An NAD⁺-dependent metabolic checkpoint regulates hematopoietic stem cell activation and aging

Abstract

How hematopoietic stem cells (HSCs) maintain metabolic homeostasis to support tissue repair and regeneration throughout the lifespan is elusive. Here, we show that CD38, an NAD⁺-dependent metabolic enzyme, promotes HSC proliferation by inducing mitochondrial Ca²⁺ influx and mitochondrial metabolism in young mice. Conversely, aberrant CD38 upregulation during aging is a driver of HSC deterioration in aged mice due to dysregulated NAD⁺ metabolism and compromised mitochondrial stress management. The mitochondrial calcium uniporter, a mediator of mitochondrial Ca²⁺ influx, also supports HSC proliferation in young mice yet drives HSC decline in aged mice. Pharmacological inactivation of CD38 reverses HSC aging and the pathophysiological changes of the aging hematopoietic system in aged mice. Together, our study highlights an NAD⁺ metabolic checkpoint that balances mitochondrial activation to support HSC proliferation and mitochondrial stress management to enhance HSC self-renewal throughout the lifespan, and links aberrant Ca²⁺ signaling to HSC aging.

Older humans are prone to many pathophysiological conditions in the hematopoietic system, such as decreased competence of the adaptive immune system¹. The maintenance of the hematopoietic system throughout adult life relies on the persistence of hematopoietic stem cells (HSCs). HSCs are capable of self-renewal to give rise to daughter stem cells and of differentiation to give rise to all the blood cell types, including myeloid and lymphoid lineages. The regenerative capacity of HSCs diminishes with age^2–4. HSC aging is a major cause of the pathophysiological conditions in the aging hematopoietic system. Mitochondrial metabolism is essential to support HSC function^5,6. Concomitant to mitochondrial activation is mitochondrial stress, which leads to loss of HSC maintenance^7–11. It is unclear how HSCs balance mitochondrial activation to support proliferation and mitochondrial stress management to support self-renewal throughout the lifespan.

NAD⁺ is not only a coenzyme for metabolic reactions but also possesses signaling functions to regulate diverse cellular processes, including mitochondrial stress management mediated by members of the sirtuin family of NAD⁺-dependent deacetylases^{2,7,10,12–14}. CD38, a transmembrane glycoprotein, catalyzes the formation of cyclic ADP-ribose (cADPR), ADP-ribose (ADPR) and nicotinic acid adenine dinucleotide phosphate (NAADP) from NAD⁺ (refs. 15,16). cADPR, ADPR and NAADP mobilize Ca²⁺ from intracellular Ca²⁺ stores^16,17. CD38 regulates immune responses in lymphocytes, oxytocin secretion and social behavior, metabolic homeostasis, and is used as a malignancy marker in leukemia and a target for treating multiple myeloma^18–21. Its expression in HSCs is low²², suggesting that CD38 activity needs to be kept in check for HSC maintenance. In this study, using mice in which the CD38 gene was deleted, we showed that CD38 supports Ca²⁺ signaling, mitochondrial activity and proliferation of HSCs at a young age. However, aberrant upregulation of CD38 in HSCs during aging results in accumulation of mitochondrial stress, with profound pathophysiological changes reminiscent of HSC aging. Thus, CD38 governs an NAD⁺ metabolic checkpoint that balances mitochondrial activity to support HSC proliferation and mitochondrial stress management to support HSC self-renewal.

CD38 is dispensable for HSCs under homeostasis at a young age

We characterized the phenotypically defined enriched HSCs (Lin⁻c-Kit⁺Sca1⁺) and highly enriched HSCs (Lin⁻c-Kit⁺Sca1⁺CD150 ⁺CD48⁻) in young (3-month-old) female CD38 knockout (KO) mice and their wild type (WT) controls. Under homeostatic condition, about 95% of HSCs are quiescent. There was no difference in the number of HSCs in the bone marrow (BM) (Extended Data Fig. 1a–c), and the percentage of Ki-67⁺ (Extended Data Fig. 1d) or 7-AAD⁺ HSCs (Extended Data Fig. 1e), indicating that CD38 inactivation does not affect HSC survival and proliferation under homeostatic condition. There was no difference in lineage differentiation in the peripheral blood (Extended Data Fig. 1f). Complete blood count analyses showed no difference (Extended Data Fig. 1g,h). BM cellularity was not altered (Extended Data Fig. 1i). Thus, CD38 is not required for HSC maintenance and hematopoiesis under homeostatic condition at a young age.

CD38 is required to support HSC proliferation at a young age

We tested whether CD38 is required for HSC activation on proliferation. To stimulate HSC proliferation, HSCs derived from young WT and CD38 KO mice were cultured in the presence of cytokines known to induce HSC proliferation and commitment for the physiological stimulation of blood production²³. After 18 h of culture, there were fewer CD38 KO HSCs than WT HSCs (Fig. 1a). Bromodeoxyuridine (BrdU) labeling showed reduced proliferation for CD38 KO HSCs (Fig. 1b), while there was no difference in HSC survival, as indicated by 7-AAD staining (Fig. 1c). HSC culture in the presence of polyvinyl alcohol (PVA) allows expansion of functional HSCs over a long time²⁴. After 7 days of culture in the presence of PVA, the number of cells derived from CD38 KO HSCs (Extended Data Fig. 2a) was reduced.

Fig. 1 |. CD38 is required to support HSC proliferation at young age.

a–c, Enriched HSCs isolated from 3-month-old WT and CD38 KO mice were cultured in the presence of cytokines. The number of HSCs (P = 0.0113) (a), BrdU incorporation (P = 0.0215) (b) and 7-AAD staining (P = 0.3356) (c) of HSCs were analyzed using flow cytometry. a,b, n = 6. c, n = 6 and 5. d–f, Competitive transplantation using HSCs isolated from 3-month-old WT and CD38 KO mice as donors. The percentage of donor-derived HSCs in the BM of the recipients (P = 0.0144) (d), donor-derived lineage differentiation in the peripheral blood (P = 0.2442) (e) and the percentage of donor-derived cells in the peripheral blood of the recipients (4 weeks, P < 0.0001; 8 weeks, P = 0.0017; 12 weeks, P = 0.0018; 16 weeks, P = 0.0039) (f) were analyzed using flow cytometry. d, n = 5 and 6. e,f, n = 11 and 12. g,h, Noncompetitive transplantation using whole-BM cells isolated from 3-month-old WT and CD38 KO mice as donors. Ki-67 staining (P = 0.0442) (g) and 7-AAD staining (P = 0.1802) (h) of donor-derived HSCs were analyzed using flow cytometry 1 month after transplantation. g, n = 5. h, n = 6. i, The homing ability of BM cells from 3-month-old WT and CD38 KO mice 16 h after transplantation was analyzed using flow cytometry (P = 0.2659). n = 6. Data are presented as the mean ± s.e. *P < 0.05, **P < 0.01, ***P < 0.001, NS P > 0.05. A one-sided Student’s t-test was used. MNC, mononuclear cell.

