier hematopoietic waves61. Interestingly, delayed initiation of NAC treatment, from 48-120 hpf, had little effect on CD41 expression in the KM (Fig. 1H), suggestive of physiological activation of ROS regulatory factor(s) as embryos mature. Together, these data indicate ROS levels must be precisely controlled in HSPCs as development progresses to properly seed terminal niches and maintain balanced differentiation capacity.

scRNAseq analysis shows bnip3lb expression increases in developing HSPCs
To understand how ROS transitions from boosting HSPC numbers in the VDA to reducing them in the CHT and KM, we performed comparative single-cell RNA sequencing (scRNAseq) analyses. Using established Tg(kdrl:Has.HRAS-mCherry)s916 vascular reporter embryos62, we microdissected and disaggregated trunk/tail regions, then collected Kdrl+ cells, including hemogenic endothelium and newly formed HSPCs, at 36 hpf and 48 hpf by FACS. Both datasets were processed using a standard Seurat workflow, including: log-normalization, selection of variable features, scaling of all genes, principal component analysis (PCA) on variable genes, construction of a shared nearest-neighbor graph, clustering, and UMAP embedding (Fig. 2A, 36 hpf; Fig. 2B, 48 hpf) using the first ten principal components63–65. Clusters were recoded into biologically interpretable labels: Endothelial Cells (EC), Hemogenic Endothelial Cells (HECs), Hematopoietic Stem Cells (HSCs), Erythroid (Ery), Macrophage (Mac), Mesenchymal Cell (MSC), Neutrophil (Neut), Neural progenitor (NP) based on differentially expressed genes and detection of established HSPC, endothelial, myeloid, and erythroid markers (e.g., cxcr4b, gata2b, itga2b, runx1, spi1b, lyz, mpx, gata1a, hbbe3, klf1, cdh5, kdrl, tie1) (Supplementary Fig. 2a–f). To quantify lineage-related programs, we computed module (activity) scores for three curated gene sets: endothelial (cdh5, kdrl, tie1), HSPC (cxcr4b, gata2b, itga2b, meis1b, mpl, myb, notch1b, runx1, spi1b), and myeloid (lyz, mpx, spi1b) (Supplementary Fig. 2c, d).

We performed differential expression analysis at 36 hpf between HSPCs vs HECs [cluster HSPC1 vs HECs] and HSPCs vs ECs [clusters HSPC1 vs EC1] and at 48 hpf between HSPCs vs ECs [clusters HSPC1, 2, and 3 merged vs EC1] using Seurat’s FindMarkers with default parameters. Gene-level statistics were then used as ranked input for GSEA, performed with clusterProfiler’s gseGO against Gene Ontology Biological Process terms using the zebrafish annotation database (org.Dr.eg.db). To harmonize library sizes between HSPCs from the two datasets, we downsampled raw RNA counts in selected HSPC clusters using the DropletUtils downsampleMatrix to a target of ~4600 UMIs per cell. Comparisons at both time points revealed a distinct change in metabolic and redox stress signatures as the HECs transitioned to HSPCs (Fig. 2C, D). Compared to either ECs or HECs, HSPCs show enrichment for “Cellular Respiration”, “Oxidative phosphorylation”, and “Mitochondrial ATP Synthesis Coupled to Electron Transport”, consistent with prior reports10,27,55. Furthermore, there is a strong upregulation of ROS-associated networks, such as “Reactive Oxygen Species Metabolic Process”, “Hydrogen peroxide metabolic/catabolic process”, and “Cell Redox homeostasis” in HSPCs compared to HECs. While at 48 hpf we can no longer detect a distinct transitional HEC fraction, when comparing mature HSPCs to ECs, upregulation of mitochondria-driven processes was maintained as a distinguishing feature. Notably, GO terms related to vascular differentiation and cell adhesion were significantly downregulated from ECs to HECs to HSPCs, consistent with their established developmental trajectory66 and supporting the physical and transcriptional changes driving EHT.

As mitophagy is an established mechanism to control oxidative phosphorylation and excess ROS production42, which can impact the balance between HSC maintenance and differentiation in the adult49,52, we directly examined expression of mitophagy-/hypoxia-associated signaling components and transcriptional regulators in the embryonic HSPC cluster at both 36 hpf and 48 hpf67–69. Interestingly, in contrast to adult HSCs, receptor-mediated mitophagy genes (bnip3, bnip3la, bnip3lb, fundc1) were generally expressed at a higher level within the HSPC subcluster than damage-responsive genes (pink1, parkin) at 36 hpf (Fig. 2E). Furthermore, while expression of mitophagy genes increased overall between 36 and 48 hpf, bnip3lb and fundc1 remained more highly expressed than pink1a and parkin in HSPCs at both timepoints (Fig. 2F). Prior reports have demonstrated direct transcriptional regulation of receptor-mediated mitophagy genes, including BNIP3 and BNIP3l by HIF1a42. Expression of the zebrafish paralogs hif1aa and hif1ab was detected in the HSPC fraction, with a strong upregulation noted in the latter across developmental time (Fig. 2F), consistent with previously reported roles in developmental HSC regulation across species51 and response to ROS stimulation52. Together, these gene expression studies are supportive of a role for directed upregulation of mitophagy in HSPCs during or immediately after EHT in response to physiological shifts in embryonic metabolic state and ROS production.

Bnip3lb is expressed in the HSPCs within the CHT, coinciding with increased mitophagy
To document the spatio-temporal onset of mitophagy as it relates to definitive hematopoiesis, we utilized two recently developed fluorescent in vivo mitophagy-reporter zebrafish lines, Tg(ubi:mito-Keima) and Tg(ubi:mito-GR)70. Both tools exploit the change in pH resulting from lysosomal fusion to identify cells undergoing active mitophagy. The Mito-Keima line expresses a single pH-sensitive fluorescent protein, Keima, targeted to mitochondria71; it appears green at neutral pH in healthy mitochondria and shifts to red when exposed to the acidic pH of the lysosomes, as illustrated schematically (Fig. 3A, left). The Mito-GR line uses a fluorescent cassette72, which tandemly expresses both GFP and mCherry in the mitochondria; when undergoing mitophagy, the GFP is quenched by the low pH of the fused lysosome, leaving puncta where the pH-resistant mCherry alone is visible (Fig. 3A, right). After crossing each line to a blue Tg(kdrl:tagBFP)mu293 endothelial reporter line73, mitochondrial dynamics of EHT were examined by live whole mount confocal imaging at 36 hpf (Fig. 3B, C). As anticipated given the previously defined role of BNIP3l-mediated mitophagy in erythroid maturation2,43, mitophagy (false-colored yellow to highlight) was detected in red blood cells in circulation in the dorsal aorta and posterior cardinal vein of Mito-Keima embryos; mitophagy was also visible in discrete cells throughout the niche, including muscle, floor plate, and a subset of ECs, which based on their rounded morphology and position at the VDA/PCV junction (pink arrowhead), were likely HECs or newly specified HSPCs (Fig. 3B). Indeed, time-lapse imaging revealed mitophagy puncta emerging immediately after select Kdrl+ cells appeared to physically undergo EHT (Fig. 3B, inset). To corroborate these findings, we repeated this analysis with Mito-GR embryos (Fig. 3C); notably, by Imaris imaging analysis of the BFP+ population, we calculated a mitophagy index score (ratio of red to green in defined tissue70) which revealed significantly more mitophagy in Kdrl+ cells in the CHT compared to VDA (Fig. 3D). To further validate this observation and eliminate potential confounding analysis of non-hemogenic ECs, we created an HSC-specific Tg(6.0itga2b:EGFP:mito-GR) referred to as Tg(CD41:mito-GR), confirming high levels of mitophagy in HSPCs localized within the vascular plexus of the CHT niche (Supplementary Fig. 3a).

As the scRNAseq revealed a strong metabolic shift toward oxidative phosphorylation and ROS production during the transition from EC to HSPC state, we next sought to understand the connection between ROS levels and mitophagy in the hematovascular niche. Using an established Tg(ubb:HyPer) ROS reporter line74, ROS levels were unexpectedly found to be significantly decreased by confocal microscopy and flow cytometry within HSPCs in the CHT compared to the VDA at 48 hpf, as marked by the Tg(Runx1+23:mCherry) reporter75 (Supplementary Fig. 3c, d). As this finding was inversely correlated with the presence of mitophagy in wild-type embryos, we next assessed the impact of ROS elevation on Mito-GR reporter activity: increased ROS production mediated by global prdx1 knockdown significantly enhanced mitophagy in Kdrl+ cells, inclusive of EC and newly specified HSPCs, in the CHT at 48 hpf (Fig. 3E and Supplementary Fig. 3b), implying that mitophagy is directly activated in response to ROS production. As our scRNAseq data indicated the presence of multiple mitophagy-associated genes during HSPC development, to further confirm and localize expression, we utilized Tg(cmyb:EGFP;kdrl:Has.HRAS-mCherry) embryos76 to perform qPCR analysis for mitophagy-related genes on FACS-purified subfractions: Kdrl-/cMyb+ (myeloid), Kdrl+/cMyb- (EC), Kdrl+/cMyb+ (HSPC), and Kdrl-/cMyb- (double negative control, inclusive of the remainder of live cells at 48 hpf). While most mitophagy genes were expressed in the cMyb+ myeloid fraction, inclusive of progenitors and primitive myeloid cells (Supplementary Fig. 3e–j); only bnip3lb and pink1a were enriched in Kdrl+cMyb+ HSPCs (Fig. 3F and Supplementary Fig. 3e). Whole embryo qPCR analysis revealed that bnip3lb expression increased significantly between 36 and 72 hpf (Fig. 3G), coinciding with VDA to CHT transition, and cellular localization of bnip3lb by fluorescent WISH (FISH) showed co-expression with runx1 within the CHT vascular plexus (Supplementary Fig. 3k). Furthermore, while pink1a expression was unchanged by qPCR at 48 hpf (Fig. 3H), bnip3lb was increased in prdx1 morphants (Fig. 3I) and decreased in response to NAC treatment (10 µM, 36–48 hpf) (Fig. 3J), indicating precise regulation in response to ROS level.

Whereas many mitophagy-associated genes are regulated post transcriptionally in response to danger or stress cues, including oxidative stress, hypoxia, and/or DNA damage, our data underscore transcriptional upregulation of bnip3lb expression during embryonic hematopoiesis (Fig. 2E). BNIP3l/NIX has an established role in developmentally programmed mitophagy, regulated by metabolism-responsive transcription factors such as FOXO367 and HIF1α68. In line with our scRNAseq data, hif1aa and hif1ab are both present in the cMyb+ myeloid fraction by qPCR (Fig. 3K and Supplementary Fig. 3j), and hif1ab shows additional enrichment in the HSPC cluster, similar to bnip3lb. Importantly, hif1ab is the primary Hif1α paralog driving HSC formation in zebrafish52 and a previously established regulator of bnip3lb expression42. While MO-mediated knockdown of hif1ab77 alone was insufficient to alter bnip3lb expression by qPCR on tail isolates at 72 hpf (Supplementary Fig. 3l), indicative of redundant or alternative routes of genomic regulation, induction of Hif1α transcriptional activity via exposure to the chemical hypoxia mimetic DMOG78 (10 µM, 48–72 hpf) significantly increased expression of bnip3lb (Fig. 3L), but not that of pink1a (Fig. 3M), in the CHT. Together, this data implies that ROS-mediated Hif1α activation drives onset of Bnip3lb-associated developmentally programmed mitophagy in newly specified HSPCs.

Bnip3lb knockdown reduces embryonic HSPC and lymphoid populations post EHT
Given the developmentally synchronized induction of mitophagy and bnip3lb expression, we sought to determine if Bnip3lb-associated mitophagy is necessary for the establishment and/or maintenance of the embryonic HSPC pool. Using a previously validated splice junction-targeting MO for bnip3lb, bnip3lb-MO70, we established an effective dose to knockdown expression (Fig. 4A) with no observable effect on vascular or primitive erythrocyte development (Supplementary Fig. 4a, b). Knockdown of bnip3lb significantly reduced mitophagy puncta within budding Kdrl+ cells in the VDA compared to matched controls (Fig. 4B, C). Consistent with the developmental timing of bnip3lb expression and Mito-GR mitophagy reporter activity, HSPC marker expression in the VDA at 36 hpf was relatively unchanged to modestly enhanced in morphants (Fig. 4D, E); however, by 72 hpf, bnip3lb proved essential for maintaining runx1/cmyb expression in the CHT (Fig. 4F, G). A significant reduction in CD41lo HSPCs was confirmed by flow cytometry at 72 hpf (Fig. 4H) and replicated using an alternative MO targeting the ATG region of bnip3lb (Fig. 4I; Supplementary Fig. 4c). Importantly, MO-mediated targeting of pink1a, bnip3la, bnip3 or fundc1 (Supplementary Fig. 4d), had no effect on CD41lo HSPC number by FACS at 72 hpf.

To verify the impact of Bnip3lb receptor-mediated mitophagy on HSPC development, we generated a bnip3lb mutant line, as previously described70. Mutants showed an unexpected increase in HSPCs in the CHT by runx1/cmyb WISH at 72 hpf (Supplementary Fig. 4e, f). Notably, however, mutant embryos demonstrated significant upregulation of other mitophagy related genes (Supplementary Fig. 4g). In addition, the impact of bnip3lb loss was strongly mitigated by combinatorial injection of morpholino against the bnip3lb homolog, bnip3la (Supplementary Fig. 4e, f), suggesting that the absence of an HSPC effect in bnip3lb mutant embryos was likely due to genetic compensation, as reported for other gene families79–81. To further validate the impact of bnip3lb knockdown, we successfully generated bnip3lb crispant embryos by targeting sequences in exons 1–3 using published protocols82 and verified through Sanger sequencing (Supplementary Fig. 4h); in line with our morphant analysis, crispant embryos exhibited significant reductions in CD41lo HSPCs by FACS at 72 hpf (Fig. 4J). Interestingly, at 48 hpf during VDA to CHT transition, runx1/cmyb expression in the aortic niche was noticeably enhanced in bnip3lb morphants (Supplementary Fig. 4i, j), in line with prior reports50, indicating that elevated ROS levels may continue to stimulate HSPCs undergoing EHT at this time point and/or suggesting that initiation of mitophagy may play a role in HSPC egress or migration to the CHT, consistent with the scRNAseq data. Furthermore, evaluation of hematopoietic development at later embryonic timepoints revealed that reduced mitophagy in bnip3lb morphants was associated with decreased numbers of lymphoid progenitors, as marked by WISH for rag1 in the thymus at 120 hpf and confirmed by flow cytometry using the established Tg(rag2:gfp) lymphoid reporter (Fig. 4K–M)59. A corresponding increase in myeloid fate, as determined by WISH for mpx and flow cytometry with Tg(mpx:GFP) neutrophil reporter line (Supplementary Fig. 4k–m), was also noted, suggestive of lineage skewing and/or the failure of progenitors to migrate to or effectively seed subsequent niches.

