Dietary restriction mitigates 5-fluorouracil-induced thrombocytopenia in aged mice via mitochondrial potentiation in hematopoietic stem cells and megakaryocyte progenitors

Xingxing Qiu; Duozhuang Tang; Jiahao Lin; Zhiyang Chen; Rongrong Qiu; Qiao Wang; Siyao He; Shaotong Zhang; Qiang Zhan; Zhendong Tao; Liu Zhang; Si Tao

doi:10.1016/j.stemcr.2026.102869

. 2026 Apr 2;21(4):102869. doi: 10.1016/j.stemcr.2026.102869

Dietary restriction mitigates 5-fluorouracil-induced thrombocytopenia in aged mice via mitochondrial potentiation in hematopoietic stem cells and megakaryocyte progenitors

Xingxing Qiu ^1,^2,⁸, Duozhuang Tang ^3,^4,⁸, Jiahao Lin ⁵, Zhiyang Chen ⁵, Rongrong Qiu ¹, Qiao Wang ¹, Siyao He ¹, Shaotong Zhang ⁵, Qiang Zhan ⁵, Zhendong Tao ⁶, Liu Zhang ⁷, Si Tao ^1,^2,^9,^∗

PMCID: PMC13083804 PMID: 41932340

Summary

With population aging, chemotherapy-induced thrombocytopenia (CIT) is a severe complication in elderly cancer patients, yet effective preventive and therapeutic strategies remain limited. Here, we demonstrate that dietary restriction (DR) significantly mitigates 5-fluorouracil (5-FU)-induced thrombocytopenia and promotes platelet recovery in both young and aged mice. Mechanistically, DR improves mitochondrial homeostasis in hematopoietic stem and progenitor cells and enhances their hematopoietic reconstitution capacity. This preconditioning facilitates mitochondrial activation after chemotherapy, thereby promoting megakaryocytic lineage recovery. Pharmacological mitochondrial activation in ad libitum-fed mice mimics the protective effects of DR, whereas mitochondrial inhibition in DR-treated mice markedly attenuates these benefits. Clinically, cancer patients with lower pre-chemotherapy body mass index ([BMI] 18.5–22.95 kg/m²) showed a lower incidence of CIT following 5-FU treatment than those with higher BMI. Together, we show that short-term DR significantly mitigates CIT and that targeting mitochondria may represent a novel therapeutic strategy for CIT in elderly cancer patients.

Keywords: dietary restriction, chemotherapy, aging, thrombocytopenia, mitochondria

Graphical abstract

Highlights

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DR mitigates 5-FU-induced thrombocytopenia
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DR improves HSPC mitochondrial homeostasis and reconstitution capacity
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DR facilitates post-chemotherapy PLT recovery by promoting HSPC mitochondrial activation
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BMI is negatively correlated with nadir PLT counts and can independently predict CIT risk

Chemotherapy-induced thrombocytopenia (CIT) is a major challenge in aging cancer care. In this article, Tao et al. demonstrate that dietary restriction (DR) significantly mitigates CIT. DR improves mitochondrial homeostasis in hematopoietic stem and progenitor cells and enhances their reconstitution capacity. This preconditioning promotes mitochondrial activation after chemotherapy, thereby accelerating platelet recovery.

Introduction

Population aging remains a major challenge faced by many countries today (Wang et al., 2024; Mózes et al., 2025). Elderly individuals are more susceptible to a range of chronic diseases, particularly cardiovascular disorders and malignancies (Pacifici et al., 2019). By 2030, over 70% of patients diagnosed with malignant tumors are projected to be older than 65 years (Kadambi et al., 2020). However, elderly cancer patients are often less tolerant of standard-dose chemotherapy and more susceptible to severe complications (Soto-Perez-de-Celis et al., 2018). Chemotherapy-induced thrombocytopenia (CIT) is a common and serious complication in cancer patients undergoing chemotherapy (Xie et al., 2022). CIT has been reported in 10%–80% of patients, depending on the chemotherapy regimen (André et al., 2004; Weycker et al., 2019). Elderly patients often have comorbidities such as hypertension and cerebral infarction, placing them at higher risk of bleeding once CIT occurs (Kuter 2022). Therefore, effective management of CIT in older cancer patients undergoing chemotherapy is of particular importance.

Due to the complex mechanisms underlying CIT, along with the lack of reliable predictive models and standardized treatments, the outcomes of CIT are often worse than those of chemotherapy-induced leukopenia (Al-Samkari 2024). Current therapeutic strategies for CIT include platelet (PLT) transfusion, recombinant human thrombopoietin, thrombopoietin receptor agonists, and recombinant human interleukin-11 (Xie et al., 2022). However, these therapeutic approaches are often associated with several potential limitations that compromise their clinical efficacy, including transfusion-related allergic reactions and infections, delayed onset of action, suboptimal therapeutic responsiveness, and substantial economic burden (Vadhan-Raj et al., 2000). Thus, identifying a strategy that enhances chemotherapy tolerance, minimizes adverse effects, and is cost-effective, especially for elderly patients, remains an urgent clinical need.

Dietary restriction (DR), which involves reducing food intake by 10%–30% without causing malnutrition, has garnered attention for its broad health benefits (Tang et al., 2016). Numerous studies have shown that DR, as a non-pharmacological approach, can delay and reduce the incidence of age-related diseases in mammals, including cancer, type 2 diabetes, and cardiovascular diseases (Shimokawa et al., 1993; Mattson 2005; Cohen et al., 2009; Colman et al., 2009; Omodei and Fontana, 2011; Mattison et al., 2012; Qiu et al., 2023). Previous studies have demonstrated that DR markedly delays the aging of hematopoietic stem cells (HSCs) and promotes the maintenance of balanced lineage differentiation, enhanced hematopoietic reconstitution capacity, and improved self-renewal potential (Ertl et al., 2008; Maharajan et al., 2019; Tao et al., 2020, 2025). These effects may be closely associated with the attenuation of inflammatory signaling in HSCs and reduced circulating levels of insulin-like growth factor-1 (Cheng et al., 2014; Tang et al., 2016; Rasa et al., 2022). Previous studies have shown that mitochondrial regulation plays a central role in determining the fate of HSCs (Ansó et al., 2017; Takihara et al., 2019; Hinge et al., 2020). In the HSC compartment, subsets of HSCs with stronger self-renewal capacity are typically enriched in mitochondria (Totani et al., 2025), yet maintain relatively low mitochondrial membrane potential (MMP) (Liang et al., 2020). This characteristic is closely related to the fact that HSCs predominantly rely on glycolytic metabolism to sustain their self-renewal ability (Vannini et al., 2016; Yang et al., 2024), while increased mitochondrial metabolic activity is often accompanied by reduced self-renewal capacity (Hinge et al., 2020). However, autophagy can maintain HSCs in a quiescent state and preserve their self-renewal potential by clearing active or damaged mitochondria and inhibiting metabolic activity (Warr et al., 2013; Ho et al., 2017; Fang et al., 2020). Recent research has also demonstrated that short-term fasting followed by refeeding induces transient autophagy, maintains mitochondrial homeostasis, and thereby markedly enhances the regenerative potential of aged HSCs (Dellorusso et al., 2024). In the context of chemotherapy, intermittent fasting has been reported to mitigate cyclophosphamide-induced immunosuppression (Cheng et al., 2014). Furthermore, we have shown that short-term DR pre-chemotherapy significantly increased the abundance of Bacteroides in the gut, reduced intestinal injury induced by high-dose methotrexate and 5-fluorouracil (5-FU), prevented bacterial translocation, and markedly improved survival in aged mice (Tang et al., 2020, 2024). These findings suggest that caloric restriction (CR) exerts protective effects on HSC function and mitigates chemotherapy-induced intestinal injury and leukopenia. However, the potential of DR to modulate chemotherapy-induced hematologic toxicity, particularly CIT, has not been explored. Whether DR can mitigate CIT, a major clinical challenge, and offer a viable preventive or therapeutic approach remains unknown.

Chemotherapy affects multiple stages of PLT production, including the differentiation of hematopoietic stem and progenitor cells (HSPCs), the proliferation and maturation of megakaryocyte (MK) progenitors (MkPs), and PLT shedding and survival (McManus and Weiss, 1984; Xie et al., 2022). 5-FU and its analog capecitabine are cornerstone chemotherapeutic agents used widely for various solid tumors, such as colorectal, gastric, breast, and lung cancers (Tang et al., 2024). Approximately 19% of patients receiving 5-FU-based chemotherapy are reported to develop CIT (André et al., 2004). Previous studies suggest that 5-FU mainly induces CIT by inhibiting HSPC and MkP differentiation, thereby reducing PLT production (Kuter 2015; Lee et al., 2018; Zhu et al., 2018). However, the underlying mechanisms of CIT induction and effective interventions, particularly in the elderly, remain poorly understood.

This study established a 5-FU-induced thrombocytopenia model in young and aged C57BL/6J mice and implemented a 1-month 30% DR prior to chemotherapy. Our study demonstrates that short-term DR prior to chemotherapy effectively mitigates CIT and improves mitochondrial homeostasis in HSPCs and enhances their hematopoietic reconstitution capacity. This preconditioning facilitated mitochondrial activation in following chemotherapy, thereby promoting megakaryocytic lineage recovery. These findings suggest that mitochondria may serve as a potential therapeutic target for preventing and treating CIT in elderly cancer patients, offering new insights for clinical intervention.

Results

DR mitigates CIT

To investigate the effects of DR on PLT following chemotherapy, we utilized 12-week-old (approximately equivalent to 20-year-old humans) and 20- to 22-month-old mice (approximately 67- to 72-year-old humans) (Dutta and Sengupta, 2016). Mice were exposed to either a 30% DR or ad libitum (AL) diet for 30 days before intraperitoneal 5-FU injection, which was performed daily for 5 days (Figure 1A). Based on our previous findings, aged mice exhibit reduced tolerance to chemotherapeutic regimens compared with young mice (Tang et al., 2024). Therefore, in accordance with our prior studies, the chemotherapy dose for aged mice was reduced by 20% to better evaluate the protective effects of DR. As previously observed, both young and aged AL mice showed a sharp decline in survival following chemotherapy. In young AL mice, survival dropped to 33.3% by day 8 post-chemotherapy. In aged AL mice, survival remained at 86.7% on day 4 (13 mice alive), then sharply declined to 26.7% by day 5 (4 mice alive), and further to 6.7% by day 7, indicating a significantly reduced survival rate in aged mice. Notably, DR markedly improved survival compared to AL mice. In DR groups, survival in young mice reached a nadir of 86.7% on day 8, while in aged mice, the lowest survival rate was 80.0% on day 10, remaining stable until the end of the 30-day post-chemotherapy observation period. These results demonstrate that DR significantly enhanced survival in aged mice (Figures S1A and S1B). To further assess overall health status, body weights were monitored post-chemotherapy. Consistent with survival data, AL mice showed significant weight loss following 5-FU treatment, while body weight remained stable in DR mice. Of note, on day 4 post-chemotherapy, the surviving aged AL mice retained 88.09% ± 3.94% of their pre-chemotherapy body weight, with no individual losing more than 20% body weight or showing evident health deterioration (Figures S1C and S1D). However, by day 5, aged AL mice exhibited pronounced health decline and weight loss exceeding 20%, necessitating humane euthanasia. To avoid end-stage bias, day 4 post-chemotherapy was selected as the endpoint for subsequent analyses in aged AL mice.

