Non-necroptotic MLKL function damages mitochondria and promotes hematopoietic stem cell aging

Domain: chemistry
Published: 2026-04-06
Editorial outcome: minor_revision

Abstract
Hematopoietic stem cells (HSCs) survive many types of cellular stress but often lose their regenerative and lymphopoietic capacities as a result. Such functional decline also occurs with age, and dysfunctional HSCs with impaired mitochondria accumulate during aging. However, the molecular link between HSC stress response and age-related functional decline remains poorly understood. Here we show that multiple stress responses converge on the RIPK3-MLKL axis to induce age-related changes in HSCs. The necroptosis effector MLKL is readily activated by inflammation and replication stress and accumulates in HSC mitochondria. Consequently, activated MLKL does not cause cell death but impairs HSC self-renewal and lymphoid differentiation. Such MLKL-mediated functional decline also occurs in HSCs during organismal aging, with activated MLKL primarily mediating age-related mitochondrial damage and reduced glycolytic flux. Collectively, our results establish the RIPK3-MLKL axis as a key mediator of HSC aging and identify a necroptosis-independent role of MLKL in mitochondrial damage.

Introduction
Aging deteriorates hematopoiesis and increases one’s susceptibility to hematologic disorders. Although lifelong production of the blood and immune cells relies on the self-renewing ability and multipotency of hematopoietic stem cells (HSCs)1, compelling evidence suggests that HSCs with reduced regenerative and lymphopoietic potential accumulate with age2. Such age-related HSC dysfunction is associated with deleterious changes in their organelles, including accumulation of DNA damage, altered epigenetic regulation, reduced autophagic flux, and impaired mitochondria3–7, as well as environmental changes such as inflammation and alteration of the HSC-supporting niche cells in the bone marrow (BM)8–10. We and others recently showed that restoration of the young niche is insufficient to rejuvenate aged HSC function, highlighting a key role for age-associated cell-intrinsic defects in HSC aging11,12. Indeed, recent studies suggest that altered mitochondrial function and metabolism cause functional impairment of aged HSCs7,13. Moreover, repeated exposure to inflammatory and replication stress causes an irreversible and age-related deficit in HSC function14,15. However, a causal relationship between the cell-intrinsic and cell-extrinsic changes remains undefined.

HSCs are molecularly wired to prevent apoptosis unless otherwise required16–18, and their apoptosis resistance is further enhanced through age-dependent changes in the environment19. We and others previously showed that necroptosis, a form of programmed necrosis that is mediated by the receptor-interacting protein kinase 3 (RIPK3)–mixed lineage kinase like (MLKL) axis and typically occurs in apoptosis-resistant cells, can occur in HSCs and limit their regenerative potential after exposure to proinflammatory ligands and myeloablative stress20–23. Indeed, HSCs upregulate the necroptosis effector MLKL via post transcriptional modification upon inflammation24, and recent studies suggest a critical role for the RIPK3–MLKL axis in mediating HSC dysfunction after certain stresses and during aging25–28. Although these studies attributed RIPK3–MLKL-mediated cell death to functional decline in aged HSCs, dysfunctional HSCs should survive and accumulate in the BM during aging29, leaving the mechanism whereby the RIPK3–MLKL axis mediates age-related HSC changes unclear. Intriguingly, emerging evidence suggests that MLKL activation does not always induce necroptotic cell death30–32 but rather can impair membrane-bound organelles, including the endosome, autophagosome, and mitochondria, presumably through disruption of membrane integrity31,33–35.

Here, we sought to clarify the role of the necroptosis pathway in age-related HSC functional decline. We show that the RIPK3–MLKL axis is readily activated in a subset of HSCs after age-related stresses and reduces HSC self-renewal and lymphopoietic potential without affecting HSC survival.

Results
MLKL is preferentially activated in HSCs upon inflammation
Previous studies show that MLKL protein is abundantly expressed in murine immature hematopoietic cells36 (Supplementary Fig. 1a, b) and the expression level is elevated in HSCs upon polyinosinic-polycytidylic acid (pIC)-induced inflammation24. To further assess the activation status of the necroptosis pathway in individual HSCs and other hematopoietic progenitors upon inflammation, we used SMART-Tg mice that ubiquitously express a Förster resonance energy transfer (FRET)-based biosensor that monitors MLKL activation and translocation to the cell membrane37. Consistent with our previous observation that necroptosis is selectively activated in HSCs but not in granulocyte/monocyte progenitors during pIC-induced inflammation22, the SMART signal was increased in HSCs and multipotent progenitors (MPPs) but not in myeloid or lymphoid progenitors upon inflammation induced by administration of pIC, lipopolysaccharide (LPS), or tumor necrosis factor α (TNF-α) (Fig. 1a, b and Supplementary Fig. 1c). This was unexpected, as our previous data indicate that pIC injections induce TNF-α- and nuclear factor κB (NF-κB)-dependent pro-survival signals that protect HSCs from necroptotic death22. The FRET signal was increased on day 1 independently of biological sex but disappeared on day 7 post-administration of the proinflammatory ligands (Supplementary Fig. 1d, e). Moreover, the acute FRET activation was significantly diminished in a RIPK3-deficient (Ripk3–/–) background (Fig. 1c and Supplementary Fig. 1f), suggesting that the necroptotic RIPK3–MLKL axis is transiently activated in HSCs in response to inflammation. We transplanted HSCs with high and low FRET signals after LPS treatment (FRET/CFPhi and FRET/CFPlo) and observed lower engraftment potential and a tendency toward reduced B-lymphopoietic potential of FRET/CFPhi HSCs compared to FRET/CFPlo HSCs (Fig. 1d–g and Supplementary Fig. 1g, h). Collectively, these results indicate that the necroptosis pathway is selectively and transiently activated in HSCs upon inflammation, and its activation status is closely associated with their functional decline.

A non-necroptotic role of MLKL impairs HSC function
The best-known function of MLKL is to induce necroptotic cell death by plasma membrane rupture. To study the effect of inflammation-induced MLKL activation on HSCs, we assessed changes in HSC numbers in pIC-treated MLKL-deficient (Mlkl–/–) mice. Unexpectedly, phosphatidylserine exposure, a hallmark of the loss of plasma membrane integrity observed prior to cell rupture in necroptosis and apoptosis, was unchanged between wild-type (WT) and Mlkl–/– HSCs, indicating that the effect of inflammation-activated MLKL on HSC survival is negligible (Fig. 2a, b). Accordingly, the absolute numbers of HSCs were comparable between pIC-treated WT and Mlkl–/– mice (Fig. 2c and Supplementary Fig. 2a). As expected, pIC-induced inflammation also caused HSCs to exit quiescence38 and shift from an endothelial protein C receptor-expressing (EPCR+) self-renewing state39 to a CD41+ myeloid-primed state40, but MLKL deficiency had no apparent impact on these changes (Supplementary Fig. 2b–d). However, upregulation of another age-related myeloid-biased HSC marker, neogenin-1 (NEO-1)41 was significantly attenuated by MLKL deficiency (Fig. 2d). To further assess whether activation of the RIPK3–MLKL axis affects age-related HSC subpopulations, we utilized MLKL-SA2 knock-in (MlklSA2) mice, where MLKL cannot be activated by RIPK3-dependent phosphorylation due to the two serine-to-alanine substitutions at codons 345 and 347 of the Mlkl locus42, and treated them with PBS or pIC. In addition to NEO-1, we also assessed surface expression of P-selectin and GPR183, which were reported to enrich myeloid-biased43 and less differentiating44 subpopulations in aged HSCs, respectively. The results revealed that inactivation of the RIPK3–MLKL axis significantly attenuated pIC-induced upregulation of NEO-1, and to a lesser extent GPR183, but not P-selectin, on HSCs (Fig. 2e and Supplementary Fig. 2e, f). As the total number of HSCs did not differ before and after acute pIC injection (Fig. 2c), this indicates that the RIPK3-MLKL axis promotes transition from NEO-1– lineage-balanced to NEO-1+ myeloid-biased states.

In line with the above MLKL-mediated phenotypic changes, HSC transplantation experiments revealed that whereas MLKL deficiency alone conferred a selective advantage to phosphate-buffered saline-treated control HSCs, MLKL deficiency significantly alleviated a pIC-induced reduction in engraftment and B-lymphopoietic potential (Fig. 2f, g and Supplementary Fig. 2g, h). Moreover, in vivo blockade of RIPK3 kinase activity prior to pIC injection via administration of UH15-3845 revealed a trend toward ameliorating pIC-induced decline in HSC regenerative capacity and B-lymphoid differentiation potential (Supplementary Fig. 3a–d), suggesting that RIPK3 acts upstream of MLKL to mediate HSC functional decline. Transplantation of NEO-1– and NEO-1+ HSCs isolated from pIC-treated WT and MlklSA2 mice further revealed a trend—despite the limited number of mice—toward improved engraftment of MlklSA2 NEO-1– HSCs, but not NEO-1+ HSCs, compared to WT counterparts (Supplementary Fig. 3e-i), indicating selective protection of NEO-1– lineage-balanced HSCs by blockade of the RIPK3–MLKL axis. This MLKL-mediated effect seems largely cell-intrinsic and independent of environmental necroptosis, as MLKL deficiency had a minor effect on pIC-induced BM inflammation (Supplementary Fig. 3j, k), and pIC-induced suppression of ex vivo EPCR+ HSC expansion was largely attributed to MLKL activity in hematopoietic cells (Supplementary Fig. 3l).

To further investigate the cell death-independent role of MLKL in HSC function, we performed HSC transplantation experiments using Mlkl–/– HSCs reconstituted with N-terminal FLAG-tagged MLKL (FLAG-MLKL), which cannot induce cell death but can oligomerize and translocate to the cell membrane32,46 (Fig. 2h and Supplementary Fig. 3m). Consistent with the cell-intrinsic, necroptosis-independent regulation of HSCs by MLKL, reconstitution of Mlkl–/– HSCs with FLAG-MLKL significantly reduced their engraftment potential (Fig. 2i–l and Supplementary Fig. 3n). These data collectively indicate that inflammation-induced MLKL activation does not cause HSC death but impairs HSC regenerative and lymphopoietic potential independently of MLKL’s necroptotic function.

MLKL mediates age-related HSC changes driven by replication and oncogenic stress
Replication stress is implicated as the driving force of HSC aging4,15. Interestingly, MLKL is also preferentially activated in HSCs and MPPs but not in myeloid or lymphoid progenitors upon administration of 5-fluorouracil (5-FU) (Fig. 3a, b and Supplementary Fig. 4a), which induces HSC proliferation and age-related alterations when serially injected15. Again, blockade of the RIPK3–MLKL axis did not change HSC death or their absolute numbers during 5-FU-induced myeloablation (Fig. 3c and Supplementary Fig. 4b–e). Thus, to assess the relevance of MLKL activation in replication-induced HSC aging, we injected WT and Mlkl–/– mice with serial rounds of 5-FU (Fig. 3d). Although hematopoietic recovery after 5-FU injections was comparable between WT and Mlkl–/– mice (Supplementary Fig. 4f), we observed marked attenuation of age-related hematopoietic changes such as myeloid-skewed hematopoiesis (Fig. 3d) and expansion of phenotypic HSCs in 5-FU-treated Mlkl–/– mice (Supplementary Fig. 4g, h). Moreover, HSC transplantation experiments revealed that MLKL deficiency alleviated the 5-FU-induced decrease in HSC engraftment and lymphopoietic potential (Fig. 3e, f and Supplementary Fig. 4i, j). During serial HSC transplantation, which also induces extensive HSC proliferation in vivo15, MLKL deficiency in HSCs attenuated a progressive decline in their regenerative potential and lymphoid cell production (Fig. 3g and Supplementary Fig. 5a–c).