Bnip3lb protects HSPCs from ROS-driven apoptosis in the CHT
We next sought to understand the mechanism by which loss of Bnip3lb-driven mitophagy impacted HSC fate. While mitophagy is regulated by multiple pathways via complex and compensatory mechanisms, canonical pathways converge on LC3-dependent autophagosome formation and lysosomal fusion for degradation of targeted components39. Chemical inhibition of the final stage of mitophagy, via blocking lysosomal acidification with bafilomycin (50 µM) treatment, mimicked the effect of bnip3lb knockdown, resulting in a dramatic reduction in runx1 and cmyb expression in the CHT and KM (Supplementary Fig. 5a–d), emphasizing the importance of mitophagy in secondary hematopoietic niches. Based on reports in adult stem cells34, and our analysis of the impact of spatio-temporal metabolic flux on HSPC development, we hypothesized that Bnip3lb-associated mitophagy is responsible for regulating ROS levels to promote survival and balance developmental lineage commitment with HSPC expansion to allow for production of the lifelong HSPC pool. To begin to validate this conjecture, the impact of bnip3lb knockdown was assessed by flow cytometry for CellROX in CD41:EGFP reporter embryos: while ROS was not statistically elevated in the embryo as a whole at this timepoint (Fig. 5A, top), it was significantly increased within the CD41lo HSPC population (Fig. 5A, bottom). A similar trend for ROS in HSPCs was observed using an independent Runx1+ reporter line, Tg(Mmu.Runx1:EGFP) (Supplementary Fig. 5e)75. To determine if increased ROS was responsible for the reduction of HSPCs in the CHT of bnip3lb morphants, we conducted an epistasis analysis with or without NAC treatment (48–72 hpf). Knockdown of bnip3lb reduced HSPCs in the CHT at 72 hpf, while NAC enhanced their appearance, as expected (Fig. 5B, C); furthermore, NAC-mediated ROS reduction was sufficient to rescue HSPC gene expression after bnip3lb knockdown by WISH (Fig. 5B, C). This epistatic relationship with NAC was confirmed and quantified by flow cytometry of CD41lo HSPCs at 72 hpf using either the splice- (Fig. 5D) or ATG-targeted bnip3lb-MOs (Supplementary Fig. 5f), indicating that ROS is the primary driver of the observed HSPC phenotypes following loss of developmentally programmed mitophagy.

We next aimed to clarify the cellular mechanisms by which unregulated ROS impacts the embryonic HSPC population. ROS can promote or impede cell cycling, depending on the cellular context and dose2. Using a FACS-based EdU incorporation assay, we found that whereas the number of dividing cells in the whole embryo remained relatively constant (Fig. 5E), there was a significant reduction in the proportion of dividing CD41lo HSPCs in bnip3lb morphants (Fig. 5F). While reduced proliferation and alterations in lineage commitment (see Fig. 4) might fully explain the impact of loss of mitophagy on HSPC maintenance in the CHT, high and/or chronic ROS stimulation can also drive apoptosis51. Analysis of a panel of apoptotic genes by qPCR at 72 hpf revealed a strongly significant response to bnip3lb knockdown overall, with significantly increased expression of the pro-apoptotic gene, caspase casp2, as well as noxa1, a gene critical for mitochondria-mediated apoptosis83, in particular (Fig. 5G). To explore a potential role for cell death in HSPC depletion in bnip3lb morphants, we employed Acridine Orange, a marker for dead and dying cells, in Tg(Mmu.Runx1:NLS-mCherry)cz201 embryos. While the overall number of Runx1+ HSPCs was reduced, acridine orange staining revealed a pronounced enhancement in the fraction of dual-labeled HSPCs in the CHT of bnip3lb morphants compared to controls, indicative of increased apoptosis (Fig. 5H, I); notably, this effect was rescued by NAC exposure, resulting in less cell death in bnip3lb morphants. Taken together, Bnip3lb-associated mitophagy appears to act to maintain the number of newly derived HSPCs in the developing embryo by allowing for proliferative expansion, while protecting cells from ROS-induced apoptosis and/or premature lineage commitment.

Mitophagy activation enlarges the HSPC population and promotes lymphoid development
To further validate the role of mitophagy in the establishment and/or maintenance of the multipotent HSPC population during embryonic development, we aimed to understand whether stimulation of mitophagy could further boost HPSC numbers in vivo. While Bnip3lb-mediated mitophagy appears to be the developmentally programmed pathway initiated during EHT to regulate ROS, expression of alternative mitophagy mediators was noted to increase over the VDA to CHT hematopoietic transition by scRNAseq (see Fig. 2E, F), suggesting that alternative pathways of activation may become functional in HSPCs. To this end, we exploited an established ΔOTC construct, which employs a clinically-relevant mutated form of mitochondrial matrix protein ornithine transcarbamylase to induce mitophagy via the misfolded protein response84–86. We engineered this gene to be expressed under the heat-shock promoter to form an inducible Tg(hsp70:ΔOTC) mitophagy activator line. Following heat-shock stimulation at 48 hpf, dual transgenic Tg(hsp70:ΔOTC;ubi:mito-GR) embryos showed an increase in overall mitophagy reporter expression within the CHT compared to matched controls (Fig. 6A, B). Using Tg(hsp70:ΔOTC;CD41:EGFP) embryos, ΔOTC induction (at 48 hpf) drove a significant expansion in CD41lo HSPCs by fluorescence microscopy and flow cytometry at 72 hpf (Fig. 6C, D). This positive outcome for mitophagy stimulation was noted in the CHT as early as 48 hpf by WISH for runx1/cmyb (Supplementary Fig. 6a–d). Furthermore, ΔOTC-stimulated mitophagy appeared to promote lymphoid lineage development, whereby a considerable increase in Rag1/2+ lymphoid cells was noted by WISH and flow cytometry at 120 hpf (Fig. 6E–G), without affecting the Mpx+ myeloid population (Supplementary Fig. 6e–g).

To confirm and extend these observations to facilitate potential translational application, we examined the impact of compound-mediated induction of mitophagy on HSPC development. Several classes of chemicals have been previously documented to stimulate mitophagy in vitro and/or in vivo via differing routes of action, including: carbonyl cyanide-trifluoromethoxyphenyl hydrazone (FCCP), which depolarizes mitochondrial membranes to engage the pink1-parkin pathway87; Nicotinamide riboside (NR), a NAD+ precursor which promotes transcriptional upregulation of mitophagy associated genes88,89; and AMPK activator CAS-84927-81-7 (referred to as CAS-849), which upregulates mitophagy via the mTOR pathway90,91. Notably, all three compounds stimulated mitophagy in vivo in Tg(ubi:mito-GR) embryos at 48 hpf (Supplementary Fig. 6h–j). In line with our spatio-temporal observations for bnip3lb loss, we saw little impact for stimulation of mitophagy on specification of HSPCs in the VDA at 36 hpf (Fig. 6H (upper), I). In contrast, runx1/cmyb expression was significantly increased in the CHT by WISH at 72 hpf following NR, FCCP or CAS-849 treatment (Fig. 6H (lower), J), which was confirmed as an increase in CD41lo HSCs by flow cytometry (Fig. 6K). Again, a positive impact for mitophagy activation was observed as early as 48 hpf, as HSPCs migrate from the VDA to CHT, by WISH (Supplementary Fig. 6c, d), suggesting that compound-driven stimulation of mitophagy may be a viable avenue to promote HSPC expansion following EHT.

NR-driven mitophagy sustains colony-forming potential in hiPSC-derived hematopoietic progenitors
As a well-established issue with production or maintenance of HSPCs in culture is limited multipotency and/or self-renewal capabilities, we next aimed to explore whether targeted stimulation of mitophagy might be functionally beneficial ex vivo. In addition to being the most consistent of the tested compounds in enhancing mitophagy in zebrafish embryos, NR is well tolerated in vivo and in vitro88,89. To confirm that the effects of NR stimulation on HSPC formation were due to mitophagy, we employed bafilomycin epistasis analysis. As anticipated, NR treatment significantly increased the CD41lo HSPC population in the CHT by flow cytometry at 72 hpf; however, NR was unable to mediate this effect in the setting of simultaneous exposure to bafilomycin (Fig. 7A). To validate a stimulatory role for mitophagy, more generally, and not alternative NR-dependent processes mediated by NAD+ flux, we repeated the epistasis experiment with FCCP and found similar results (Fig. 7B). Interestingly, NR stimulation was also less effective in the setting of bnip3lb knockdown, implying that receptor-mediated mitophagy remains essential in expanding HSPCs and cannot be completely compensated for by activation of alternative pathways (Fig. 7C). Finally, sustained exposure to NR stimulation (48–120 hpf) continued to positively influence HSPCs colonizing the KM niche by WISH for cmyb (Fig. 7D, E), and significantly increased CD41lo HSPCs and the Rag2+ lymphoid progenitor population in the thymus by FACS (Fig. 7F, G). We therefore focused on the use of NR and FCCP to stimulate mitophagy for translational studies.

In our primary analysis, Human iPSCs were differentiated toward hematopoietic fate using a modified version of the established “Keller” differentiation protocol7 and treated with NR (1 mM) in the media during the EHT stage (stating on day 8) (Supplementary Fig. 7a). Comparable to the lack of impact on HSPC specification in the zebrafish VDA, no significant differences were found in the number or proportions of CD34+CD45+ multipotent hematopoietic progenitors initially generated after 5 days of culture with NR (Supplementary Fig. 7b). However, NR was noted to reduce intracellular ROS levels in all cultured cells as measured by CellROX at this time point (Supplementary Fig. 7c). Following flow cytometric profiling, control and NR-treated cells were collected for assessment of hematopoietic progenitor potential via a standard methylcellulose colony forming (CFU) assay92,93. At day 14 of the primary CFU, there was a dramatic increase in all colony types scored for cells previously exposed to NR-stimulation compared to controls; of note, this included the presence of multipotent CFU-GEMM colonies, which were not observed in controls (Supplementary Fig. 7d). To assess potential long-term impact on HSPC production and/or function, cells were harvested and serially replated at equal doses into secondary CFU culture: at 28 days from termination of NR stimulation and initial harvest, increased production of all colony types detected was retained in NR-treated samples (Supplementary Fig. 7e), indicating that induction of mitophagy promotes the production and maintenance of de novo-derived human multipotent hematopoietic progenitors in vitro.

Given recent advances in the efficiency and reproducibility of iPSC to HSC differentiation protocols, we sought to validate and extend our findings using an alternative approach which more closely resembles phenotypic HSC production using the “Loh” protocol94. Human iPSCs were differentiated toward hematopoietic fate using a modified version of the Loh protocol95 and treated with NR (1 mM) or FCCP (5 µM) on day 8 of differentiation (Fig. 7H), which correlates with EHT. At day 11, compound activators were removed, and the floating, non-adherent cell population was collected from control, NR- and FCCP-treated samples at day 11 for assessment of HSPC frequency and CFU potential. At day 11, flow cytometry analysis showed a statistically significant increase in CD34+CD45+ HSPCs in NR-treated samples and no change in HSPC numbers for FCCP-treated samples (Fig. 7I and Supplementary Fig. 7f). As seen with the Keller protocol, at day 14 of the primary CFU assay, there was a dramatic increase in all colony types scored following prior NR- or FCCP-stimulation compared to controls, including expansion of multipotent CFU-GEMM colony number (Fig. 7J, left). To assess impact on HSC production and/or function, equal doses of cells were serially replated into secondary CFU culture: at 28 days from cessation of mitophagy stimulation and initial harvest, NR and FCCP-treated samples continued to exhibit enhanced CFU potential in vitro (Fig. 7J, right), indicating that induction of mitophagy, whether through NR or FCCP, promotes maintenance of de novo-derived human multipotent hematopoietic progenitors in vitro regardless of differentiation platform utilized.

Altogether, our data indicate that activation of developmentally programmed mitophagy via Bnip3lb function is an essential component of HSPC development in the vertebrate embryo, which regulates ROS levels to allow for expansion, controlled differentiation, and survival of the stem cell pool. Furthermore, our in vivo and in vitro data indicate that exogenous activation of mitophagy is sufficient to support HSC expansion and maintenance in culture, which has direct therapeutic implications for enhancing the efficiency and self-renewal capacity of HSCs derived de novo in vitro from human iPSCs.

Methods
Ethics declarations and approval for animal experiments
Zebrafish were maintained and utilized in accordance with protocols approved by Boston Children’s Hospital and Beth Israel Deaconess Medical Center Institutional Animal Care and Use Committees (Protocol #: 0002155 (BCH) and 045-2024 (BIDMC)).

Zebrafish husbandry
Adults were maintained in a standard circulating water facility on a live brine shrimp diet supplemented with Gemma 300 pellets (Skretting). Wild-type embryos used were of the AB strain; transgenic and/or mutant lines utilized are detailed in the Key Resources Table (see Supplementary Table 1). Mutants were genotyped by PCR on tail-clip samples (see Supplementary Table 1 for primers). After screening for developmental abnormalities, healthy embryos were maintained in E3 fish medium in a 28°C incubator until use. Zebrafish sex is indeterminate until ~1 month of age; as all samples used in this study were 5 days post fertilization or less, sex was not determined and unable to be considered as a variable in analysis.

hiPSC culture
Wild-type hiPSC line #1157.2 was generated with informed consent (Boston Children’s Hospital IRB protocol #: 09-02-0068) from PBMCs of a healthy adult donor, as previously described97. hiPSCs were cultured in Stemflex medium (Thermo Fisher Scientific) in 37°C, 20% O2, 5% CO2 incubators using matrigel-coated wells, with clump passaging (ReleSR, Stem Cell Technologies). Karyotyping and mycoplasma testing were performed as quality control measures to verify chromosomal integrity and ensure lines were free of mycoplasma contamination before use.

Chemical treatments
Embryos were exposed to N-acetyl-L-Cysteine (NAC) (10 µM), AMPK activator CAS849727-81-7 (CAS-849) (0.25 µM), Nicotinamide Riboside (NR) (10 µM), FCCP (0.01 µM), DMOG (10 µM), H202 (0.05%), Glucose (1%) or equivalent percentage of vehicle (DMSO) for sibling controls in E3 (fish) water.

Flow cytometry and cell sorting
Embryos (pools of 8 embryos/sample) were dissociated in 500 µL LiberaseTM solution (75 µg/mL in 1xPBS/1mM EDTA) at 34°C for 45 min. Samples were dissociated by pipetting and filtered. SYTOX Red viability stain was added to a concentration of 5 nm; most figures display the proportion of GFP positive cells as a percentage of cells negative for the SYTOX red dye. All reagents can be found in the Key Resources Table (see Supplementary Table 1). Flow cytometry was performed on an LSR Fortessa (BD) and analyzed by FlowJo software. For ROS detection, CellROX orange dye (1:1000 dilution) was added to samples 30 min prior to analysis and measured using 561 nm laser and PE filter. For cell sorting, embryos were bisected along the transverse plane of the trunk at the point where the yolk sac narrows, and tail sections were dissociated as above. Sorting for live cells was performed on a FACSAria (BD).

In situ hybridization
Whole-mount in situ hybridization (WISH) was performed as described (http://zfin.org/ZFIN/Methods/ThisseProtocol.html). For each condition, three clutches of ≥20 stage-matched embryos were fixed in 4% paraformaldehyde for processing. Fixed embryos were treated with 0.8% KOH/0.9% H2O2 to remove pigment, then dehydrated in methanol. Once rehydrated in PBS 0.1% Tween (PBST), embryos were permeabilized using 10 µg/mL Proteinase K, re-fixed in 4% paraformaldehyde, then washed again in PBST. Hybridization with digoxigenin-UTP-labeled riboprobes was performed in hybridization buffer containing 50% formamide at 70°C overnight. Published probes are shown in the Key Resources Table (see Supplementary Table 1); the bnip3lb probe was made from cDNA of the coding sequence using primers shown in the Key Resources Table. Embryos were washed in sodium citrate buffer/0.1% Tween and incubated overnight with Anti-dioxigenin-AP, Fab fragments. After washing in PBST, antibodies were visualized by incubation in Tris buffer (pH 8.5) with BCIP/NBT. The intensity of expression was qualitatively categorized as high, medium, or low relative to the median intensity of the control group for a given clutch. For runx1/cmyb expression, a score of “high” typically reflects continuous aortic signal with robust thickness, indicating numerous cell clusters; “medium” denotes moderate, yet continuous signal, with occasional budding cells; and “low” indicates spotty/discontinuous signal. At least 20 embryos from a minimum of two independent clutches were analyzed per treatment condition and/or genotype, with phenotype scoring done blinded and independently confirmed. The scoring distribution of each sample set was averaged across each independent clutch, and control and treatment groups compared and analyzed with a chi-squared test (p-value < 0.05 is considered significant). Representative images were captured using a Zeiss Axio Imager A1/Axio Cam MRC and Axiovision LE software. Fluorescent WISH was performed as described94, by a procedure comparable to conventional WISH, but utilizing Anti-dioxigenin-POD Fab fragments to stain the bnip3lb probe, Anti-Fluorescein-POD Fab fragments for the runx1 probe, and Rabbit anti-GFP TP401 and Goat anti-Rabbit IgG Alexa Fluor 647 to detect Kdrl:GFP103. Fluorescent WISH images were captured using confocal microscopy as described below.