Next, we analyzed peripheral hematological parameters following chemotherapy. In young mice, 5-FU significantly reduced PLT counts in AL mice to 271.4 ± 95.5 ×10³/μL on day 3 post-treatment (31.1% of baseline), reaching a nadir of 116.8 ± 47.6 10³/μL (13.4% of baseline) on day 6, after which recovery commenced. In contrast, DR mice showed significantly higher PLT counts: 576.0 ± 182.2 ×10³/μL (59.3% of baseline) on day 3 and 261.8 ± 51.6 ×10³/μL (26.9% of baseline) on day 6. PLT recovery was significantly accelerated in DR mice compared to AL mice (Figure 1B). Consistent with our previous findings (Tang et al., 2016), DR pre-chemotherapy reduced peripheral white blood cell (WBC) and lymphocyte counts in young mice. However, post-chemotherapy reductions in WBC and lymphocyte counts were more pronounced in AL mice than in DR mice counterparts (Figure S1E). Although both red blood cells (RBCs) and hemoglobin (HGB) declined significantly following 5-FU treatment, DR mice consistently showed higher levels than AL mice up to day 12 (Figure S1F). In aged mice, 5-FU treatment caused a marked reduction in PLT counts in AL mice as early as day 1, with further decline by day 4 (395.40 ± 185.50 ×10³/μL; 33.47% of baseline). Interestingly, PLT counts in DR mice were significantly higher than in AL mice throughout days 1–4 (815.80 ± 116.70 ×10³/μL; 73.28% of baseline) and increased substantially between days 6 and 15 post-chemotherapy. Interestingly, aged mice exhibited earlier PLT recovery after chemotherapy compared with young mice (Figure 1C). Similar to findings in young mice, DR significantly reduced pre-chemotherapy peripheral WBC and lymphocyte counts in aged mice. However, from day 1 to 4 post-chemotherapy, WBC and lymphocyte declines were more severe in AL mice than in DR mice. Notably, from days 6–15, WBC and lymphocyte counts in the DR group significantly increased (Figures S1G and S1H). Compared to young mice, aged mice showed a milder decline in HGB and RBC post-chemotherapy, with DR still providing a protective effect (Figure S1I). These findings suggest that peripheral blood (PB) cell dynamics following 5-FU treatment in aged mice resemble those observed in young mice.

Bone marrow (BM) cell counts showed that chemotherapy led to a significant decrease in both young and aged AL mice. In young AL mice, BM cell counts reached the nadir on day 6 after chemotherapy, while in aged AL mice, the nadir was observed on day 4 during the observation period. Notably, DR conferred a significantly protective effect on BM cells. In young mice, BM cell counts in the DR group on day 6 after chemotherapy were significantly higher than in the AL group (young AL vs. young DR: 8.16 ± 1.00 vs. 16.32 ± 2.49 million per mouse) and recovery by day 9 was more rapid compared with AL mice (Figure 1D). In aged mice, under steady-state conditions, DR markedly reversed the aging-induced increase in BM cells. However, compared to AL mice, DR mice showed a significantly attenuated decline in BM cells, with levels on day 4 post-chemotherapy reaching nearly twice those of AL mice (AL vs. DR: 39.60 ± 5.18 vs. 80.40 ± 6.07 million per mouse) (Figure 1F). Additionally, by quantifying MKs in femoral sections, we found that under steady-state conditions, DR significantly reduced the number of MKs in young mice, including both thrombocytogenic MKs (characterized by large cell diameter, abundant cytoplasm, and clearly lobulated nuclei) and non-thrombocytogenic MKs (characterized by small cell diameter, sparse cytoplasm, and unlobulated or minimally lobulated nuclei). However, on day 3 post-chemotherapy, DR mice exhibited significantly higher MK counts compared to AL mice, particularly in the non-thrombocytogenic MK population, and showed a more rapid recovery by days 6 and 9 post-chemotherapy (Figure 1E). Consistent with observations in young mice, aged DR mice also demonstrated a significant reduction in MK numbers under steady-state conditions. Notably, on day 1 post-chemotherapy, MK counts in AL mice were significantly lower than those in DR mice, especially among thrombocytogenic MKs. On day 4 post-chemotherapy, both AL and DR mice exhibited a significant recovery of thrombocytogenic MKs. However, in AL mice, this recovery appeared to be achieved at the expense of depleting the limited pool of non-thrombocytogenic MKs, leading to a further reduction in this cell subset. Notably, non-thrombocytogenic MKs in DR mice recovered rapidly, with counts significantly higher than those in AL mice (Figure 1G). These results indicate that DR significantly promotes the recovery of MKs following chemotherapy.

To investigate the effect of DR on PLT and MK apoptosis, we examined the binding of PLT to Annexin V. Phosphatidylserine (PS) exposure is a typical marker of apoptosis in nucleated cells. However, PS exposure in PLT usually requires stronger stimuli (Leytin 2012; Tseng et al., 2014). We found that 5-FU significantly induced PS exposure in PLT, but no significant difference was observed between aged AL and DR mice, suggesting that DR did not have a direct protective effect on PLT (Figure S1J). Further TUNEL staining of MKs also showed that DR did not have significant protective effects on apoptosis in MKs in aged mice (Figure S1K). These results were not surprising, as previous studies have shown that 5-FU treatment primarily induces apoptosis in MkPs, leading to a reduction in MKs and PLT (Zeuner et al., 2007).

DR protects HSPCs against 5-FU

To understand the underlying mechanisms of the impact of DR on PLTs following chemotherapy, we next investigated changes in the HSPCs population, as HSPCs are usually upstream cell origins of MkPs and MKs and are also the primary target of 5-FU treatment (Randall and Weissman, 1997). Although c-Kit⁺ lineage⁻ cells are considered a reliable marker for identifying HSPCs, 5-FU administration appeared to alter the c-Kit expression pattern and significantly reduce the number of these cells (Umemoto et al., 2018, 2022), thereby making post-chemotherapy identification of HSPCs more challenging. Therefore, we used EPCR as a marker to define HSCs, a method for identifying long-term HSCs ([LT-HSCs], CD48⁻ CD150⁺ EPCR⁺ lineage⁻ = CD48⁻ CD150⁺ LE), short-term HSCs ([ST-HSCs], CD48⁻ CD150⁻ LE), and multipotent progenitors ([MPPs], CD48⁺ LE) that does not rely on Sca-1 or c-Kit (Umemoto et al., 2018, 2022). Flow cytometry analysis revealed that in young mice, DR significantly reduced the numbers of LT-HSCs and ST-HSCs under steady-state conditions. However, on day 1 post-chemotherapy, both AL and DR groups exhibited a significant decrease in LT-HSCs, ST-HSCs, and MPPs. Notably, the AL group showed no significant recovery of these cell populations by day 6 post-chemotherapy. In contrast, the DR group demonstrated a significant increase in LT-HSCs, ST-HSCs, and MPPs starting from day 3 post-chemotherapy, with levels stabilizing between days 6 and 9 (Figures 1H and S2A). Consistent with the observations in young mice, DR effectively mitigated the aging-associated expansion of the HSC compartment in aged mice under steady-state conditions. Notably, on day 1 post-chemotherapy, the absolute numbers of LT-HSCs, ST-HSCs, and MPPs were significantly higher in the DR group compared with the AL group. Furthermore, by day 4 post-chemotherapy, these progenitor populations remained substantially enriched in the DR cohort relative to their AL counterparts, underscoring the sustained protective effect of DR on hematopoietic recovery (Figure 2I). Additionally, we observed that while the proportion of c-Kit^high lineage⁻ cells significantly decreased post-chemotherapy, the proportion of c-Kit^low lineage⁻ cells significantly increased (Figure S2B). Katama N. et al. suggest that post-chemotherapy, HSPCs primarily reside within the c-Kit^low lineage⁻ population, followed by the c-Kit^high lineage⁻ population, while the c-Kit⁻ lineage⁻ population does not contain HSPCs with hematopoietic reconstitution capacity (Katayama et al., 1993). Therefore, we further used c-Kit^high+low lineage⁻ to define LT-HSCs (CD48⁻ CD150⁺ c-Kit^high+low Sca-1⁺ lineage⁻ = CD48⁻ CD150⁺ LSK^hi+lo), ST-HSCs (CD48⁻ CD150⁻ LSK^hi+lo), and MPPs (CD48⁺ LSK^hi+lo) in aged mice. Flow cytometry results using this gating strategy were consistent with those based on EPCR⁺ lineage⁻ markers, further confirming the protective effect of DR on HSCs (Figure S2C). These results indicate that DR significantly promotes the recovery of HSCs following chemotherapy.

DR preserves HSC reconstitution capacity after 5-FU

(A) Schematic diagram of spleen colony-forming assay in aged mice.

(B) Number of CMPs, MEPs and MkPs per million BM cells in LK^lo+hi on day 1 after 5-FU treatment.

(C) CFU-Spleen number and representative photographs (one-way ANOVA).

(D and E) Flow cytometry analysis of CFU-Spleen and representative photographs.

(F) Schematic diagram of transplantation in aged mice.

(G) Number of LT-HSCs LSK^lo+hi per million BM cells on day 1 after 5-FU treatment.

(H and I) Chimerism of donor-derived cells in the PB and BM at 4 months after primary transplantation and representative photographs.