As an increased propensity to myelodysplastic syndrome (MDS) is another hallmark of aging in the hematopoietic system47, we next subjected WT and Mlkl–/– HSCs to oncogenic stress by introducing a C-terminal truncation mutant of Runt-related transcription factor 1 (RUNX1), which mimics MDS in mice by causing myeloid-skewed differentiation and ineffective hematopoiesis48. Transplantation of mutant RUNX1–transduced WT or Mlkl–/– HSCs showed that MLKL deficiency in hematopoietic cells did not affect HSC engraftment but attenuated the development of lethal MDS (Fig. 3h and Supplementary Fig. 5d). Further analyses of the recipient mice revealed that MLKL deficiency had no detectable impact on mutant RUNX1-induced dysplasia in hematopoietic cells (Supplementary Fig. 5e) but ameliorated lethality mainly due to severe defects in erythropoiesis (Supplementary Fig. 5f). These data support the idea that age-related HSC changes driven by replication and oncogenic stress are mediated at least partly through cell-intrinsic activity of MLKL.

MLKL mediates HSC dysfunction during organismal aging
The above findings prompted us to investigate whether MLKL mediates the functional decline in HSCs during organismal aging. We analyzed SMART signals in 3- and 12-month-old WT and Ripk3–/– mice and found that the frequency of FRET/CFPhi HSCs increased with age in a RIPK3-dependent manner, particularly in male mice (Fig. 4a–c and Supplementary Fig. 6a). Furthermore, BM analyses of 3- and 18-month-old WT and Mlkl–/– mice showed that MLKL deficiency largely rescued myeloid-skewed hematopoiesis (Fig. 4d, e) and loss of BM common lymphoid progenitors (Lin–/c-Kitlo/Sca-1lo/Flk2+/IL-7Rα+) (Supplementary Fig. 6b, c). Although BM HSC numbers and cell death frequency were largely unaffected in aged Mlkl–/– mice (Supplementary Fig. 6b–d), the accumulation of γH2AX foci, a hallmark of HSC aging, was significantly alleviated in aged Mlkl–/– HSCs (Fig. 4f and Supplementary Fig. 6e), indicating that the aging process is attenuated in Mlkl–/– HSCs. Accordingly, transplantation experiments using HSCs of 3- and 18-month-old WT or Mlkl–/– mice revealed better regenerative and B/T-lymphopoietic potential of aged Mlkl–/– HSCs compared to aged WT HSCs (Fig. 4g, h and Supplementary Fig. 6f–h), further demonstrating that age-related HSC functional decline is mediated at least partly through an MLKL-dependent mechanism.

MLKL can promote inflammation by releasing damage-associated molecular patterns21. Thus, the observed attenuation in HSC aging could be due to the secondary effects of necroptosis-mediated inflammation. To assess this possibility, we examined the levels of various inflammatory cytokines in the BM of 3- and 18-month-old WT and Mlkl–/– mice. Luminex multiplex cytokine analysis showed no apparent changes in age-associated inflammatory cytokines and chemokines, including interleukin-1 (IL-1) and regulated upon activation normal T cell expressed and secreted (RANTES), which are implicated in myeloid skewing of HSCs49,50 (Fig. 5a and Supplementary Fig. 7a). In line with this, reciprocal transplantation of 3-month-old WT BM cells into 18-month-old WT and Mlkl–/– mice revealed no apparent impact of aged Mlkl–/– environment on myeloid differentiation of WT BM cells (Fig. 5b), suggesting that MLKL minimally affects age-associated BM inflammation.

To further assess the mechanism whereby MLKL mediates HSC aging, we analyzed age-related transcriptomic changes in WT and Mlkl–/– HSCs (Fig. 5c). Surprisingly, although we could detect differentially expressed genes in WT HSCs with age, including Selp, Mt1, and Nupr1 as the top three aging signature genes43 (Supplementary Fig. 7b, c and Supplementary Data 1–3), few genes were significantly altered between aged WT and Mlkl–/– HSCs (Fig. 5d–f and Supplementary Data 4). As epigenetic changes can influence HSC fate without transcriptomic changes51, we also assessed age-related changes in chromatin accessibility of WT and Mlkl–/– HSCs (Fig. 5g). Again, although many genomic loci showed altered accessibility in WT HSCs with age (Supplementary Fig. 7d, e and Supplementary Data 5–7), few loci were detected as differentially accessible between aged WT and Mlkl–/– HSCs (Fig. 5h–j and Supplementary Data 8), indicating that MLKL minimally affects the HSC transcriptome and chromatin accessibility during aging.

Together, these results indicate that MLKL mediates HSC dysfunction during organismal aging, likely through mechanisms independent of BM inflammation and transcriptional regulation.

MLKL mediates age-related mitochondrial damage in HSCs
Despite minimal changes in the transcriptome, gene set enrichment analyses identified several candidate pathways potentially altered in aged Mlkl–/– HSCs but not in young Mlkl–/– HSCs (Supplementary Fig. 7f, g). One such candidate was mitochondrial oxidative phosphorylation, which was upregulated in HSCs during aging and closely associated with functional decline in aged HSCs5. Indeed, transmission electron microscopy showed that elongated and swollen mitochondria with disorganized cristae, a hallmark of dysfunctional mitochondria observed in HSCs with a history of replication stress52, were enriched in aged WT but not in aged Mlkl–/– HSCs (Fig. 6a, b). Consistent with this observation, immune electron microscopy analysis and proximity ligation assays independently revealed an accumulation of active, phosphorylated MLKL at serine 345 (p-MLKL) in the mitochondria of aged HSCs (Fig. 6c, d). Notably, inflammatory exposure and FLAG-MLKL overexpression also induced accumulation of p-MLKL in HSC mitochondria (Supplementary Fig. 8a, b). As previously reported, the level of reactive oxygen species (ROS) in HSCs was reduced with age5, but no changes in ROS levels were observed between aged WT and Mlkl–/– HSCs (Supplementary Fig. 8c). Autophagosome frequency and starvation-induced autophagy flux were also similar between aged WT and Mlkl–/– HSCs (Supplementary Fig. 8d and e), suggesting that MLKL minimally affects autophagy in aged HSCs. However, the age-related decrease in mitochondrial membrane potential was significantly alleviated in aged Mlkl–/– HSCs (Fig. 6e). Seahorse analyses showed that these changes were associated with improved mitochondrial ATP production and glycolytic flux in aged Mlkl–/– HSCs compared to aged WT HSCs (Fig. 6f, g and Supplementary Fig. 8f). Although the rarity of HSCs in young animals precluded us from directly comparing mitochondrial metabolism between young and aged Mlkl–/– HSCs, this could reflect the attenuation of age-related metabolic rewiring from glycolysis to oxidative phosphorylation5. In vitro experiments using isolated mitochondria and recombinant MLKL protein revealed that mitochondrial membrane potential was decreased by N-terminal four-helix bundle domain of recombinant mouse MLKL protein (MLKL-NTD) that oligomerizes and forms pores on the cell membrane without stimulation, and to a lesser extent by N-terminal FLAG-tagged MLKL-NTD (FLAG-MLKL-NTD) that retains oligomerization capacity but cannot perform pore formation on the cell membrane, but not by C-terminal pseudokinase domain of recombinant mouse MLKL protein (MLKL-CTD) that does not oligomerize or form pores on the cell membrane53 (Fig. 6h and Supplementary Fig. 8g), indicating that activated MLKL directly impairs mitochondrial membrane potential via mechanisms both dependent on and independent of its pore-forming capacity. Together, these results identify MLKL as a key mediator of age-related mitochondrial dysfunction and metabolic rewiring in HSCs.

Methods
Ethics
All animal experiments comply with all relevant ethical regulations and were performed in accordance with protocols approved by the Animal Care and Use Committee at the Institute of Medical Science, University of Tokyo (protocol number: PA19-07) and the St. Jude Children’s Research Hospital Institutional Animal Care and Use Committee (protocol number: 3252).

Reagents and resources
Details of key reagents and resources used in this study are listed in Supplementary Table 1.

Mice
WT CD45.2 and CD45.1 C57BL/6 mice were purchased from Japan SLC and Sankyo Labo Services, respectively. Mlkl–/– mice were obtained from Dr. M. Pasparakis (University of Cologne)68, and WT and Ripk3–/– SMART-Tg mice were obtained from Dr. H. Nakano (Toho University)37. MlklSA2 mice were purchased from the Jackson Laboratory42. WT and Mlkl–/– mice were bred and aged in-house at the Institute of Medical Science, University of Tokyo, WT and Ripk3–/– SMART-Tg mice at Toho University, and WT, SMART-Tg, Mlkl–/–, and MlklSA2 mice at St. Jude Children’s Research Hospital. At the time of analysis, young mice were 6–12 weeks of age, and aged mice were 18–20 months of age unless otherwise specified. For transplantation experiments, 8–12-week–old CD45.1 C57BL/6 mice were used as recipients. Respective littermates or age-matched mice were used as controls. No specific randomization or blinding protocol was used, and both male and female animals were used indiscriminately in this study. All mice were bred and maintained on a 12 h light cycle in temperature- and humidity-controlled specific pathogen-free mouse facilities.