To stimulate HSC proliferation in vivo, we performed a competitive BM transplantation assay using HSCs isolated from young WT and CD38 KO mice as donors. Sorted HSCs from CD45.2 donor mice were mixed with CD45.1 B6.SJL competitor BM cells and were injected into lethally irradiated B6.SJL recipient mice. CD38 KO HSCs showed reduced engraftment in the BM of recipient mice (Fig. 1d), comparable lineage differentiation in peripheral blood (Fig. 1e) and reduced capacity to reconstitute the blood system of lethally irradiated recipient mice (Fig. 1f). Staining of donor-derived HSCs in the BM of recipient mice showed a reduced frequency of Ki-67⁺ CD38 KO HSCs (Fig. 1g), while the percentage of 7-AAD⁺ HSCs was comparable between the two genotypes (Fig. 1h), suggesting that reduced CD38 KO HSC engraftment was due to reduced proliferation but not cell death. There was no difference in homing ability (Fig. 1i). As an alternative approach to stimulate HSC proliferation in vivo, we treated mice with poly(I:C)²⁵. The frequency of Ki-67⁺ HSCs was reduced in CD38 KO mice on poly(I:C) treatment (Extended Data Fig. 2b). Together, these data indicate that CD38 is required to support HSC proliferation at a young age. Consistently, although the CD38^lo population is more enriched for colony-forming cells compared to the CD38^hi population, the CD38^hi population proliferates faster, forms larger colonies and supports long-term reconstitution²⁶.

To determine whether CD38 regulates HSC function autonomously, we performed a competitive BM transplantation assay using CD38 KO HSCs with or without CD38 overexpression as donors. CD38 overexpression resulted in increased capacity to reconstitute the blood system of lethally irradiated recipient mice (Extended Data Fig. 2c), suggesting that CD38 regulates HSC function autonomously.

CD38 promotes mitochondrial activity on HSC proliferation

On cytokine stimulation, RNA sequencing (RNA-seq) analyses of HSCs isolated from young WT and CD38 KO mice showed differentially expressed genes related to cell cycle and mitochondria (Fig. 2a–c and Supplementary Table 1). In contrast, HSCs derived from young WT and CD38 KO mice under homeostatic condition did not show these changes (Supplementary Table 1). To explore the metabolic effects of CD38 in HSCs on proliferation, we administered [U-¹³C]glucose to WT and CD38 KO HSCs ex vivo in the presence of cytokine stimulation. Metabolomics revealed that the total levels of the tricarboxylic acid (TCA) cycle intermediates succinate and fumarate were reduced in CD38 KO HSCs (Fig. 2d,e). CD38 KO HSCs had reduced incorporation of [U-¹³C]glucose into the TCA cycle intermediates (Fig. 2f,g), which is consistent with reduced mitochondrial metabolism.

Fig. 2 |. CD38 regulates Ca²⁺ signaling and mitochondrial activity on HSC proliferation.

a–l, Enriched HSCs isolated from 3-month-old WT and CD38 KO mice were cultured in the presence of cytokines. a–c, RNA-seq analyses of enriched HSCs. a, Volcano plot summarizing differentially expressed genes (red, upregulated in CD38 KO; blue, downregulated in CD38 KO. P_adj < 0.1. Benjamini–Hochberg-corrected). Genes related to HSC proliferation are labeled. b,c, Gene set enrichment analysis (GSEA) for mitochondria (b) and cell cycle (c). n = 2. d–g, [U-¹³C]glucose was administered to the HSC culture. Relative abundances of total metabolites (succinate, P = 0.0399 (d); fumarate, P = 0.0273 (e)) and ¹³C-labeled metabolites (malate, P = 0.0362 (f); fumarate, P = 0.0076 (g)) in enriched HSCs were measured using liquid chromatography–mass spectrometry (LC–MS). n = 4. h, ATP levels in enriched HSCs (P = 0.0193). n = 3. i–l, The staining for Fluo-4 (P = 0.0071) (i), Rhod-2 (P = 0.0245) (j), TMRM (P = 0.0027) (k) and ROS (P = 0.0272) (l) of HSCs was analyzed using flow cytometry. i,k,l, n = 5 and 6. j, n = 6. Data are presented as the mean ± s.e. *P < 0.05. **P < 0.01. A one-sided Student’s t-test was used. MFI, mean fluorescence intensity; NES, normalized enrichment score.

HSCs reside in a hypoxic niche in vivo and primarily rely on glycolysis for energy production²⁷. To further explore the metabolic effects of CD38 in HSCs on proliferation in vivo, we performed metabolomics of HSCs isolated from mice transplanted with WT and CD38 KO HSCs 1 month after transplantation. The levels of fructose 6-phosphate and lactate were increased in HSCs derived from mice transplanted with CD38 KO HSCs (Extended Data Fig. 2d,e), which is consistent with reduced mitochondrial metabolism and compensatory glycolysis. Cellular ATP levels were reduced in CD38 KO HSCs on either cytokine-stimulated proliferation (Fig. 2h) or transplantation-stimulated proliferation (Extended Data Fig. 2f). Together, these data suggest that in HSCs, CD38 is required for mitochondrial metabolism and proliferation.

Mitochondrial Ca²⁺ influx supports HSC proliferation

On HSC transition from quiescence to proliferation, cytosolic and mitochondrial Ca²⁺ levels increased²⁸. Staining with Rhod-2, a fluorescent indicator for mitochondrial Ca²⁺ levels, and Fluo-4, a fluorescent indicator that monitors cytoplasmic Ca²⁺ levels, showed that both cytosolic and mitochondrial Ca²⁺ levels were reduced in CD38 KO HSCs on cytokine-stimulated proliferation (Fig. 2i,j) and transplantation-stimulated proliferation (Extended Data Fig. 2g–j). Mitochondrial activity and stress, as indicated by staining for tetramethylrhodamine methyl ester (TMRM) that detects mitochondrial membrane potential and MitoSOX mitochondrial superoxide indicator, were decreased in CD38 KO HSCs under these proliferation conditions (Fig. 2k,l and Extended Data Fig. 2k–n). Additionally, CD38 KO HSCs also showed reduced Ca²⁺ levels, mitochondrial membrane potential and stress under poly(I:C)-induced proliferation (Extended Data Fig. 3a–c). CD38^lo HSCs had lower mitochondrial membrane potential and stress than CD38^hi HSCs on cytokine-stimulated proliferation (Extended Data Fig. 3d–f). In contrast, there was no difference in Ca²⁺ levels, mitochondrial membrane potential and stress between quiescent WT and CD38 KO HSCs under homeostatic condition (Extended Data Fig. 4a–h).