(J–L) Chimerism of donor-derived cells in the PB and BM at 4 months after secondary transplantation and representative photographs.

n = 5 mice per group randomly picked from two independent experiments. Results were displayed as mean ± SD by Unpaired two-tailed Student’s t test. ^∗p < 0.05; ^∗∗p < 0.01; ^∗∗∗p < 0.001; ^∗∗∗∗p < 0.0001; ns, not significant. Tx: transplantation; FCM: flow cytometry.

Next, we assessed the alterations in hematopoietic progenitor cell (HPC) populations following chemotherapy. Flow cytometry results showed that, under steady-state conditions in young mice, the numbers of common myeloid progenitors ([CMPs], CD16/32⁻ CD34⁺ c-Kit^high+low Sca-1⁻ lineage⁻ = CD16/32⁻ CD34⁺ LK^hi+lo) and MK-erythroid progenitors ([MEPs], CD16/32⁻ CD34⁻ LK^hi+lo) of the DR group were significantly higher than those in the AL group, whereas MkPs (CD41⁺ CD150⁺ LK^hi+lo) showed no significant difference. Chemotherapy markedly reduced the numbers of CMPs, MEPs, and MkPs in both AL and DR groups, with recovery beginning at day 7 post-treatment. Notably, from days 7 to 9 after chemotherapy, the recovery of these progenitor populations was significantly more rapid in the DR group compared to the AL group (Figures 1J and S2A). In aged mice, under steady-state conditions, DR only significantly increased the number of MEPs compared to the AL group, while CMPs and MkPs showed no significant differences. On day 1 post-chemotherapy, the numbers of CMPs, MEPs, and MkPs in the BM of DR mice were significantly higher than those in AL mice, and by day 4 post-treatment, the recovery of these progenitor populations was significantly more rapid in DR mice compared to AL mice (Figure 1K). Interestingly, at the same chemotherapy dose as aged mice (40 mg/kg/day), DR still significantly mitigated CIT and promoted HSPC recovery in young mice. The reduced-dose regimen (40 vs. 50 mg/kg/day previously) also improved survival in young AL mice. Notably, aged DR mice exhibited an earlier recovery of CMPs, MEPs, and MkPs compared with young DR mice, suggesting that hematopoietic repair in aged mice may respond more rapidly to DR intervention (Figure S3).

To investigate whether DR affects HSPC apoptosis in young and aged mice, we assessed the apoptosis of LT-HSCs and MkPs using Annexin V staining. Flow cytometric analysis indicated that 5-FU treatment markedly induced apoptosis in HSCs and MkPs. In young mice, DR did not appreciably attenuate the proportion of apoptotic HSCs and MkPs. By contrast, in aged mice, DR significantly reduced the apoptotic rate of LT-HSCs and MkPs (Figures S2D and S2E). Furthermore, we assessed LT-HSC proliferation using Ki-67 staining. Flow cytometry analysis revealed that, under steady-state conditions, the quiescent fraction of LT-HSCs was significantly higher in aged mice than in young mice and DR further enhanced HSC quiescence in both age groups. On day 1 post-chemotherapy, the proportion of proliferating LT-HSCs in both young and aged DR mice was significantly higher than in AL controls. Notably, in young mice, nearly half of LT-HSCs were proliferating at steady state and 5-FU treatment appeared to preferentially deplete these proliferating cells. In contrast, approximately 90% of LT-HSCs in aged mice remained quiescent under steady-state conditions. However, on day 1 after chemotherapy, aged mice mounted a more rapid proliferative response to stress compared with young mice, with LT-HSC proliferation increasing sharply, and this response was further potentiated by DR. However, during the later phase post-chemotherapy, the proliferation of LT-HSCs in both young and aged mice increased further (Figures S2F and S2G). These results indicate that DR primarily mitigates chemotherapy-induced BM suppression by reducing apoptosis in HSPCs and promoting the proliferation of LT-HSCs, especially in aged mice.

Clinically, severe infection can often lead to a reduction in PLT counts (Fogagnolo et al., 2022). In our previous study, we have shown that 5-FU could induce a significant increase of gut opportunistic pathogens and their translocation, which caused death of these mice and could be rescued by DR (Tang et al., 2024). To investigate whether the protective effect of DR on PLTs was mediated indirectly through modulation of the gut microbiota, we administered broad-spectrum antibiotics (Abx) to 5-FU-treated aged mice and examined their impact on PLTs (Figure S4A). The results showed that AL mice continued to exhibit significant PLT loss, whereas DR effectively rescued this phenotype. Notably, although Abx treatment reduced mortality in AL mice (Figure S4B), DR exerted a more pronounced effect by markedly promoting PLT recovery (Figure S4C). BM cell counts demonstrated findings consistent with those observed in young mice. 5-FU treatment induced a significant depletion of BM cells in AL mice, with cell numbers reaching their nadir on day 6 post-treatment (AL vs. DR: 23.60 ± 4.34 vs. 72.00 ± 13.64 million per mouse). In contrast, BM cell counts in DR mice exhibited a gradual recovery beginning as early as day 1 post-chemotherapy and remained consistently elevated relative to AL mice throughout the observation period (Figure S4D). Subsequent flow cytometric analyses further revealed that DR markedly accelerated the recovery of LT-HSCs, ST-HSCs, MPPs, CMPs, MEPs, and MkPs compared to AL mice (Figures S4E and S4F). These findings indicate that 5-FU alone can significantly reduce PLT counts. This phenotype can be rescued by DR and was not a result of elimination of infection from gut pathogens.

DR preserves HSC reconstitution capacity after 5-FU

To assess the effect of DR on the function of aged HPCs following chemotherapy, colony-forming unit-spleen (CFU-Spleen) assays were conducted using BM cells isolated from aged mice sacrificed on day 1 post-chemotherapy (Figure 2A). We first assessed the proportions of each cell population within a defined number of cells. Flow cytometric analysis revealed no significant differences in the numbers of CMPs and MkPs between the transplanted BM cells from AL and DR mice, but the number of MEPs was significantly higher in the DR group compared to the AL group (Figure 2B). 0.5 million BM cells from AL and DR mice were transplanted into recipient mice via tail vein injection. On day 7 post-transplantation, we performed CFU-Spleen counting. The results showed that there was no significant difference in the number of CFU-Spleen from AL and DR mice that were previously treated with saline control. However, 5-FU treatment markedly reduced the number of CFU-Spleen colonies formed by BM cells from AL mice compared with those from DR mice (AL vs. DR: 1.00 ± 0.71 vs. 17.80 ± 4.92) (Figure 2C). Further flow cytometric analysis of CFU-Spleen revealed that DR donor-derived BM cells exhibited higher outputs in CMPs, MEPs, and MkPs (Figures 2D and 2E), indicating that DR significantly improved the hematopoietic reconstitution capacity of HPCs.

To further investigate the effects of DR following chemotherapy on the long-term self-renewal capacity and hematopoietic reconstitution potential of aged HSCs, we performed a competitive transplantation assay. BM cells were harvested from AL or DR mice on day 1 after 5-FU treatment, mixed at a 1:1 ratio with BM cells from Ly5.1/Ly5.2 mice, and transplanted into lethally irradiated primary recipient mice via tail vein injection (Figure 3F). Pre-transplantation, flow cytometric analysis revealed no significant difference in the number of LT-HSCs between AL and DR donor BM cells (Figure 3G). Four months post-transplantation, we analyzed the chimerism of donor-derived BM and PB cells and performed serial transplantation. DR did not improve the reconstitution of WBCs, lymphocytes, and myeloid cells in PB and BM in the primary transplantation (Figure 3H). Intriguingly, the chimerism of DR donor-derived BM cells was significantly higher in LT-HSCs, ST-HSCs, CMPs, MEPs, and MkPs compared to AL donor-derived BM cells (Figure 3I). In the serial transplantation, compared to the primary transplantation, DR donor-derived BM cells showed a higher output of WBCs, lymphocytes, and myeloid cells in PB and BM (Figures 3J and 3K). The chimerism of DR donor-derived BM cells in LT-HSCs, ST-HSCs, MPPs, CMPs, MEPs, and MkPs was also significantly higher than that of AL donor-derived BM cells, and AL-donor derived HSCs were nearly exhausted, thereby failing to reconstitute hematopoiesis (Figure 3L).

DR enhances mitochondrial function in HSCs and MkPs after 5-FU

(A) BM cells were harvested from saline AL, saline DR, 5-FU AL, and 5-FU DR aged mice. HSPCs (Lin−c-Kit^lo+hi) were collected by flow cytometric sorting then subjected to 10× Genomics Chromium capture platform.

(B and C) Volcano plot for DEGs in HSCs and CMPs/MkPs.

(D–I) Representative GO analysis results enriched in upregulated and downregulated DEGs of HSCs and CMPs/MkPs.

(J) Expression levels of autophagy-related proteins in HSPCs.

(K) Analysis of mitochondrial superoxide level in LT-HSCs and MkPs.

(L–P) Analysis of MMP (TMRE and JC-1) and mitochondrial mass (JC-1 Green MFI) in LT-HSCs and MkPs and representative photographs.

(Q) Representative transmission electron microscopy images of HSPCs illustrating mitochondrial structure. Mitochondrial mass per cell and the mitochondrial-to-cell area ratio were quantified (n = 3 mice per group).

(R) Intracellular ATP content in HSPCs.

n = 5 mice per group randomly picked from two independent experiments. Results were displayed as mean ± SD by unpaired two-tailed Student’s t test (J) and one-way ANOVA (K–R). ^∗p < 0.05; ^∗∗p < 0.01; ^∗∗∗p < 0.001; ^∗∗∗∗p < 0.0001; ns, not significant.

Consistent with the results observed in aged mice, the primary transplantation in young mice demonstrated that BM cells derived from DR donors produced significantly higher outputs of WBCs, lymphocytes, and myeloid cells in both PB and BM compared to those from AL donors. Additionally, the chimerism of DR donor-derived BM cells in LT-HSCs, ST-HSCs, MPPs, CMPs, MEPs, and MkPs was significantly greater than that of AL donor-derived BM cells (Figures S2H–S2L). Sequential transplantation further confirmed this trend, revealing a markedly increased disparity in chimerism between AL- and DR donor-derived BM cells across these cell populations, with AL donor-derived BM cells exhibiting a near-complete loss of hematopoietic reconstitution capacity (Figures S2M–S2O). These results suggest that DR significantly improves the long-term self-renewal and hematopoietic reconstitution capacity of HSCs in young and aged mice following chemotherapy.