Confocal imaging
Confocal imaging was performed on a Ti2 (Nikon) scope using an iXon Life 888 EMCCD (ANDOR) camera and analyzed using Imaris software (Oxford Instruments) and Python. Embryos were treated with 0.003% PTU at 24 hpf to inhibit pigmentation. For live imaging, embryos were mounted on glass-bottom dishes in 1% low-melting point agarose in E3 media containing 0.2 mg/ml Tricaine. For acridine orange experiments, 72 hpf embryos were exposed to 10 µg/mL of acridine orange in E3 media for 1 hour while protected from light, then washed in E3 three times before live imaging. See Key Resource Table (Supplementary Table 1) for more information regarding reagents.
Adult Tg(kdrl:TagBFP)mu293 and Tg(ubi:mito-Keima) or Tg(ubi:mito-GR) zebrafish were crossed, and embryos were maintained in E3 at 28.5°C until imaging. For imaging of MitoKeima embryos, laser settings were: 405 nm (50%, 100 ms), 488 nm (50%, 70 ms), and 594 nm (50%, 70 ms), all at 540 MHz and 16-bit dual gain 1/4. For Mito-GR embryos, images were captured using 488 nm laser light and 561 nm laser light with an ET FITC/TRITC dual emission filter (Chroma) in triggered acquisition mode. For both lines, a 3-h time-lapse was acquired at 5-minute intervals (37 loops). Z-stacks were collected with a NIDAQ piezo (1.2 µM steps) spanning -30 to +30 µM (51 slices). For all quantitative experiments, images from all samples were collected with identical laser power, camera gain and exposure time.
Images were de-noised with the bm3d algorithm where indicated and are otherwise unaltered. Raw images were collected in Nikon Elements (.nd2) format and a ratiometric image representing mitophagy (visualized as yellow in images) was generated in Python by converting the image into 64-bit and applying the following equation: (mCherry channel)/(GFP channel + 150). The 150 constant is higher than background signal ensuring that areas of low signal from both mCherry and GFP channels are excluded70. Images were exported in .tiff format and converted into .ims files for matus using the Imaris file converter software for display. HyPer relative fluorescence was quantified by masking on mCherry and taking the average values of pixels in the HyPer channel. For quantitative purposes, mitophagy puncta were defined by applying a threshold (identical across all samples) to the radiometric mitophagy channel. For experiments where it is indicated that the mitophagy index was calculated within Kdrl:BFP+ cells, a threshold (identical across all samples) was applied to mask only the BFP positive pixels. The mitophagy index was calculated by dividing the sum of all mitophagy puncta volumes by the total tissue volume. The results were averaged to give final values reported, with statistical significance determined by Students t test.

qPCR
RNA was extracted from mechanically homogenized pooled embryos (n=20/condition) using RNAEasy Micro Kits according to the manufacturer’s instructions. Reverse transcription of 1 µg of RNA was performed using a High-Capacity cDNA Reverse Transcription Kit according to the manufacturer’s instructions. SYBR Green PCR Master Mix was used for qPCR with an ABI7900 (Applied Biosystems) according to the manufacturer’s instructions. Relative expression was calculated using the ΔΔCT method104 See Key Resources Table (Supplementary Table 1) for qPCR primer sequences.

scRNAseq
Two replicate samples of pooled dissected tail preps of 48 hpf Tg(kdrl:Has.HRAS-mCherry)s916 embryos (300 embryos/condition) and two replicate samples of pooled whole 36 hpf Kdrl+ embryos (300 embryos/condition) were isolated by FACS as described above. Samples were processed on a Chromium Next GEM Single Cell 3’ GEM, Library & Gel Bead Kit v3.1 and sequenced on a Novaseq 6000 system (Illumina).
Downstream analysis was performed in R using Seurat (v4.4)63. After filtering for low-quality cells, datasets were log-normalized, variable features were identified, data were scaled, and PCA was computed on variable genes. A shared-nearest-neighbor graph was constructed for clustering, and UMAP embeddings were generated using the first 10 principal components. Clusters were annotated based on differentially expressed genes and canonical hematovascular markers.
Differential expression analysis between HECs, ECs, and HSPCs was performed using Seurat’s Wilcoxon test on SCT-normalized data. Ranked gene lists were used as input for clusterProfiler (v4.0)64 for Gene Ontology Biological Process enrichment (org.Dr.eg.db). Differential Expression Analysis of genes detected in the RNAseq dataset generated in this manuscript from FACS-isolated 36 hpf Tg(kdrl:Has.HRAS-mCherry)s916 embryos, comparing HSPCs vs ECs and HSPCs vs HECs, is included in Supplementary Data File 1 and Supplementary Data File 2, respectively. Differential Expression Analysis of genes detected in the RNAseq dataset generated in this manuscript from FACS-isolated 48 hpf Tg(kdrl:Has.HRAS-mCherry)s916 embryos comparing HSPCs vs ECs is indicated in Supplementary Data File 3. Fold increase, p-adjusted value, and normalized reads are denoted for each gene. For comparisons between 36 hpf and 48 hpf datasets, HSPC libraries were harmonized via UMI downsampling using DropletUtils65.

Morphants
To create morphants, 1–3 nl of a given morpholino oligonucleotide was injected into the yolk ball of embryos at the 1-cell stage at concentrations shown in the Key Resources Table (see Supplementary Table 1) using a microinjector (Narishige) and pulled glass needle. Uninjected sibling embryos were used as matched controls.

Transgenesis
Tg(6.0itga2b:EGFP:mito-GR), referred to as Tg(CD41:mito-GR), was generated using tol2 transgenesis as described by Wrighton et al.5. Briefly, gateway cloning using LR Clonase II was used to make a pTol2-Cd41:mitoGR;cryaa:cerulean vector, from p5E-CD41, pME-mitoGR, p3E-polyA and pDestTol2pA2 according to the manufacturer’s protocol. This vector along with tol2 transposase mRNA, made using the mMessage mMachine SP6 in vitro transcription kit, was injected directly into fertilized embryos at the 1-cell stage. F0 embryos were screened for cerulean fluorescent lenses and outcrossed as adults. Potential founders were screened for brightness, stable integration and copy number. Tg(hsp70:ΔOTC) was generated as above using p5E-hsp70, pME-ΔOTC, p3E-polyA and pDestTol2pA2 as shown in Key Resources table (see Supplementary Table 1).

Mutants and crispants
For production of mutant and crispant embryos, dgRNPs were designed against the coding sequence of bnip3lb, using the CHOPCHOP crispr design tool99, mixed and complexed with tracrRNA and Cas9 protein (IDT DNA), then injected into the cytoplasm of 1-cell stage embryos according to the methods laid out by Quick et al.78. Efficacy was validated by gel electrophoresis of PCR products of genomic DNA surrounding the sites, with visible recombination in >70% of tested embryos and confirmed by Sanger sequencing (see Supplementary Fig. 6h). crRNA sequences and genotyping primers are shown in the Key Resources Table (see Supplementary Table 1). For the creation of bnip3lb mutants, we targeted sequences in exon 1 (CGGTGGCACTGAAGCGCTGGAGG) and exon 2 (ATCCATTCACAATGGCGACATGG). For crispants, an additional sequence in exon 3 was targeted (TAAGAGTCACTGAACGCCGTGGG).

EdU assay
Larvae were exposed to 500 µM EdU in E3 water (10% DMSO) for 1 hour at 4°C. Pooled larvae were dissociated and filtered as above, then cells were fixed, permeabilized and stained as per the Click-iT™ EdU Alexa Fluor™ 647 Flow Cytometry Assay Kit. Flow cytometric data were collected on LSR Fortessa (BD) and analyzed using FlowJo.

hiPSC differentiation
Loh Protocol: Human 1157.2 iPSCs97 were differentiated into hematopoietic progenitors using a 2D adherent culture, where each step of differentiation is defined. Differentiation was performed based on previously published protocols95,96 with slight modifications, including an extra day of culture, extending the differentiation from 10 to 11 days, and the addition of hematopoietic cytokines SCF (100 ng/mL), TPO (30 ng/mL), and FLT3L (10 ng/mL) from day 8-11 of differentiation. An additional modification to the published protocol was the inclusion of retinyl acetate (RETA) at 50 nM from days 1 to 2, 2 µM from days 2 to 4, and 100 nM from days 4 to 11. In brief and per Zheng et al.95, STAR Protocols, iPSCs were dissociated with Accutase, counted, and seeded at single-cell density on day -1 in the presence of Thiazovivin. On day 0, cells were washed with DMEM/F12, and a chemically defined medium was added to begin differentiation. Specific media conditions driving stepwise differentiation are described in Fowler et al.96. On day 11, non-adherent hematopoietic progenitors were harvested, washed, counted, and plated for CFU assays.
Keller Protocol: Human 1157.2 iPSCs97 were maintained on Matrigel matrix using Stemflex media and differentiated into CD34+45+ HSPCs cells using a previously published protocol105. Briefly, iPSC colonies were scraped, then cultured in ultra-low attachment plates with aggregation media containing BMP4 (10 ng/ml, days 0–2), bFGF (5 ng/ml, days 1–2), CHIR99021 (3 μM, day 2), and SB431542 (6 μM, day 2), VEGF (15 ng/ml, days 3–8), bFGF (5 ng/ml, days 3–8), SCF (50 ng/ml, days 6–8), EPO (days 6–8), IL-6 (days 6–8), IL-11 (days 6–8), and IGF-1 (days 6–8) to allow the formation of embryoid bodies (EBs). On day 8, EBs were dissociated using an EB Dissociation kit (Miltenyi), and CD34+ hemogenic endothelium (HE) cells were isolated by magnetic-activated cell sorting (MACS) using the human CD34 Microbead Kit (Miltenyi). CD34+ HE cells were treated with DMSO or 1 mM Nicotinamide Riboside (NR) and cultured for 5 days in StemPro-34 media (Invitrogen) supplemented with BMP4, bFGF, IL3, IL6, IL11, IGF-1, VEGF, SCF, EPO, TPO, FLT-3L, and SHH according to standard protocols100 to allow for the formation of non-adherent CD34+45+ HSPCs.

Hematopoietic colony-forming unit (CFU) assay
CFU assays were performed using MethoCult H4636 methylcellulose media following the manufacturer’s instructions (see Supplementary Table 1). 5k cells were plated for the primary CFU assay. Colonies were counted after 14 days of culture; cells were then collected, washed with PBS and replated at 10K cells/well for the Loh protocol and 30k cells/well for the Keller protocol in secondary CFU assays.