To determine whether mitochondrial Ca²⁺ influx is required to support HSC proliferation, a loxP allele for the mitochondrial calcium uniporter (MCU) located in the organelle’s inner membrane²⁹, was crossed with Vav-iCre transgenic mice to delete MCU in the HSCs. HSCs in Vav-iCre^+/−;MCU^loxP/loxP mice (referred to as MCU KO mice below) and Vav-iCre^+/− mice (referred to as WT mice below) were characterized. Under homeostatic condition, there was no difference in the number of HSCs in the BM (Extended Data Fig. 5a) and in the percentage of 7-AAD⁺ HSCs (Extended Data Fig. 5b). There was no difference in lineage differentiation in peripheral blood (Extended Data Fig. 5c). BM cellularity was unchanged (Extended Data Fig. 5d). Thus, MCU is not required for HSC maintenance and hematopoiesis under homeostatic condition. However, under transplantation-stimulated proliferation, MCU KO HSCs showed reduced engraftment in the BM of recipient mice (Extended Data Fig. 6a) and reduced capacity to reconstitute the blood system of lethally irradiated recipient mice (Extended Data Fig. 6b). BrdU labeling showed reduced proliferation of MCU KO HSCs compared to WT HSCs on cytokine-stimulated proliferation (Extended Data Fig. 6c). The frequency of 7-AAD⁺ HSCs was comparable between the two genotypes (Extended Data Fig. 6d). MCU KO HSCs had reduced Ca²⁺ signaling (Extended Data Fig. 6e), mitochondrial membrane potential (Extended Data Fig. 6f) and stress (Extended Data Fig. 6g). Together, these data suggest that mitochondrial Ca²⁺ influx is required for HSC proliferation.

To determine whether CD38 regulates HSC function by Ca²⁺ signaling and mitochondrial activation, we cultured HSCs from 3-month-old WT and CD38 KO mice treated with or without MCU short hairpin RNA (shRNA) for 2 days in the presence of cytokines. While CD38 KO HSCs showed reduced mitochondrial Ca²⁺ levels, mitochondrial membrane potential, number of HSCs and colony-forming ability compared to WT HSCs in the absence of MCU shRNA, these differences were blunted in the presence of MCU shRNA (Extended Data Fig. 7a–d), suggesting that CD38 regulates HSC function via Ca²⁺ signaling and mitochondrial activation.

CD38 is upregulated in HSCs during aging

NAD⁺ levels are reduced in HSCs during aging³⁰. The cellular NAD⁺ pool is maintained by several biosynthesis pathways from NAD⁺ precursors (the Preiss–Handler, de novo biosynthesis and salvage pathways) (Extended Data Fig. 8a)³¹. The major NAD⁺-consuming enzymes include sirtuins, poly (ADP-ribose) polymerases (PARPs) and CD38. We examined the expression of NAD⁺ metabolic enzymes in HSCs of young (3-month-old) and aged (24-month-old) WT mice. The expression of several enzymes for NAD⁺ biosynthesis (NAMPT, NAPRT, NMNAT3) and consumption (PARP1) was unchanged (Extended Data Fig. 8b). The expression of several sirtuins was reduced in aged HSCs^2,7,10, while the expression of quinolinate phosphoribosyltransferase (QPRT) was slightly increased in aged HSCs (Extended Data Fig. 8b). Because sirtuins are NAD⁺-consuming enzymes and QPRT is a key enzyme in de novo NAD⁺ biosynthesis, these changes are unlikely to contribute to the reduced NAD⁺ levels during aging. The expression of CD38, an NAD⁺-consuming enzyme, was increased in aged HSCs (Extended Data Fig. 8b–d), suggesting that increased CD38 expression may contribute to a decline in NAD⁺ levels in HSCs during aging. Aged HSCs had increased Ca²⁺ signaling (Extended Data Fig. 8e), reduced mitochondrial membrane potential (Extended Data Fig. 8f) and increased mitochondrial stress (Extended Data Fig. 8g) on cytokine-stimulated proliferation, which is consistent with CD38 activation during HSC aging.

CD38 upregulation is a driver of HSC aging

We sought to determine whether CD38 upregulation is a driver of HSC aging. HSC aging is characterized by myeloid-biased differentiation, reduced regenerative capacity per cell and defective hematopoiesis^2,32–36. Compared to age-matched WT controls, aged (24-month-old) CD38 KO mice had increased NAD⁺ levels in HSCs (Fig. 3a), increased white blood cell count and lymphocyte count in the peripheral blood (Fig. 3b,c), and alleviated myeloid-biased differentiation in peripheral blood (Fig. 3d). BM cellularity was increased in aged CD38 KO mice (Fig. 3e). In a competitive transplantation assay, HSCs from aged CD38 KO mice exhibited improved engraftment in the BM of recipient mice (Fig. 3f) and improved regenerative capacity to reconstitute the blood system of lethally irradiated mice (Fig. 3g). The recipient mice of old CD38 KO HSCs showed ameliorated donor-derived, myeloid-biased differentiation in peripheral blood (Fig. 3h). Thus, aged CD38 KO mice are protected from HSC aging. RNA-seq analyses of HSCs derived from aged WT and CD38 KO mice showed reduced expression of ribosomal proteins in aged CD38 KO HSCs (Extended Data Fig. 9a,b and Supplementary Table 1). Increased expression of ribosomal proteins is a molecular signature of HSC aging³⁷. Therefore, reduced expression of ribosomal proteins in aged CD38 KO HSCs is consistent with the reversal of HSC aging.

Fig. 3 |. CD38 inactivation prevents HSC aging.

a–h, Comparison of 24-month-old WT and CD38 KO mice. a, NAD⁺ levels in enriched HSCs (P = 0.0454). n = 5 and 6. b,c, White blood cell (WBC) (b) and lymphocyte (c) analyses (P = 0.0182 (b); P = 0.0193 (c)). n = 8 and 7. d, Lineage differentiation in peripheral blood was analyzed using flow cytometry (P = 0.0392). n = 6 and 7. e, BM cellularity normalized according to body weight (P = 0.0472). n = 6 and 8. f–h, Competitive transplantation using HSCs isolated from 24-month-old WT and CD38 KO mice as donors. Donor-derived HSC engraftment in the BM (P = 0.0311) (f), the percentage of total donor-derived cells in peripheral blood (4 weeks, P = 0.0647; 8 weeks, P = 0.0313; 12 weeks, P = 0.0090; 16 weeks, P = 0.0104) (g) and donor-derived lineage differentiation in the peripheral blood (P = 0.0107) (h) of the recipients were analyzed using flow cytometry. f, n = 6. g,h, n = 12 and 14. Data are presented as the mean ± s.e. *P < 0.05. **P < 0.01. A one-sided Student’s t-test was used.

We further took a pharmacological approach to determine the role of CD38 in HSC aging. We treated 24-month-old WT mice with 78c, a small-molecule inhibitor of CD38 (refs. 21,38), for 8 weeks (Fig. 4a). Treatment with 78c increased the NAD⁺ levels in HSCs (Fig. 4b), increased the white blood cell and lymphocyte count in peripheral blood to the levels equivalent to those of young mice (Fig. 4c–h), increased BM cellularity (Fig. 4i), reversed myeloid-biased differentiation in peripheral blood (Fig. 4j), decreased the frequency of myeloid-biased HSCs in the BM (Fig. 4k) and increased the number of colony-forming cells in the BM (Fig. 4l,m). The mitochondrial membrane potential and stress in HSCs were reduced by 78c treatment (Fig. 4n,o). Collectively, these data indicate that CD38 upregulation drives HSC aging and the pathophysiological changes of the aging hematopoietic system.