DR enhances mitochondrial pathways in HSCs and MkPs after 5-FU

To systematically elucidate the underlying mechanisms by which DR mitigates CIT, we performed single-cell RNA sequencing (scRNA-seq) on HSPCs from AL and DR mice (Figure 3A). Since DR exhibited similar PLT rescue effects in both young and aged mice, subsequent analyses were conducted only in aged mice to reduce animal usage. We sorted c-Kit^high+low lineage⁻ cell populations by flow cytometry and analyzed the proportion of each cell subtype within this population. The results indicated that a large portion of the cells were classified as HSPCs, primarily including HSCs, MPPs, CMPs, MkPs, MEPs, and granulocyte-macrophage progenitors (GMPs, CD16/32⁺ CD34⁺ LK^hi+lo). Pre-chemotherapy, both AL and DR mice predominantly contained GMPs, followed by MPPs, CMP/MkPs, and MEPs, with HSCs representing the smallest proportion. Post-chemotherapy, the proportion of MEPs notably increased, and DR mice exhibited a significantly higher proportion of MEPs compared to AL mice. Meanwhile, the proportions of MPPs, CMPs/MkPs, and GMPs were significantly reduced, with no substantial differences between AL and DR mice. Notably, the proportion of HSCs significantly increased post-chemotherapy, but DR mice exhibited a significantly lower proportion of HSCs compared to AL mice (Figure S5A). This outcome was anticipated, as previous studies have shown that HSCs, being one of the most quiescent cell types in BM, exhibit considerable resistance to chemotherapy (Zhao et al., 2019).

Using the 10X Genomics capture platform, we obtained a total of 48,266 high-quality cells (including 10,808 saline AL cells, 12,571 saline DR cells, 12,396 5-FU AL cells, and 12,491 5-FU DR cells), with each cell averaging 3,070 genes. These cells were clustered into 11 distinct cell types. We annotated each cluster based on previously reported classical marker genes (Konturek-Ciesla et al., 2023; Zeng et al., 2023; Poscablo et al., 2024): HSCs (Hlf, Mecom, Ly6a, and Procr), MPPs (Flt3, Cd34, and Gcnt2), CMPs/MkPs (Angpt1, Pf4, Cd9, Itga2b, and Gata2), MEPs (Cd34-, Ly6a-, Fcgr3-, Gata1, Tfrc, and Klf1), GMPs (F13a1, Irf8, Mpo, Csf1r, and Fcgr3), PreNeutrophil (Cebpe, Gfi1, Prtn3, and Elane), MatureNeutrophil (Ly6g, Ly6c2, Cebpe, and Mpo), dendritic cells (Irf8 and Klf4), plasma cells (Jchain and Cd74), mast cell/basophil/eosinophil progenitors (Ms4a2, Fcer1a, Prss34, and Mpo), and natural killer/T cells (Lck and Ncr1) (Figures S5B and S5C). In line with the flow cytometry analysis, both AL and DR mice predominantly contained GMPs, with HSCs comprising the smallest proportion under homeostasis. Post-chemotherapy, both AL and DR mice exhibited an increase in the proportion of HSCs and MEPs, with DR mice showing a significantly higher proportion of MEPs compared to AL mice, while the proportion of HSCs in DR mice was lower than in AL mice (Figure S5D).

In the early phase following chemotherapy, HSCs and MkPs play a critical role in the recovery and maintenance of peripheral PLT levels (Poscablo et al., 2024; Tang et al., 2025). To further elucidate the potential mechanisms by which DR mitigates CIT, we analyzed the transcriptomic changes in HSCs and CMPs/MkPs. In HSCs, 1,188 differentially expressed genes (DEGs) were identified between saline- and 5-FU-treated AL mice and 1,689 DEGs were identified between 5-FU-treated AL and DR mice. In contrast, only 577 DEGs were found between saline-AL and DR mice (Figure 3B). In CMPs/MkPs, 1,144 DEGs were identified between saline- and 5-FU-treated AL mice and 570 DEGs between 5-FU-treated AL and DR mice, while only 228 DEGs were identified between saline-AL and DR mice (Figure 3C). These results indicate that DR significantly alters the transcriptomic profiles of HSCs and CMPs/MkPs following chemotherapy. Interestingly, gene Ontology (GO) analysis revealed that, compared to saline-treated AL mice, 5-FU-treated AL mice exhibited enriched upregulated pathways related to mitochondrial structure and function, as well as cell proliferation, in both HSCs and CMPs/MkPs (Figures 3D and 3E; Table S1). Notably, compared to 5-FU-treated AL mice, 5-FU-treated DR mice showed further enrichment of upregulated pathways associated with mitochondrial structure and function, cell proliferation, and DNA repair in HSCs and CMPs/MkPs, along with enrichment of downregulated pathways related to inflammation and apoptosis. Moreover, PLT function-related pathways were enriched in CMPs/MkPs from 5-FU-treated DR mice compared to 5-FU-treated AL mice (Figures 3F and 3G). In addition, compared with saline-AL mice, saline-DR mice exhibited a significant downregulation of pathways associated with reactive oxygen species (ROS) response, inflammatory response, and apoptosis in HSCs and CMPs/MkPs (Figures 3H and 3I). These results indicate that, prior to chemotherapy, DR conditions maintain HSCs and CMPs/MkPs in a protective homeostatic state characterized by low oxidative stress, reduced inflammation, and resistance to apoptosis.

DR enhances mitochondrial function in HSCs and MkPs after 5-FU

Recent studies have shown that during stress-induced hematopoiesis, HSCs are forced to exit their quiescent state, and mitochondrial activation plays a crucial role in driving HSC differentiation and self-renewal following chemotherapy (Ho et al., 2017; Umemoto et al., 2018; Takihara et al., 2019). Additionally, the MMP and mitochondria mass are essential for preserving HSC self-renewal, regenerative potential, and quiescence under homeostatic conditions (Ito et al., 2012; Maryanovich et al., 2015; Mansell et al., 2021). However, the role of mitochondria in HSCs and MkPs in the context of CIT is unknown. Since scRNA-seq revealed substantial enrichment of mitochondria-related pathways in HSCs and MkPs from DR mice following 5-FU treatment, we first evaluated mitochondrial function. Western blot analysis of HSPCs showed that, compared with the saline-AL group, saline-DR mice exhibited significantly increased LC3B-II/LC3B-I ratio, accompanied by reduced p62 levels (Figure 3J), indicating that DR robustly enhances autophagic activity in HSPCs. Consistent with previous reports (Ho et al., 2017; Fang et al., 2020), activation of autophagy in HSPCs is typically accompanied by reductions in mitochondrial ROS (MitoROS) and MMP (Figures 3K–3N). These findings suggest that, under homeostatic conditions, DR markedly improves mitochondrial function. However, following 5-FU treatment, both AL and DR mice displayed significant increases in MMP and mitochondrial mass in HSCs and MkPs. Notably, compared with AL mice, DR further enhanced MMP and mitochondrial mass in HSCs and MkPs after chemotherapy (Figures 3L–3Q), concomitant with a significant elevation in ATP levels (Figure 3R), while MitoROS levels did not show a corresponding increase (Figure 3K). Collectively, these data indicate that DR markedly activates mitochondrial function after chemotherapy, thereby promoting hematopoietic recovery.

Mitochondrial potentiation rescues PLT and HSC regeneration in AL mice after 5-FU

To further investigate whether improving mitochondrial function in LT-HSCs and MkPs post-chemotherapy may play a significant role in mitigating CIT, we administered mitoquinol (MQ) via intraperitoneal injection to AL mice for 5 consecutive days (Figure 4A). MQ, a mitochondrial-targeted coenzyme-Q10 (Kelso et al., 2001; James et al., 2004, 2005; Murphy and Smith, 2007), has been shown in vivo to improve mitochondrial function in aged HSCs (Murphy and Smith, 2007; Mansell et al., 2021). The results show that there was a trend toward enhanced MMP and mitochondrial mass in LT-HSCs and MkPs following MQ injection in AL mice pre-chemotherapy, although the differences were not statistically significant. However, MQ significantly enhanced the MMP and mitochondrial mass in LT-HSCs and MkPs compared to AL mice post-chemotherapy (Figures S6A–S6C). Peripheral hematological parameter analysis indicated that MQ significantly protected the PLT in AL mice post-chemotherapy (Figure 4B). Additionally, BM cell counts from MQ-treated mice were significantly higher than those in vehicle-control treated AL mice (Figure 4C). Analysis of BM sections revealed that on day 1 post-chemotherapy, the number of thrombocytogenic MKs was markedly higher in MQ-treated mice compared to AL mice. By day 4, non-thrombocytogenic MKs were significantly increased, thereby promoting MKs recovery (Figure 4D). Further flow cytometric analysis showed that on day 1 post-chemotherapy, MQ-treated AL mice had significantly higher numbers of HSPCs, including LT-HSCs, ST-HSCs, MPPs, CMPs, MEPs, and MkPs, compared to vehicle-control treated AL mice. On day 4, MQ treatment further promoted the recovery of these cells (Figures 4E and 4F). These results indicate that MQ significantly enhances hematopoietic reconstitution in AL mice following chemotherapy, especially the regeneration of PLTs. We further examined the apoptosis of LT-HSCs and MkPs using Annexin V staining. Flow cytometry revealed that the apoptosis rate of LT-HSCs and MkPs in the MQ group was significantly reduced, suggesting that the increase in MMP significantly decreased apoptosis in these cells (Figure 4G). Collectively, these results demonstrate that mitochondrial potentiation by MQ markedly promotes hematopoietic recovery in AL mice after chemotherapy.

To investigate the impact of MMP upregulation by MQ on colony forming capacity of HPCs, we performed spleen colony-forming assays and measured the proportion of HPCs in the transplanted BM cells (Figure 4H). Flow cytometric analysis revealed no significant differences in the numbers of CMPs and MkPs between the transplanted BM cells from AL and MQ mice, but the number of MEPs was significantly higher in the MQ group compared to the AL group on day 1 post-chemotherapy (Figure 4I). On day 7 post-transplantation, we performed CFU-Spleen counting. The results showed that there was no significant difference in the number of CFU-Spleen from vehicle-control and MQ-treated mice without 5-FU treatment. However, chemotherapy significantly reduced the number of CFU-Spleen formed by both vehicle-control and MQ treated mice. Notably, the number of CFU-Spleen derived from MQ mouse BM cells was significantly higher than that derived from vehicle-control mouse BM cells (Figure 4J). Further flow cytometric analysis of CFU-Spleen formed by BM cells on the first day after chemotherapy revealed that MQ donor-derived BM cells exhibited higher outputs in CMPs, MEPs, and MkPs (Figure 4K), indicating that MQ significantly improved the colony-forming capacity of HPCs post chemotherapy.