In vivo assays
For pIC-induced inflammation, mice were injected intraperitoneally with 5 mg/kg pIC (Cytiva) in phosphate-buffered saline (PBS) once or 7 times every other day. For LPS-induced inflammation, mice were injected intraperitoneally with 0.4 mg/kg LPS (InvivoGen) in PBS once. For TNF-α-induced inflammation, mice were injected retro-orbitally with 2 µg TNF-α (Genentech) in PBS once. For repeated 5-FU treatment, mice were injected intraperitoneally with 150 mg/kg 5-FU (Kyowa Kirin) in PBS once a month for 3 months (3 times). For in vivo blockade of RIPK3 kinase activity, mice were injected intraperitoneally with 30 mg/kg UH15-38 (Bio-Techne) in PBS once every day for 4 consecutive days before pIC injection. For HSC transplantation, recipient mice (CD45.1) were lethally irradiated (9.5 Gy, delivered in split doses 3 h apart) using an X-ray irradiator (MBR-1520R; Hitachi) and retro-orbitally injected with 250 donor HSCs (CD45.2+) along with 2 × 105 BM cells (CD45.1+) within the next 6 h. When necessary, 2 × 106 BM cells were pooled from each recipient at 4 months and retro-orbitally injected into freshly prepared, lethally irradiated secondary and tertiary recipients. For BM chimeras, recipient mice were lethally irradiated and retro-orbitally injected with 2 × 106 BM cells isolated from WT and Mlkl–/– mice. For reciprocal transplantation, 18-month-old WT and Mlkl–/– recipient mice were lethally irradiated and retro-orbitally injected with 2 × 106 BM cells isolated from 3-month-old WT mice. For transplantation of WT and Mlkl–/– HSCs after RUNX1S291fs transduction, lethally irradiated recipients were retro-orbitally injected with 1000 GFP+ transduced HSCs along with 5 × 105 Sca-1-depleted helper BM cells (CD45.1+). For transplantation of Mlkl–/– HSCs reconstituted with N-terminal FLAG-MLKL, lethally irradiated recipients were retro-orbitally injected with 1000 GFP+ transduced HSCs along with 2 × 105 BM cells (CD45.1+). Transplanted mice received antibiotic water containing Baytril (Bayer) for 4 weeks. Peripheral blood (PB) was analyzed monthly and collected via retro-orbital bleeding in 4 mL of ACK lysis buffer (150 mM NH4Cl and 10 mM KHCO3) containing 10 mM EDTA for flow cytometry analyses, and BM was analyzed at 4 months post-transplantation.

Flow cytometry
Single-cell suspensions of BM cells were obtained in PBS containing 2% heat-inactivated fetal bovine serum (FBS) (Sigma-Aldrich) either by crushing femurs, tibiae, pelvises, humeri, and the sternum or by flushing femurs and tibiae. Erythrocytes were removed by ACK lysis, and contaminating bone fragments were further removed by centrifugation on a Ficoll gradient (Histopaque-1119; Sigma-Aldrich). BM cellularity was determined by using a Vi-CELL-XR or Vi-CELL-BLU automated cell counter (Beckman-Coulter). For immature cell sorting, BM cells were pre-enriched for c-Kit+ cells with c-Kit microbeads and LS columns (Miltenyi Biotec). For immature cell analyses and sorting, a lineage (Lin) cocktail of CD3ε-PECy5 (1:200), CD4-PECy5 (1:800), CD5-PECy5 (1:800), CD8α-PECy5 (1:800), B220-PECy5 (1:800), Mac-1-PECy5 (1:800), Gr-1-PECy5 (1:800), and Ter119-PECy5 (1:400) was used to define immature BM cells. For immature cell sorting, c-Kit-enriched BM cells were stained with Lin-PECy5, c-Kit-APC (1:800), Sca-1-PECy7 (1:800), Flk2-BV421 (1:50), CD48-APCeF780 (1:400), and CD150-PE (1:400). For immature cell analyses, unfractionated BM cells were stained with Lin-PECy5, c-Kit-APC, Sca-1-PECy7, Flk2-Bio (1:100)/SA-BV605 (1:400), CD48-AF700 (1:400), CD150-BV650 (1:200), CD34-FITC (1:50), FcγR-BV510 (1:800), IL-7Rα-APCCy7 (1:100), CD41-BV421 (1:400), and EPCR-PE (1:800). For NEO-1 expression, cells were incubated with anti-FcγR (1 µg per 106 cells) for 10 min and stained with Lin-PECy5, c-Kit-APC, Sca-1-PECy7, CD48-APCeF780, CD150-PE and anti-NEO-1 (1:13)/anti-goat IgG-AF488 (1:200). For P-selectin and GPR183 expression, cells were stained with Lin-PECy5, c-Kit-APC, Sca-1-PECy7, CD48-APCeF780 together with either CD150-BV650 and P-selectin-PE (1:200) or CD150-PE and GPR183-FITC (1:50). For PB analyses, cells were stained with Ter119-PECy5, Gr-1-eF450 (1:800), Mac-1-APC (1:800), B220-BV605 or -AF700 (1:800), and CD3ε-PECy7 (1:200). For analyses of FRET in immature SMART-Tg cells, unfractionated BM cells were stained with Lin-PECy5, c-Kit-APC, Sca-1-PECy7, CD48-APCeF780, and CD150-PE. For transplantation experiments, PB cells were stained with Ter119-PECy5, Gr-1-eF450, Mac-1-APC, B220-BV605, CD3ε-PECy7, CD45.1-APCeF780 (1:200), and CD45.2-FITC or -BV786 (1:200), and BM cells were stained with Lin-PECy5, c-Kit-APC, Sca-1-PECy7, Flk2-Bio/SA-BV605, CD48-AF700, CD150-BV650, IL-7Rα-PE, CD45.1-APCeF780, and CD45.2-FITC or -BV786. For ex vivo expanded HSC analyses, cells were stained with CD41-FITC (1:800), Mac-1-Bio (1:800)/SA-BV605, FcγR-BV510, Sca-1-PECy7, c-Kit-APC, CD150-BV650, CD48-AF700, and EPCR-PE. For staining with CellROX, unfractionated BM cells were stained with Lin-PECy5, c-Kit-APC, Sca-1-PECy7, Flk2-BV421, CD48-AF700, and CD150-BV650 and incubated with 5 µM CellROX Green in 2% FBS/PBS for 30 min at 37°C, 5% CO2. For staining with JC-1, unfractionated BM cells were stained with Lin-PECy5, c-Kit-APC, Sca-1-PECy7, Flk2-BV421, CD48-APCeF780, CD150-BV650 and incubated for 30 min at 37°C, 5% CO2 with 2 µM JC-1 in Iscove’s modified Dulbecco’s media (IMDM) supplemented with 5% FBS, 1× penicillin–streptomycin– L-glutamine (Fujifilm Wako), 0.1 mM non-essential amino acids (Gibco), 1 mM sodium pyruvate (Gibco), and 50 μM 2-mercaptoethanol (Sigma-Aldrich). All antibody staining was performed on ice for 45 min when a CD34 antibody was included or 30 min otherwise, and stained cells were finally resuspended in 2% FBS/PBS containing 1 µg/ml propidium iodide (PI) to exclude dead cells. For Annexin V staining, unfractionated BM cells stained with Lin-PECy5, c-Kit-APC, Sca-1-PECy7, CD48-APCeF780, CD150-PE, or retrovirally transduced GFP+ cells were washed in 1× Binding buffer and incubated with Annexin V-FITC or -APC (1:20) in 1× Binding buffer for 15 min at room temperature. Stained cells were finally resuspended in 1× Binding buffer containing DAPI (1 µg/ml), SYTOX Blue, or PI (1 µg/ml) before analysis. For intracellular Ki-67/DAPI staining, unfractionated BM cells were stained with Lin-PECy5, c-Kit-APC, Sca-1-PECy7, CD48-APCeF780, and CD150-PE, and then fixed and permeabilized with Cytofix/Cytoperm buffer for 20 min on ice. After washing with Perm/Wash (BD Biosciences), cells were stained with anti-Ki-67-FITC (1:100) in Perm/Wash for 30 min on ice, washed with Perm/Wash and then resuspended in Perm/Wash containing 1 µg/ml DAPI before analysis. Cell sorting was performed on FACSAria IIIu or Fusion (Becton Dickinson). All data were collected on FACS Aria IIIu, Celesta, or LSRFortessa (Becton Dickinson) and analyzed with FlowJo (BD Biosciences, v10.10.0). For detection of FRET from ECFP to Ypet in the SMART probe, ECFP and FRET signals excited by a 405 mm laser were detected with 450/40 and 530/30 filters, respectively, on a FACSAria IIIu or LSRFortessa. Cells with an increase in FRET/CFP ratio and a decrease in CFP signals were gated as FRET/CFPhi.

Ex vivo assays
For ex vivo HSC expansion, 500 HSCs were directly sorted per well of a 96-well flat-bottom plate in 200 µL of Ham’s F-12 media (Gibco) supplemented with 0.1% polyvinyl alcohol, 10 mM HEPES (Gibco), 1× penicillin–streptomycin– L-glutamine, 1× insulin–transferrin–selenium–ethanolamine (Gibco), 10 ng/mL mouse SCF (BioLegend), and 100 ng/mL mouse TPO (PeproTech) as described previously69. After 7 days, cells were collected and counted on Vi-CELL XR, and the frequency of EPCR+ HSCs (CD41–/Mac-1–/FcγR–/c-Kit+/Sca-1+/CD48–/CD150+/EPCR+) was evaluated by flow cytometry. For autophagy flux analyses, HSCs were sorted and incubated with DALGreen (1 µM) for 30 min at 37°C in 2% FBS/PBS. After being washed twice with 2% FBS/PBS, 1000–2000 stained cells were seeded per well of 96-well flat bottom plate, incubated for 8 h at 37°C, 5% CO2 in 200 µL of cytokine-free IMDM supplemented with 5% FBS, 1× penicillin–streptomycin– L-glutamine, 0.1 mM non-essential amino acids, 1 mM sodium pyruvate, and 50 μM 2-mercaptoethanol with or without 5 nM bafilomycin A1, and analyzed by flow cytometry.

Retroviral transduction of HSCs
For transduction of N-terminal FLAG-MLKL, 3×FLAG was added in frame at the N-terminus of mouse Mlkl cDNA and cloned into the pMY-IRES-GFP retroviral vector. The pMY-IRES-GFP and pMY-3×FLAG-MLKL-IRES-GFP were transfected into Plat-E packaging cells with FuGENE HD, and the culture supernatants were obtained at 40–48 h and filtered through 0.45 µm PVDF filters. For transduction of the C-terminal truncation mutant of RUNX1, the culture supernatant of 293GPG packaging cells stably transfected with the pMY-RUNX1S291fs-IRES-GFP retroviral vector was collected at 48–60 h after tetracycline withdrawal and filtered through a 0.45 µm PVDF filter. For HSC transduction, 2000 WT and Mlkl–/– HSCs (CD45.2+) were directly sorted per well of a 96-well flat bottom plate pre-coated with RetroNectin (Takara Bio) in 200 µL SF-O3 media (EIDIA) supplemented with 0.2% BSA, 100 ng/mL mouse SCF, and 100 ng/mL human TPO, incubated overnight at 37°C, 5% CO2, and subjected to magnetofection for 1 h in the retroviral supernatants with a 96-well magnetic plate and ViroMag R/L. The medium was then replaced with fresh cytokine-containing media, and cells were incubated for 48 h before the re-isolation of live PI–/GFP+ transduced cells for transplantation experiments.