Fig. 4 |. 78c treatment reverses HSC aging.

a–o, Twenty-four-month-old WT mice were treated with or without 78c for 8 weeks. a, Schematic illustration of the experimental setup and timeline. b, NAD⁺ levels in enriched HSCs (P = 0.0002). n = 6. c–h, Lymphocyte and WBC analyses of young and aged mice before treatment (c,f) (P = 0.0009 (c); P = 0.0561 (f)) and aged mice after treatment for 5 weeks (d,g) (P = 0.0387 (d); P = 0.0888 (g)) or 8 weeks (e,h) (P = 0.0142 (e); P = 0.0459 (h)). c,f, n = 10 and 25. d,g, n = 9. e,h, n = 10 and 9. i, BM cellularity normalized according to body weight (P = 0.0485). n = 6. j, Lineage differentiation in peripheral blood was analyzed using flow cytometry (P = 0.0474). n = 9. k, Lineage-biased HSCs in the BM were analyzed using flow cytometry. Myeloid-biased HSCs: Lin⁻c-Kit⁺Sca1⁺CD135⁻CD34⁻CD150^hi. Lineage-balanced HSCs: Lin⁻c-Kit⁺Sca1⁺CD135⁻CD34⁻CD150^lo (P = 0.0029). n = 6. l,m, Colonies formed in a colony-forming assay using BM cells (l) and quantification (P = 0.0038) (m). n = 6 and 5. n,o, Staining for TMRM (P = 0.0490) (n) and ROS (P = 0.0362) (o) of HSCs was analyzed using flow cytometry. n, n = 5 and 6. o, n = 5. Data are presented as the mean ± s.e. *P < 0.05, **P < 0.01, ***P < 0.001. A one-sided Student’s t-test was used. Ctrl, control.

Alleviated mitochondrial stress in HSCs of aged CD38 KO mice

We investigated whether CD38 upregulation drives HSC aging due to reduced NAD⁺ metabolism. Several members of the sirtuin family of NAD ⁺-dependent deacetylases (SIRT2, SIRT3 and SIRT7) are critical regulators of HSC aging that govern mitochondrial stress management programs and prevent HSCs from undergoing caspase 1-mediated cell death^2,7,10. Aged CD38 KO HSCs showed reduced mitochondrial membrane potential and oxidative stress (Extended Data Fig. 9c,d), reduced expression of HSP60, a marker of mitochondrial protein folding stress (Extended Data Fig. 9e), reduced percentage of 7-AAD⁺ HSCs (Extended Data Fig. 9f), reduced percentage of activated caspase 1⁺ HSCs (Extended Data Fig. 9g) and comparable HSC proliferation (Extended Data Fig. 9h), which is consistent with elevated NAD⁺ signaling and sirtuin activities.

Protection of HSC aging in MCU KO mice

To determine the contribution of Ca²⁺ signaling to HSC aging, we characterized 24-month-old MCU KO mice. Aged MCU KO mice showed increased white blood cell and lymphoid cell count in peripheral blood (Extended Data Fig. 10a,b) and increased BM cellularity (Extended Data Fig. 10c). In a competitive transplantation assay, HSCs from aged MCU KO mice showed improved regenerative capacity to reconstitute the blood system of lethally irradiated recipient mice (Extended Data Fig. 10d) and ameliorated myeloid-biased differentiation (Extended Data Fig. 10e). These data suggest that reducing mitochondrial Ca²⁺ signaling protects against HSC aging.

Discussion

Our study established CD38 as a critical regulator of HSC fate decision balancing NAD⁺ metabolism and Ca²⁺ signaling to fine-tune mitochondrial activity and stress management (Extended Data Fig. 10f). Because female mice show a more pronounced myeloid-biased differentiation with aging, we primarily characterized female mice in this study. Mitochondrial activation and stress management were thought to be regulated independently. Our findings suggest that these two processes are more closely linked than previously thought. The existence of such a mechanism in HSCs highlights the importance of fine-tuning mitochondrial activity in HSC fate decisions. CD38 is required for Ca²⁺ signaling, mitochondrial activity and proliferation of HSCs at a young age. However, when CD38 is upregulated in old age, it tips the balance between Ca²⁺ signaling-mediated mitochondrial activity and NAD⁺ signaling-mediated mitochondrial stress management governed by sirtuins, leading to aging-associated HSC decline and pathophysiological changes of the aging hematopoietic system. Thus, CD38 must be kept in check to support HSC maintenance, which is in keeping with the observation that the CD38^lo cell population is enriched for HSCs²².

A recent study examined CD38 in dormant HSCs³⁹. Ibneeva et al.³⁹ used CD38 KO mice and their results were consistent with our findings, that is, CD38 KO HSCs showed reduced reconstitution on BM transplantation and there was no difference in HSC proliferation between young WT and CD38 KO mice under homeostatic conditions. However, most experiments in the study by Ibneeva et al. were based on comparison of CD38^hi and CD38^lo HSCs, which may differ in more than just CD38 expression. Indeed, Ibneeva et al. showed that while there was no difference in HSC proliferation between young WT and CD38 KO mice under homeostatic conditions, CD38^lo HSCs were more proliferative than CD38^hi HSCs under the same condition, indicating that the difference in proliferation between CD38^lo and CD38^hi HSCs is not due to CD38 expression. They further showed that CD38^lo HSCs were more proliferative than CD38 KO HSCs, again suggesting that increased proliferation of CD38^lo HSCs is not due to CD38 expression.

The age-dependent effects of CD38 and MCU on HSCs are striking. The sharp contrast in the requirement of CD38 and MCU for supporting HSC function between young and old ages is consistent with the notion that mitochondrial metabolism is essential to support HSC proliferation; yet, increased mitochondrial stress in old HSCs drives their deterioration^5,6. These observations also beg the question as to when and how the metabolic checkpoint becomes dysregulated along the lifespan. Importantly, pharmacological inhibition of CD38 reverses HSC aging and pathophysiological changes of the aging hematopoietic system, giving hope for the therapeutic potential of the NAD⁺ metabolic checkpoint regulating HSC activation and aging. Deletion of MCU is sufficient to prevent HSC aging, linking Ca²⁺ signaling to HSC aging and providing a nodal control point for intervention.

Our findings have important implications in understanding the current CD38-targeting treatments and informing potential CD38-targeting treatments. Anti-CD38 antibody therapy is transforming the treatment of multiple myeloma because of its anti-myeloma efficacy and manageable safety profile²⁰. Our findings suggest that anti-CD38 antibody treatment could reverse HSC aging and the pathophysiological changes of the aging immune system, contributing to its anti-myeloma efficacy and safety profile. CD38 is a malignancy marker for leukemia¹⁸. Increased expression of CD38 in aged HSCs may account for increased susceptibility of older individuals to leukemia.