To further explore the effect of MMP upregulation on the hematopoietic reconstitution function of BM cells, we transplanted BM cells from vehicle-control and MQ-treated mice on day 1 post-chemotherapy mixed with competitive cells into recipient mice (Figure 4L). Flow cytometric analysis of the donor BM cells pre-transplantation showed no significant difference in the number of LT-HSCs between the two groups (Figure 4M). MQ donor-derived BM showed significantly higher outputs of peripheral WBCs, B cells, and myeloid cells in both the PB and BM (Figure 4N). Intriguingly, the chimerism of MQ donor-derived BM cells, including LT-HSCs, ST-HSCs, MPP, CMPs, MEPs, and MkPs, was also significantly higher than that of vehicle-control donor-derived BM cells (Figure 4O). These results further demonstrate that enhancing MMP with MQ significantly improves the hematopoietic reconstitution ability of HSCs in AL mice post-chemotherapy, thus promoting the recovery of the hematopoietic system.

Mitochondrial dysfunction impairs PLT protection and hematopoietic function in DR mice after 5-FU

To further investigate the impact of mitochondrial function on the effect of DR in hematopoietic recovery following chemotherapy, we administered carbonyl cyanide 3-chlorophenylhydrazone (CCCP) via intraperitoneal injection to the DR mice pre-chemotherapy. CCCP is a protonophore that increases the permeability of the mitochondrial inner membrane to protons, leading to mitochondrial dysfunction (Jiao et al., 2021; Mansell et al., 2021). Mice were fed an AL or DR diet for 30 days, and then DR mice received a single dose of CCCP or saline as control. Afterward, mice were treated with 5-FU or saline (Figure 5A). In the non-chemotherapy groups, we observed that CCCP tended to reduce MMP in LT-HSCs and MkPs before chemotherapy, but had no significant effect on mitochondrial mass. Interestingly, CCCP significantly reduced both MMP and mitochondrial mass in LT-HSCs and MkPs in DR-CCCP mice post-chemotherapy, compared to the DR-saline group (Figures S6D–S6F). Intriguingly, on day 1 post-chemotherapy, we observed a significant reduction in PLT in the CCCP group, with a further decrease on day 4 (Figure 5B). Additionally, the BM cell counts in the DR-CCCP group were significantly lower than in the DR-saline group (Figure 5C). Hematoxylin and eosin (H&E) staining on BM sections showed that CCCP significantly decreased the number of thrombocytogenic MKs in DR mice on day 1 post-chemotherapy, and on day 4, the number of non-thrombocytogenic MKs further decreased, preventing the recovery of MKs in the BM (Figure 5D). Flow cytometric analysis revealed that on day 1 post-chemotherapy, the numbers of HSPCs, including LT-HSCs, ST-HSCs, MPPs, CMPs, MEPs, and MkPs, in the DR-CCCP group were significantly lower than in the DR-saline group. On day 4, CCCP significantly inhibited the recovery of HSPCs (Figures 5E and 5F). To further investigate apoptosis in LT-HSCs and MkPs, we used Annexin V staining. The results revealed that the apoptosis rate of LT-HSCs and MkPs in the DR-CCCP group was significantly higher than in the DR-saline group. These results indicate that the reduction in MMP significantly promoted apoptosis in LT-HSCs and MkPs (Figure 5G). In summary, these results indicate that CCCP-induced mitochondrial dysfunction impairs DR-mediated hematopoietic recovery and PLT regeneration after chemotherapy.

To examine the functional impact of MMP downregulation on HPCs, we performed spleen colony-forming assays (Figure 5H). We also measured the proportion of HPCs in the transplanted BM cells, which indicated that CCCP significantly reduced the number of MEPs, although CMPs and MkPs were not significantly affected on day 1 post-chemotherapy (Figure 5I). On day 7 post-transplantation, we performed CFU-Spleen counting. Results showed no significant difference in CFU-Spleen numbers among AL, DR-saline, and DR-CCCP mice in non-chemotherapy-treated mice. However, chemotherapy markedly reduced the number of CFU-Spleen formed by BM cells derived from all three groups. Notably, BM cells from the DR-CCCP group generated significantly fewer CFU-Spleen numbers compared to those from the DR group (Figure 5J). Further flow cytometric analysis of CFU-Spleen derived from BM cells on day 1 post-chemotherapy revealed that CCCP donor-derived BM cells exhibited lower output in CMPs, MEPs, and MkPs (Figure 5K). These results indicate that CCCP significantly impairs the number and function of HPCs.

To further explore the effect of MMP downregulation on the hematopoietic regeneration function of BM cells, we transplanted BM cells from AL, DR-saline or DR-CCCP mice on day 1 post-chemotherapy along with competitive cells into recipient mice (Figure 5L). Flow cytometric analysis of BM cells collected on day 1 post-chemotherapy from AL, DR-saline, and DR-CCCP mice pre-transplantation showed no significant differences in LT-HSC counts (Figure 5M). Four months post-transplantation, we analyzed the chimerisms of donor-derived BM cells and PB cells. There were no significant differences in the chimerisms of WBCs, lymphocytes, and myeloid cells in PB and BM (Figure 5N). Interestingly, the chimerisms of DR-CCCP donor-derived BM cells, including LT-HSCs, ST-HSCs, CMPs, MEPs and MkPs, were significantly lower than that of DR-saline donor-derived BM cells (Figure 5O). These results further confirm that inhibiting MMP with CCCP significantly impairs the hematopoietic cell reconstitution ability of DR mice post-chemotherapy.

DR delays tumor growth without impairing its efficacy in mitigating CIT

To explore the impact of tumors on chemotherapy after DR, 12-week-old female C57BL/6J mice were subcutaneously injected with MC38 colon cancer cells to establish a colon cancer mouse model. After successful tumor growth, the mice underwent one month of DR treatment. After 1 month, we administered 50 mg/kg/day of 5-FU for 5 consecutive days to the AL and DR mice and euthanized the mice on day 6 post-chemotherapy (Figure 6A). By measuring tumor volume and mass, we found that DR significantly delayed tumor growth, which was consistent with previous findings (Mukherjee et al., 2002; Lee et al., 2012; Lien et al., 2021; Pomatto-Watson et al., 2021). We also observed that, although the tumor volume and mass in DR-treated chemotherapy mice showed a decreasing trend, the differences were not statistically significant (Figures 6B and 6C).

To further investigate whether the tumor affected the protective effect of DR on chemotherapy-induced BM suppression, we measured PB PLT. In line with previous results, DR mice had significantly higher PLT than AL mice on day 6 post-chemotherapy (Figure 6D). Compared to DR mice, AL mice had significantly lower BM cell counts post-chemotherapy (Figure 6E). MK counting results showed that under homeostasis, DR significantly reduced the number of both thrombocytogenic and non-thrombocytogenic MKs. However, both thrombocytogenic and non-thrombocytogenic MK counts in AL mice were significantly lower than in DR mice on day 6 post-chemotherapy (Figures 6F and 6G). Further flow cytometry analysis revealed that the numbers of LT-HSCs, ST-HSCs, and MPPs in AL mice were significantly lower than those in DR mice post-chemotherapy (Figure 6H). These results indicate that DR not only significantly delays tumor growth but also preserves its protective effect against chemotherapy-induced BM suppression.

PLT count inversely correlates with BMI in 5-FU-treated cancer patients

Previous studies have reported that individuals undergoing CR typically have a lower body mass index ([BMI] 19.7 ± 1.8 kg/m²) (Walford et al., 1992). Accordingly, we retrospectively analyzed 100 patients with gastrointestinal malignancies treated with 5-FU– or capecitabine-based chemotherapy and assessed the association between pre-chemotherapy BMI and the overall nadir PLT count observed during the first eight chemotherapy cycles (Figure 7A). Receiver operating characteristic (ROC) curve analysis was performed to evaluate the predictive value of BMI for CIT occurrence and to determine the optimal cutoff value. CIT was defined as a post-chemotherapy PLT count <100 × 10⁹/L, which served as the dependent variable, with BMI as the predictor. The analysis identified an optimal BMI cutoff value of 22.95 kg/m², with an area under the curve (AUC) of 0.8559 (95% confidence interval [CI]: 0.7766–0.9352), indicating that BMI has a strong predictive value for CIT occurrence (Figure 7B). Furthermore, correlation analysis revealed a moderate negative correlation between BMI and post-chemotherapy nadir PLT count (r = −0.446, p < 0.001), suggesting that higher BMI is associated with an increased risk of CIT (Figure 7C). Patients were then stratified into two groups based on the BMI cutoff value of 22.95 kg/m². The comparison of nadir PLT counts between the two groups revealed that patients with a BMI >22.95 kg/m² had significantly lower nadir PLT counts than those with a BMI of 18.5–22.95 kg/m² (Figure 7D). However, within the same patient cohort, no significant differences in BMI were observed between patients who developed chemotherapy-induced neutropenia or anemia and those who did not (Figure 7E).

To further clarify whether BMI is an independent risk factor for CIT, a multivariate logistic regression model was constructed, adjusting for potential confounders including age, sex, baseline PLT, liver function (aspartate aminotransferase [AST] and alanine aminotransferase [ALT]), renal function (urea, creatinine [Cre], estimated glomerular filtration rate [eGFR]), tumor type, tumor stage, chemotherapy regimen, and dose intensity. The results demonstrated that BMI remained independently associated with the occurrence of CIT (odds ratio [OR] = 1.68, 95% CI: 1.32–2.25), indicating that for each 1 kg/m² increase in BMI, the odds of developing CIT increased by approximately 68% (Figure 7F; Table S2). To further assess potential confounding effects, patients were stratified according to a BMI of 18.5–22.95 kg/m² and BMI >22.95 kg/m². The two groups showed no significant differences in age, sex distribution, dose adjustments, or treatment interruption rates. In addition, none of the patients received antiplatelet or anticoagulant therapy during chemotherapy, suggesting good comparability between the two groups. These findings enhance the reliability of the observed association between BMI and CIT (Figure 7G; Table S2). Collectively, these results demonstrate that BMI serves not only as a robust predictive marker but also as an independent risk factor for CIT, thereby providing further evidence supporting the potential clinical applicability of CR as a preventive strategy against CIT.