Immunofluorescence and proximity ligation assay
A total of 2000 HSCs were sorted and pipetted onto MAS adhesive glass slides (Matsunami TF1205M) or µ-Slide 18 Well (ibidi), incubated for 30 min at 4°C, fixed with 4% paraformaldehyde for 10 min at room temperature, and permeabilized in 0.3% Triton X-100/PBS for 2 min at room temperature. For γH2AX immunofluorescence, slides were blocked in 2% BSA/PBS for 1 h at 4°C and incubated overnight at 4°C in 2% BSA/PBS with anti-γH2AX antibody (1:500). Slides were then washed 3 times in PBS and incubated for 1 h at room temperature in 2% BSA/PBS with AF488-conjugated goat anti-mouse IgG (1:200). For FLAG-MLKL immunofluorescence, slides were blocked in 2% BSA/PBS for 1 h at 4°C and incubated overnight at 4°C in 2% BSA/PBS with anti-MLKL (1:500) and anti-FLAG (1:500) antibody. Slides were then washed 3 times in PBS and incubated for 1 h at room temperature in 2% BSA/PBS with AF647-conjugated goat anti-mouse IgG (1:500) and AF594-conjugated goat anti-rabbit IgG (1:500). Slides were then washed 3 times in PBS, incubated with 1 mg/ml DAPI/PBS for 5 min, and washed twice in PBS. Slides were finally mounted with coverslips by using ProLong Glass or Gold (Invitrogen). Cells were imaged on an A1Rsi inverted confocal microscope with a ×100 objective (Nikon) or an LSM 980 Airyscan inverted confocal microscope with a ×63 objective (Zeiss). For the proximity ligation assay, slides were blocked in Duolink blocking solution (Sigma-Aldrich) and incubated overnight at 4°C in Duolink antibody diluent (Sigma-Aldrich) with primary antibodies against COX-IV (1:400) and p-MLKL (1:1600). Slides were washed twice in 1× Duolink wash buffer A (Sigma-Aldrich) and incubated for 1 h at 37°C in Duolink antibody diluent with PLUS and MINUS probes (1:5; Sigma-Aldrich). Slides were then washed twice in 1× Duolink wash buffer A, incubated for 30 min at 37°C in 1× Duolink ligation buffer with ligase (1:40, Sigma-Aldrich), washed twice in 1× Duolink wash buffer A, and incubated for 100 min at 37°C in 1× Duolink amplification buffer with polymerase (1:80; Sigma-Aldrich). After being washed twice in 1× and once in 0.01× Duolink wash buffer B (Sigma-Aldrich), slides were finally mounted with coverslips by using Duolink in situ mounting medium with DAPI (1:80; Sigma-Aldrich), and cells were imaged on an N-SIM super-resolution confocal microscope with a 100× objective (Nikon) or an LSM 980 Airyscan inverted confocal microscope with a ×63 objective (Zeiss). For quantification, cells were randomly captured, and γH2AX or PLA foci were counted by eye using an NIS-Elements Viewer (Nikon, v4.11.0) or ImageJ (v1.53c).

Cytokine profiling
For collecting BM fluids, two femurs and two tibiae per mouse were flushed out with the same 200 µL of 2% FBS/PBS by using a 1 mL syringe with a 23 G needle and were spun down at 300×g for 5 min to remove BM cells. Supernatants were further clarified by centrifugation at 12,000×g for 10 min and stored at –80°C until use. 50 µL of 2× diluted samples were analyzed with a Bio-Plex Pro mouse cytokine 23-Plex panel on a Bio-Plex 200 analyzer (Bio-Rad) according to the manufacturer’s protocol. Cytokine concentrations were calculated using standard curves. Morpheus was used to generate heatmaps showing normalized cytokine levels with hierarchical clustering of samples and cytokines with Euclidean distance.

RNA-seq
RNA was purified from 10,000 HSCs isolated from WT and Mlkl–/– mice with the RNeasy Plus Micro Kit. Double-stranded cDNA was generated using the SMART-Seq HT Kit and fragmented using the M220 Focused ultrasonicator (Covaris). Sequencing libraries were prepared using the NEBNext Ultra DNA Library Prep Kit (New England Biolabs). Different index primers were used for multiplexing samples in one lane, and pooled libraries were sequenced on NextSeq2000 (Illumina) with single-read 70 base pairs. Data quality was verified by FastQC (v0.12.0), and demultiplexing was performed with bcl2fastq (v2.20). Sequencing reads were mapped to the mouse reference genome (mm10) with HISAT2 (v2.2.1) and quantified with StringTie (v2.2.3). Normalization and pairwise differential expression analyses were performed using edgeR (v3.30.3). Principal component analysis was performed using standard packages in R (v4.0.2), and plots were generated using the first 2 principal components. The Benjamini–Hochberg method was used to correct type I errors for multiple gene-level comparisons, with Wald tests applied and a q-value cutoff of 0.01 set to define differentially expressed genes (DEGs). Identified DEGs were visualized in volcano plots and area-proportional Venn diagrams using R. Gene set enrichment analysis (v4.3.0) was performed using the 50 “MH” orthology-mapped hallmark gene sets in mouse MSigDB collections, and only those with P < 0.05 were displayed as significantly enriched gene sets.

ATAC-seq
For each biological replicate, 10,000 HSCs were isolated from six young WT mice, six young Mlkl–/– mice, three aged WT mice, and three aged Mlkl–/– mice. All HSC replicates were prepared and processed on the same day to avoid potential batch effects. At harvest, cells were lysed and immediately transposed as described previously. In brief, HSCs were resuspended in cold lysis buffer (10 mM Tris–HCl, pH 7.4, 10 mM NaCl, 3 mM MgCl2, and 0.1% IGEPAL CA-630) and incubated for 10 min on ice. Samples were then spun down at 600×g for 10 min at 4°C and incubated for 35 min at 37°C in 50 μL of transposase reaction mix consisting of 25 μL Tagment DNA buffer (Illumina), 2.5 μL Tagment DNA enzyme (Illumina), and 22.5 μL nuclease-free water. Transposed fragments were purified with MinElute PCR Purification Kit (QIAGEN), eluted in 14 µL of nuclease-free water, and stored at –20°C until library preparation. After the optimization of PCR cycle number, libraries were semi-quantified with SYBER Green I Nucleic Acid gel Stain (Takara Bio) to optimize the number of PCR cycles and then amplified using NEBNext High Fidelity 2× PCR Master mix (New England Biolabs) and index primers. Amplified libraries were purified with the MinElute PCR Purification Kit, size-selected between 240 and 360 base pairs by using Blue Pippin (Sage Science), and sequenced with NextSeq 500 (Illumina) with 70 bp single-read length. Sequencing reads were mapped to the mouse reference genome (mm10) with Bowtie 2 (v2.5.3). Peaks were called with MACS2 (v2.2.7.1) using the nomodel function, with a q-value cutoff of 0.001 set to define accessible peaks in each sample. The catalog of all peaks called in any samples was generated by merging all called peaks that overlapped by at least one base pair by using the bedtools (v2.31.0) merge function. As a result, a total of 76,286 merged peaks were detected and used as a map file for downstream processing. Reads at each peak in the catalog were quantified with the bedtools map function by using bed files of each sample. Differentially accessible regions (DARs) were detected using edgeR with a q-value cutoff of 0.01 and visualized as volcano plots and area-proportional Venn diagrams using R.

Transmission electron microscopy
A total of 100,000 HSCs were sorted in 2% FBS/PBS, spun down at 300×g for 5 min at 4°C, and fixed for 2 h at room temperature with 1% glutaraldehyde in 0.1 M sodium phosphate buffer pH 7.4. After fixation, cells were rinsed and post-fixed on ice for 2 h with 2% osmium tetroxide in 0.1 M sodium phosphate buffer, pH 7.4. Cells were then washed, dehydrated in a graded series of ethanol, and embedded in Epon 812 resin mixture (TAAB). Semi-thin sections of about 0.7 µm thickness were cut on an EM UC7 ultramicrotome (Leica), stained with 0.2% toluidine blue, and examined under an ECLIPSE Si microscope (Nikon). Ultra-thin sections were cut, stained for 5 min at room temperature with uranyl acetate and lead citrate, and imaged on a JEM-1400Flash electron microscope (JEOL). Mitochondria and autophagosomes were counted by eye, and mitochondrial morphology was analyzed with ImageJ.

Immunoelectron microscopy
A total of 100,000 HSCs were sorted in 2% FBS/PBS, spun down at 300×g for 5 min at 4°C, and fixed for 30 min on ice with 4% paraformaldehyde and 0.2% glutaraldehyde in 0.1 M sodium phosphate buffer pH 7.4. After fixation, cells were rinsed, dehydrated in a graded series of ethanol, and embedded in LR white medium-grade resin (Electron Microscopy Sciences). Semi-thin sections of about 0.7 µm thickness were cut on an EM UC7 ultramicrotome, stained with 0.2% toluidine blue, and examined under an ECLIPSE Si microscope. Ultra-thin sections were cut and blocked with 1% BSA/PBS for 10 min at room temperature. The sections were then incubated for 1 h at room temperature in 1% BSA/PBS with a primary antibody against p-MLKL (1.0 µg/mL). Sections were washed 3 times in PBS and incubated for 1 h at room temperature in 1% BSA/PBS with 5 nm gold-conjugated secondary antibody against rabbit IgG (1×). Sections were washed 3 times in water, post-fixed in 1% glutaraldehyde for 10 min, and stained with 1% uranyl acetate for 3 min at room temperature. Cells were imaged on a JEM-1400Flash electron microscope (JEOL).

Recombinant protein expression and purification
N-terminal four-helix bundle domain of recombinant mouse MLKL protein (residues 1–169; MLKL-NTD), N-terminal FLAG-tagged MLKL-NTD (FLAG-MLKL-NTD), and C-terminal pseudokinase domain of recombinant mouse MLKL protein (residues 179–464; MLKL-CTD) were expressed and purified from BL21 (DE3)-RIPL Escherichia coli strains. Briefly, cells were grown in 2XTY medium (20 g/L tryptone, 10 g/L yeast extract, 5 g/L NaCl, 50 µg/ml kanamycin, 30 µg/ml chloramphenicol) to an optical density at 600 nm of 0.8 and induced with 0.5 mM isopropylthio-β-galactoside for 16 h at 22°C. Pellets were resuspended in buffer (25 mM Tris–HCl pH 7.4, 500 mM NaCl, 1 mM TCEP, 10% glycerol, 0.05% Triton-X 100, 25 mM imidazole), lysed at high pressure, clarified by centrifuge, subjected to Ni2+ column, washed and eluted with buffer (25 mM Tris–HCl pH 7.4, 500 mM NaCl, 1 mM TCEP, 10% glycerol, 0.05% Triton-X 100, 250 mM imidazole). Elution was collected and incubated with TEV protease at 4°C overnight to cleave off the His tag, followed by a reverse Ni2+ column to remove the TEV protease and any undigested His tag protein. The digested proteins were further polished by Superdex S200 gel filtration column pre-equilibrated with gel filtration buffer (20 mM Tris pH 7.4, 200 mM NaCl, 1 mM TCEP, 1 mM EDTA). Selected fractions were pooled and concentrated by centrifugal ultrafiltration to 1 mg/ml and dialyzed in the buffer (10 mM Tris–HCl pH 7.5, 10 mM KCl, 250 mM sucrose, 1.5 mM MgCl₂).