Methods

Mice

CD38 KO mice (C57BL/6) have been described previously⁴⁰. B6.SJL-Ptprc^a Pepc^b/BoyJ mice, Vav-iCre mice (C57BL/6) and MCU^loxP/loxP mice (C57BL/6) were obtained from the Jackson Laboratory. For experiments using young and aged WT mice, C57BL/6 mice ranging from 3 to 24 months old were obtained from the National Institute on Aging. Female mice show more pronounced myeloid-biased differentiation with aging and were used in the study. All mice were housed on a 12–12 h light–dark cycle at 25 °C. Mice were provided with standard chow (PicoLab Rodent Diet 20, cat. no. 5053, LabDiet) and water ad libitum. Humidity was kept at 30–70%. On-site veterinarians provided health status checks. Animal procedures were performed in accordance with the University of California, Berkeley animal care committee. Mice were injected intraperitoneally with 5 mg kg⁻¹ poly(I:C) (Sigma-Aldrich) and were analyzed 16 h later.

Flow cytometry and cell sorting

BM cells were obtained by crushing the long bones with staining medium (sterile PBS without calcium and magnesium supplemented with 2% FCS). Lineage staining contained a cocktail of anti-mouse antibodies against Mac-1 (CD11b), Gr-1 (Ly-6G/C), Ter119 (Ly-76), CD3, CD4, CD8a (Ly-2) and B220 (CD45R) (all BioLegend) at 4 °C for 20 min. For detection or sorting of HSCs, we used Lin-APC-Cy7, c-Kit-APC, Sca-1-Pacific Blue, CD48-FITC and CD150-PE antibodies (all BioLegend). For congenic strain discrimination, anti-CD45.1-PerCP and anti-CD45.2-PE-Cy7 antibodies (both from BioLegend) were used. Antibodies were diluted 1:100. To assess cell death and cell cycle, 7-AAD or Ki-67 (both from BioLegend) staining were performed according to the manufacturer’s recommendations after cell surface staining. For intracellular activated caspase-1 staining, fluorescently labeled inhibitors of caspase probe staining (ImmunoChemistry Technologies) was performed as described previously¹⁰.

For intracellular calcium, mitochondrial calcium, mitochondrial membrane potential and mitochondrial superoxide levels, cells were incubated with 1 μM Fluo-4, AM (Thermo Fisher Scientific), 1 μM Rhod-2, AM (Thermo Fisher Scientific), 1 μM TMRM (Thermo Fisher Scientific) or 5 μM MitoSOX Red Mitochondrial Superoxide Indicator (Thermo Fisher Scientific), respectively, for 30 min at 37 °C in the dark after HSC surface staining. Cells were washed with staining medium before flow cytometry analysis. Data were collected on a Fortessa analyzer with the FACSDiva software (BD Biosciences); data analyses were performed with FlowJo v.10.4 (FlowJo LLC). For cell sorting, c-Kit enrichment was performed according to the manufacturer’s instructions (Miltenyi Biotec). Cells were sorted using an Aria Sorter (BD Biosciences).

HSC culture

For cytokine-stimulated HSC proliferation, enriched HSCs were cultured with StemSpan SFEM (STEMCELL Technologies) supplemented with 10% ES-Cult FCS (STEMCELL Technologies), 1% penicillin-streptomycin (Invitrogen), interleukin-3 (IL-3) (20 ng ml⁻¹), interleukin-6 (IL-6) (20 ng ml⁻¹), thrombopoietin (TPO) (50 ng ml⁻¹), Flt3l (50 ng ml⁻¹) and stem cell factor (SCF) (100 ng ml⁻¹) (PeproTech). For HSC proliferation, cells were pulsed with 10 μM BrdU (Invitrogen) for 18 h at 37 °C. BrdU-pulsed cells were resuspended in 100 μl staining medium. After HSC surface staining, cells were washed with staining medium, resuspended in 0.5 ml of fixation buffer (BioLegend) and incubated at room temperature for 20 min. Cells were washed with 1× perm/wash buffer (BioLegend). Then, 30 μl DNase I were added to 170 μl resuspended cells and incubated at 37 °C for 60 min. Cells were washed with 1× perm/wash buffer. Then, 2 μl of diluted anti-BrdU antibody (1:25 dilution of stock in 1× perm/wash buffer) was added to 100 μl of resuspended cells and incubated at room temperature for 30 min. Cells were washed with 1× perm/wash buffer before flow cytometry analysis.

Long-term HSC culture with PVA was performed as described elsewhere²⁴. Briefly, 50 HSCs (Lin⁻c-Kit⁺Sca1⁺CD150⁺CD34⁻) were sorted into a fibronectin-treated 96-well plate (Corning) and cultured with F-12 medium (Thermo Fisher Scientific), 1% ITSX (Thermo Fisher Scientific), 1% P/S/G, 10 mM HEPES, 100 ng ml⁻¹ mouse TPO, 10 ng ml⁻¹ mouse SCF and 0.1% PVA (Sigma-Aldrich). Complete medium was changed on day 5. Cells were counted using a hemocytometer after 7 days.

Transplantation

For noncompetitive transplantation, 2 × 10⁶ BM cells from CD45.2 donors were injected into lethally irradiated (950 Gy) CD45.1 B6.SJL recipient mice. The BM cells of recipient mice were analyzed 1 month after transplantation.

For competitive transplantation, 250 freshly sorted HSCs from CD45.2 donor mice were mixed with 5 × 10⁵ CD45.1 B6.SJL competitor BM cells and injected into lethally irradiated B6.SJL recipient mice. To assess the multilineage reconstitution of transplanted mice, peripheral blood was collected every month for 4 months. Red blood cells were lysed using FACS lysing solution (BD Biosciences) for 5 min at room temperature, washed with PBS and stained with an antibody cocktail consisting of CD45.2-PE-Cy7, CD45.1-PerCP-Cy5.5, Mac1-PE, Gr1-FITC, B220-APC and CD3-Pacific Blue (BioLegend) for 20 min at 4 °C. Cells were washed with PBS before the flow cytometry analyses. BM cells were analyzed 4 months after transplantation.

Homing

A total of 2 × 10⁶ BM cells from CD45.2 donors were injected into lethally irradiated (950 Gy) CD45.1 B6.SJL recipient mice; 16 h after transplantation, the BM cells of recipients were stained with CD45.2-PE-Cy7 antibody before the flow cytometry analyses.