Discussion

This study systematically investigates the dynamic changes in PLT developmental-related cells, including upstream HSPCs, MkPs, and MKs and downstream PLTs, in both young and old mice following 5-FU treatment. We found that 5-FU significantly reduced the abundance of these cell populations in AL mice and severely impaired the hematopoietic reconstitution capacity of HSPCs. For the first time, we observed that a 1-month period of DR prior to chemotherapy, a non-pharmacological intervention, effectively mitigated CIT in both young and old mice. DR enhanced the hematopoietic reconstitution function of HSPCs, thereby promoting the recovery of peripheral PLTs post-chemotherapy. Mechanistically, DR improved mitochondrial homeostasis in HSPCs and enhanced their hematopoietic reconstitution capacity. This preconditioning facilitated mitochondrial activation following chemotherapy, thereby promoting the recovery of the megakaryocytic lineage. In AL mice, drug-induced mitochondrial activation mimicked the protective effects of DR, whereas mitochondrial inhibition in DR mice significantly attenuated these benefits. In conclusion, short-term DR effectively alleviates CIT, and targeting mitochondria may offer a novel approach for preventing and treating CIT in elderly cancer patients.

Globally, many countries are facing rapidly aging populations, leading to an increased proportion of elderly patients with malignancies. It is projected that by 2030, over 70% of cancer patients will be over 65 years old (Kadambi et al., 2020). Elderly cancer patients frequently have underlying conditions such as hypertension and stroke, which predispose them to an elevated risk of bleeding in the context of CIT (Takihara et al., 2019). Therefore, managing CIT in elderly cancer patients undergoing chemotherapy is particularly critical. Current thrombopoietic agents have delayed onset and notable side effects and chemotherapy interruptions due to CIT may reduce therapeutic efficacy, negatively impacting patient outcomes (Vadhan-Raj et al., 2000; Mansell et al., 2021). There is thus a substantial unmet clinical need for effective prevention and treatment of CIT in elderly patients. Our study identifies DR as a promising non-pharmacological intervention that significantly mitigates 5-FU-induced thrombocytopenia in young and aged mice. Single-cell transcriptomic analysis revealed that 5-FU upregulates mitochondrial-related pathways in HSCs and MkPs, which are further enhanced by DR. Moreover, we used MQ, a clinically available drug known to improve mitochondrial function, and found that it significantly mitigated CIT in aged AL mice. Previous studies have shown that MQ enhances mitochondrial function in aged HSCs, thereby reducing ROS levels, facilitating DNA repair, and reversing myeloid bias (Mansell et al., 2021). However, whether MQ promotes PLT recovery post-chemotherapy remains unknown. Our findings suggest that MQ holds potential as a clinical therapeutic to facilitate PLT regeneration following chemotherapy.

Previous studies have shown that mitochondrial regulation plays a central role in determining the fate of HSCs (Ansó et al., 2017; Takihara et al., 2019; Hinge et al., 2020). In the HSC compartment, subsets of HSCs with stronger self-renewal capacity are typically enriched in mitochondria (Totani et al., 2025), yet maintain relatively low MMP (Liang et al., 2020). This characteristic is closely related to the fact that HSCs predominantly rely on glycolytic metabolism to sustain their self-renewal ability (Vannini et al., 2016; Yang et al., 2024). Other reports suggest that MMP activation and increased mitochondrial mass after chemotherapy are crucial for hematopoietic recovery (Umemoto et al., 2018). Additionally, mitochondrial dysfunction in PLTs after chemotherapy may increase the risk of CIT (Baaten et al., 2018). To date, effective non-pharmacological strategies for modulating mitochondrial function in HSCs are lacking and the effects of MMP and mitochondrial mass on aged HSCs and MkPs post-chemotherapy remain poorly understood. In aged HSCs, a subset maintains regenerative capacity through high autophagy activity and low metabolic status, while autophagy-deficient cells accumulate dysfunctional mitochondria (Ho et al., 2017; Fang et al., 2020; Borsa et al., 2024). Moreover, the persistent accumulation of chronic inflammation and ROS exacerbates the decline in autophagic capacity in aged HSCs (Ho et al., 2017; Fang et al., 2020). Our previous research indicates that DR can reverse the aging-associated phenotypes of HSCs and reduce chronic inflammation, thereby enhancing their self-renewal and hematopoietic reconstitution capacity (Tang et al., 2016; Tao et al., 2020, 2025; Rasa et al., 2022). Although previous studies found that intermittent fasting can attenuate cyclophosphamide-induced immunosuppression and promote lymphocyte recovery (Cheng et al., 2014), the impact on PLTs post-chemotherapy remains unaddressed and the research was limited to young mice. The effects of DR on hematopoiesis in aged mice post-chemotherapy remain unexplored. Our study demonstrates that short-term DR prior to chemotherapy effectively mitigates CIT and improved mitochondrial homeostasis in HSPCs and enhanced their hematopoietic reconstitution capacity. This preconditioning facilitated mitochondrial activation following chemotherapy, thereby promoting the recovery of the megakaryocytic lineage.

Previous studies have demonstrated that MQ, as a dietary supplement, can inhibit tumor proliferation and metastasis and reduce recurrence in breast cancer (Cheng et al., 2023). Moreover, in glioblastoma, MQ acts as a lipophilic antioxidant, suppressing tumor cell invasiveness and chemoresistance through free radical scavenging (Burić et al., 2019). Mansell et al. demonstrated that MQ can enhance mitochondrial function in aged HSCs, reduce ROS levels, promote DNA repair, and restore lineage bias (Mansell et al., 2021). However, its potential role in mitigating hematologic toxicity remains unclear. Our findings indicate that MQ can significantly mitigate CIT and promote hematopoietic recovery. These results suggest that the application of MQ for the prevention and management of CIT in cancer patients is worthy of further investigation.

In clinical settings, sex- and age-related biological differences among cancer patients are well documented. In our study, only aged female C57BL/6J mice were used due to known sex-based variations in physiology, metabolism, immunity, and hematopoiesis. Studies have shown that female hormones (such as estrogens) can protect the HSC pool, thereby alleviating damage caused by radiotherapy and chemotherapy and promoting hematopoietic recovery (Shaikh et al., 2016; Oguro et al., 2017; Chapple et al., 2018; Velardi et al., 2018; Fananas-Baquero et al., 2021). Clinically, females tend to experience more adverse reactions and lower tolerance to chemotherapy compared to males (Unger et al., 2022). Therefore, aged female mice were selected to minimize sex-related variability. While an ideal approach would involve establishing a tumor-bearing chemotherapy model in aged mice to better mimic clinical scenarios, this is hindered by the high cost, and reduced tolerance to chemotherapy in aged tumor-bearing animals, which makes experimental implementation considerably challenging. Although we did not conduct related studies in aged tumor-bearing mice, we observed that DR markedly mitigates CIT and promoted PLT recovery in healthy aged mice. To address these limitations, we used young tumor-bearing mice for substitution. Results showed that DR not only inhibited tumor growth but also mitigated CIT and promoted hematopoietic recovery. Previous studies also support that CR suppresses tumor growth and enhances chemosensitivity, regardless of diet composition (Lee et al., 2012; Brandhorst et al., 2013; Ma et al., 2018; Lien et al., 2021).

To date, the effects of CR on the human body remain poorly understood due to theoretical and practical limitations. A study on strict CR showed that a 30% reduction in energy intake led to a 15% body weight decrease, with BMI dropping from 23.9 to 19.7 kg/m². Participants exhibited physiological adaptations similar to CR rodents, including reductions in insulin, cholesterol, triglycerides, and WBCs, along with elevated cortisol (Most et al., 2017). Another study on CR with optimal nutrition (CRON) in individuals adhering to CR for 15 years (∼1,800 kcal/day; BMI: 19.7 ± 1.8 kg/m²) revealed metabolic and molecular changes distinct from those on a normal diet, akin to long-lived CR animal models (Walford et al., 1992). Consequently, we used BMI as a projection of DR to assess its impact on PLT counts after 5-FU-based chemotherapy. Our results showed that cancer patients with lower pre-chemotherapy BMI (20.61 ± 1.31 kg/m², close to the BMI of previously reported CR volunteers) had a significantly lower incidence of CIT compared to those with higher BMI (>22.95 kg/m²), suggesting that pre-treatment BMI could be an important predictor of CIT risk in chemotherapy. Previous studies have also shown that obese cancer patients exhibit a significantly higher incidence of CIT compared with patients of normal weight (Gutierrez et al., 2016; Ando et al., 2020). It should be noted that BMI has certain limitations in reflecting the state of DR. As an indirect measure of DR, BMI cannot accurately reflect short-term changes in energy intake and the metabolic remodeling processes that follow. Therefore, in this study, BMI is used only as an initial clinical-related measure to explore the relationship between DR-like states and PLT changes, rather than directly quantifying the physiological effects of DR. In future clinical prospective studies, systematically collecting data on weight fluctuations, dietary energy intake, and nutritional status will be crucial for more accurately characterizing the metabolic state induced by DR.

In conclusion, our findings demonstrate that short-term DR prior to chemotherapy effectively mitigates CIT and improved mitochondrial homeostasis in HSPCs and enhanced their hematopoietic reconstitution capacity. This preconditioning facilitated mitochondrial activation following chemotherapy, thereby promoting the recovery of the megakaryocytic lineage. Given the high incidence of CIT in cancer patients undergoing chemotherapy, DR presents a promising low-cost, low-toxicity non-pharmacological strategy for the prevention and management of CIT.

Methods

Mice and diets

Twelve-week-old and 20- to 22-month-old C57BL/6J female mice were obtained from Hunan SJA Laboratory Animal Co., Ltd., and maintained in the animal facilities of Nanchang Royo Biotech under pathogen-free conditions on a 12-h light/12-h dark cycle at 23°C–25°C. The mice were housed individually and received a regimen of either an AL diet (fed an unlimited amount of food) or a DR diet (fed daily with 70% of the food intake of body weight- and sex-matched AL mice). The food amount provided remained constant over the entire DR period. The mice were fed the Mouse Growth and Reproduction Formula Feed (Tianjin Keshi Feed Co., Ltd., product no.101670654803973120). The nutritional composition of this feed is as follows: 24.02% protein, 12.95% fat, and 63.03% carbohydrates, with a total metabolizable energy of 3.44 kcal/g. The Animal Experimental Ethical Inspection of Nanchang Royo Biotech Co., Ltd., (RYE2022041301) approved all mouse experiments.