Isolation and treatment of mitochondria
Mitochondria were isolated from mouse liver tissue using the Mitochondria Isolation Kit for Cultured Cells (Abcam) according to the manufacturer’s protocol. Briefly, ~50 mg of liver tissue was dissected, immediately transferred into 500 µL of Reagent A pre-dispensed in the Dounce homogenizer tube, and homogenized by 30 strokes with pestle A. Homogenate was centrifuged at 1000×g for 10 min at room temperature, and the supernatant (SN1) was collected. The pellet was then resuspended in 500 µL of Reagent B, homogenized by pipetting up and down 30 times using a P1000 micropipette, and the supernatant was collected (SN2) after centrifugation at 1000×g for 10 min at room temperature. SN1 and SN2 were combined, and mitochondrial fractions were pelleted by centrifugation at 12,000×g for 15 min at room temperature. The mitochondrial pellet was resuspended in 200 µL of mitochondrial buffer (10 mM Tris–HCl pH 7.5, 10 mM KCl, 250 mM sucrose, 1.5 mM MgCl₂), and 2.5 µL of mitochondria were incubated in 87.5 µL of mitochondrial buffer for 45 min at 37°C with 20 µM recombinant MLKL-NTD, 20 µM recombinant FLAG-MLKL-NTD, 20 µM recombinant MLKL-CTD, or 50 µM CCCP (Thermo Fisher). Treated mitochondria were stained in mitochondrial buffer containing 500 nM TMRM (Invitrogen) and 200 nM MitoTracker Green (Thermo Fisher) for 30 min at 37°C before flow cytometry analysis. All procedures were performed without vortexing to avoid mechanical disruption of mitochondria and preserve mitochondrial membrane potential.

Seahorse assays
A total of 50,000–100,000 HSCs were sorted in SF-O3 and spun down at 300×g for 5 min at 4°C. Cells were resuspended in Seahorse XF basic DMEM (Agilent) supplemented with 10 mM glucose, 1 mM pyruvate, and 2 mM L-glutamine, seeded in a cell-culture plate pre-coated with Cell-Tak (Corning) at a density of 50,000–70,000 cells per well, and spun down by centrifugation at 400×g for 5 min at room temperature. Oxygen consumption rate and extracellular acidification rate were measured at baseline and after sequential injections of 1 μM oligomycin, 2 μM FCCP, 0.5 μM rotenone plus 0.5 μM antimycin, and 50 mM 2-deoxy-D-glucose on a Seahorse XFe96 Extracellular Flux Analyzer (Agilent) according to the manufacturer’s instructions as described previously70. Data were normalized to seeded cell numbers, analyzed with Seahorse Analytics (v1.0.0-520), and exported to GraphPad Prism (v10.4.0).