Quantification and statistical analysis
Data analyses were performed using GraphPad Prism and Excel. Statistical significance was determined by a two-tailed unpaired t test with Welch’s correction, by Pearson’s chi-square (two-sided), or a two-tailed ANOVA calculation. Error bars indicate the mean and standard error of the mean (SEM). P value < 0.05 was considered significant. Statistical significance was calculated to the 4th decimal place. Values of greater significance are reported as P < 0.0001.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In the manuscript by Meader et al. entitled “Bnip3lb-driven mitophagy maintains fate of the embryonic hematopoietic stem cell pool,” the authors describe how ROS-mediated mitophagy can increase HSPCs numbers in the CHT expansion niche in zebrafish. In particular, they identified a role for receptor-mediated mitophagy via Bnip3lb rather than pink/parkin stress-induced mitophagy in the process. While the data provided mostly support the authors conclusions, some facets require further substantiation.\nMajor points:\n1) The authors present data in Figures 2 and 3 to build an argument that mitophagy is a programmed component of HSPC during the transition from the specification to expansion niche. Some of the data are difficult to interpret. For example, in figure 2G and H, they claim to have found a strong bias toward increased expression of mitophagy-associated TF and effectors in the HSPC fraction. The strong bias is not very apparent from the data presented. Can statistics be applied? Violin plots with accompanying adj. p-values showing the difference in expression between the HE and HSPC clusters could better illustrate the point. Additionally, in figure 3, it is extremely difficult to see what cells were analyzed in the mitoQC reporter assays, as the reporter is expressed everywhere. It is unclear how they ensured the signal quantified was only in kdrl+ cells? Flow cytometry might be more sensitive to detect the mcherry-to-gfp ratio in the kdrl:BFP+ cells to ensure the signal quantified is in the right cells. The quantification using the cd41:mitoQC partially alleviates these concerns.\n2) In the human iPSC experiments presented in Figure 7, the authors show less ROS in the cells treated with NR. This phenotype is the opposite of what is expected from the literature and their work in zebrafish. Is it possible that altering NAD+ levels is working via a non-ROS and non-mitophagy means? To link these data to the findings in the rest of the paper, the authors need to demonstrate whether the NR effect is mitophagy/ROS independent or dependent.\nAdditionally, in Figure 7G, the secondary plating effect is quite promising, but it looks mostly like an expansion of CFU-G not the multipotent GEMMs. These data suggest a role for NR on differentiation rather than HSPC maintenance/self-renewal. Please clarify.\n3) While the use of a grading system to define the amount of ISH signal as a means to quantify the amount of HSPC or differentiated blood cells within zebrafish embryos/larvae is common practice, more robust quantification is warranted. To further support their claims, the authors need to use appropriate statistics such as Chi-squared tests for all graphs denoting phenotypic distributions.\n4) The majority of the experiments testing bnip3lb in zebrafish are performed using a single morpholino. The authors did assess HSPC levels in an established bnip3lb mutant but instead of lower levels as seen in the morphants, the mutants showed higher levels of HSPC in the CHT. They argue this is due to genetic compensation, but the data are not so clear. The observed that knockdown of bnip3la in bnip3lb mutants lowered HSPC levels to near wildtype levels, but it did not result in diminished HSPC levels similar to what was observed in the morphants. Another way to illustrate that morpholino effects that are different from mutants are still from on-target effects on the gene of interest is to inject the morpholino into mutants. If the morpholino effect is on-target, then the morpholino phenotype should be suppressed in the mutants. Also, there is an incomplete description of the bnip3lb mutant in the manuscript or in the cited paper from Wrighton et al. Please add more information on the type of mutation in the mutant.\nIt is also important to provide an independent validation of the ROS-mediated effects by bnip3lb. Performing the NAC epistasis experiment using an independent bnip3lb morpholino would further support this major conclusion of the manuscript. This is especially important as elevated apoptosis and diminished proliferation are implicated in the diminished HSPC levels in bnip3lb morphants. Both of these cellular processes can be mediated by Tp53, a pathway known to be aberrantly upregulated by some morpholinos.\nMinor points\n1) To help orient the reader, please add experimental schemas and time points to the figures. This especially important for figures with multiple developmental stages and different treatment paradigms.\n2) Related to Figure 1B- the main text says 48 hpf but the figure says 72 hpf. Please make sure they match and list the correct developmental time point when the experiment was performed.\n3) In Figure 1E, please provide example flow cytometry plots to illustrate how low vs high cd41:gfp was defined.\n4) In Figure 1G, it is hard to see the KM in the images so please provide better images. Also quantification should be included to support the claim.\n5) In Figure S1E, please denote the specific transgenic line used to assess neutrophils.\n6) In the main text on page 6, the authors state, “GO terms related to biosynthesis, perhaps reflecting the beginning of their rapid expansion phase.” This statement is quite speculative. It is also possible that the change reflects the change from stationary to migratory cells ate this transition. Please soften or remove the statement\n7) In the main text, the authors state “Importantly, by confocal mitophagy of the Tg(ubb:HyPer) in vivo.” Please correct mitophagy to microscopy.\n8) Figure 2F- definition of color scale is missing. Unclear what the comparison is and if the genes in green are more or less represented in the newly formed HSPC. It is also unclear if the numbers in the boxes represent fold change or log2 fold change. Please clarify.\n9) Critical statistics are missing in several locations. For example, Figure 3G- stats should be included for NAC bnip3lb qPCR; Figure 4A- stats are missing in the comparison of the mitophagy reporter signal in bnip3lb knockdown compared to controls. Please ensure statistics are included for all quantitative data.\n10) In Supp Figure 4B, it is very difficult to see the o-dianisidine staining in the larvae. Please use better images.\n11) In Figure 4G, the cmyb ISH signal in the pink1 MO is out of focus. Please provide a better image.\n12) In Figure 5H, the number of cells/embryos analyzed should be included on the figure.", "reviewer_2": "In this study, Meader et al. investigate the regulatory mechanisms of embryonic hematopoietic stem and progenitor cells (HSPCs) in maintaining multipotency during rapid proliferation. They demonstrate that in zebrafish embryos, ROS levels drive initial HSPC specification in the dorsal aorta, but these levels need subsequent control to prevent apoptosis and differentiation. Bnip3lb-mediated mitophagy emerges as a key mechanism in this process, reducing ROS during HSPCs' endothelial-to-hematopoietic transition (EHT) and in the caudal hematopoietic tissue (CHT). Knockdown of Bnip3lb in zebrafish impairs mitophagy, leading to increased ROS, apoptosis, and skewed differentiation toward myeloid lineages, effects reversible by antioxidant treatment. Further, activation of mitophagy in human iPSC-derived HSPCs increases their long-term hematopoietic potential, suggesting therapeutic implications for improving stem cell transplant efficacy. This study underscores the potential significance of developmentally programmed mitophagy in stem cell maintenance and suggests strategies to boost HSPC production ex vivo through controlled mitophagy.\nDespite these intriguing implications, the study seems significantly under developed. Notable limitations include: (1) an over-reliance on qualitative data, (2) insufficient controls for morpholino studies and lack of alternative methods, and (3) limited validation in the human system, despite the claims made. Overall, it is unclear whether the central tenet of this paper is adequately supported by the data presented.\nMajor comments:\n1) Throughout the manuscript (e.g. Fig 1D, 1J, 4C, 4E, 4H, 4J, 4L, etc), the authors rely on qualitative metrics rather than quantitative metrics. They frequently bin data into “high”, “medium”, or “low” based on subjective scoring. Additionally, sample sizes (n) are not provided, nor are statistical tests applied to demonstrate significant differences between conditions. There is also no indication that these categories were graded in a blinded manner.\nMany to most of these studies should be quantified. For example, WISH data could be quantified – either through alternative approaches (e.g. CD41:GFP in the VDA in Fig 1C) and/or through direct quantitation of signal in regions of interest.\n2) Genetic rescue experiments were not performed, nor genetic alternatives used to corroborate the effects of any morpholinos.\n3) Fig 3B. This figure appears critical, yet it is unclear how well it supports the hypothesis or aligns with its description in the text. How was the strength of the mCherry signal in Kdrl+ cells evaluated? To what was it compared? Quantification is lacking. Furthermore, the only cells presented (Fig 3 and S3) seem to come from the same image/movie of a single sample. This analysis needs to be quantified and repeated across multiple embryos to support the claim robustly. More specific markers for cell types of interest could also improve clarity.\n4) Mitophagy was fundamentally mischaracterized in several places:\na) “Pink1 is then ubiquitinated by Parkin” – please provide a reference for this, as most studies do not support this claim.\nb) “Loss of outer membrane integrity leads to PINK1 translocation from the matrix to the mitochondria’s surface” – Please provide a reference; most studies suggest a failure to import from the cytosol (e.g., due to the ANT complex, PMID: PMC6858570), rather than retrotranslocation from the matrix.\nc) “[Mitophagy] is initiated when proteins on the outer mitochondrial membrane are tagged to mark the organelle for destruction” – This is accurate for PINK1/Parkin-mediated mitophagy, but the initiating signal for FUNDC1/BNIP3/NIX is less clear. Phosphorylation mediates some cases but is not an absolute switch, as suggested here. It may be that general expression levels of these receptors, driven by specific transcription factors, control mitophagy induction.\nd) mitoQC was attributed to Zhao et al (ref 64). I believe this is a spurious reference. If so, please correct. (suggested PMID: 24176932 or similar).\ne) Fig 3A misrepresents the mitoQC reporter. It should display both red and green before the lysosome, and red only in the lysosome. Speculation may be helpful for why this does not seem to be the case in the zebrafish model.\n5) The application of these findings to hiPSCs is potentially impactful, but needs to be solidified. Importantly,\na. Given the broad and potentially varied effects of NR, the authors should confirm that NR treatment increases mitophagy in hiPSCs.\nb. Moreover, the beneficial effects of NR in Fig 7 should be BNIP3 dependent to support this model.\nc. Does a general decrease in ROS (e.g. NAC treatment) similarly improve the secondary CFUs of hematopoetic progenitors, or is there something special about mitophagy’s role?\nMinor comments:\n1) Fig 1B and related box plots (e.g. 5A, 7F). While these are “representative examples,” it is unclear whether the statistical test was performed across three biological replicates or only on the single sample shown. This should be assessed across biological replicates (N=3), with information on the number of cells analyzed per sample. Box plots are an atypical choice for this data; a histogram or violin plot might be more appropriate. At a minimum, include a description of the box plot structure (e.g., inner/outer quartiles [box] and 1.5*IQR [whiskers]).\n2) Is OTC-mediated mitophagy BNIP3Lb-dependent? If not, does the form of mitophagy matter?\n3) Fig 1C. At 72h, are HSPCs in prdx1-MO animals still stuck in the VDA, suggesting they fail to migrate?\n4) Fig 2F. Please provide a reference for the mitophagy reactome annotated genes. Mitophagy is generally not recognized to have a specific transcriptional signature.\n5) Fig 2H. How is “enrichment” determined? PINK1, FUNDC1, and BNIP3 do not appear notably enriched. Please specify the unbiased metric(s) used to support enrichment claims.\n6) Pg 12. “…ATG5-dependent autophagy is essential for EHT, highlighting the importance of multiple autophagic pathways in embryonic HSC production.” Does this imply that BNIP3lb-mediated mitophagy and Atg5-mediated autophagy are distinct? If so, what evidence suggests that BNIP3lb-mediated mitophagy is ATG5-independent?\n7) Pg 5. For clarity, please introduce the CD41 marker at its first mention (Fig 1E) rather than its second (Fig 1G, H).", "reviewer_3": "In the manuscript, the authors describe the role of Bnip3lb-regulated mitophagy in maintaining the embryonic hematopoietic stem cell pool in zebrafish. The authors elaborate on how spatiotemporal control of ROS is important for generating and regulating the HSPC pool. First, the authors show the essentiality of ROS for HSPC specification in the ventral-dorsal aorta (VDA) niche but not for HSPC specification in the caudal hematopoietic tissue (CHT) niche. Next, they claim that Bnip3lb-driven mitophagy regulates ROS within the CHT. Knocking down Bnip3lb causes a reduction of embryonic HSPC and increased ROS at CHT, which can be rescued by treatment with NAC. Finally, they present data on how NR-induced mitophagy can be leveraged to form hiPSC-derived hematopoietic progenitors. This is a potentially very interesting manuscript, but the data are unfortunately not of sufficient quality to conclude that Bnip3lb-regulated mitophagy is responsible for the observed phenotypes.\nMajor Points\n1. In Figures 3-4, 6A and S3 the authors use the Ubi:mito-QC transgenic zebrafish line to study mitophagy in zebrafish. The authors describe the GFP to be mitochondria and the magenta (mCherry) to be mitophagy, which is misleading as the reporter is a tandem-tagged (mCherry-EGFP) mitochondrial protein. Thus, mitochondria are labeled both green and red (should be seen as yellow or white) in normal situations, while upon induction of mitophagy (and transfer of the reporter to acidic lysosomes), the green signal quenches and thus only red/magenta-only puncta can be seen. For quantification of mitophagy, it is therefore important to adjust the microscopy settings so that mitochondria are seen as yellow/white (optimal is to treat with a lysosomal inhibitor). There is no information about how they have done the quantification of mitophagy in their samples, other than (p.14) “ a ratio of the mCherry/GFP values for each pixel was taken, and the results were averaged to give the final value”. Thus, if the microscopy settings are off (as indicated by the authors as well on p.6, “GFP signal being bright for mitochondria”) these data cannot be evaluated properly.\n2. The authors used morpholino (MO) to deplete various genes (including Bnip3lb), but there is no validation of the knockdown efficiencies. Moreover, as only one MO/gene is used, the authors should either use another MO and/or CRISPR-Cas9. In addition, it would be advisable to rescue the Bnip3l phenotype.\n3. Whole mount in situ hybridization (WISH). Throughout the manuscript, they do WHISH for runx1/cmyb together. Any reason why it is not done individually? The negative controls (using the sense probe) are missing, and the WHISH staining varies a lot from image to image (e.g. control in Figure 4D vs 4G). The latter is particularly problematic as they quantify this staining throughout the manuscript and use this as a basis for their phenotype distribution. The phenotypic distribution charts are also confusing. How do they score high, medium, and low? They have not mentioned the number of larvae used and whether the results are significant or not.\nMinor points:\n1.Text on page 5 line 8 says –“flow cytometry of disaggregated 48 hpf embryos” whereas in figure 1b it is written 72 hpf.\n2.Figure 1B, 5A og 7F; definition of X-axis units are missing\n3.Figure 1G, both for CHT and KM panels, the fluorescent signal is not adequate to see any difference. They should include an inset (zoomed image) of the region indicated in the KM panel.\n4.Both rag1 and rag2 are used without clear elaboration. WISH was done for rag1 (Figure 1i) while FACS was done on cells from transgenic line Tg(rag2:GFP). Why the switch?\n5.FISH for bnip3lb staining looks unspecific (Figure 3 D). They should include a negative control? Is the strong staining observed in the vertebral column true or autofluorescence?\n6.Can the bnip3lb-driven mitophagy-dependent HSC specification in CHT be compensated by FUNDC1? Both Pink1 and Fundc1 were expressed at low levels in both subclusters in Figure 2H. The authors checked for the effect of pink1 knockdown in HSPC gene expression at 72 hpf (Figure 4G) but Fundc1 was not investigated. Can fundc1 have a role similar to bnip3lb in HSPC specification via mitophagy/ROS axis since both are induced upon hypoxia?\n7.For the FACS analysis figures they use “% Live Cells” for the y-axis. This is confusing, as it is not well described.\n8.The pie chart shown in Figure 5H of dead and alive cells is a bit vague. They should describe how the analysis was done. The legend says “Pie charts show dead Runx1+ cells (magenta) as a proportion of the total, n=3/condition” Is this 3 larvae per condition?\n9.On p.9 the authors state “we utilized an established ΔOTC construct, which employs a clinically-derived mutated form of mitochondrial matrix protein ornithine transcarbamylase to induce mitophagy via the misfolded protein response64”. This is misleading as the referred paper does not show that this mutant induces mitophagy. Moreover, the current manuscript does not show an effect on mitophagy, as only one image with no quantification is shown in Figure 6A.\n10.Use of the mitophagy-inducing drug NR shows an increase in HSPC specification in CHT along with Cas849. Have the authors tested DMOG or DFP?", "reviewer_4": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": "Response to Reviewer Comments:\nReviewer #1 (Remarks to the Author):\nIn the manuscript by Meader et al. entitled “Bnip3lb-driven mitophagy maintains fate of the\nembryonic hematopoietic stem cell pool,” the authors describe how ROS-mediated mitophagy\ncan increase HSPCs numbers in the CHT expansion niche in zebrafish. In particular, they\nidentified a role for receptor-mediated mitophagy via Bnip3lb rather than pink/parkin stress-\ninduced mitophagy in the process. While the data provided mostly support the author's\nconclusions, some facets require further substantiation.