Chemical treatment of mice

5-FU treatment

5-FU (Jinyao Pharmaceutical Co., Ltd., China, purity: 99%) was diluted in saline for injection. Mice were administered 5-FU via intraperitoneal injection for 5 days. The daily dose of 5-FU was 50 mg/kg/day for young mice. To increase safety and tolerance, a 20% dose reduction was applied in the aged mice (40 mg/kg/day) as in clinical practices.

Abx treatment

Abx treatment was performed as previously described (Tang et al., 2024). Mice were orally gavaged with 0.2 mL of an antibiotic cocktail (10 mg/mouse/day) containing ampicillin sodium salt (Solarbio, A8180), neomycin sulfate (Solarbio, N8090), metronidazole (Solarbio, M8060), and vancomycin hydrochloride (Sangon Biotech, A600983-0001) for 5 days to ensure equivalent dosing. Thereafter, antibiotics were provided in the drinking water (ampicillin, neomycin, and metronidazole, 1 g/L each; vancomycin, 0.5 g/L) until the end of the experiment to minimize stress associated with long-term gavage. Drinking water was replaced twice weekly. Control mice received saline by gavage in parallel and were provided with regular drinking water.

MQ and CCCP treatment

MQ was administered through intraperitoneal injection (2 mg/kg/day, Cayman, 89950) for 5 days. CCCP (1 mg/kg/day, Sigma, C2759) was administered through a single intraperitoneal injection. All chemicals were administered in saline.

PB cell counting

PB cells were collected from the orbital venous plexus, placed in 5 μL 0.5 M EDTA, and counted on an automated hematology analyzer (Sysmex, XS-500i) according to the manufacturer’s instructions.

H&E staining

Femurs from AL and DR mice were collected and fixed in 4% paraformaldehyde for 24 h. Fixed samples were processed and embedded in paraffin using standard protocols, and 5-μm paraffin sections were prepared. Bone section histology was assessed by H&E staining (Leagene, DH0001 and DH0053) according to the manufacturer’s instructions. Representative areas were photographed using an Olympus IX73 microscope (Japan). With the assistance of pathologists, 9–12 fields were randomly selected from each sample for MK counting. Based on their morphology, MKs were classified as non-thrombopoietic MKs (small cell diameter, sparse cytoplasm, and unlobulated or minimally lobulated nucleus) and thrombopoietic MKs (large cell diameter, abundant cytoplasm, and clearly lobulated nucleus).

TUNEL staining

According to the manufacturer’s instructions, TUNEL staining was performed sequentially using a TUNEL assay kit (Abcam, ab206386) to evaluate MK apoptosis, and periodic acid-Schiff Stain Kit (Abcam, ab150680) to stain the cytoplasm of MKs, allowing for their distinction from other cell types. Representative areas were photographed using an OLYMPUS IX73 microscope (Japan). In each sample, 80–120 MKs were examined, and apoptotic MKs were quantified using the TUNEL assay.

PS exposure assay

For PLT PS exposure analysis, PB cells were collected from the orbital venous plexus and placed in 5 μL 0.5 M EDTA. PLTs were isolated from PB using differential centrifugation. PLTs were washed with CGS buffer (Leagene, CZ0055) and resuspended in modified Tyrode’s buffer (Pricella, PB180340). PLTs were sequentially stained with anti-CD42b antibody (emfret, M040-2) at room temperature for 15 min and Annexin V (BD, 556570) at room temperature for 15 min. After staining, cells were analyzed using a flow cytometer (FACSCanto II; BD), and data were analyzed using FlowJo software (v.10.8).

Flow cytometry

BM cells were obtained by crushing the hind limbs and pelvis in sterile phosphate-buffered saline (PBS), followed by filtration through a 40-μm cell strainer. The cells were then resuspended in red cell lysis buffer (BD, 555899) and incubated at room temperature for 5 min to lyse RBCs. Afterward, the cells were washed, counted, and incubated with flow cytometric antibodies on ice for 30 min (including a 90-min incubation with anti-CD34 antibody). LT-HSC, ST-HSC, and MPP were detected using a lineage cocktail (biotinylated anti-TER-119, -Gr-1, -B220, -CD11b, -CD3, -CD4, and -CD8a), streptavidin-APC-Cy7, EPCR-PE (eBioscience), c-Kit-APC, Sca-1-PE-Cy7 (BD), CD150-PerCP-Cy5.5, and CD48-BV510. CMPs, MKMEPs, MkPs, and GMPs were detected using a lineage cocktail, streptavidin-APC-Cy7, c-Kit-APC, Sca-1-PE-Cy7 (BD), CD34-AF700 (eBioscience), and CD16/32-BV421, CD41-BV605, and CD150-PerCP-Cy5.5. For chimerism analyses, LT-HSC, ST-HSC, and MPP were detected using a lineage cocktail, streptavidin-APC-Cy7, c-Kit-APC, Sca-1-PE-Cy7 (BD), CD150-PerCP-Cy5.5, and CD48-BV510, CD45.1-PE, and CD45.2-FITC. CMPs, MEPs, and MkPs were detected using a lineage cocktail, streptavidin-APC-Cy7, c-Kit-APC, Sca-1-PE-Cy7 (BD), CD34-AF700 (eBioscience), and CD16/32-BV421, CD41-BV605, CD150-PerCP-Cy5.5, CD45.1-PE, and CD45.2-FITC. Differentiated BM cells were detected using CD45.1-PE, CD45.2-FITC, B220-PE-Cy7, and CD11b-APC-Cy7. PB cells were detected using CD45.1-PE, CD45.2-FITC, B220-PE-Cy7, CD11b-APC-Cy7, CD4-APC, and CD8a-PerCP-Cy5.5. After staining, cells were analyzed on a flow cytometer (FACSCanto II; BD), and data were analyzed using FlowJo software (v.10.8). All antibodies were obtained from BioLegend unless otherwise noted.

Cell cycle and apoptosis assay

For cell cycle analysis, surface-stained cells were incubated with the Cytofix/Cytoperm Fixation and Permeabilization Solution Kit (BD, 554722) at room temperature for 20 min and then washed with 1X washing buffer (BD, 554723). The cells were then incubated with Ki67-FITC antibody (BioLegend, 151204) for 1 h on ice. DAPI was added and incubated at room temperature for 15 min before analysis. Apoptosis assays were performed using the Annexin V Apoptosis Detection Kit (BD, 556570). For Annexin V staining, cells were stained with surface markers, and 5 μL of Annexin V antibody was added and incubated at room temperature for 15 min in 1X binding buffer, followed by the DAPI incubation before analysis. After staining, cells were analyzed using a flow cytometer, and data were analyzed using FlowJo software.

CFU-spleen assay

0.5 million BM cells from AL or DR mice were injected via tail vein into lethally irradiated (9 Gy X-ray) 12-week-old Ly5.1 recipient mice. On day 7 post-transplantation, recipient mice were euthanized, and their spleens were collected and fixed in Davidson’s fixative (Sigma, H0290) for CFU-Spleen counting under a microscope (Olympus SZ51). Spleen cells were obtained by crushing the spleen in sterile PBS and filtering through a 40-μm cell strainer. The cells were then resuspended in RBC lysis buffer (BD, 555899) and incubated at room temperature for 5 min to lyse RBCs. Afterward, the cells were washed, counted, and incubated with flow cytometric antibodies on ice for 90 min. CMPs, MEPs and MkPs were detected using a lineage cocktail, streptavidin-APC-Cy7, c-Kit-APC, Sca-1-PE-Cy7 (BD), CD34-AF700 (eBioscience), and CD16/32-BV421, CD41-BV605, and CD150-PerCP-Cy5.5. After staining, cells were analyzed using a flow cytometer, and data were analyzed using FlowJo software.

Transplantation

For the primary BM transplantation, 1 million BM cells from young or aged mice subjected to AL or DR on day 1 post-chemotherapy along with 1 million BM cells from 12-week-old Ly5.1/Ly5.2 mice (competition cells) were injected via tail vein into lethally irradiated (X-ray, 9 Gy) 12-week-old Ly5.1 recipient mice. BM and PB analyses were performed at 4 months post-transplantation. For the secondary transplantation, 10 million BM cells from the post-transplant mice were injected via tail vein into lethally irradiated 12-week-old Ly5.1 recipient mice. BM and PB analyses of the secondary transplantation were performed at 4 months after transplantation.

scRNA-seq

Sample preparation and library construction

BM cells were harvested from four experimental groups: saline AL, saline DR, 5-FU-treated AL, and 5-FU-treated DR. For each group, BM cells from five individual mice were pooled to generate a single composite sample, in order to minimize individual variability and ensure consistency across samples for downstream analyses. HSPCs (Lin⁻ c-Kit^lo+hi) were collected by flow cytometric sorting (FACSAria III; BD) and then subjected to 10× Genomics Chromium capture platform. scRNA-seq libraries were prepared using the Chromium Single Cell 3ʹ Reagent Kits v.3 (10× Genomics), according to the manufacturer’s instructions. Briefly, approximately 2 × 10⁵ FACS-sorted cells were washed three times with 0.04% BSA in Dulbecco’s phosphate-buffered saline and resuspended to a final concentration of 700–1,200 cells/μL (viability ≥85%) prior to downstream processing. Cells were captured in droplets at a targeted cell recovery of cells. After the reverse transcription step, emulsions were broken and barcoded-cDNA was purified with Dynabeads, followed by PCR amplification. Amplified cDNA was then used for 3′ gene expression library construction. For gene expression library construction, 50 ng of amplified cDNA was fragmented and end-repaired, double-size selected with SPRIselect beads, and sequenced on a NovaSeq platform (Illumina) to generate 150-bp paired-end reads.

Single-cell RNA data processing and analysis

Raw sequencing reads were demultiplexed and aligned to the reference genome (mm10) using the 10× Genomics Cell Ranger (v.8.0.1) pipeline with default parameters. All downstream analyses were performed in R (v.4.3.3). Filtered feature-barcode matrices were loaded into Seurat (v.4.4.0), and quality control filters were applied: genes detected in fewer than three cells, cells with fewer than 200 or more than 7,500 transcripts, and cells with >25% mitochondrial gene expression were excluded. The retained data were normalized and scaled following Seurat’s standard workflow. Differential expression analysis was performed using Seurat’s FindMarkers function, and genes with p < 0.05 and fold change >2 were deemed significantly differentially expressed.