Quantification and statistical analysis
Data are represented as mean ± standard error (s.e.m.) unless otherwise specified in the figure legends. Statistical analysis was performed using R for RNA-seq and ATAC-seq data and Prism for all the other data. The statistical test used, the number of biological replicates, and the definition of biological replicates are indicated in the figure legends. Briefly, two-tailed Student’s t-test and Mann–Whitney U-test were used when two groups were compared, and one-way ANOVA, two-way ANOVA, Welch and Brown–Forsythe test, and Kruskal–Wallis test were used when three or more groups were compared. For multiple comparisons, p-values were adjusted either by Šídák correction, Fisher’s least significant difference (LSD) test, or the false discovery rate of 5% based on the two-stage linear step-up procedure of Benjamini, Krieger, and Yekutieli, and adjusted p-values were subsequently used to indicate significance. For transplantation experiments, outliers were excluded before statistical analyses based on the PB donor chimerism at 4 months post-transplantation by using the ROUT method with a Q value set to 1%. Recipient mice with <1% PB donor chimerism at 4 months post-transplantation were excluded from the analyses of donor lineage distribution. Survival rates were compared using the Mantel–Cox log-rank test. Statistical significance is reported using exact P values.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In their study, Yamada et al. investigated the role of the necroptosis factor MLKL in HSC aging. Using SMART-Tg mice they demonstrate an increase in MLKL, measured through FRET, upon treatment with inflammation-inducing agents (poly-IC, LPS and TNF-alpha), as well as following replication stress and in aged mice. This increase in MLKL was correlated with poorer HSC reconstitution, which was also observed in aged mice. Utilizing an MLKL knockout strain, the authors showed that MLKL deficiency partially rescued the phenotype of poorer HSC reconstitution and other aging-associated features. Furthermore, Yamada et al. discovered that activated MLKL accumulated in mitochondria leading to mitochondrial dysfunction in aged HSCs, which could partially be corrected by MLKL deficiency.\nThe study is very comprehensive and well-executed, presenting intriguing findings about the potential role of MLKL beyond necroptosis. However, my concern is that the cell death-independent role of MLKL is based solely on Figure 2B, which includes only four mice from two experiments, showing quite some variation, especially in the WT pIC group. I recommend including more replicates in Figure 2B, and incorporating cell death analysis as validation in subsequent figures, such as in the Figure 2F experiment. Another concern is that the authors have not conclusively shown that MLKL underlies aging, as stated in the title and throughout the manuscript. While it is evident that MLKL contributes to or even promotes aging, I disagree to the statement that MLKL is the driver of aging considering that the authors only demonstrate partial restoration when MLKL is knocked out (e.g. Figure 4G, Figure 6F).\nFurther points for clarification:\n1. Line 88 states that necroptosis is selectively and transiently activated in HSCs. However, the data clearly show activation in both HSCs and multipotent progenitors upon treatment with pIC, LPS and TNFa (Figure 1B). Was lymphoid progenitors analyzed as well?\n2. In Figure 1D, FRET high and low cells were sorted and transplanted. It would be informative to show the sorting gates for FRET high and low. Does FRET low indicate “negative” cells or truly low cells? Would the truly FRET negative cell not be a better control than PBS control?\n3. In transplantation experiments it would be informative to see the total reconstituted levels of each blood lineage (i.e. out of total blood cells) rather than only relative reconstitution frequencies within the donor-derived compartment. This is especially relevant if total donor reconstitution levels vary a lot between recipient mice.\n4. Could the absence of significant transcriptional differences between WT and MLKL-/- be due to the bulk RNA-seq mode, which might omit differences that would otherwise be detected with single cell RNA-seq?\n5. Why were young mice not included in Figure 6E, G, H for comparison?\n6. Do MLKL-/- mice live longer?\n7. Have you checked for any differences between females and males in inflammation-induced MLKL activation and in aging?", "reviewer_2": "The work by Yamada et al. investigates a cell death-independent role of MLKL in HSC inflammation and aging, which does not regulate the transcriptome; instead, it impairs mitochondrial fitness. This work is well-conducted and controlled; the phenotype is robust; and the scientific rigor is high. One of several experiments I appreciate a lot is the N-flag MLKL rescue one. As N-flag MLKL cannot induce necroptosis but can still rescue the MLKL KO HSC phenotype, I believe the \"cell-death-independent role\" of MLKL in HSC is very solid. Although the overall quality of this work is high, some minor concerns can be addressed to enhance its excellence further.", "reviewer_3": "The manuscript by Yamada et al. describes the role of Mlkl in hematopoietic stem cells (HSC). Authors showed that Mlkl is activated in HSC and multipotent progenitor (MPP) pool upon inflammation using FRET based sensor that monitor MLKL translocation to membrane. However, they showed that this activation does not lead to cell death of HSCs, as expected by the known function of Mlkl in necroptosis. Next, authors performed series of experiments on Mlkl-/- mice, and demonstrated that such mice do not show many established hallmarks of HSC aging or strong inflammation-driven exhaustion of HSC function. Finally, the authors showed the link of Mlkl with age-related changes in HSCs and demonstrated that Mlkl-/- HSCs has more ATP production and sustained glycolytic flux.\nSignificance and innovation:\nIn the field of HSC aging there are many phenotypes reported that show premature aging of HSC. However, there very few studies demonstrating phenotypes that protect HSCs from acquiring aging hallmarks, like myeloid-biased or stem cell pool expansion with reduced reconstitution potential. Therefore, in my opinion the presented work is significant contribution to the field. While I am not an expert in the field of programmed cell death and necroptosis, the observation of the Mlkl translocation to membrane without induction of necroptosis seem to be not well studied, with some prior papers (eg. Vucur et al Immunity 2023) cited.\nMy major question/remarks:\nFirst is the conclusion about “sublethal necroptosis” stated within the title and text. While the term “sublethal necroptosis” has been used in field previously, in my opinion it might be misleading – either there is necroptosis, and the cell is dying or there is just necroptosis-independent role of Mlkl. The fact that the used FRET sensor shows Mlkl translocation to the membranes may not necessarily means it is involved in membrane rupture. Now, given the novel link with mitochondrial energy production this together may indicate an unrecognized function of Mlkl, that is not related to necroptosis. eg. membrane trafficking. Therefore, I am not fully convinced with the term sublethal necroptosis.\nSecond, I think it is not logically correct to state that “Sublethal necroptosis underlies hematopoietic stem cell aging” based on the observation of Mlkl deficiency result is resistance to age-related changes in HSC. I think, based on what is presented I would state that loss of MLKL protects against hallmarks of HSC aging, what is still an important observation.\nThe article open next important questions. How the Mlkl affects mitochondria, what it is role here? However I fully understand that this a wide research question and may be beyond the current study and was also stated by the authors in the discussion. But one thing I would suggest for consideration is to add that at this point the observed reversal of mitochondrial function may be the reason or the effect of the aging protection.\nFinally, I find very interesting that the protection of aging in Mlkl-/- mice is not linked to reversal of expansion of the expansion of myeloid-biased HSC defined as CD41+. Now, given the authors transplanted the whole pool of HSCs that means that either Mlkl-deficiency protects the “balanced” fraction of HSCs during aging or reverse the myeloid and aging phenotype of “myeloid-biased” HSCs. Given it is possible now to prospectively transplant myeloid-biased HSCs (work by Ross et al. using Neo-1 as marker, Nature 2024, Weissman lab) would it be possible to answer this question?\nResearch strategy:\nIn my opinion all HSC assays are well and robustly performed, well-presented and I have no doubts about the clear and significant effect of the Mlkl-deficiency on HSC aging protection. There are few places where I would suggest some more precise description (below).\nOther comments:\n- Extended Fig 1 C – the quality of the blot is poor, not sure if it is needed to include it – maybe the IHC is more informative\n- Authors wrote observed lower engraftment potential and a tendency toward reduced lymphopoietic potential of FRET/CFPhi HSCs compared to FRET/CFPlo HSCs. (Fig 1d-f) But this mostly B lineage at the cost of myelo, T cells are not changed here.\n- In pIC experiments authors stated that Mlkl deficiency significantly sustained the lymphoid potential, but to be precise is still only B lineage, and T lin even decreased.\n- Fig. 1 – it is stated that Mlkl activation is preferential in HSC, but the data shows it is also observed in MPPs to the same extend.\n- The same is in the Fig. 3a – statement that the activation is preferential to HSCs would require comparison to more populations (eg. MPPs) not only to MP\n- Authors discussed the changes of the mitochondria with aging as “HSC memory”. While I found the concept of HSC memory very valid and interesting, in my opinion the term memory makes sense when the memory gouvern some advantage to secodanry stimulai/challenge. Otherwise for me it is just another aging hallmark/feature. I can’t see what kind of “memory advantage” would the observed decline in mitochondria decline would provide."}, "author_response": "Point-by-point responses to reviewers' comments:\nWe wish to start by expressing our sincere appreciation to the reviewers for their constructive comments,\nwhich have helped us significantly improve our manuscript. The revision experiments have led us to\ndiscover two major findings: (1) the RIPK3-MLKL axis promotes age-related HSC functional decline\nprimarily via depleting NEO-1– lineage-balanced functional HSCs, and (2) active MLKL directly impairs\nmitochondrial membrane potential via mechanisms dependent on and independent of its pore-forming\nfunction. We have also revised the title and text in accordance with the reviewers’ suggestions as well as\nthe revised conclusions. Below, we provide a point-by-point response to each of the reviewer’s comments.\nReviewer #1\nWe thank the reviewer for her/his pertinent feedback on our manuscript. We are encouraged by the\nreviewer’s recognition of the scope and rigor of our study, and her/his acknowledgment of the potential\nrole of MLKL beyond necroptosis. The updated results now more convincingly support cell death-\nindependent role of the RIPK3-MLKL axis in age-related HSC functional decline. We have changed the\ntitle and revised the text in accordance with the reviewer’s suggestion along with the new experimental\ndata. We have also addressed the other concerns raised by the reviewer by performing additional\nexperiments and reanalyzing existing data.\nComment 1. In their study, Yamada et al. investigated the role of the necroptosis factor MLKL in HSC\naging. Using SMART-Tg mice they demonstrate an increase in MLKL, measured through FRET, upon\ntreatment with inflammation-inducing agents (poly-IC, LPS and TNF-alpha), as well as following\nreplication stress and in aged mice. This increase in MLKL was correlated with poorer HSC reconstitution,\nwhich was also observed in aged mice. Utilizing an MLKL knockout strain, the authors showed that MLKL\ndeficiency partially rescued the phenotype of poorer HSC reconstitution and other aging-associated\nfeatures. Furthermore, Yamada et al. discovered that activated MLKL accumulated in mitochondria\nleading to mitochondrial dysfunction in aged HSCs, which could partially be corrected by MLKL\ndeficiency. The study is very comprehensive and well-executed, presenting intriguing findings about the\npotential role of MLKL beyond necroptosis. However, my concern is that the cell death-independent role\nof MLKL is based solely on Figure 2B, which includes only four mice from two experiments, showing quite\nsome variation, especially in the WT pIC group. I recommend including more replicates in Figure 2B, and\nincorporating cell death analysis as validation in subsequent figures, such as in the Figure 2F experiment.\nResponse. We have added additional experimental mice to the results shown in Fig. 2b, which now include\n7 biological replicates from 3 independent experiments and support our initial conclusion that MLKL\nactivation does not mediate HSC death (Fig. 2b). We have also confirmed that overexpression of N-\nterminal FLAG-tagged MLKL does not increase HSC death (Fig. 2i) but decreases their regenerative\npotential with additional experiments (Fig. 2j-l) To assess the impact of 5-FU-induced activation of the\nRIPK3-MLKL axis on HSC death, we utilized MLKL-SA2 knock-in mice (JAX #039024) where MLKL\ncannot be activated by RIPK3-dependnet phosphorylation due to the two serine-to-alanine substitutions\nat codons 345 and 347 of the Mlkl locus (Yeap HW et al. EMBO J 2025, PMID: 40128366). Due to the\nsubstantial reduction of c-Kit+ expression on functional HSCs early after 5-FU administration, we omitted\nc-Kit from our analysis panel and instead added EPCR to enrich HSC populations as described previously\n(Umemoto T et al. J Exp Med 2018, PMID: 29946000). While MLKL activation was observed in HSCs\nat 16 h post-5-FU injection, cell death frequencies in Lin–/CD48–/CD150+/Sca-1+/EPCR+ HSCs were kept\nas low as ~5% at that time (Fig. 3c, Supplementary Fig. 4d). Moreover, even though the HSC death\nfrequencies elevated up to ~35% at 48 h post-5-FU administration, inactivation of the RIPK3-MLKL axis\nhad no significant impact on the 5-FU-induced HSC death (Fig. 3c, Supplementary Fig. 4d), confirming\nthat 5-FU-induced MLKL activation does not cause HSC death.