\nAuthor response: Thank you for taking the time to review our manuscript. We appreciate the\nthoughtful critiques and recommendations, and hope that you find the changes adequate.\nMajor points:\n1) The authors present data in Figures 2 and 3 to build an argument that mitophagy is a\nprogrammed component of HSPC during the transition from the specification to expansion niche.\nSome of the data are difficult to interpret. For example, in figure 2G and H, they claim to have\nfound a strong bias toward increased expression of mitophagy-associated TF and effectors in the\nHSPC fraction. The strong bias is not very apparent from the data presented. Can statistics be\napplied? Violin plots with accompanying adj. p-values showing the difference in expression\nbetween the HE and HSPC clusters could better illustrate the point.\nAuthor response: We have significantly expanded and adapted our analysis to provide a more\nrobust, statistically grounded demonstration of mitophagy-related transcriptional upregulation in\nembryonic HSPCs. Our original submission presented findings from a 36hpf scRNAseq dataset,\nduring the peak of HSPC specification in the aorta. We have now expanded this analysis in the\nrevised submission, by adding another dataset from a later time point, 48hpf, to further underscore\nthe developmentally-timed metabolic switch and its functional consequences as HSPCs bud from\nthe ventral dorsal aorta (VDA) and seed the caudal hematopoietic tissue (CHT). We have also\nsignificantly updated our analysis of each dataset: both the 36hpf and 48hpf scRNAseq datasets\nwere reprocessed using a standard and fully documented Seurat workflow, including log-\nnormalization, selection of variable features, scaling, Principal Component Analysis, construction\nof a shared nearest-neighbor graph, clustering, and UMAP embedding (Figure 2a-b). Clusters\nwere recoded into biologically interpretable labels (EC, HEC, HSPC, Ery, Mac, MSC, Neut, NP)\nbased on differentially expressed genes (Supp. Figure 2a-b), and validated with canonical\nHSPC, myeloid, erythroid and endothelial lineage markers, including: cxcr4b, gata2b, itga2b,\nrunx1, spi1b, lyz, mpx, gata1a, hbbe3, klf1, cdh5, kdrl, and tie1 (Supp. Figure 2e-f). To quantify\nlineage-related transcriptional programs, we computed module scores (Figure 2c-d) for curated\nendothelial (cdh5, kdrl, tie1), HSPC (cxcr4b, gata2b, itga2b, meis1b, mpl, myb, notch1b, runx1,\nspi1b), and myeloid (lyz, mpx, spi1b) gene sets using Seurat’s AddModuleScore. (Supp. Figure\n2c-d). We performed differential expression analyses using FindMarkers in Seurat (default\nWilcoxon test) comparing HSPC clusters against HECs or ECs. GSEA was performed using\nclusterProfiler’s gseGO with the zebrafish Gene Ontology annotation (org.Dr.eg.db). Expression\nof mitophagy- and hypoxia-related genes (e.g., bnip3, bnip3la, bnip3lb, fundc1, pink1, park2,\nhif1aa, hif1ab) was directly examined within the HSPC cluster at each timepoint (Figure 2e-\nf). To harmonize raw UMI counts between datasets, we downsampled RNA counts from selected HSPC\nclusters using DropletUtils downsampleMatrix to a target of ~4,600 UMIs per cell.\nAt 36hpf, in addition to downregulation of genes associated with endothelial identity/function when\nHSPCs are compared to HECs or ECs by GSEA pathway enrichment, we see a significant\nenrichment in metabolic signatures and ROS/redox related pathways (Figure 2c). By 48hpf, we\nno longer detect a distinct HEC fraction, matching the kinetics of hematopoietic development\nwhere endothelial identity is further downregulated as HSPCs bud from the aorta and transit to\nseed the secondary “expansion” niche in the CHT.1 Interestingly, while the enrichment in\nmetabolic/mitochondrial pathways (Figure 2d) is retained, we no longer see a strong signature\nfor ROS/redox response, suggesting that as HSPCs mature, they may be inducing pathways to\ncontrol environmental “stressors” like ROS. Due to the pronounced increase in metabolic state\nand mitochondria-related pathways, we profiled mitophagy-related genes within the HSPC cluster\nto determine if the machinery was present and therefore potentially able to contribute to the\nregulation of ROS/stress response within the HSPC pool. Specifically, we looked at the\nexpression of pink1, parkin, fundc1, bnip3, bnip3la, and bnip3lb within our datasets (Figure 2e-\nf), as well as related transcriptional regulators hif1aa and hif1ab. While expression of several\nmitophagy related genes increased between 36 and 48hpf, we found that bnip3lb was the highest\nexpressed within the HSPC fraction. To validate these observations, we performed qPCR analysis\nfor each gene in sorted populations from Tg(cmyb:EGFP;kdrl:Has.HRAS-mCherry) transgenic\nreporter embryos, inclusive of Endothelial (Kdrl+/Myb-), Myeloid (Kdrl-/Myb+) and HSPC\n(Kdrl+/Myb+) fractions, finding the strongest enrichment for bnip3lb and pink1a in HSPCs (Figure\n3f, Supp. Figure 3f-k). Through modulation of Hif1α activity and ROS levels (Figure 3k-m, Supp.\nFigure 3m), we identified Bnip3lb as the more physiologically relevant regulator of mitophagy\nduring developmental hematopoiesis, and confirmed a distinct regulatory role for bnip3lb (vs\npink1) through MO knockdown studies for impact on CD41lo HSPC formation (Figure 4h-j, Supp.\nFigure 4d). We believe that the combination of our expanded bioinformatics analysis and\nvalidation studies now more definitively supports the spatio-temporal induction of mitophagy\nduring HSC formation.\nAdditionally, in figure 3, it is extremely difficult to see what cells were analyzed in the mitoQC\nreporter assays, as the reporter is expressed everywhere. It is unclear how they ensured the\nsignal quantified was only in kdrl+ cells? Flow cytometry might be more sensitive to detect the\nmcherry-to-gfp ratio in the kdrl:BFP+ cells to ensure the signal quantified is in the right cells. The\nquantification using the cd41:mitoQC partially alleviates these concerns.\nAuthor Response: We concur that using either the MitoKeima and MitoGR reporter lines, “red-\nonly” expression indicative of mitophagy is present in other cells within the VDA region\n(representative sample for: MitoKeima - Figure 3b; MitoGR - Figure 3c); this includes nearby\nfloor plate and muscle, as well as red blood cells in circulation within the lumen of the aorta and\nvein, which have been previously shown to utilize developmental programmed mitophagy during\ntheir maturation, and serve as a positive control for the tool at this timepoint.2 To that end, we\nutilized an established Tg(kdrl:TagBFP) pan-endothelial cell reporter3 to aid identification of\nendothelial cell (EC) fractions within the VDA region, inclusive of hemogenic endothelium (HEC)\nand newly specified HSPCs; we have now included a detailed description in the methods for the\n“masking” performed in Imaris using the BFP+ marker in the confocal images to localize\nevaluation of mitophagy to the EC/HEC/HSPC fraction (see page 17). Interestingly, in regard to\nthe ECs lining the dorsal aorta and posterior cardinal vein, we only observed the presence of\nmitophagy puncta in rounded or “budding” cells along the ventral wall of the dorsal aorta across\nmultiple independent biological samples using two distinct reporter lines (5 replicates for\nMitoKeima (Figure 3b); 4 replicates for MitoGR (Figure 3c)), which are presumptive HEC/HSPCs\nbased on their location and morphology.1 While these images were only intended to illustrate\nlocalization of mitophagy positive cells within the hemogenic niche, in the prior submission and\nrevised figure (Figure 3d), we did employ confocal-based analysis for mitophagy activation using\nthe MitoGR tool in the Kdrl:BFP+ population to distinguish the proportion of mitophagy positive\ncells in the VDA versus CHT, showing a statistically significant increase in the latter. The\n“mitophagy index” shown on y-axis of that graph was calculated using the same mathematical\nmethod as that published previously by our collaborators4; for clarity, we now directly indicate it is\nan assessment of the red/green ratio within the BFP+ fraction to help define the methodology in\nthe figure, legend and text. Please note, to enhance consistency with the prior publication and\nconstruct design, we adjusted the MitoQC label to MitoGR throughout.\nAs the previously established in vivo mitophagy reporter systems utilize both GFP and mCherry\nfluorescent channels, we were unfortunately unable to specifically co-localize the presence of\nactive mitophagy with well-characterized HSPC reporter lines, such as the Runx1+23 transgenic\nreporter5 - a highly conserved marker for newly specified HSCs in the dorsal aorta niche at this\nstage of development - for which we have only red and green tools available. Given that caveat,\nand our reliance on Kdrl:BFP-labeling, it was challenging to definitively quantify differences in\nECs/HECs versus budding HSPCs in the VDA at 36hpf by confocal using the mitophagy index\nmethodology, as described in Wrighton et al.4, which is essentially a ratio of puncta volume/\ntissue volume. Following presentation of refined localization of mitophagy gene expression to the\nHSPCs versus ECs by qPCR in subsequent figure panels (Figure 3f, Supp. Figure 3f-k), we\nprovide quantification of changes in MitoGR reporter activity within presumptive HSPCs after\ngenetic or compound-mediated perturbation compared to matched controls using the mitophagy\nindex score (Figure 6a-b) or manual counts of mitophagy positive puncta (Figure 4b-c).\nAs suggested in the review, we did attempt to perform FACS analysis using the CD41-driven\nMitoGR line that we developed to further assess numbers of HSPCs undergoing mitophagy at\n48hpf. However, the low number of total CD41 positive cells in the whole embryo (using 10 pooled\nembryos per sample) prohibited clear visualization of individual gated red and/or green\npopulations at sufficient signal strength over background to calculate a statistically meaningful\nmitophagy ratio. Therefore, we believe our quantitative analysis of the confocal imaging is the\nbest way to illustrate changes in HSPC-related mitophagy in vivo between treatment groups.\n2) In the human iPSC experiments presented in Figure 7, the authors show less ROS in the cells\ntreated with NR. This phenotype is the opposite of what is expected from the literature and their\nwork in zebrafish. Is it possible that altering NAD+ levels is working via a non-ROS and non-\nmitophagy means? To link these data to the findings in the rest of the paper, the authors need to\ndemonstrate whether the NR effect is mitophagy/ROS independent or dependent.\nAuthor Response: Based on prior publications6,7, we expected to see a decrease in ROS after\ntreatment with the mitophagy-stimulating agent NR. There are a number of ways that NR has\nbeen shown to impact ROS levels, including NAD+ modulation (as indicated) and upregulation of\nSIRT1, impacting antioxidant gene expression.8 To help clarify this point for the reader, we have\nchanged the text on page 12 to say “However, NR was noted to reduce intracellular ROS levels\nin all cultured cells as measured by CellROX at this timepoint (Supp. Figure 7c).“\nTo further address the role of mitophagy in NR-driven impact on ROS-mediated HSC regulation,\nwe performed an additional set of epistasis experiments in zebrafish embryos. Bafilomycin\ntreatment can be used to block the lysosomal fusion that is necessary for completion of\nmitophagy.9,10 Embryos were treated with NR, Bafilomycin, or a combination of both agents from\n48-72hpf; we found that when progression of mitophagy was disrupted by Bafilomycin exposure,\nNR stimulation was no longer able to increase HSPC numbers in the CHT (Figure 7a). We\nrepeated this experiment with an alternative mitophagy-stimulating agent, FCCP, which is thought\nto act via a distinct method from that of NR (via alterations in proton gradients11), and found\nessentially the same results (Figure 7b). Together, this indicates that ROS-associated impact on\nHSC number can be regulated by modulation of mitophagy in general. While we cannot formally\nrule out that NR treatment would not also enhance relevant antioxidant gene expression to help\nmaintain or amplify regulatory impact on ROS, we did find that NR stimulation was unable to\nincrease HSPCs over wild-type levels in the setting of bnip3lb knockdown in zebrafish, suggestive\nof redundant and/or compensatory balance.\nTo further document that the effect of NR on HSPCs is mediated via mitophagy in the culture dish,\nwe have now additionally utilized FCCP during in vitro iPSC to HSPC generation. For direct side-\nby-side comparison of impact, we utilized both NR and FCCP treatment in replicate studies,\nemploying a newly published and refined iPSC to HSC differentiation platform (termed the “Loh”\nprotocol) in which phenotypic HSPCs can be reliably produced over a 10-day protocol. NR and\nFCCP were applied in the media when “floating cells” (indicative of EHT) began to appear in the\nculture to most closely phenocopy the dosing window in fish (Figure 7h). Once “budding” HSPCs\nwere generated, they were isolated from the media and utilized in serial CFU assays to assess\ntheir self-renewal and differentiation capacity. We found that NR and FCCP treatment each\nenhanced serial CFU production (Figure 7j), in line with our prior observations for NR, with a\nsignificant increase in the multipotent GEMMs noted for both treatments. Consistent with our\nzebrafish work, this indicates the effect on HSPC expansion is mediated through mitophagy more\nbroadly, and not just an effect of NR treatment. Of note, as the “Keller” protocol utilized in the prior\nsubmission is now understood to primarily produce cells with short-term HSPC-like function, that\ndata has been moved to Supplemental Figure 7.\nAdditionally, in Figure 7G, the secondary plating effect is quite promising, but it looks mostly like\nan expansion of CFU-G not the multipotent GEMMs. These data suggest a role for NR on\ndifferentiation rather than HSPC maintenance/self-renewal. Please clarify.\nAuthor response: Thank you for this important point of clarification. We did not continue to\nexpose the cells undergoing the serial CFU assay to the mitophagy-stimulating compounds after\nthe initial dose during differentiation culture. As such, in the secondary replating, we would not\nnecessarily expect to see either maintenance or further expansion of multipotent GEMMs, which\nare presumably being stimulated to differentiate via the cytokines included in the CFU media.\nHowever, if transient NR exposure during hematopoietic specification and EHT successfully\nexpanded production and/or maintained GEMMs through the first round of plating, those cells\nwould then be present in sufficient number to functionally read out as increased hematopoietic\ncolony output in the second round of plating compared to matched control, which is indeed what\nwe observed (Supp. Figure 7d-e). Importantly, in the revised submission, we observed similar\neffects on total colonies and GEMMs for both NR and FCCP treatment when we independently\nreplicated the serial replating assay using mitophagy-stimulated cells derived from the Loh-\nderived protocol (Figure 7j), underscoring the utility of mitophagy in HSPC maintenance.\n3) While the use of a grading system to define the amount of ISH signal as a means to quantify\nthe amount of HSPC or differentiated blood cells within zebrafish embryos/larvae is common\npractice, more robust quantification is warranted. To further support their claims, the authors need\nto use appropriate statistics such as Chi-squared tests for all graphs denoting phenotypic\ndistributions.\nAuthor response: We have now performed Chi-square analysis of the phenotypic distributions\nfor all WISH data presented in this manuscript as suggested. Furthermore, we made sure to\nfurther support conclusions drawn from WISH analysis through use of a corresponding\nquantitative method such as flow cytometry or qPCR to increase the robustness of our claims.\n4) The majority of the experiments testing bnip3lb in zebrafish are performed using a single\nmorpholino. It is also important to provide an independent validation of the ROS-mediated effects\nby bnip3lb. Performing the NAC epistasis experiment using an independent bnip3lb morpholino\nwould further support this major conclusion of the manuscript. This is especially important as\nelevated apoptosis and diminished proliferation are implicated in the diminished HSPC levels in\nbnip3lb morphants. Both of these cellular processes can be mediated by Tp53, a pathway known\nto be aberrantly upregulated by some morpholinos.\nAuthor response: To address potential concerns regarding utilization of a single bnip3lb MO,\nwe have now included analyses for several of the foundational experiments, including effects on\nHSPCs at 72hpf and epistasis with NAC, with a second independent bnip3lb MO, as requested\n(Figure 4I, Supp. Figure 4c, Supp. Figure 5f). We also generated bnip3lb crispant embryos to\nfurther validate effects on HSCs (Figure 4j, Supp. Figure 4h); we see corresponding decreases\nin HSCs whether using the splice or ATG bnip3lb MO, or the crispant embryos, each of which are\nnow included in the revised manuscript in Figure 4h-j for comparison.\nThe authors did assess HSPC levels in an established bnip3lb mutant but instead of lower levels\nas seen in the morphants, the mutants showed higher levels of HSPC in the CHT. They argue\nthis is due to genetic compensation, but the data are not so clear. The observed that knockdown\nof bnip3la in bnip3lb mutants lowered HSPC levels to near wildtype levels, but it did not result in\ndiminished HSPC levels similar to what was observed in the morphants.\nAuthor response: Thank you for pointing out this discrepancy between the data shown in the\nfigure and our claims in the text. A review of the relevant data revealed a transposition error in\nthe final version of the graph utilized in Supplemental Figure 4 for the bnip3lb mutant. After\nreanalysis of the data from this experiment and an additional repetition for appropriate statistical\napplication of the requested chi-square analysis (Supp. Figure 4e-f), as well as qPCR analysis\nfor expression of alternative mitophagy mediators (Supp. Figure 4g), we are now able to better\ndemonstrate the suspected gene compensation, consistent with our prior statement.\nAnother way to illustrate that morpholino effects that are different from mutants are still from on-\ntarget effects on the gene of interest is to inject the morpholino into mutants. If the morpholino\neffect is on-target, then the morpholino phenotype should be suppressed in the mutants.