Enrichment analysis

Enrichment analyses of Kyoto Encyclopedia of Genes and Genomes pathways and GO terms were performed using the Metascape web platform (Metascape provides a biologist-oriented resource for the analysis of systems-level datasets; [http://metascape.org]). Parameters were set as follows: minimum overlap = 3, p value cutoff = 0.01, and minimum enrichment = 1.5. The resulting enrichment profiles were then re-visualized using ggplot2.

Western blotting

HSPCs were lysed in RIPA buffer (ServiceBio, G2002-100ML) supplemented with protease inhibitors (ServiceBio, G2006-250UL). Protein concentration was determined using a BCA assay kit (ServiceBio, G2026-200T). Equal amounts of protein were separated by SDS-PAGE and transferred onto PVDF membranes (ServiceBio, WGPVDF45). Membranes were blocked with a protein-free rapid blocking buffer (ServiceBio, G2052-500ML) and incubated overnight at 4°C with primary antibodies against LC3B (Abcam, ab192890, 1:2,000) and p62 (Abcam, ab109012, 1:10,000). β-actin (ServiceBio, GB15003, 1:1,000) was used as a loading control. After washing, membranes were incubated with HRP-conjugated goat anti-rabbit secondary antibody (ServiceBio, GB23303, 1:3,000). Protein bands were visualized using ECL reagents (ServiceBio, G2020-25ML) and quantified by densitometry.

Electron microscopy

HSPCs were collected and fixed in electron microscopy fixative for 4 h at 4°C. After fixation, the cells were washed three times with 0.1 M phosphate buffer (pH 7.4). They were then post-fixed in 1% osmium tetroxide in phosphate buffer for 2 h at room temperature, followed by three additional washes with phosphate buffer. The samples were dehydrated through a graded series of ethanol concentrations (50%, 70%, 80%, 90%, 95%, and 100%; 15 min each), followed by a 15-min treatment with 100% acetone. The dehydrated cells were embedded in an appropriate resin and polymerized at 60°C for 48 h. Ultrathin sections (∼60 nm) were cut using a Leica UC6 microtome, mounted onto copper grids, stained with lead citrate, and examined using a Hitachi HT7650 transmission electron microscope.

ATP content

Intracellular ATP content in HSPCs was determined using an ATP assay kit (ServiceBio, G3409) following the manufacturer’s instructions. Briefly, cells were resuspended in lysis buffer (50 μL per 0.5 million cells) and homogenized for 5 min. The lysates were centrifuged at 10,000 × g for 10 min, and the supernatants were collected. Aliquots of the supernatant were mixed with the ATP detection working solution, and luminescence was measured using a microplate reader (BioTek, Epoch).

Mitochondrial membrane potential, mitochondrial mass, and mitochondrial superoxide

After cell surface staining, cells were washed and incubated for 30 min at 37°C in 1 mL of FACS buffer (PBS +2% fetal bovine serum) containing one of the following mitochondrial dyes: TMRE (100 nM, Abcam, ab113852), MitoProbe JC-1 Assay Kit (2 μM, Invitrogen, M34152), or MitoSOX Green Mitochondrial Superoxide Indicator (5 μM, Invitrogen, M36006), according to the manufacturer’s instructions. Verapamil (50 μM, Sigma, V4629) was included in all staining conditions. These dyes were used to assess MMP, mitochondrial mass, and mitochondrial superoxide levels, respectively. After staining, cells were analyzed using a flow cytometer, and data were analyzed using FlowJo software.

Cell lines and tumor model

The MC38 murine colon carcinoma cell line was obtained from Bluefbio (Shanghai, China; BFN60808402) on October 1, 2024. Authentication was conducted using isoenzyme analysis and DNA fingerprinting. Mycoplasma testing was performed upon receipt, and the cell line was confirmed to be free of contamination prior to use in experiments. The cell line was cultured in an incubator set at 37°C in 5% CO₂ and 95% atmospheric air, maintained with DMEM (Gibco, 11965118) supplemented with 10% fetal bovine serum (Gibco, 16140071) and antibiotic-antimycotic (Gibco, 15240112). The cell line was assessed regularly to ensure it was free of Mycoplasma contamination. 0.5 million MC38 cells resuspended in 100 μL PBS were injected subcutaneously into the right flank of 12-week-old C57BL/6J female mice. Mice were acquired from Hunan SJA Laboratory Animal Co., Ltd., and a minimum of five mice were used per group. Assessment of tumor size began 6–8 days after cell inoculation and was performed once every 2–4 days using calipers. Tumor volume was calculated with the formula V = (d²×D)/2, where d stood for minor tumor axis and D for major tumor axis, and the data were presented as mean ± SD in mm³.

Clinical data collection

This retrospective study included clinical data from 100 patients diagnosed with gastric, colon, or rectal cancer who received chemotherapy with either 5-FU or capecitabine. The data were collected between January 2023 and December 2024 at the Second Affiliated Hospital of Nanchang University. All patients underwent at least 8 cycles of chemotherapy. Pretreatment BMI was recorded, and patients with BMI<18.5 were excluded. The lowest PB values observed during the 8 cycles of chemotherapy were collected, including complete blood count, age, sex, BMI, liver function (AST and ALT), renal function (urea, creatinine, eGFR), tumor type, tumor stage, chemotherapy regimen, dose intensity, number of chemotherapy cycles, dose adjustments, treatment interruptions, and use of anticoagulants or antiplatelet agents. The use of human data in this study was approved by the Institutional Ethics Committee of the Second Affiliated Hospital of Nanchang University. All procedures involving human participants were conducted in accordance with the Declaration of Helsinki (as revised in 2013). The requirement for informed consent was waived by the Ethics Committee due to the retrospective nature of the study.

Statistical analysis

GraphPad Prism software (v.9.5.1) was used for statistical analysis. To calculate p values, the unpaired two-tailed Student’s t test was used for two-group datasets and one-way ANOVA was used for multigroup (more than two groups) datasets. Gehan-Breslow-Wilcoxon test was used for survival rate analysis. All results were displayed as mean ± SD. ^∗p < 0.05; ^∗∗p < 0.01; ^∗∗∗p < 0.001; ^∗∗∗∗p < 0.0001; ns, not significant.

Resource availability

Lead contact

Requests for further information and resources should be directed to and will be fulfilled by the lead contact, Si Tao (ndefy11188@ncu.edu.cn).

Materials availability

This study did not generate new unique reagents.

Data and code availability

Raw sequence data have been deposited in the Genome Sequence Archive (GSA) at the National Genomics Data Center, China National Center for Bioinformation/Beijing Institute of Genomics, Chinese Academy of Sciences, under the accession number GSA: CRA031165 and are publicly available as of the date of publication (https://ngdc.cncb.ac.cn/gsa).

Acknowledgments

This work was supported by the National Natural Science Foundation of China (NSFC-82360287 and NSFC-82160029) and the Jiangxi Provincial Natural Science Foundation (20232ACB206014).

Author contributions

S.T., X.Q., and D.T. conceived and designed the experiments. S.T. and D.T. developed the methodology and secured funding. X.Q. and D.T. performed and analyzed the majority of the experiments. R.Q., Q.W., S.H., Z.T., and L.Z. assisted with H&E staining, imaging, and MK quantification. J.L., Z.C., S.Z., and Q.Z. provided suggestions and assisted with single-cell data analysis. The manuscript was written by S.T. and X.Q., with comments and revisions from all authors.

Declaration of interests

The authors declare no competing interests.

Published: April 2, 2026

Footnotes

Supplemental information can be found online at https://doi.org/10.1016/j.stemcr.2026.102869.

Supplemental information

Document S1. Figures S1–S6

mmc1.pdf^{(11.4MB, pdf)}

Table S1. Quality control metrics and mitochondrial-related genes in scRNA-seq data.

mmc2.xlsx^{(87.8KB, xlsx)}

Table S2. Baseline Clinical Characteristics of Patients.

mmc3.xlsx^{(248.9KB, xlsx)}

Document S2. Article plus supplemental information

mmc4.pdf^{(27.4MB, pdf)}

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Document S1. Figures S1–S6

mmc1.pdf^{(11.4MB, pdf)}

Table S1. Quality control metrics and mitochondrial-related genes in scRNA-seq data.

mmc2.xlsx^{(87.8KB, xlsx)}

Table S2. Baseline Clinical Characteristics of Patients.

mmc3.xlsx^{(248.9KB, xlsx)}

Document S2. Article plus supplemental information

mmc4.pdf^{(27.4MB, pdf)}

Data Availability Statement

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PERMALINK

Dietary restriction mitigates 5-fluorouracil-induced thrombocytopenia in aged mice via mitochondrial potentiation in hematopoietic stem cells and megakaryocyte progenitors

Xingxing Qiu

Duozhuang Tang

Jiahao Lin

Zhiyang Chen

Rongrong Qiu

Qiao Wang

Siyao He

Shaotong Zhang

Qiang Zhan

Zhendong Tao

Liu Zhang

Si Tao

Summary

Graphical abstract

Highlights

Introduction

Results

DR mitigates CIT

Figure 1.

DR protects HSPCs against 5-FU

Figure 2.

DR preserves HSC reconstitution capacity after 5-FU

Figure 3.

DR enhances mitochondrial pathways in HSCs and MkPs after 5-FU

DR enhances mitochondrial function in HSCs and MkPs after 5-FU

Mitochondrial potentiation rescues PLT and HSC regeneration in AL mice after 5-FU

Figure 4.

Mitochondrial dysfunction impairs PLT protection and hematopoietic function in DR mice after 5-FU

Figure 5.

DR delays tumor growth without impairing its efficacy in mitigating CIT

Figure 6.

PLT count inversely correlates with BMI in 5-FU-treated cancer patients

Figure 7.

Discussion

Methods

Mice and diets

Chemical treatment of mice

5-FU treatment

Abx treatment

MQ and CCCP treatment

PB cell counting

H&E staining

TUNEL staining

PS exposure assay

Flow cytometry

Cell cycle and apoptosis assay

CFU-spleen assay

Transplantation

scRNA-seq

Sample preparation and library construction

Single-cell RNA data processing and analysis

Enrichment analysis

Western blotting

Electron microscopy

ATP content

Mitochondrial membrane potential, mitochondrial mass, and mitochondrial superoxide

Cell lines and tumor model

Clinical data collection

Statistical analysis

Resource availability

Lead contact

Materials availability

Data and code availability

Acknowledgments

Author contributions

Declaration of interests

Footnotes

Supplemental information

References

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

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RESOURCES

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