\nComment 2. Another concern is that the authors have not conclusively shown that MLKL underlies aging,\nas stated in the title and throughout the manuscript. While it is evident that MLKL contributes to or even\npromotes aging, I disagree to the statement that MLKL is the driver of aging considering that the authors\nonly demonstrate partial restoration when MLKL is knocked out (e.g. Figure 4G, Figure 6F).\nResponse. We apologize for any unintended consequences arising from the manuscript’s title and wording,\nwhich may have been misleading. We totally agree that the partial restoration of aging hallmarks in Mlkl–\n/– HSCs rather indicates the RIPK3-MLKL axis is one of the mechanisms that mediate HSC aging, and\nthis was indeed the conclusion we highlighted in the Discussion section of the original manuscript. As\ndetailed in our response to reviewer #2, comment 3, we have performed additional experiments using\nisolated mitochondria and recombinant MLKL protein, and demonstrated that MLKL directly impairs\nmitochondrial function (Fig. 6h). To better reflect these notions, we have now changed the title to “non-\nnecroptotic MLKL function damages mitochondria and promotes hematopoietic stem cell aging” and\nreplaced the term “driver” by “mediator” throughout the manuscript.\nComment 3. Line 88 states that necroptosis is selectively and transiently activated in HSCs. However, the\ndata clearly show activation in both HSCs and multipotent progenitors upon treatment with pIC, LPS and\nTNFa (Figure 1B). Was lymphoid progenitors analyzed as well?\nResponse. We agree that MPP2 and MPP3/4 also showed FRET activation and have included this point\nin the text (lines 75–76). Regarding the FRET analysis on CLP, the key marker IL-7α could not be included\nin the original flow panel designed for identification of HSCs and MPPs and used in Fig. 1b, due to the\nlimited numbers of fluorescent channels available after allotting three colors required for the FRET\nanalysis (i.e., CFP, FRET, YPet). Thus, we have performed additional experiments with the new flow\npanel focusing on the CLP and demonstrated that while the FRET signal was significantly increased in\nLin–/Sca-1+/c-Kit+ cells that consist of HSCs and MPPs, no such increase was observed in CLPs\n(Supplementary Fig. 1c). Similarly, FRET signals were selectively elevated in HSCs/MPPs but not in\nCLPs or MPs upon 5-FU treatment (Fig. 3b, Supplementary Fig. 4a). These results consolidate our\nconclusion that the RIPK3-MLKL axis is preferentially activated in HSCs and MPPs.\nComment 4. In Figure 1D, FRET high and low cells were sorted and transplanted. It would be informative\nto show the sorting gates for FRET high and low. Does FRET low indicate “negative” cells or truly low\ncells? Would the truly FRET negative cell not be a better control than PBS control?\nResponse. We have now explicitly shown the sorting gates to isolate LPS-treated FRET/CFPhi and\nFRET/CFPlo HSCs for transplantation studies (Fig. 1e). Using the FRET signals in PBS-treated HSCs, we\nfirst defined the gates to select cells with the top 10% of FRET/CFP ratio as FRET/CFPhi (which normally\ncorresponds to cells showing decrease in CFP intensity due to FRET) and those with the bottom 30% as\nFRET/CFPlo and then applied the same gates to LPS-treated HSCs. As a result, we selected cells with the\nbottom 4% of FRET/CFP ratio as FRET/CFPlo in LPS-treated HSCs, which we believe selects truly FRET-\nlow cells. As the MLKL activation status is considered the sole difference between LPS-treated\nFRET/CFPhi and FRET/CFPlo HSCs, we indeed used FRET/CFPlo HSCs as the control to dissect the effect\nof LPS-induced MLKL activation on HSCs as explained in the original manuscript (lines 84–86).\nHowever, the relatively similar engraftment levels between PBS-treated HSCs and LPS-treated\nFRET/CFPlo HSCs (Fig. 1f) also indicates that LPS-treated HSCs without apparent MLKL activation\nmaintains similar levels of engraftment potential compared to the steady-state HSCs, which we think\nwould be also informative to the readers and thus be kept in the revised manuscript.\nComment 5. In transplantation experiments it would be informative to see the total reconstituted levels of\neach blood lineage (i.e. out of total blood cells) rather than only relative reconstitution frequencies within\nthe donor-derived compartment. This is especially relevant if total donor reconstitution levels vary a lot\nbetween recipient mice.\nResponse. We have reanalyzed the existing transplantation data to show donor chimerism within each\nlineage of peripheral blood leukocytes. The results were consistent with our conclusions, showing that\nMLKL deficiency improved the reconstitution levels of lymphoid cell lineages by pIC-treated\n(Supplementary Fig. 2g), 5-FU-treated (Supplementary Fig. 4i), serially transplanted (Supplementary Fig.\n5b), and aged HSCs (Supplementary Fig. 6g).\nComment 6. Could the absence of significant transcriptional differences between WT and MLKL-/- be due\nto the bulk RNA-seq mode, which might omit differences that would otherwise be detected with single cell\nRNA-seq?\nResponse. As the reviewer pointed out, we cannot formally exclude the possibility that MLKL might\naffect a specific HSC subpopulation that could not be detected by bulk RNA-seq. Unfortunately, due to\nour recent move to St. Jude, we no longer have access to aged Mlkl–/– mice to perform single cell RNA-\nseq. However, in response to the Reviewer #3 comment 4, we have found that blockade of the RIPK3-\nMLKL axis largely prevents pIC-induced conversion from neogenin-1-negative (NEO-1–) lineage-\nbalanced HSCs to NEO-1+ myeloid-biased HSCs (Fig. 2d-e). While we could not detect Neo-1 gene\nexpression changes between WT and Mlkl–/– aged HSCs in the bulk RNA-seq, the results support the\nnotion that activation of the RIPK3-MLKL axis shifts NEO-1– lineage-balanced HSCs toward NEO-1+\nmyeloid-biased HSCs. Thus, we have ameliorated the corresponding statement (lines 199–200) and\ndiscussed the limitation of our bulk approaches in the Discussion section (line 285–286).\nComment 7. Why were young mice not included in Figure 6E, G, H for comparison?\nResponse. Fig. 6d (i.e., Fig. 6e of the original manuscript) has now included quantitative data and\nstatistical analyses for young and aged Mlkl–/– HSCs. For Fig. 6f,g (i.e., Fig. 6g,h of the original\nmanuscript), the seahorse assay requires ~280,000 HSCs (i.e., at least 70,000 cells per well, with a\nminimum of four biological replicates for reliable measurement). While we could obtain ~100,000 HSCs\nfrom single aged mice due to their age-dependent expansion, it is extremely challenging to obtain enough\nnumbers of HSCs from young mice, as we could typically obtain 5,000 HSCs per mouse and would require\n>50 mice per experiment. We have now clarified this limitation in the Result section (lines 220–221).\nComment 8. Do MLKL-/- mice live longer?\nResponse. Although we did not formally analyze the lifespan of Mlkl–/– mice, we did not recognize any\napparent difference in terms of their morbidity and mortality, in line with the prior observation (Tovey\nCrutchfield EC et al. Cell Death Differ 2023, PMID: 36755069). We have included this point in the\nDiscussion section (lines 293–294).\nComment 9. Have you checked for any differences between females and males in inflammation-induced\nMLKL activation and in aging?\nResponse. We have reanalyzed the existing data of SMART-Tg mice to assess potential sex differences in\nMLKL activation. The results indicate that inflammation-induced (Supplementary Fig. 1e) and age-related\nMLKL activation (Supplementary Fig. 6a) was generally conserved between male and female mice,\narguing for sex-independent MLKL activation in HSCs.\nReviewer #2\nWe greatly appreciate the reviewer’s positive and thoughtful evaluation of our study. We are pleased that\nthe reviewer values the robustness of our findings and the scientific rigor of our approach, in particular\nthe N-terminal FLAG-tagged MLKL (FLAG-MLKL) rescue experiments. The reviewer raised important\nquestions as to whether RIPK3 acts upstream of MLKL to promote HSC aging, and the mechanism\nwhereby MLKL impairs mitochondrial function in HSCs. We have performed a series of experiments to\naddress these questions and revised the title to better align with the conclusions.\nComment 1. The work by Yamada et al. investigates a cell death-independent role of MLKL in HSC\ninflammation and aging, which does not regulate the transcriptome; instead, it impairs mitochondrial\nfitness. This work is well-conducted and controlled; the phenotype is robust; and the scientific rigor is\nhigh. One of several experiments I appreciate a lot is the N-flag MLKL rescue one. As N-flag MLKL cannot\ninduce necroptosis but can still rescue the MLKL KO HSC phenotype, I believe the \"cell-death-\nindependent role\" of MLKL in HSC is very solid. Although the overall quality of this work is high, some\nminor concerns can be addressed to enhance its excellence further.\nResponse. As described above, we have performed additional experiments to address the concern and\nstrengthen our work. The detailed responses are shown below.\nComment 2. As the authors do have the RIP3 KO mice, it would be great to test whether RIP3 KO can\nphenocopy MLKL KO, in assays such as Fig. 2d, 3d and 4g. If the aged RIP3 KO mice are not available,\nthen just discuss this as a limitation of this study, as I don't want to delay the publication for too long. This\nRIP3 KO mice work will complete the story and add more impact to this study.\nResponse. We agree that testing whether RIPK3 is the upstream regulator of MLKL in the observed HSC\nchanges would further strengthen our work. Unfortunately, our recent relocation from Japan to the US\nprevented our access to the RIPK3–/– mice. To circumvent this issue, we have instead taken advantage of\nUH15-38, a RIPK3 inhibitor recently developed for in vivo use (Gautam A et al. Nature 2024, PMID:\n38600381). We found that administration of UH15-38 prior to pIC injection showed trend towards\nameliorating pIC-induced decline in HSC regenerative capacity and lymphoid differentiation potential\n(Supplementary Fig. 3a-d), which is reminiscent of the phenotype observed in Mlkl–/– HSCs. To further\naddress the role of RIPK3-dependent MLKL activation in HSC changes, we utilized MLKL-SA2 knock-\nin mice (JAX #039024), where MLKL cannot be activated by RIPK3-dependnet phosphorylation due to\nthe two serine-to-alanine substitutions at codons 345 and 347 of the Mlkl locus (Yeap HW et al. EMBO J\n2025, PMID: 40128366). We found that MLKL-SA2 and MLKL deficiency similarly attenuate pIC-\ninduced upregulation of age-related marker NEO-1 on HSCs (Fig. 2d-e). These data, together with the\ndata shown in Fig. 2f, further consolidate our conclusion that the RIPK3-MLKL axis promotes age-related\nfunctional decline in HSCs. As the inhibitor is not suitable for long-term inhibition of RIPK3 in vivo, we\ncould not apply this approach to other aging model such as sequential 5-FU treatment or organismal aging,\nand the limitation of our study regarding RIPK3 requirement has been discussed in the Discussion section\n(lines 267–270).\nComment 3. For the N-flag-MLKL rescue experiments, it would be interesting to know whether N-flag-\nMLKL can still target the mito (via PLA) or impair mito function. This is important because N-flag-MLKL\ncan still localize to the plasma membrane but cannot perform pore formation on the plasma membrane\n(https://www.nature.com/articles/cdd201570). So this work would be informative.\nResponse. We thank the reviewer for this insightful suggestion that gave us an opportunity to gain\nmechanistic insights as to how MLKL impairs mitochondrial function. We first confirmed the expression\nof FLAG-MLKL by immunofluorescence (Supplementary Fig. 3m). We then performed proximity\nligation assay (PLA) with p-MLKL and COX-IV antibodies to examine whether FLAG-MLKL can still\ntarget mitochondria. The results have convincingly shown induction of PLA foci in FLAG-MLKL-\ntransduced HSCs, supporting mitochondrial localization of N-FLAG-MLKL (Supplementary Fig. 8b). To\ndetermine whether MLKL directly impairs mitochondrial membrane potential, mitochondria were isolated\nfrom mouse liver and incubated in vitro with either (i) N-terminal four-helix bundle domain of\nrecombinant mouse MLKL protein (MLKL-NTD) that oligomerizes and forms pores on the cell\nmembrane without stimulation, (ii) N-terminal FLAG-tagged MLKL-NTD (FLAG-MLKL-NTD) that\nretains oligomerization capacity but cannot perform pore formation on the cell membrane, or (iii) C-\nterminal pseudokinase domain of recombinant mouse MLKL protein (MLKL-CTD) that does not\noligomerize or form pores on the cell membrane (Hildebrand JM et al. PNAS 2014, PMID: 25288762).\nMitochondria were then stained with Tetramethylrhodamine methyl ester (TMRM) as well as MitoTracker\ngreen, and TMRM signals within MitoTracker-positive population were analyzed by flow cytometry.\nTMRM signals in mitochondria were significantly reduced by incubation with MLKL-NTD, and to a\nlesser extent by FLAG-MLKL-NTD, but not by MLKL-CTD (Fig. 6h). These results indicate that\nactivated MLKL directly impairs mitochondrial membrane potential via mechanisms both dependent and\nindependent of its pore-forming capacity.\nComment 4. For the title, since it is not \"sub-lethal\", it is \"necroptosis-independent\", please revise it to\nalign with the current abstract. And I would also suggest that include \"inflammation\" on the title as I think\nthis work is more than just studying \"aging\".\nResponse. We agree that our results support a necroptosis-independent role of the RIPK3-MLKL axis\nrather than the effect of its sublethal activation. While we also agree that inflammation is another keyword\nto consider in the title, we think inflammation is just one of the stressors that induce age-related HSC\nchanges via non-necroptotic MLKL activation in HSCs. Now that we have established the causality\nbetween MLKL and mitochondrial impairment by the above experiments, we feel that including this\nnotion helps the title better represent the scope of the revised manuscript. Thus, we have changed the title\nto “non-necroptotic MLKL function damages mitochondria and promotes hematopoietic stem cell aging”\nand revised the text accordingly.\nReviewer #3\nWe thank the reviewer for the positive and insightful comments. We appreciate the recognition of our\nwork’s significance in revealing a non-canonical role of MLKL in HSC regulation. We have carefully\nconsidered all points to further strengthen the manuscript, with particular emphasis on additional analyses\nof NEO-1, including transplantation experiments.\nComment 1. The manuscript by Yamada et al. describes the role of Mlkl in hematopoietic stem cells (HSC).