\nAuthor response: As indicated in the response above, we have concerns regarding probable\ngenetic compensation in the bnip3lb mutant embryos. For that reason, we prioritized the design\nand application of a second MO and the production of new crispant embryos, which together\nconsistently demonstrated the same HSPC phenotype (Figure 4h-j) and further support the\nvalidity and specificity of our findings for Bnip3lb-mediated mitophagy in this process.\nAlso, there is an incomplete description of the bnip3lb mutant in the manuscript or in the cited\npaper from Wrighton et al. Please add more information on the type of mutation in the mutant.\nAuthor response: We apologize for this oversight and are happy to provide additional information\nregarding the bnip3lb mutant line, which was initially developed for use in the investigation by\nWrighton et al.4 The method used for the creation and validation of each of the mitophagy mutant\nlines was identical to those described in the text for bnip34, based on a previously published high-\nefficiency Crispr genome editing protocol.12 While the targeting sequences for bnip3 and bnip3la\nwere listed in the published supplementary table of the Wrighton paper, the bnip3lb mutants were\nnot yet established as a stable line and therefore not included in the final publication. The targeting\nsequences for creation of the bnip3lb mutant allele are now indicated in the methods and\nMethods Key Resources table; successful mutation was validated by Sanger sequencing as\npreviously described13.\nMinor points\n1) To help orient the reader, please add experimental schemas and time points to the figures.\nThis especially important for figures with multiple developmental stages and different treatment\nparadigms.\nAuthor response: Thank you for this helpful suggestion. We have now added a more informative\nschematic in Figure 1a. This figure now includes colored arrows to indicate the relevant treatment\nwindows. To help orient the reader throughout the paper, we have now included corresponding\ncolored text to any figure with precisely timed dosing to better showcase the distinct window(s) of\napplication. Furthermore, we have explicitly stated dosing windows for each experiment in the\nresults and figure legends to aid experimental clarity and interpretation.\n2) Related to Figure 1B- the main text says 48 hpf but the figure says 72 hpf. Please make sure\nthey match and list the correct developmental time point when the experiment was performed.\nAuthor response: This was an unintentional formatting hold-over from a prior draft of Figure 1.\nWe have corrected the main text to indicate “72hpf” to match the labeling and CellROX data\npresented in the final version of Figure 1b, as well as ensured that it is indicated correctly in the\ncorresponding figure legend (page 5, first paragraph).\n3) In Figure 1E, please provide example flow cytometry plots to illustrate how low vs high cd41:gfp\nwas defined.\nAuthor response: We have added representative flow plots for the CD41:GFP analysis (Supp.\nFigure 1d), as well as the first use of other flow cytometry analyses employed throughout the\npaper [i.e. Rag2:GFP (Supp. Figure 1e) and Mpx:GFP (Supp. Figure 1i)] to Supplemental\nFigure 1. The same gating strategies were applied for all zebrafish flow cytometry data presented\nin subsequent figures employing each of these same transgenic lines.\n4) In Figure 1G, it is hard to see the KM in the images so please provide better images. Also\nquantification should be included to support the claim.\nAuthor response: We have replaced these panels with images taken by confocal microscopy to\nincrease resolution (see updated Figure 1g). Due to its complex positioning at the head/body\njunction and lack of a secondary reporter line to clearly delineate the tissue boundaries (vs cells\nnearby in circulation), we did not take direct measurements of the fluorescent patterning in the\nKM by confocal microscopy. Rather, we performed further quantification of the HSPC effect via\nflow cytometric analysis using Tg(cmyb:eGFP) embryos, which are presented as Figure 1h.\n5) In Figure S1E, please denote the specific transgenic line used to assess neutrophils.\nAuthor response: We have added the transgenic tool designation for the myeloperoxidase line,\nTg(mpx:gfp), to the text on page 5 in the second paragraph: “Notably, prdx1 morphants showed\nminor changes in expression of the neutrophil marker mpx at this timepoint by WISH or via flow\ncytometry on Tg(mpx:GFP) embryos”. The appropriate reference is indicated here14 and in the\nsupplemental methods.\n6) In the main text on page 6, the authors state, “GO terms related to biosynthesis, perhaps\nreflecting the beginning of their rapid expansion phase.” This statement is quite speculative. It is\nalso possible that the change reflects the change from stationary to migratory cells ate this\ntransition. Please soften or remove the statement\nAuthor response: We completely revised the scRNAseq analysis, as indicated in the response\nabove, and have removed this statement as suggested.\n7) In the main text, the authors state “Importantly, by confocal mitophagy of the Tg(ubb:HyPer) in\nvivo.” Please correct mitophagy to microscopy.\nAuthor response: Thank you for noting that error. We have corrected this mistake (page 8, first\nparagraph), so the sentence now reads “Using an established Tg(ubb:HyPer) ROS reporter line,\nROS levels were unexpectedly found to be significantly decreased by confocal microscopy and\nflow cytometry within HSPCs in the CHT, …”.\n8) Figure 2F- definition of color scale is missing. Unclear what the comparison is and if the genes\nin green are more or less represented in the newly formed HSPC. It is also unclear if the numbers\nin the boxes represent fold change or log2 fold change. Please clarify.\nAuthor response: We have eliminated this figure in favor of a more informative analysis and\npresentation in the current version of the manuscript.\n9) Critical statistics are missing in several locations. For example, Figure 3G- stats should be\nincluded for NAC bnip3lb qPCR; Figure 4A- stats are missing in the comparison of the mitophagy\nreporter signal in bnip3lb knockdown compared to controls. Please ensure statistics are included\nfor all quantitative data.\nAuthor response: We have added the statistical analysis and notation for these figures, which\ncan now be found in Figure 3j (bnip3lb in con vs NAC), and Figure 4b (mitophagy puncta in\ncontrol vs bnip3lb MO), respectively.\n10) In Supp Figure 4B, it is very difficult to see the o-dianisidine staining in the larvae. Please use\nbetter images.\nAuthor response: We have repeated this experiment and retaken representative photos of the\nembryos as requested. The new images are now included in Supp. Figure 4b.\n11) In Figure 4G, the cmyb ISH signal in the pink1 MO is out of focus. Please provide a better\nimage.\nAuthor response: As data was added to illustrate the specificity of impact of loss of bnip3lb\ncompared to other mitophagy genes on HSPCs, we focused on validating effects through flow\ncytometry to better enhance quantitative evaluation and comparison. In doing so, we eliminated\nthe WISH photos/distribution for pink1a. Flow data for pink1a MO knockdown on CD41lo HSPCs\nis shown in Supp. Figure 4D, along with the data for bnipla, bnip3, and fundc1 MOs.\n12) In Figure 5H, the number of cells/embryos analyzed should be included on the figure.\nAuthor response: We have removed the pie chart distributions and added scatter plots with error\nbars with each point representing the individual samples analyzed for each condition. Please find\nthe updated visual display of the “% Apoptotic HSPCs” included as Figure 5h-i.\nReviewer #2 (Remarks to the Author):\nIn this study, Meader et al. investigate the regulatory mechanisms of embryonic hematopoietic\nstem and progenitor cells (HSPCs) in maintaining multipotency during rapid proliferation. They\ndemonstrate that in zebrafish embryos, ROS levels drive initial HSPC specification in the dorsal\naorta, but these levels need subsequent control to prevent apoptosis and differentiation. Bnip3lb-\nmediated mitophagy emerges as a key mechanism in this process, reducing ROS during HSPCs'\nendothelial-to-hematopoietic transition (EHT) and in the caudal hematopoietic tissue (CHT).\nKnockdown of Bnip3lb in zebrafish impairs mitophagy, leading to increased ROS, apoptosis, and\nskewed differentiation toward myeloid lineages, effects reversible by antioxidant treatment.\nFurther, activation of mitophagy in human iPSC-derived HSPCs increases their long-term\nhematopoietic potential, suggesting therapeutic implications for improving stem cell transplant\nefficacy. This study underscores the potential significance of developmentally programmed\nmitophagy in stem cell maintenance and suggests strategies to boost HSPC production ex vivo\nthrough controlled mitophagy.\nDespite these intriguing implications, the study seems significantly underdeveloped. Notable\nlimitations include: (1) an over-reliance on qualitative data, (2) insufficient controls for morpholino\nstudies and lack of alternative methods, and (3) limited validation in the human system, despite\nthe claims made. Overall, it is unclear whether the central tenet of this paper is adequately\nsupported by the data presented.\nAuthor response: Thank you for your thorough reading of our manuscript. We have aimed to\naddress the broader concerns raised above along with the specific comments listed below. To\naddress potential over-reliance on qualitative data, we have added both statistical analysis and\nan additional quantitative method to confirm phenotypes found in each qualitative experiment. To\naddress concerns regarding insufficient controls for MO studies and lack of alternative methods,\nwe have now expanded our controls and incorporated alternative tools to validate our claims.\nFinally, we have expanded our assessment of relevance in the human system. The specific\nchanges are reflected in the manuscript and addressed in more detail below.\nMajor comments:\n1) Throughout the manuscript (e.g. Fig 1D, 1J, 4C, 4E, 4H, 4J, 4L, etc.), the authors rely on\nqualitative metrics rather than quantitative metrics. They frequently bin data into “high”, “medium”,\nor “low” based on subjective scoring. Additionally, sample sizes (n) are not provided, nor are\nstatistical tests applied to demonstrate significant differences between conditions. There is also\nno indication that these categories were graded in a blinded manner.\nAuthor response: We appreciate your concerns regarding the qualitative nature of the WISH\ndata. Given that zebrafish are not inbred animals nor artificially synchronized in their development,\na bell-shaped distribution of expression levels is typically observed in untreated, wild-type\nembryos, particularly across replicate clutches, hence the utility of scoring animals into these\nthree relative categories (high/medium/low). Stained embryos were directly compared to stage-\nmatched sibling controls across multiple clutches, with the exception of embryos from\nhomozygous mutant incrosses, which were compared alongside matched wild-type clutches as\nindicated. At least 20 embryos from a minimum of two independent clutches were analyzed per\ntreatment condition and/or genotype, with blinded phenotype scoring independently assessed by\nat least two lab members. Images were selected to represent the median expression pattern\nobserved for each treatment/condition. We have expanded this information in the methods to\nprovide further clarification for the reader (page 16). We have also added the appropriate chi-\nsquared statistical analyses to support claims regarding comparisons between experimental\nconditions and updated the figure legends to include specific sample sizes for each experimental\nvariable, rather than the previously reported n-value ranges. Additionally, for each major claim of\nthe paper we have ensured that there is an independent quantitative method of evaluation (qPCR,\nFACS or confocal) utilized to support the validity of our conclusions.\nMany to most of these studies should be quantified. For example, WISH data could be quantified\n– either through alternative approaches (e.g. CD41:GFP in the VDA in Fig 1C) and/or through\ndirect quantitation of signal in regions of interest.\nAuthor response: Where independent validation by qPCR, FACS or confocal was not already\nincluded, we have now added corresponding quantitative methods to support each relevant\nqualitative study presented in the manuscript. Of note, the CD41:GFP reporter is not strongly\ndetected at early timepoints in the VDA without the use of confocal imaging, however this line can\nbe easily utilized to enumerate HSPCs via flow cytometry at 48 or 72hpf. As alluded to in the\nresponse above, due to the inherent experimental variability in WISH stain intensity across\nreplicates from alternate dates (hence comparisons to matched controls from the same cohort)\nand difficulties resolving individual cells in the absence of a cell border marker, we prefer not to\nmeasure or enumerate 2D area of RNA signal and apply binned normalization to “quantify” data,\nwhich is then independently validated by an alternative method as indicated above.\n2) Genetic rescue experiments were not performed, and genetic alternatives were not used to\ncorroborate the effects of any morpholinos.\nAuthor response: As indicated above in response to Reviewer 1, we have performed the base\nexperiments, including validation of the effects on HSPCs at 72hpf and epistasis analysis with\nNAC, with a second bnip3lb MO. We have also generated a bnip3lb crispant tool to evaluate\nimpact on HSCs by an alternative method. We find a corresponding HSPC decrease whether\nusing a splice- or ATG-targeted bnip3lb MO or the crispant as shown in Figure 4h-j.\n3) Fig 3B. This figure appears critical, yet it is unclear how well it supports the hypothesis or aligns\nwith its description in the text. How was the strength of the mCherry signal in Kdrl+ cells\nevaluated? To what was it compared? Quantification is lacking. Furthermore, the only cells\npresented (Fig 3 and S3) seem to come from the same image/movie of a single sample. This\nanalysis needs to be quantified and repeated across multiple embryos to support the claim\nrobustly. More specific markers for cell types of interest could also improve clarity.\nAuthor Response: Thank you for this important comment. The goal of this image was to visually\nconfirm where the mitophagy was occurring within the endothelial population lining the dorsal\naorta and posterior cardinal vein, marked by Kdrl:BFP expression; notably, we only observed the\npresence of mitophagy puncta in rounded or “budding” cells in the VDA. The data shown in Figure\n3b using the MitoKeima reporter is a representative sample from a series of 5 independent\nbiological samples analyzed. We have now also included a representative image from analysis of\nthe MitoGR line, from a series of 4 independent biological samples in Figure 3c; this line was\npreviously utilized to calculate the mitophagy index score quantification shown in Figure 3d,\nwhich indicated significantly more mitophagy was present within the BFP+ fraction inclusive of\nEC/HEC/HSPCs within the CHT compared to the VDA, in line with our hypothesis.\nOne limitation of this analysis, is that each of the previously established MitoKeima and MitoGR\nreporter systems utilize both the green/GFP and red/mCherry fluorescent channels, making it\ndifficult to co-localize mitophagy with well characterized HSC marker lines such as Runx1\ntransgenic reporter - a highly conserved marker for HSCs in the AGM at this stage of development\n- for which we have only red and green transgenic tools available. While crossing this line to a\n“blue” endothelial cell reporter Tg(kdrl:TagBFP) allowed us to identify putative budding HSPCs\nbased on morphology and location in the niche, we were unable to definitively quantify mitophagy\nin HSPCs vs non-HSPCs within the labeled Kdrl+ endothelium using just the previously published\n‘mitophagy index’ methodology (as described in Wrighton et al4), which is essentially a ratio of\npuncta volume/ tissue volume, given that the “tissues” are not distinguished by a definitive label.\nNevertheless, as indicated above, while primarily using these panels to illustrate the presence of\nmitophagy within the Kdrl+ fraction, we were able to quantify the fraction of Kdrl+ cells showing\nactive mitophagy by confocal imaging, revealing a statistically significant increase in co-\nexpression when comparing cells from the VDA and CHT (Figure 3d). Furthermore, we now also\nprovide qPCR analysis demonstrating enrichment for mitophagy related genes, bnip3lb and pink1,\nwithin the Kdrl+cMyb+ HSPC fraction, as opposed to non-hemogenic Kdrl+ cells (Figure 3f,\nSupp. Figure 3f-j). With HSPC co-localization more firmly established, in subsequent figures, we\npresent claims regarding alterations in MitoGR-labeled mitophagy within HSPCs following gene\nknockdown or induction, utilizing a count of individual mitophagy positive puncta (Figure 4b-c)\nor the recommended “mitophagy index” quantification (Figure 6a-b) to support our claims.\n4) Mitophagy was fundamentally mischaracterized in several places:\na) “Pink1 is then ubiquitinated by Parkin” – please provide a reference for this, as most studies\ndo not support this claim.\nAuthor response: We apologize that the abbreviated description of mitophagy initiation and\nprogression inadvertently came off as a mischaracterization, and have amended the text to more\naccurately reflect the steps of the process. The sentence on page 3 in the second paragraph\nindicated above now reads: “Pink1 then phosphorylates Parkin, which draws in the autophagy\nmachinery35–37” with appropriate citations indicated.\nb) “Loss of outer membrane integrity leads to PINK1 translocation from the matrix to the\nmitochondria’s surface” – Please provide a reference; most studies suggest a failure to import\nfrom the cytosol (e.g., due to the ANT complex, PMID: PMC6858570), rather than\nretrotranslocation from the matrix.\nAuthor response: We have amended the text to better reflect current understanding of the\nprocess of mitophagy. The sentence on page 3 in the second paragraph now reads: “Loss of\nmembrane integrity leads to Pink1 accumulation on the mitochondrial outer surface35. “\nc) “[Mitophagy] is initiated when proteins on the outer mitochondrial membrane are tagged to mark\nthe organelle for destruction” – This is accurate for PINK1/Parkin-mediated mitophagy, but the\ninitiating signal for FUNDC1/BNIP3/NIX is less clear. Phosphorylation mediates some cases but\nis not an absolute switch, as suggested here. It may be that general expression levels of these\nreceptors, driven by specific transcription factors, control mitophagy induction.