Lentiviral production

CD38 was cloned into the pFUGw lentiviral construct. MCU shRNA was cloned into the pLKO.5 lentiviral construct (TRCN0000251263, Sigma-Aldrich). 293T cells were transfected with lentiviral packaging plasmids together with the pFUGw or pLKO.5 vectors using PEI Max (Polysciences) according to the manufacturer’s protocol. Then, 24 h after transfection, cells were changed to fresh medium. Forty-eight hours after transfection, the virus was collected and filtered through a 45-μm syringe filter. Fresh medium was added back to cells; the second round of virus collection was conducted 72 h after transfection. Lentivirus was concentrated using centrifugation at 17,900g, at 4 °C for 90 min and resuspended in HSC medium (StemSpan SFEM, STEMCELL Technologies) supplemented with 10% ES-Cult FCS (STEMCELL Technologies), 1% penicillin-streptomycin, IL-3 (20 ng ml⁻¹), IL-6 (20 ng ml⁻¹), TPO (50 ng ml⁻¹), Flt3l (50 ng ml⁻¹) and SCF (100 ng ml⁻¹) (PeproTech).

Lentiviral transduction of HSCs

Freshly isolated HSCs were cultured with HSC medium for 3 h at 37 °C. Lentiviral medium was added to HSCs in a 96-well round-bottom plate and centrifuged for 90 min at 270g in the presence of 10 μg ml⁻¹ polybrene. After centrifugation, the lentiviral medium was replaced by fresh HSC medium. This process was repeated 24 h later.

78c treatment

78c was administered to mice via intraperitoneal injection (10 mg kg⁻¹ per dose) twice daily over a period of 8 weeks. Body weight was measured weekly. Control mice received vehicle (5% dimethylsulfoxide, 15% polyethylene glycol 400, 80% of 15% hydroxypropyl-γ-cyclodextrin (in citrate buffer, pH 6.0)) injection.

mRNA analysis

RNA was isolated from cells using TRIzol reagent (Invitrogen) according to the manufacturer’s instructions. Complementary DNA was generated using the qScript cDNA SuperMix (Quanta Biosciences) according to the manufacturer’s instructions. Gene expression was determined using real-time PCR with the Eva qPCR SuperMix Kit (BioChain Institute) on an ABI StepOnePlus system. The following primers were used: CD38, forward: 5′-GGACGAAACACCGGCCATGTAAC-3′; reverse: 5′-AACCTTCCCGACTTAGGGGC-3′; HSP60, forward, 5′-ACCTGTGACAACCCCTGAAG; reverse: 5′-TGACACCCTTTCTTCCAACC-3′; PARP1, forward: 5′-GGCAGCCTGATGTTGAGGT-3′; reverse: 5′-GCGTACTCCGCTAAAAAGTCAC-3′; NAMPT, forward: 5′-GCAGAAGCCGAGTTCAACATC-3′; reverse: 5′-TTTTCACGGCATTCAAAGTAGGA-3′; NAPRT, forward: 5′-TGCTCACCGACCTCTATCAGG-3′; reverse: 5′-CGAAGGAGCCTCCGAAAGG-3′; NMNAT3, forward: 5′-CCTGTGGTTCCTTCAACCCC-3′; reverse: 5′-AGATGATGCCCTCAATCACCT-3′; and QPRT, forward: 5′-CCGGGCCTCAATTTTGCATC-3′; reverse: 5′-GGTGTTAAGAGCCACCCGTT-3′.

RNA-seq analysis

The total RNA of sorted enriched HSCs was isolated with the RNeasy Plus Micro Kit (QIAGEN) according to the manufacturer’s instructions. RNA-seq libraries were prepared using QuantSeq 3′ mRNA-Seq Library Prep Kit FWD (Lexogen); sequencing was performed on an HiSeq 4000 system. Data analysis was processed within the Galaxy public server (https://usegalaxy.org/). Briefly, after removing low-quality reads and adapter sequences, data were mapped against the mouse genome (mm10) using HISAT2 (Galaxy v.2.2.1). Gene expression levels were measured from feature counts using HTSeq (Galaxy v.0.9.1). Differential gene expression was analyzed using DESeq2 (Galaxy v.1.34.0). Differentially expressed genes (P_adj < 0.1, Benjamini–Hochberg-corrected) between CD38 KO and WT samples were uploaded to the online Database for Annotation, Visualization and Integrated Discovery v.6.8 (https://david.ncifcrf.gov/) for functional annotations. Differentially expressed genes were also subjected to GSEA using the GSEA v.4.3.2 software (https://www.gsea-msigdb.org/gsea/index.jsp).

Metabolomics sample preparation

In vivo.

BM cells were obtained by crushing the long bones with ice-cold sterile PBS without calcium and magnesium and were subjected to lysis using ice-cold ACK lysis buffer for 5 min on ice. This was followed by c-Kit enrichment and HSC surface staining as described above. After staining, the cell pellet was washed with ice-cold PBS and resuspended in 1 ml ice-cold saline (9 g l⁻¹ NaCl in deionized water) to prevent phosphate-induced ion suppression in MS. The sorter was configured with a 70-μm nozzle tip and the sheath fluid consisted of 5 g l⁻¹ NaCl in deionized water. A total of 5 × 10⁴ enriched HSCs were sorted directly into 50 μl ice-cold acetonitrile (ACN) for rapid quenching and to extract metabolites. All procedures were carried out at 4 °C to minimize metabolic changes during sample preparation.

Ex vivo.

A total of 5 × 10⁴ enriched HSCs were cultured in glucose-free Roswell Park Memorial Institute 1640 medium (Thermo Fisher Scientific) supplemented with 10% dialyzed FCS (Cytiva), 1% penicillin-streptomycin, IL-3 (20 ng ml⁻¹), IL-6 (20 ng ml⁻¹), TPO (50 ng ml⁻¹), Flt3l (50 ng ml⁻¹), SCF (100 ng ml⁻¹) (PeproTech) and 25 mM [U-¹³C]-glucose (cat. no. CLM-1396–2, Cambridge Isotope Laboratories) at 37 °C for 20 h. After HSC surface staining, cells were washed with ice-cold PBS and resuspended in 0.5 ml ice-cold saline (9 g l⁻¹ NaCl in deionized water). The sorter was configured with a 70-μm nozzle tip and the sheath fluid consisted of 5 g l⁻¹ NaCl in deionized water. A total of 5 × 10⁴ enriched HSCs were sorted directly into 50 μl ice-cold ACN.

Metabolite measurements using LC–MS

Samples were centrifuged at 16,000g for 10 min at 4 °C; 30 μl of supernatant were transferred to MS vials. A quadrupole orbitrap mass spectrometer (Q Exactive, Thermo Fisher Scientific) operating in negative ion mode was coupled to a Vanquish UHPLC system (Thermo Fisher Scientific) with electrospray ionization and used to scan from m/z 70 to 300 at 2 Hz, with a 140,000 resolution. LC separation was achieved on an XBridge BEH Amide Column (2.1 × 150 mm², 2.5-μm particle size, 130-Å pore size, Waters Corporation) using a gradient of solvent A (95:5 water: ACN with 20 mM ammonium acetate and 20 mM ammonium hydroxide, pH 9.45) and solvent B (ACN). The flow rate was 150 μl min⁻¹. The LC gradient was: 0 min, 85% B; 2 min, 85% B; 3 min, 80% B; 5 min, 80% B; 6 min, 75% B; 7 min, 75% B; 8 min, 70% B; 9 min, 70% B; 10 min, 50% B; 12 min, 50% B; 13 min, 25% B; 16 min, 25% B; 18 min, 0% B; 23 min, 0% B; 24 min, 85% B; and 30 min, 85% B. The autosampler temperature was 5 °C and the injection volume was 15 μl. Data were analyzed using the maven software (build 682; http://maven.princeton.edu/index.php). Natural isotope correction was performed with AccuCor2 R code (https://github.com/wangyujue23/AccuCor2).