\nAuthors showed that Mlkl is activated in HSC and multipotent progenitor (MPP) pool upon inflammation\nusing FRET based sensor that monitor MLKL translocation to membrane. However, they showed that this\nactivation does not lead to cell death of HSCs, as expected by the known function of Mlkl in necroptosis.\nNext, authors performed series of experiments on Mlkl-/- mice, and demonstrated that such mice do not\nshow many established hallmarks of HSC aging or strong inflammation-driven exhaustion of HSC\nfunction. Finally, the authors showed the link of Mlkl with age-related changes in HSCs and demonstrated\nthat Mlkl-/- HSCs has more ATP production and sustained glycolytic flux. Significance and innovation: In\nthe field of HSC aging there are many phenotypes reported that show premature aging of HSC. However,\nthere very few studies demonstrating phenotypes that protect HSCs from acquiring aging hallmarks, like\nmyeloid-biased or stem cell pool expansion with reduced reconstitution potential. Therefore, in my opinion\nthe presented work is significant contribution to the field. While I am not an expert in the field of\nprogrammed cell death and necroptosis, the observation of the Mlkl translocation to membrane without\ninduction of necroptosis seem to be not well studied, with some prior papers (eg. Vucur et al Immunity\n2023) cited. My major question/remarks: First is the conclusion about “sublethal necroptosis” stated\nwithin the title and text. While the term “sublethal necroptosis” has been used in field previously, in my\nopinion it might be misleading – either there is necroptosis, and the cell is dying or there is just\nnecroptosis-independent role of Mlkl. The fact that the used FRET sensor shows Mlkl translocation to the\nmembranes may not necessarily means it is involved in membrane rupture. Now, given the novel link with\nmitochondrial energy production this together may indicate an unrecognized function of Mlkl, that is not\nrelated to necroptosis. eg. membrane trafficking. Therefore, I am not fully convinced with the term\nsublethal necroptosis.\nResponse. We appreciate the reviewer’s thoughtful comments. We agree that our results support a\nnecroptosis-independent role of the RIPK3-MLKL axis rather than the effect of its sublethal activation.\nTo address this and other points raised by the reviewer, we have now revised the title to “non-necroptotic\nRIPK3-MLKL damage mitochondria and promotes hematopoietic stem cell aging by damaging\nmitochondria.”\nComment 2. Second, I think it is not logically correct to state that “Sublethal necroptosis underlies\nhematopoietic stem cell aging” based on the observation of Mlkl deficiency result is resistance to age-\nrelated changes in HSC. I think, based on what is presented I would state that loss of MLKL protects\nagainst hallmarks of HSC aging, what is still an important observation.\nResponse. We apologize for any unintended consequences arising from the manuscript’s title and wording,\nwhich may have been misleading. We totally agree that the partial restoration of aging hallmarks in Mlkl–\n/– HSCs rather indicates the RIPK3-MLKL axis is one of the mechanisms that mediate HSC aging, and\nthis was indeed the conclusion we highlighted in the Discussion section of the original manuscript. As\ndetailed below in our response to the reviewer’s comment 3, we have performed additional experiments\nusing isolated mitochondria and recombinant MLKL protein, and demonstrated that MLKL directly\nimpairs mitochondrial function (Fig. 6h). To better reflect these notions, we have now changed the title to\n“non-necroptotic MLKL function damages mitochondria and promotes hematopoietic stem cell aging”\nand revised the text accordingly.\nComment 3. The article open next important questions. How the Mlkl affects mitochondria, what it is role\nhere? However I fully understand that this a wide research question and may be beyond the current study\nand was also stated by the authors in the discussion. But one thing I would suggest for consideration is to\nadd that at this point the observed reversal of mitochondrial function may be the reason or the effect of\nthe aging protection.\nResponse. We thank the reviewer for this insightful suggestion that gave us an opportunity to gain\nmechanistic insights as to how MLKL impairs mitochondrial function. We first confirmed that N-terminal\nFLAG-tagged MLKL (FLAG-MLKL), which lacks the capacity to form pores on the cell membrane, can\nstill target mitochondria in HSCs using proximity ligation assay PLA (Supplementary Fig. 8b). To\ndetermine whether pore-forming function of MLKL directly impairs mitochondrial membrane potential,\nmitochondria were isolated from mouse liver and incubated in vitro with either (i) N-terminal four-helix\nbundle domain of recombinant mouse MLKL protein (MLKL-NTD) that oligomerizes and forms pores\non the cell membrane without stimulation, (ii) N-terminal FLAG-tagged MLKL-NTD (FLAG-MLKL-\nNTD) that retains oligomerization capacity but cannot perform pore formation on the cell membrane, or\n(iii) C-terminal pseudokinase domain of recombinant mouse MLKL protein (MLKL-CTD) that does not\noligomerize or form pores on the cell membrane (Hildebrand JM et al. PNAS 2014, PMID: 25288762).\nMitochondria were then stained with Tetramethylrhodamine methyl ester (TMRM) as well as MitoTracker\ngreen, and TMRM signals within MitoTracker-positive population were analyzed by flow cytometry.\nTMRM signals in mitochondria were significantly reduced by incubation with MLKL-NTD, and to a\nlesser extent by FLAG-MLKL-NTD, but not by MLKL-CTD (Fig. 6h). These results indicate that\nactivated MLKL directly impairs mitochondrial membrane potential via mechanisms both dependent and\nindependent of its pore-forming capacity.\nComment 4. Finally, I find very interesting that the protection of aging in Mlkl-/- mice is not linked to\nreversal of expansion of the expansion of myeloid-biased HSC defined as CD41+. Now, given the authors\ntransplanted the whole pool of HSCs that means that either Mlkl-deficiency protects the “balanced”\nfraction of HSCs during aging or reverse the myeloid and aging phenotype of “myeloid-biased” HSCs.\nGiven it is possible now to prospectively transplant myeloid-biased HSCs (work by Ross et al. using Neo-\n1 as marker, Nature 2024, Weissman lab) would it be possible to answer this question?\nResponse. We thank the reviewer for this important comment. Additional experiments with PBS- or\npolyinosinic-polycytidylic acid (pIC)-treated Mlkl–/– mice revealed that MLKL deficiency significantly\nattenuates pIC-induced NEO-1 upregulation on HSCs (Fig. 2d). To further assess whether activation of\nthe RIPK3-MLKL axis affects age-related HSC subpopulations, we utilized MLKL-SA2 knock-in mice\n(JAX #039024) where MLKL cannot be activated by RIPK3-dependnet phosphorylation due to the two\nserine-to-alanine substitutions at codons 345 and 347 of the Mlkl locus (Yeap HW et al. EMBO J 2025,\nPMID: 40128366), and treated them with PBS or pIC. In addition to NEO-1, we also assessed surface\nexpression of P-selectin and GPR183, which were reported to enrich myeloid-biased (Flohr Svendsen A\net al. Blood 2021, PMID: 33876187) and less differentiating (Totani H et al. Nat Aging 2025; PMID:\n40050412) subpopulations in aged HSCs, respectively. The results revealed that inactivation of the\nRIPK3-MLKL axis significantly attenuated pIC-induced upregulation of NEO-1, and to a lesser extent\nGPR183, but not P-selectin, on HSCs (Fig. 2e, Supplementary Fig. 2e-f). As the total number of HSCs did\nnot differ before and after acute pIC injection (Fig. 2c), this indicates that the RIPK3-MLKL axis promotes\ntransition from NEO-1– to NEO-1+ states. Transplantation of NEO-1– and NEO-1+ HSCs isolated from\npIC-treated WT and MlklSA2 mice further revealed a trend toward improved engraftment of MlklSA2 NEO-\n1– HSCs, but not NEO-1+ HSCs, compared to WT counterparts (Supplementary Fig. 3e-i), indicating\nselective protection of NEO-1– HSCs by blockade of the RIPK3-MLKL axis. Thus, we concluded that the\nRIPK3-MLKL axis primarily promotes age-related functional decline in NEO-1– lineage-balanced HSCs.\nComment 5. Research strategy: In my opinion all HSC assays are well and robustly performed, well-\npresented and I have no doubts about the clear and significant effect of the Mlkl-deficiency on HSC aging\nprotection. There are few places where I would suggest some more precise description (below).\nResponse. We thank the reviewer for the positive assessment of our research strategy and HSC assays.\nWe have revised the manuscript according to the following comments.\nOther comments:\nComment 6. Extended Fig 1 C – the quality of the blot is poor, not sure if it is needed to include it – maybe\nthe IHC is more informative\nResponse. In accordance with the reviewer’s suggestion, we have removed the Western blot result and\ndeleted the corresponding text.\nComment 7. Authors wrote observed lower engraftment potential and a tendency toward reduced\nlymphopoietic potential of FRET/CFPhi HSCs compared to FRET/CFPlo HSCs. (Fig 1d-f) But this mostly\nB lineage at the cost of myelo, T cells are not changed here.\nResponse. We have revised the text accordingly (line 86).\nComment 8. In pIC experiments authors stated that Mlkl deficiency significantly sustained the lymphoid\npotential, but to be precise is still only B lineage, and T lin even decreased.\nResponse. We have revised the text accordingly (lines 115–116).\nComment 9. Fig. 1 – it is stated that Mlkl activation is preferential in HSC, but the data shows it is also\nobserved in MPPs to the same extend.\nResponse. We have revised the text accordingly (lines 75–76).\nComment 10. The same is in the Fig. 3a – statement that the activation is preferential to HSCs would\nrequire comparison to more populations (eg. MPPs) not only to MP\nResponse. Additional analysis revealed that FRET signals were significantly elevated in MPP2 and\nMPP3/4 but not in CLP upon 5-FU treatment (Fig. 3b, Supplementary Fig. 4a). We have revised the text\nto reflect the results (lines 137–139).\nComment 11. Authors discussed the changes of the mitochondria with aging as “HSC memory”. While I\nfound the concept of HSC memory very valid and interesting, in my opinion the term memory makes sense\nwhen the memory gouvern some advantage to secodanry stimulai/challenge. Otherwise for me it is just\nanother aging hallmark/feature. I can’t see what kind of “memory advantage” would the observed decline\nin mitochondria decline would provide.\nResponse. We agree that the term “memory” could imply an adaptation mechanism that is primarily\ndesigned for cells to improve their outcome in response to secondary stimuli. While accumulating\ndamaged mitochondria could be an adaptive HSC response to prevent their unlimited self-renewal as\ndiscussed elsewhere (Hinge A et al. Cell Stem Cell 2020, PMID: 32059807), this concept has not formally\nbeen proven yet. We have thus rephrased the text in Supplementary Fig. 8h and the Discussion section\n(lines 245–253) to infer cumulative cellular damage that limits HSC function after stress."}, {"round": 2, "reviewer_comments": {"reviewer_1": "In the revised study by Yamada et al., the authors have comprehensively addressed the concerns raised during the previous review round by incorporating additional experiments and new analyses. These additions substantially strengthen the manuscript and enhance the clarity of its conclusions. Notably, the inclusion of experiments investigating MLKL in mitochondrial function and the role of RIPK3–MLKL in age-related HSC functional decline, through the analysis of NEO-positive and NEO-negative HSCs, represents a significant improvement.\nOverall, the revisions have increased the quality and impact of the study. I am satisfied with the authors’ responses and the new data presented.", "reviewer_3": "The authors have addressed all of my comments. I find the observation of reversed Neo-1 expression and the analysis of myeloid-biased HSCs following pIC treatment particularly interesting and representing an added value to the manuscript. I also acknowledge that the authors performed Neo-1⁺/Neo-1⁻ HSC transplantation experiments, which support the conclusion that RIPK3–MLKL blockade acts on the Neo-1 fraction; however, the number of mice included in these experiments is relatively limited.\nNevertheless, in my opinion, the revision is comprehensive and has significantly improved the manuscript."}, "author_response": "Point-by-point responses to reviewers' comments:\nWe thank the editor and reviewers for the in-principle acceptance of our manuscript. As no further\nrevisions were requested at this stage, we confirm that no additional changes have been made beyond\nthose already incorporated during the previous revision cycle, except for minor data corrections and\nbrief textual clarifications. We are grateful for the constructive feedback that strengthened the study.\nReviewer #1\nComment. In the revised study by Yamada et al., the authors have comprehensively addressed the\nconcerns raised during the previous review round by incorporating additional experiments and new\nanalyses. These additions substantially strengthen the manuscript and enhance the clarity of its\nconclusions. Notably, the inclusion of experiments investigating MLKL in mitochondrial function and\nthe role of RIPK3–MLKL in age-related HSC functional decline, through the analysis of NEO-\npositive and NEO-negative HSCs, represents a significant improvement. Overall, the revisions have increased\nthe quality and impact of the study. I am satisfied with the authors’ responses and the new data\npresented.\nResponse. We sincerely thank the reviewer for the positive assessment and for recognizing the added\nexperiments and analyses, including the MLKL mitochondrial function assays and RIPK3–MLKL\nevaluations in Neo-1⁺ and Neo-1⁻ HSCs.\nReviewer #2\nComment. Accept\nResponse. We thank the reviewer for her/his support of the manuscript.\nReviewer #3\nComment. The authors have addressed all of my comments. I find the observation of reversed Neo-1\nexpression and the analysis of myeloid-biased HSCs following pIC treatment particularly interesting\nand representing an added value to the manuscript. I also acknowledge that the authors performed\nNeo-1⁺/Neo-1⁻ HSC transplantation experiments, which support the conclusion that RIPK3–MLKL\nblockade acts on the Neo-1 fraction; however, the number of mice included in these experiments is\nrelatively limited. Nevertheless, in my opinion, the revision is comprehensive and has significantly\nimproved the manuscript.\nWe thank the reviewer for the constructive feedback and positive evaluation. We acknowledge the\ncomment regarding the limited number of mice in the transplantation experiments and have added a\nbrief clarification in the Results section (line 122)."}]

Decision letter