\nAuthor response: We appreciate the need for this important distinction and have amended the\npage 3 text to say: “The latter, a specialized form of cellular autophagy, is initiated when proteins\non the outer membrane of a mitochondrion are tagged to mark the organelle for destruction or via\ncontext-driven transcriptional regulation of mitophagy receptor expression...” We hope this\nrevision now acceptably captures both routes of activation as they are currently understood.\nd) mitoQC was attributed to Zhao et al (ref 64). I believe this is a spurious reference. If so, please\ncorrect. (suggested PMID: 24176932 or similar).\nAuthor response: Thank you for spotting this reference error. We have now corrected the text\nto include the citation utilized previously in the Wrighton paper (Rojansky et al., 2016).15\ne) Fig 3A misrepresents the mitoQC reporter. It should display both red and green before the\nlysosome, and red only in the lysosome. Speculation may be helpful for why this does not seem\nto be the case in the zebrafish model.\nAuthor response: We apologize that our simplified schematic of the in vivo mitophagy reporter\ntool shown in the prior version of Figure 3a was perceived to inadvertently mischaracterize the\nMitoQC reporter line. The illustration was meant to describe the MitoKeima tool where the “red”\ncolor becomes detectable in the presence of mitophagy due to the pH change resulting from\nlysosomal fusion; however our description in the text did not adequately describe the behavior of\nthe single pH sensitive fluorophore used in the representative images in Figure 3b. In regard to\nthe MitoGR reporter (please note, to enhance consistency with the prior publication and construct\ndesign, we have adjusted the MitoQC label to MitoGR throughout the paper text and figures), we\nagree with the reviewer’s statements on expected baseline versus mitophagy active fluorophore\nexpression: as demonstrated in the data in Figure 3c, as well as the prior report by Wrighton et\nal4, both red and green proteins are expressed at baseline; each color is detected separately by\nconfocal microscopy, and then visualized as varying degrees of yellow (to white) in the resulting\nmerged images based on signal strength. Upon fusion with the lysosome, due to the Ph change,\nthe green GFP signal is quenched, allowing only the more pH resistant red mCherry signal to\nremain detectable in select cells. In the revised version of the schematic, we have now added a\nsecond series of images to highlight the differences between the two tools, including re-coloring\nthe initial image of MitoGR mitochondria to more clearly indicate the combined fluorescent\nlabeling at baseline, and the remaining “red” color post lysosome fusion. In our hands, during\ntimepoints relevant to HSPC development, the MitoGR line proved more readily quantifiable by\nImaris when utilized in combination with the Kdrl:BFP EC/HEC/HSPC marker, and as such was\nused for the mitophagy index calculations shown.\n5) The application of these findings to hiPSCs is potentially impactful, but needs to be solidified.\nImportantly,\na. Given the broad and potentially varied effects of NR, the authors should confirm that NR\ntreatment increases mitophagy in hiPSCs.\nAuthor response: We appreciate this concern as NR is known to impact Sirtuin activity16, which\nin addition to influencing initiation of mitophagy, can lead to regulation of antioxidant gene\nexpression8, providing an additional route to control ROS levels. To address this important point,\nwe set out to create a stable iPSC line expressing a previously described mitophagy reporter;\nunfortunately, despite several attempts at introducing the construct, we were unable to establish\na line that consistently documented mitophagy activation in response to biochemical stimulation.\nAs an alternative approach to validate the beneficial impact of mitophagy on newly specified HSCs\nin culture more generally, we exposed cells to FCCP, which stimulates mitophagy through\ndepolarization of the mitochondria membrane. Importantly, we found a similar enhancement in\nHSPC production for exposure to either NR or FCCP in iPSCs (Figure 7j) as well as in zebrafish\nembryos (Figure 7b). Furthermore, in the zebrafish model, we confirmed that both NR and FCCP\nare mediating their effect (at least in large part) through mitophagy via chemical epistasis\nexperiments with Bafilomycin, which inhibits lysosomal fusion. When progression of mitophagy is\nblocked via Bafilomycin treatment, neither NR nor FCCP stimulation can enhance CD41lo HSPCs\nnumber (Figure 7a-b). A primary role for NR-driven mitophagy was further confirmed in zebrafish\nthrough epistatic investigations with the bnip3lb MO, where NR stimulation was unable to increase\nHSPC numbers in the setting of loss of receptor-mediated mitophagy (Figure 7c). Together, our\ndata indicates that NR is most likely eliciting its primary effect on HSPCs through activation of\nmitophagy and subsequent control of ROS levels.\nb. Moreover, the beneficial effects of NR in Fig 7 should be BNIP3 dependent to support this\nmodel.\nAuthor response: We would politely disagree with this point. While our zebrafish data indicates\nthat the primary regulatory program activated in the developmental setting is Bnip3lb-related (with\nsome modest compensation and/or redundancy from other receptor subunits), we find also that\nany exogenous compound-mediated activation of mitophagy, provided that the machinery\nnecessary to initiate the mitophagic program is present (all components are expressed in iPSC\ncells according to https://anglohlabs.shinyapps.io/blood_devcell_shiny), should exhibit a\nbeneficial impact on ROS levels and maintenance of HSC numbers. To support this conjecture,\nwe now provide data for both NR and FCCP stimulation and see a similar impact in vitro (Figure\n7j). We did also conduct the requested epistasis investigation in zebrafish embryos using the\nCD41 HSPC reporter and found that loss of bnip3lb did blunt the ability of NR exposure to\nsignificantly enhance CD41lo HSPC numbers (Figure 7c).\nc. Does a general decrease in ROS (e.g. NAC treatment) similarly improve the secondary CFUs\nof hematopoetic progenitors, or is there something special about mitophagy’s role?\nAuthor response: We thank the reviewer for bringing up this important point. ROS levels have\nbeen well documented to tip the scale between cell cycling and quiescence17, as well as lineage\ncommitment and retention of adult HSC fate.18 NAC can be utilized to regulate cellular ROS levels\nmore generally, both in vivo and in the culture dish19, however, the precise level of ROS that is\nneeded to allow for HSC expansion, while maintaining multi-potency is not established. Indeed,\nprior reports have indicated that in the cell culture setting, as well as in the clinic following stem\ncell transplant, complete ROS inhibition has detrimental effects on HSC function and\nhematopoietic homeostasis.20 For that reason, we were intrigued by the potential impact of\nmitophagy in maintaining HSC state, as its function can be dependent on perceived local\nenvironmental stimulation (both in regard to gene expression of its components and cellular\nactivation). Our data indicates that transient activation of mitophagy during iPSC differentiation\nappears to be sufficient to decrease ROS to an acceptable level to promote self-renewal.\nMinor comments:\n1) Fig 1B and related box plots (e.g. 5A, 7F). While these are “representative examples,” it is\nunclear whether the statistical test was performed across three biological replicates or only on the\nsingle sample shown. This should be assessed across biological replicates (N=3), with\ninformation on the number of cells analyzed per sample. Box plots are an atypical choice for this\ndata; a histogram or violin plot might be more appropriate. At a minimum, include a description of\nthe box plot structure (e.g., inner/outer quartiles [box] and 1.5*IQR [whiskers]).\nAuthor response: We agree that the box plot display was not an informative representation of the\ncomplete analysis performed. We have adapted these figures to a bar graph form with normalized\nmedian fluorescence intensity (MFI) data displayed. Each point represents a pool of 5-9 embryos\nanalyzed with at least 5 biological replicates for each analysis. The CellROX plot in Figure 1B\nnow also directly correlates to the CD41:GFP HSPC flow plot in Figure 1F; the ROS data in\nFigure 5a and Supp. Figure 5e and Supp. Figure 7s were similarly updated.\n2) Is OTC-mediated mitophagy BNIP3Lb-dependent? If not, does the form of mitophagy matter?\nAuthor response: We agree with the reviewer that it is generally believed that delta-OTC\nassociated mitophagy is not Bnip3lb dependent, but rather stimulates the Pink/Parkin pathway\nvia damage/stress response activation.21,22 As now more accurately detailed in our analysis, we\nfind that developmentally programmed/receptor-mediated mitophagy, such as that mediated via\nBnip3lb, appears to be required during HSC development to allow for HSC expansion while\npreserving multipotency. However, as long as the pathway components are expressed in the cell\ntype of interest, we are in fact able to further stimulate HSC production through activation of any\nform of mitophagy via transgene induction or compound exposure both in vivo and in vitro.\n3) Fig 1C. At 72h, are HSPCs in prdx1-MO animals still stuck in the VDA, suggesting they fail to\nmigrate?\nAuthor response: This is an interesting question. When evaluating these samples, we always\nlook at the whole embryo in addition to locally in the hematopoietic niche of interest at the time\npoint of analysis. We did not observe HSPC retention within the VDA at 72hpf in these samples.\n4) Fig 2F. Please provide a reference for the mitophagy reactome annotated genes. Mitophagy is\ngenerally not recognized to have a specific transcriptional signature.\nAuthor response: Thank you for pointing out that oversight in the original manuscript. In the\nreanalysis of our single cell data, and with the addition of the new dataset, we have eliminated\nthis analysis altogether. We now perform GSEA and highlight key Gene Ontology - Biological\nProcess terms along with direct visualization of known mitophagy genes in the HSPC clusters.\n5) Fig 2H. How is “enrichment” determined? PINK1, FUNDC1, and BNIP3 do not appear notably\nenriched. Please specify the unbiased metric(s) used to support enrichment claims.\nAuthor response: Thank you for this important clarification question. We have now reanalyzed\nand restructured the scRNAseq expression profiling study to better support our claims regarding\nexpression within the various niche fractions, inclusive of the HSPCs; we have also incorporated\nnew data sets to provide additional insight into changes associated with maturation of EC to HSPC\nstate in the VDA and CHT niche, including a metabolic switch associated with strong upregulation\nof oxidative phosphorylation and ROS production (Figure 2c-d). Our data indicates that\nexpression of mitophagy components increases during the transition from EHT (36hpf) to HSC\nexpansion (48hpf) in the CHT (Figure 2e-f), with bnip3lb, fundc1 and pink1 showing both the\nhighest comparative expression in the HSPC fraction (bnip3la is also upregulated in the CHT).\nTo further investigate cellular specificity, we profiled expression in Kdrl/cMyb-labeled FACS isolated\ncell fractions by qPCR (Figure 3f, Supp. Figure 3f-j) and can now more confidently support the\nclaim that bnip3lb is uniquely enriched in HSPCs during this window of hematopoietic\ndevelopment in the developing embryo.\n6) Pg 12. “…ATG5-dependent autophagy is essential for EHT, highlighting the importance of\nmultiple autophagic pathways in embryonic HSC production.” Does this imply that BNIP3lb-\nmediated mitophagy and Atg5-mediated autophagy are distinct? If so, what evidence suggests\nthat BNIP3lb-mediated mitophagy is ATG5-independent?\nAuthor response: We thank the reviewer for this insightful question. While ATG5 is a core\ncomponent of the canonical autophagy machinery, several studies have shown that mitophagy\ncan occur through ATG5-independent mechanisms, depending on the molecular context and the\nreceptors involved. For example, certain forms of receptor-mediated mitophagy, including those\ninvolving BNIP3 and NIX (BNIP3L), can proceed even in the absence of ATG5, through alternative\nmacroautophagy routes utilizing other autophagy-related proteins such as ATG6/Beclin1 and LC3\nfamily members.23,24 In our system, Bnip3lb appears to act as the primary mitophagy receptor\nutilized during hematopoietic development, but our data do not exclude potential involvement of\nAtg5. Instead, our interpretation is that Bnip3lb-mediated mitophagy could function either through\ncanonical ATG5-dependent autophagy or via compensatory pathways employing other\nautophagy-related proteins (e.g., Atg6/Beclin1, as described in erythroid mitophagy). Thus, the\nrequirement for ATG5 in developmental mitophagy may vary depending on cellular context and\nautophagic flux requirements. Importantly, ATG5-dependent autophagy is not specific to\nmitochondria and encompasses multiple degradative processes24, whereas BNIP3lb-mediated\nmitophagy represents a more selective, developmentally regulated form of autophagy. Therefore,\nour data emphasize that multiple autophagic mechanisms, both general (ATG5-dependent) and\nselective (BNIP3lb-mediated), contribute to embryonic HSC emergence, underscoring the novelty\nof our findings in revealing their coordinated functional impact during EHT. We have now better\nhighlighted this interesting point in the discussion as it will be an important area for future\ninvestigation.\n7) Pg 5. For clarity, please introduce the CD41 marker at its first mention (Fig 1E) rather than its\nsecond (Fig 1G, H).\nAuthor response: Our apologies for that oversight; while the figures have been reorganized in\nthe revised submission, we have carefully amended the text as requested to ensure that the CD41\nreporter, as well as other lines utilized, are defined appropriately at first mention.\nReviewer #3 (Remarks to the Author):\nIn the manuscript, the authors describe the role of Bnip3lb-regulated mitophagy in maintaining\nthe embryonic hematopoietic stem cell pool in zebrafish. The authors elaborate on how\nspatiotemporal control of ROS is important for generating and regulating the HSPC pool. First,\nthe authors show the essentiality of ROS for HSPC specification in the ventral-dorsal aorta (VDA)\nniche but not for HSPC specification in the caudal hematopoietic tissue (CHT) niche. Next, they\nclaim that Bnip3lb-driven mitophagy regulates ROS within the CHT. Knocking down Bnip3lb\ncauses a reduction of embryonic HSPC and increased ROS at CHT, which can be rescued by\ntreatment with NAC. Finally, they present data on how NR-induced mitophagy can be leveraged\nto form hiPSC-derived hematopoietic progenitors. This is a potentially very interesting manuscript,\nbut the data are unfortunately not of sufficient quality to conclude that Bnip3lb-regulated\nmitophagy is responsible for the observed phenotypes.\nAuthor response: Thank you for this comprehensive review of the data provided in our original\nsubmission. We have addressed your comments as detailed in the point-by-point response below,\nwhich we believe help to further strengthen our conclusions in this revised manuscript.\nMajor Points\n1. In Figures 3-4, 6A and S3 the authors use the Ubi:mito-QC transgenic zebrafish line to study\nmitophagy in zebrafish. The authors describe the GFP to be mitochondria and the magenta\n(mCherry) to be mitophagy, which is misleading as the reporter is a tandem-tagged (mCherry-EGFP)\nmitochondrial protein. Thus, mitochondria are labeled both green and red (should be seen\nas yellow or white) in normal situations, while upon induction of mitophagy (and transfer of the\nreporter to acidic lysosomes), the green signal quenches and thus only red/magenta-only puncta\ncan be seen. For quantification of mitophagy, it is therefore important to adjust the microscopy\nsettings so that mitochondria are seen as yellow/white (optimal is to treat with a lysosomal\ninhibitor). There is no information about how they have done the quantification of mitophagy in\ntheir samples, other than (p.14) “ a ratio of the mCherry/GFP values for each pixel was taken, and\nthe results were averaged to give the final value”. Thus, if the microscopy settings are off (as\nindicated by the authors as well on p.6, “GFP signal being bright for mitochondria”) these data\ncannot be evaluated properly.\nAuthor response: We apologize that our description of the in vivo mitophagy reporter system\ninadvertently mischaracterized the fluorophore expression in the MitoGR reporter line (please\nnote, to enhance consistency with the prior publication and accurately reflex construct design, we\nhave adjusted the MitoQC label to MitoGR throughout the paper text and figures); due to\nred/green colorblindness amongst members of our investigational team, we intentionally false\ncolor the image panels to aid interpretation and often subsequently describe the data as an\neffective binary switch - “healthy” to “mitophagy +” - rather than more accurately describing the\ncolor-change (for MitoKeima) or relative balance (for MitoGR) of each fluorophore detected.\nWe have utilized both the MitoKeima and MitoGR lines to assess mitophagy in the VDA and CHT\nto independently validate our observations; after reviewing the confocal settings for each sample\nto confirm appropriate labeling, we now show representative image panels from a total of 5\nbiological replicates for MitoKeima (Figure 3b) and 4 for MitoGR (Figure3c). We have also\nreconstructed the schematic in Figure 3a to help distinguish between the two lines. In agreement\nwith the reviewer’s comment, the MitoGR construct is composed of a mitochondria targeting\nsequence upstream of tandem eGFP and mCherry sequence tags, driven by the Ubitiquin (ubi)\npromoter, as previously reported by Wrighton et al4, or by CD41, as created specifically for the\nHSPC analysis presented. As illustrated in the original study, both red and green proteins are\nindeed appropriately expressed at baseline; each color is detected separately by confocal\nmicroscopy, and can be visualized as varying degrees of yellow (to white) in the resulting\nmerged images dependent on signal strength. Upon fusion with the lysosome, due to the Ph change,\nthe GFP signal is quenched in the MitoGR line, allowing only the more pH resistant red signal to\nremain detectable. We have now included a more detailed description in the methods regarding\nhow the embryos were imaged, as well as how mitophagy was quantified within the VDA or CHT\nregion; of note, this work was conducted in collaboration with the Goessling lab, who published\noriginal zebrafish study, with similar methods employed and several of the authors directly\nassisting with imaging and analysis. To aid identification of mitochondrial status in the cells of\ninterest for the reader, we have now further broken down the representative image panels\nincluded to reveal both the individual and merged fluorophores, with false coloring employed to\nhighlight the relevant expression driving calculation of the mitophagy index within the Kdrl:BFP+\nEC/HEC/HSPC population.