ATP quantification

A total of 1 × 10⁴ enriched HSCs were resuspended in 100 μl PBS and equilibrated at room temperature for 30 min before the same volume of Cell Titer-Glo reagent (Promega Corporation) was added. After 10 min incubation at room temperature, the intensity of luminescence was measured using a SpectraiMax i3 Microplate Reader (Molecular Devices).

NAD⁺ quantification

A total of 1 × 10⁴ enriched HSCs were resuspended in 50 μl PBS; the same volume of NAD/NADH-Glo Detection Reagent (Promega Corporation) was added according to the manufacturer’s instructions. After 60 min incubation at room temperature, the intensity of luminescence was measured using the SpectraiMax i3 Microplate Reader.

Colony-forming assay

A total of 2 × 10⁴ BM cells were used for the colony formation assay in MethoCult GF M3434 (STEMCELL Technologies) according to the manufacturer’s instructions. The colony-forming units were quantified on day 12.

Complete blood count

Peripheral blood was collected via submandibular bleeding into EDTA-treated blood collection tubes. Complete blood count analysis was conducted at the Comparative Pathology Laboratory at the University of California, Davis.

Statistics and reproducibility

The number of mice chosen for each experiment was based on the principle that the minimum number of mice should be used to have sufficient statistical power and was comparable to the published literature for the same assays performed. No statistical method was used to predetermine sample size. Data points were excluded if they were identified as outliers using the Grubbs’ test. No randomization was used to allocate mice to the experimental groups. Analyses of mice and tissue samples were performed by investigators blinded to the treatment or genetic background of the animals. Measurements were taken from distinct samples from different mice. Statistical analysis was performed with Student’s t-tests (in Excel), unless specified otherwise. Data distribution was not tested. Data are presented as the mean while the error bars represent the s.e. In all corresponding figures, *P < 0.05, **P < 0.01, ***P < 0.001, NS P > 0.05. Information about replication is indicated in the figures and figure legends.

Extended Data

Extended Data Fig. 1 |. CD38 is not required for HSC maintenance under homeostatic condition at young age.

a-i. Comparison of 3-month-old WT and CD38 KO mice. A. Gating strategy for HSCs. Enriched HSCs: Lin⁻c-Kit⁺Sca1⁺. Highly enriched HSCs: Lin⁻c-Kit⁺Sca1⁺CD150⁺CD48⁻. B, C. The number of enriched (p = 0.4210) (B) and highly enriched (p = 0.4955) (C) HSCs in the bone marrow were analyzed via flow cytometry. n = 6. D, E. Ki-67 (p = 0.2928) (D) and 7-AAD (p = 0.2349) (E) staining of HSCs were analyzed via flow cytometry. n = 6. F. Lineage differentiation in the peripheral blood was analyzed via flow cytometry. MNCs, mononuclear cells (p = 0.4013). n = 5 and 6. G, H. Complete blood count analyses (G, p = 0.4174; H, p = 0.1768). n = 6. I. Bone marrow cellularity normalized by body weight (p = 0.0718). n = 6 and 5. Data are presented as mean values +/− SE. ns: p > 0.05. One-sided student’s t test.

Extended Data Fig. 2 |. CD38 regulates Ca²⁺ signaling and mitochondrial activity upon HSC proliferation.

a. HSCs isolated from 3-month-old WT and CD38 KO mice were cultured in the presence of PVA for 7 days. Total cell number was counted by hemocytometer (p = 0.0026). n = 9. b. Flow cytometry analyses of Ki-67 staining of HSCs isolated from 3-month-old WT and CD38 KO mice 16 hours post pIpC treatment (p = 0.0229). n = 6. c. HSCs isolated from 3-month-old CD38 KO mice were transduced with lentivirus overexpressing CD38 or control lentivirus and were used as donors in competitive transplantation. The percentage of total donor-derived cells in the peripheral blood of the recipients was analyzed by flow cytometry (4 weeks p = 0.0652; 8 weeks p = 0.0059; 12 weeks p = 0.0085; 16 weeks p = 0.0260). n = 12. d-n. Noncompetitive transplantation was performed using bone marrow cells from 3-month-old WT and CD38 KO mice as donors. Donor-derived enriched HSCs from the recipient mice were compared. d, e. Relative abundances of metabolites were measured by LC-MS (D, p = 0.0273; E, p = 0.0105). n = 6 and 7(D). n = 10 (E). f. ATP levels (p = 0.0007). n = 4. g-n. Representative histograms and quantification for Fluo-4 (p = 0.0357) (G, H), Rhod-2 (p = 0.0291) (I, J), TMRM (p = 0.0389) (K, L), and ROS (p = 0.0367) (M, N) staining analyzed via flow cytometry. n = 6 and 5 (G, H). n = 5 and 6 (I, J), n = 6 (K, L), n = 5 (M, N). Data are presented as mean values +/− SE. *: p < 0.05. **: p < 0.01. ***: p < 0.001. One-sided student’s t test.

Extended Data Fig. 3 |. CD38 regulates Ca²⁺ signaling and mitochondrial activity upon HSC proliferation.

a–c. Flow cytometry analyses of Rhod-2 (p = 0.0049) (A), TMRM (p = 0.0400) (B), ROS (p = 0.0497) (C) staining of HSCs isolated from 3-month-old WT and CD38 KO mice 16 hours post pIpC treatment. n = 5 and 6 (A, C). n = 5 (B). d. Gating strategy. e, f. Flow cytometry analyses of TMRM (p = 0.0265) (E) and ROS (p = 0.0137) (F) staining of CD38^high and CD38^low HSCs isolated from 3-month-old WT mice cultured for 18 hours in the presence of cytokines. n = 6 and 5 (E). n = 6 (F). Data are presented as mean values +/− SE. *: p < 0.05. **: p < 0.01. One-sided student’s t test.

Extended Data Fig. 4 |. Effect of CD38 on Ca²⁺ signaling and mitochondrial activity under homeostatic condition.

a-h. Comparison of enriched HSCs derived from 3-month-old WT and CD38 KO mice. A, B. Representative histograms (A) and quantification (p = 0.3041) (B) for Fluo-4 staining analyzed via flow cytometry. MFI, mean fluorescence intensity. n = 6. c, d. Representative histograms (C) and quantification (D) for Rhod-2 staining analyzed via flow cytometry (p = 0.2955). n = 6. e, f. Representative histograms (E) and quantification (F) for ROS staining analyzed via flow cytometry (p = 0.3350). n = 6. g, h. Representative histograms (G) and quantification (H) for TMRM staining analyzed via flow cytometry (p = 0.1819). n = 6. Data are presented as mean values +/− SE. ns: p > 0.05. One-sided student’s t test.