\n2. The authors used morpholino (MO) to deplete various genes (including Bnip3lb), but there is\nno validation of the knockdown efficiencies. Moreover, as only one MO/gene is used, the authors\nshould either use another MO and/or CRISPR-Cas9. In addition, it would be advisable to rescue\nthe Bnip3l phenotype.\nAuthor response: We appreciate this concern and have performed the base experiments,\nincluding analysis of effects on HSPCs at 72hpf and epistasis with NAC, with a second bnip3lb\nMO targeting the ATG site; we also generated a bnip3lb crispant as an alternative method to\nevaluate impact on HSCs. We see corresponding reductions in HSPC marker gene expression\nor HSC number by FACS whether utilizing the splice or ATG bnip3lb MO and in the crispant\nembryos (Figure 4h-j; Supp. Figure 5f). We did not perform the suggested rescue experiments\nas the bnip3lb phenotype is altered by compensation in the stable mutant line, as now more\nextensively illustrated in Supp. Figure 4e-g, and co-injection of bnip3lb RNA with either MO\nconstruct at doses eliciting phenotypic knockdown was not well tolerated by the embryo.\nNevertheless, the consistent HSPC phenotype observed across the expanded tools, as well as\nthe retention of the ROS/NAC epistatic interactions, together support the validity of our claims.\n3. Whole mount in situ hybridization (WISH). Throughout the manuscript, they do WHISH for\nrunx1/cmyb together. Any reason why it is not done individually? The negative controls (using the\nsense probe) are missing, and the WHISH staining varies a lot from image to image (e.g. control\nin Figure 4D vs 4G). The latter is particularly problematic as they quantify this staining throughout\nthe manuscript and use this as a basis for their phenotype distribution. The phenotypic distribution\ncharts are also confusing. How do they score high, medium, and low? They have not mentioned\nthe number of larvae used and whether the results are significant or not.\nAuthor response: We apologize for any confusion caused by the presentation of the WISH data\nand/or rationale for the methodology employed and evaluation of results. We have opted to utilize\ncombined application of the runx1/cmyb probe set as it helps to delineate budding HSPCs from\nother populations present within the ventral dorsal aorta. Runx1 is expressed at low levels on\nprimitive erythrocytes in circulation, flattened endothelial cells on the ventral wall of the dorsal\naorta which have initiated hemogenic gene programs (but may or may not complete budding), as\nwell as newly specified HSPCs. At this stage of development, cMyb is expressed on primitive\nmyeloid populations, and maturing HSPCs budding from the ventral wall of the aorta. As all of\nthese populations are adjacent in time and space, we generally combine the probes to better\nhighlight (amplify signal) the beads-on-a-string pattern of the “budding” runx1/cmyb double\npositive fraction representative of HSPCs, and not nearby primitive myeloid or erythroid cells or\nendothelial cells that may not complete EHT. Both probes have been widely utilized in the field\nsince the early 2000s25, such that we and others have published numerous reports with them\nindividually or together (including sense strand controls) to establish the validity of the expected\nexpression pattern and relevance to our biological process of interest. We have elected to keep\nthem combined here to more easily compare newly specified HSPCs between embryos; however,\nwe have now also provided a representative image of the individual WISH probes in the CHT at\n72hpf for comparison (Supp. Figure 1c).\nGiven that zebrafish are not inbred animals, and their development was not artificially\nsynchronized from time of aqueous fertilization, a bell-shaped distribution of expression levels is\ntypically observed in untreated, wild-type embryos, hence the utility of scoring animals into these\nthree relative expression categories (high/medium/low) across replicate clutches. Stained\nembryos were directly compared with stage-matched sibling controls within a single clutch to\njudge alterations in WISH patterning and intensity, with the exception of embryos from\nhomozygous mutant incrosses, which were compared alongside matched wild-type clutches as\nindicated. At least 20 embryos from a minimum of two independent clutches were analyzed per\ntreatment condition and/or genotype, with blinded phenotype scoring independently assessed by\nat least two lab members; numbers of individual embryos analyzed per condition are now\nindicated to replace the previously reported ranges. Images were selected to represent the\nmedian expression pattern observed for each treatment/condition, using embryos from the same\nclutch. We have added this information to the methods to provide further clarification to the reader\n(page 16). As WISH distribution is a qualitative assessment of expression levels, in addition to\nthe chi-square analysis now provided for each data set, we have included an additional\nindependent quantitative assessment by qPCR, flow cytometry and/or microscopy to verify and\nfurther enumerate the phenotypes observed and support the validity of our claims.\nMinor points:\n1.Text on page 5 line 8 says –“flow cytometry of disaggregated 48 hpf embryos” whereas in figure\n1b it is written 72 hpf.\nAuthor response: Thank you for pointing out that error, which was a holdover from an earlier\niteration of the manuscript. We have adjusted the text to match the figure, indicating 72hpf.\n2.Figure 1B, 5A og 7F; definition of X-axis units are missing\nAuthor response: We have updated the formatting of the ROS analysis to better capture\nchanges in data across conditions/treatments. The mean fluorescence intensity (MFI) of CellROX\nincorporation is now displayed as relative values to better illustrate scale and directionality of\nchanges across treatment groups and replicate runs by fold change.\n3.Figure 1G, both for CHT and KM panels, the fluorescent signal is not adequate to see any\ndifference. They should include an inset (zoomed image) of the region indicated in the KM panel.\nAuthor response: This figure panel was replaced to better visualize the fluorescent signal.\n4.Both rag1 and rag2 are used without clear elaboration. WISH was done for rag1 (Figure 1i)\nwhile FACS was done on cells from transgenic line Tg(rag2:GFP). Why the switch?\nAuthor response: Both Rag1 and Rag2 mark the same population of lymphoid cells26, with the\nlatter generally showing more dynamic turnover. The alteration in use was simply for practical\nreasons: the transgenic line utilized in the field is driven via the rag2 promoter (both lines were\noriginally created in the Zon lab27, but only the rag2 line retained heritable adult expression of the\nreporter construct), while the rag1 in situ probe27 is the more widely available tool.\n5.FISH for bnip3lb staining looks unspecific (Figure 3 D). They should include a negative control?\nIs the strong staining observed in the vertebral column true or autofluorescence?\nAuthor response: We appreciate concerns raised regarding specificity of bnip3lb expression by\nFISH (now Supp. Figure 3l), however, our data are in line with WISH patterning for Bnip3lb shown\non Zfin.org, as assessed by global expression analysis in the Thisse lab28 (Thisse et al., 2004 -\nFast Release Clones: A High Throughput Expression Analysis: https://zfin.org/ZDB-FIG-050630-\n12393) and strikingly similar to that reported for its presumptive transcriptional regulator hif1ab\n(https://zfin.org/ZDB-IMAGE-040914-1194).28 While expressed at a low level in a variety of cell\ntypes during this window of development given the increasingly metabolically active state of the\ngrowing embryo, by qPCR analysis of sorted cell fractions from manually disaggregated tail\nregions of Tg(kdrl:mCherry;cmyb:GFP) embryos, we were able to confirm bnip3lb expression in\nboth the double negative Kdrl-cMyb- fraction (inclusive of cells from the vertebral column), as well\nas in the Kdrl+cMyb+ HSPCs population (Figure 3f).\n6.Can the bnip3lb-driven mitophagy-dependent HSC specification in CHT be compensated by\nFUNDC1? Both Pink1 and Fundc1 were expressed at low levels in both subclusters in Figure 2H.\nThe authors checked for the effect of pink1 knockdown in HSPC gene expression at 72 hpf (Figure\n4G) but Fundc1 was not investigated. Can fundc1 have a role similar to bnip3lb in HSPC\nspecification via mitophagy/ROS axis since both are induced upon hypoxia?\nAuthor response: This is an important point, which we agree that we previously did not discuss\nin detail in the paper, but have now better incorporated for several key analyses. By scRNAseq,\nin addition to bnip3lb, we see expression of both pink1 and fundc1 in the HSPC cluster at 36 and\n48hpf (Figure 2e-f). Via qPCR analysis on FACS-isolated fractions of the tail region from\nTg(kdrl:mCherry;cmyb:GFP) embryos, most Hif1α-regulated, receptor-mediated mitophagy\ncomponents (fundc1, bnip, bnip3la, and bnip3lb) as well as classically Hif1α-independent (pink1a,\nparkin1) mitophagy mediators appear to be expressed in the macrophage (Kdrl-cMyb+) and/or\ndouble negative (Kdrl-cMyb-) populations at 48hpf (Figure 3f; Supp. Figure 3f-j); only pink1a\nand bnip3lb showed strong enrichment in the Kdrl+cMyb+ HSPCs. To further assess a potential\nfunctional role for any of the other mitophagy components present at this stage of development,\nwe knocked down bnip3, bnip3la, fundc1 and pink1 with their respective MOs: no change in\nCD41lo HSPC formation was observed by FACS for any of the other MOs tested (Supp. Figure\n4d ), in line with the data shown previously for pink1a in the primary submission.\n7.For the FACS analysis figures they use “% Live Cells” for the y-axis. This is confusing, as it is\nnot well described.\nAuthor response: We apologize for confusion arising from use of this abbreviated notation. The\nFACS analysis is conducted on cells falling within the “live” gated fraction of total cells, following\nlive/dead exclusion with SYTOX Red (ThermoFisher). As such the “% Live Cells” label is simply\nindicative of the population exhibiting the marker of interest indicated, usually on top of the graph,\nfrom the “live” fraction of cells within the total disaggregated cell population analyzed per sample.\nWe have now defined the term at first use in the text and methods (page 5).\n8.The pie chart shown in Figure 5H of dead and alive cells is a bit vague. They should describe\nhow the analysis was done. The legend says “Pie charts show dead Runx1+ cells (magenta) as\na proportion of the total, n=3/condition” Is this 3 larvae per condition?\nAuthor response: We have significantly altered the presentation and description of the apoptosis\ndata in Figure 5h-i. We counted the number of Runx1+ HSPCs marked with acridine orange\n(indicating cells undergoing apoptosis) per CHT region imaged, and divided this number by the\ntotal number of Runx1+ HSPCs present in the field to generate the percent of HSPCs undergoing\napoptosis. This data is now represented by a scatterplot with error bars. Each data point\nrepresents a single sample analyzed, with 3 sibling larvae examined per condition.\n9.On p.9 the authors state “we utilized an established ΔOTC construct, which employs a clinically-\nderived mutated form of mitochondrial matrix protein ornithine transcarbamylase to induce\nmitophagy via the misfolded protein response64”. This is misleading as the referred paper does\nnot show that this mutant induces mitophagy. Moreover, the current manuscript does not show\nan effect on mitophagy, as only one image with no quantification is shown in Figure 6A.\nAuthor response: We have revised the description of this construct and the referenced citations\nto highlight both the impact of the causal mutation21,22 and the previously established connection\nto mitophagy.29 The image panels included for the mitophagy reporter (now shown as Figure 6a)\nwere representative samples from 5 embryos over 2 biological replicates. As per the Reviewers’\nrecommendation, we have reanalyzed the samples as suggested to compute a “mitophagy index\nscore” to more clearly demonstrate the strong increase in mitophagy reporter expression in the\nCHT following heat-shock induction of the ΔOTC construct (Figure 6b).\n10.Use of the mitophagy-inducing drug NR shows an increase in HSPC specification in CHT along\nwith Cas849. Have the authors tested DMOG or DFP?\nAuthor response: As now indicated more clearly stated in the text, we observed a notable\nincrease in both mitophagy reporter expression (Supp Figure 6k) and HSPC production with any\nof the mitophagy activating drugs applied (Figure 6k). This is in contrast to the knockdown\nstudies, where only loss of bnip3lb (Figure 4f-j vs Supp Figure 4d) or the use of the mitophagy\nblocking agent Bafilomycin (Supp. Figure 5c-d, Figure 7a-b) had a potent impact on HSPC\nformation, suggesting that Bnip3lb-receptor mediated mitophagy is the ROS-responsive pathway\ninitiated physiologically during development, driven at least in part via transcriptional regulation\nmediated by Hif1α (Figure 3l). Altogether, the data now presented in Figure 6 implies that as long\nas the relevant mitophagy machinery is present in the target cell-type, compound-mediated\nexogenous activation of mitophagy via stimulatory route is sufficient to drive HSC production\nand/or protection from premature differentiation, as determined by increased CD41lo HSPC\nnumbers by FACS. While not investigated in depth here, we previously demonstrated that DMOG\ntreatment can enhance HSPC development in the VDA via activation of Hif1α-associated\ntranscriptional regulation19, which would presumably include the Bnip family as identified here, as\nwell as a number of other relevant hematovascular targets.30,31\nReviewer #4 (Remarks to the Author):\nI co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is\npart of the Nature Communications initiative to facilitate training in peer review and to provide\nappropriate recognition for Early Career Researchers who co-review manuscripts.\nResponse to Reviewer Comments – Second Revision\nReviewer #1 (Remarks to the Author):\nThe authors have sufficiently addressed concerns from the prior submission.\nReviewer #3 (Remarks to the Author):\nThe authors have addressed my comments and concerns in a satisfactory manner and the\nmanuscript has been substantially improved.\nAuthor response: Thank you for your assistance in reviewing and improving our manuscript.\nReviewer #2 (Remarks to the Author):\nThe authors have made meaningful improvements, including stronger quantitative analyses, the\naddition of appropriate morpholino controls (including a second morpholino and crispant\ncontrols), and more substantial validation in human systems.\nMy only remaining concerns include some generally modest effect sizes across several assays,\nas well as the lack of direct evidence that the observed phenotypes in human cells specifically\nreflect autophagy (although this was shown to be true in zebrafish, so there is precedent).\nAuthor response: We are glad to find that you are satisfied with the majority of updates and\nextensions included in the revised submission. Regarding the first point raised, while we\nappreciate that effect sizes presented may not necessarily be perceived as dramatic across all\nassays and analyses, all claims of impact were based on robust statistical comparisons. Indeed,\nin this secondary revision, we double checked all statistical tests for accuracy, ensured all\nanalyses reported were two-tailed, and adjusted each t-test to add further rigor using Welch’s\ncorrection. Additionally, we made sure to base any claims on at least two independent methods\nof analysis (WISH vs FACS vs QPCR) and/or means of targeting each gene or pathway of\ninterest. It is important to note that for each study, one data point is often representative of a\ncohort of 5-10 (qPCR, FACS) to hundreds (scRNAseq) of zebrafish embryos; likewise, iPSC\ncolony forming data is representative of 5,000 cells plated per assay. While our data does not\nindicate that HSCs are unable to form in the absence of developmentally programmed mitophagy,\nit clearly demonstrates that robust production and maintenance of HSCs both in vivo and in vitro\nis impacted – positively or negatively – by the ability to activate mitophagy.\nConcerning the second comment, as indicated in the response to the prior review, we agree that\ndue cell viability and transformation efficiency issues we were unable to directly demonstrate that\nthe observed phenotypes in human cells specifically reflect mitophagy (specialized autophagy).\nHowever, as mentioned in the earlier rebuttal, we were able to show that stimulation with either\nNR and FCCP during hematopoietic differentiation culture of iPSC elicited the same beneficial\nimpact on HSPC numbers and function. As these two compound activators are thought to act\nthrough distinct methods, with their primary commonality being the ability to drive mitophagy,\nwhile we cannot formally exclude beneficial impact(s) from additional mechanisms of action, we\nfelt confident in our claim that mitophagy is able to enhance the functional capacity and\nmaintenance of iPSC-derived HSPCs. Additionally, as indicated in the comments, whereas a\nmore comprehensive epistatic analysis was not performed in iPSCs, we were able to directly\ndemonstrate the ability of both NR and FCCP to drive mitophagy and the resultant impact on\nHSPC numbers in the zebrafish embryo, further supporting our proposed regulatory mechanism.\nWe have included statements in the discussion (page 13) to further underscore the potential for\nalternate routes of cellular regulation in the iPSC system that will need to be explored further:\n“Of note, while similar results were observed for FCCP treatment, which acts via an\nindependent mechanism to induce mitophagy, NR regulates cellular redox via multiple\npathways and is not specific to mitophagy alone; additional work will be needed to\ndistinguish the isolated and combinatorial impact of each potential arm of its regulatory\nmechanism on HSC fate.”\nMinor point: the title of Supplemental Figure 2 (“Expression of mitophagy related genes in flow\nsorted HSPC cells”) appears inaccurate, as the figure now pertains to gene expression used to\ndefine cluster identities rather than mitophagy per se.\nAuthor response: Thank you for pointing out that disconnect between the revised supplementary\nfigure and title. We have now updated it to read:\n“Supplemental Figure 2: Identification of the HSPC population within the scRNAseq\ndatasets (Related to Figure 2)”"}]

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