Extended Data Fig. 5 |. MCU is not required for HSC maintenance under homeostatic condition at young age.

a-d. Comparison of 7-month-old WT and MCU KO mice. A. HSC number in the bone marrow analyzed via flow cytometry (p = 0.2488). n = 6. B. 7-AAD staining of HSCs analyzed via flow cytometry (p = 0.2157). n = 6. C. Lineage differentiation in the peripheral blood analyzed via flow cytometry. MNCs, mononuclear cells (p = 0.2453). n = 10 and 11. D. Bone marrow cellularity normalized by body weight (p = 0.1160). n = 6. Data are presented as mean values +/− SE. ns: p > 0.05. One-sided student’s t test.

Extended Data Fig. 6 |. MCU is required to support HSC proliferation.

a, b. Competitive transplantation using HSCs isolated from 7-month-old WT and MCU KO mice as donors. The percentage of donor-derived HSCs in the bone marrow of the recipients (p = 0.0267) (A) and the percentage of donor-derived cells in the peripheral blood of the recipients (4 weeks p = 0.0425; 8 weeks p = 0.0101; 12 weeks p = 0.0142; 16 weeks p = 0.0090) (B) were quantified by flow cytometry. n = 6 (A). n = 14 (B). c-g. Enriched HSCs isolated from 3-month-old WT and MCU KO mice were cultured in the presence of cytokines. BrdU incorporation (p = 0.0477) (C), 7-AAD (p = 0.0984) (D), Rhod-2 (p = 0.0031) (E), TMRM (p = 0.0445) (F), and ROS (p = 0.0278) (G) staining of HSCs were analyzed by flow cytometry. n = 6 (C, D). n = 5 (E-G). Data are presented as mean values +/− SE. *: p < 0.05. **: p < 0.01. ns: p > 0.05. One-sided student’s t test.

Extended Data Fig. 7 |. CD38 regulates HSC function via Ca²⁺ signaling and mitochondrial activity.

a-d. HSCs from 3-month-old WT and CD38 KO mice were treated with or without MCU shRNA for 48 hours in culture in the presence of cytokines before flow cytometry analyses of Rhod2 staining (Ctrl, p = 0.0239; MCU shRNA, p = 0.4907) (A), TMRM staining (Ctrl, p = 0.0334; MCU shRNA, p = 0.0720) (B), HSC number (Ctrl, p = 0.0002; MCU shRNA, p = 0.2252) (C), and colony forming assay (Ctrl, p = 0.0250; MCU shRNA, p = 0.2180) (D). n = 3. Data are presented as mean values +/− SE. *: p < 0.05. ***: p < 0.001. ns: p > 0.05. One-sided student’s t test.

Extended Data Fig. 8 |. CD38 is upregulated in HSCs during aging.

a. Schematic illustration of the NAD⁺ metabolic pathways. b-g. Comparison of HSCs isolated from young (3 months old) and old (24 months old) WT mice. B. Quantitative real-time PCR analyses of NAD⁺ metabolic enzymes (PARP1, p = 0.4431; NAMPT, p = 0.1517; NAPRT, p = 0.1546; NMNAT3, p = 0.3542; QPRT, p = 0.0419; CD38, p = 0.0010). n = 6 and 7 (CD38). n = 3 and 4 (other enzymes). C, D. Representative histograms (C) and quantification (D) for CD38 staining analyzed via flow cytometry (p = 0.0472). n = 4. E-G. Flow cytometry analyses of Fluo-4 (p = 0.0269) (E), TMRM (p = 0.0005) (F), and ROS (p = 0.0001) (G) staining. n = 5. Data are presented as mean values +/− SE. *: p < 0.05. **: p < 0.01. ***: p < 0.001. ns: p > 0.05. One-sided student’s t test.

Extended Data Fig. 9 |. CD38 inactivation prevents HSC aging.

a-h. Comparison of HSCs isolated from 24-month-old WT and CD38 KO mice. A, B. RNA-sequencing analyses of enriched HSCs. Volcano plot summarizing the differentially expressed genes (Red: upregulated in CD38 KO. Blue: downregulated in CD38 KO. P adj<0.1. Benjamini-Hochberg procedure). Genes related to ribosomal proteins are labeled (A). Gene ontology analysis for the biological functions of differentially expressed genes (B). n = 2. C, D. Flow cytometry analyses of TMRM (p = 0.0076) (C) and ROS (p = 0.0058) (D) staining. n = 10 (C). n = 10 and 12 (D). E. Quantitative real-time PCR analysis of HSP60 (p = 0.0022). n = 3. F-H. Staining for 7-AAD (p = 0.0029) (F), activated caspase 1 (p = 0.0254) (G), and Ki-67 (p = 0.1974) (H) were analyzed by flow cytometry. n = 6 (F). n = 6 and 5 (G). n = 5 and 6 (H). Data are presented as mean values +/− SE. *: p < 0.05. **: p < 0.01. ns: p > 0.05. One-sided student’s t test.

Extended Data Fig. 10 |. MCU inactivation prevents HSC aging.

a-e. Comparison of 24-month-old WT and MCU KO mice. A, B. Complete blood count analyses (A, p = 0.0190; B, p = 0.0122). n = 8. C. Bone marrow cellularity normalized by body weight (p = 0.0181). n = 7 and 8. D, E. Competitive transplantation using HSCs isolated from 24-month-old WT and MCU KO mice as donors. The percentage of donor-derived cells in the peripheral blood of the recipients (4 weeks p = 0.0419; 8 weeks p = 0.0001; 12 weeks p = 0.0023; 16 weeks p = 0.0010) (D) and donor-derived lineage differentiation in the peripheral blood of the recipients (p = 0.0002) (E) were determined by flow cytometry. n = 12 and 9. f. A working model. CD38 catalyzes synthesis of Ca²⁺ second messengers from NAD⁺. At young age, CD38 is required for HSC proliferation through activating Ca²⁺ signaling and mitochondrial metabolism. NAD⁺ metabolism is required for activating sirtuins and suppressing mitochondrial stress. The balanced NAD⁺ metabolism and Ca²⁺ signaling ensure sufficient mitochondrial activation and adequate mitochondrial stress resistance. At old age, CD38 is upregulated, tipping the balance toward increased Ca²⁺ signaling and decreased NAD⁺ metabolism, and leading to uncontrolled mitochondrial activation and stress. Data are presented as mean values +/− SE. *: p < 0.05. **: p < 0.01. ***: p < 0.001. One-sided student’s t test.

Data availability

Sequencing data are deposited to the Gene Expression Omnibus under accession no. GSE268310. All data supporting the findings are available from the corresponding author. Source data are provided with this paper.