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The Journal of Biological Chemistry logoLink to The Journal of Biological Chemistry
. 2026 Jan 12;302(3):111143. doi: 10.1016/j.jbc.2026.111143

Starvation of leukemic cells enhances DNA damage-induced apoptosis in vitro via ROS/p38 MAPK and prevents leukemia progression in fasting xenograft mice

Ajay Yadav 1, Nina Richartz 1, Sampada Bhagwat 1, Marta Maria Burman 2, Monica Cheng Munthe-Kaas 2, Bjørn Steen Skålhegg 3, Jonas Aakre Wik 3, Soheil Naderi 4, Heidi Kiil Blomhoff 1,∗
PMCID: PMC12914434  PMID: 41534829

Abstract

Most children with acute lymphoblastic leukemia (ALL) achieve long-term survival due to intensive multimodal chemotherapy. However, the use of cytotoxic DNA-damaging agents is frequently associated with severe long-term side effects, prompting continued efforts to improve treatment strategies. This study explores the potential of starving the leukemic cells to enhance the efficacy of DNA-damaging therapy in ALL. Previous work demonstrated that cAMP signaling attenuates DNA damage-induced apoptosis in ALL cells, both in vitro and in a xenograft model. The current findings show that glucose and serum deprivation reverse the effect of cAMP, converting it from a survival factor to a promoter of DNA damage-induced apoptosis in ALL-derived cell lines and patient-derived leukemic cells in vitro. The starvation-induced sensitization was independent of p53 but was shown to require increased levels of reactive oxygen species (ROS). In turn, the elevated ROS levels enhanced the activation of the mitogen-activated protein kinase p38 (p38 MAPK). The resulting augmented cell death was inhibited both by the ROS scavenger N-acetyl cysteine and the p38 MAPK inhibitor SB 202190. The translational potential of increasing the efficacy of DNA damaging agents in starving ALL cells was supported by in vivo data showing that intermittent fasting, combined with subtherapeutic doses of irradiation, significantly inhibits the leukemia progression in a xenograft model of severe combined immunodeficiency mice.

Keywords: ALL, apoptosis, cAMP, DNA damage, fasting, p38 MAPK, ROS, Xenograft

B-cell precursor acute lymphoblastic leukemia (ALL) is the most common pediatric cancer (1). Thanks to intensive multimodal chemotherapy, typically over years, the long-term survival rate of the children with ALL is now approaching 90 % (2, 3). However, the improved survival often comes with a price in terms of long-term morbidity due to harmful side effects of the treatment (4). Current standard protocols for treatment of pediatric ALL include cytotoxic therapeutics that ultimately leads to DNA damage both in the leukemic and normal cells (5, 6). Consequently, efficient treatment of ALL may typically result in late effects such as cardiotoxicity, neurocognitive and neuroendocrine problems, as well as risk of secondary malignancies (2, 3). Hence, there is a continuous need for improved protocols that may increase the efficiency of the therapy and thereby reduce the devastating late effects of the treatment.

Inhibiting the cyclic adenosine monophosphate (cAMP) signaling pathway serves as a potential strategy for improving the efficacy of DNA-damaging treatment for ALL patients (7, 8, 9, 10). ALL develops in niches in the bone marrow, near stromal cells producing cAMP-activating compounds like prostaglandin E2 (PGE2) (11, 12, 13, 14, 15). Elevated endogenous cellular cAMP levels reduce the DNA damage-induced killing of ALL cells in vitro (7, 8, 14, 16), and we have shown that inhibiting the production of PGE2 by indomethacin in vivo delays the progression of leukemia in a xenograft model of ALL in mice (17).

High level of cAMP is generally regarded as a starvation signal downstream of glucagon receptors on target cells, activating signaling cascades that result in depletion of cellular ATP levels (18, 19). There are functional glucagon receptors on lymphoid cells and tissues (20, 21), but their roles as regulators of ATP production and metabolism in these cells are scarcely studied. There has been ample attention in recent years on the prospects of improving cancer therapy by calorie restriction and intermittent fasting, both in terms of increasing the efficiency and reducing the side effects of treatments (22, 23, 24, 25). In an interesting paper by Lu and coworkers they demonstrated that intermittent fasting of mice selectively blocks the development of ALL (26). In line with their study, Chan and colleagues showed that restricted nutrition had inhibitory effects on development of ALL but not on AML (27). Subsequent studies employing fasting-mimicking conditions in vitro and intermittent fasting in vivo have demonstrated promising effects on the development and treatment of hematological malignancies, including of ALL (12, 26, 27, 28). Fasting constitutes starvation signals that may lead to elevated levels of endogenous cAMP (19). Therefore, previous findings from our group identifying cAMP as promoter of leukemia progression (7, 8, 16) and as an inhibitor of DNA-damaging treatment efficacy in ALL (12), may appear contradictory to the reported beneficial effects of fasting on cancer therapy. Consequently, we here aimed to reveal how cAMP signaling regulates DNA-damaging responses in fasting versus non-fasting ALL cells in vitro and in vivo. Starvation of ALL cells in vitro was found to turn cAMP signaling from inhibiting to enhancing DNA damage-induced cell death. Mechanistically, this shift was shown to depend on reactive oxygen species (ROS)-mediated activation of the p38 MAPK. In vivo, we used a xenograft model of ALL in severe combined immunodeficiency (SCID) mice (26) to show that intermittent fasting significantly improved the inhibitory effect of DNA-damaging irradiation (IR) on leukemia progression. Taken together, our results support a beneficial role of fasting on targeting ALL by DNA-damaging treatment.

Results

Starvation enhances DNA damage-induced killing of ALL cells by inhibiting the cAMP-mediated protection

We investigated the effects of cAMP signaling on DNA-damage responses in fasting ALL cells in vitro. The adenylyl cyclase activator forskolin was used to activate the cAMP signaling pathway in the ALL-derived REH cells, and DNA damage was induced by exposing the cells to X-ray-mediated IR, 10 Gy or to the chemotherapeutic agent doxorubicin (300 nM). In these experiments, the ALL cells were cultured in the presence or absence of serum and/or glucose to mimic either optimal - or starving growth conditions (29, 30, 31, 32) (see Materials and Methods and Figure legends for details). Alternatively, we used Earle's Balanced Salt Solution to induce starvation of the cells (33, 34). Cell death was assessed by staining the cells with propidium iodide (PI) 24 hours post IR. In line with our previous findings (7, 9), 70 μM of forskolin inhibited IR-induced death of the ALL cells cultured in complete medium (CM) by nearly 30 % (Fig. 1A). Culturing the cells in normal RPMI medium in the absence of serum fetal bovine serum (RPMI - FBS) or in RPMI medium lacking glucose but supplemented with serum (RPMI (- Glc) + FBS) had a marginal effect on the viability of the cells, and only slightly reduced the protecting effect of forskolin on the IR-induced cell death. By contrast, RPMI medium devoid of both glucose and serum (RPMI (- Glc) - FBS); hereafter referred to as depleted medium (DM), prevented the protecting effect of forskolin on IR-induced cell death. Indeed, forskolin under these culture conditions not only markedly enhanced the IR-mediated killing of the cells, but also alone reduced the viability of the REH cells as measured after 24 hours treatment (Fig. 1A). Re-adding glucose to the cells cultured in DM (RPMI (- Glc) - FBS + Glc) partially restored the protecting effects of forskolin on IR-mediated cell death. Culturing the cells in the Earle's Balanced Salt Solution starvation medium also reduced the protecting effect of forskolin, but the effect was not as pronounced as culturing the cells in medium deprived of both glucose and serum. Based on the results shown in Figure 1A, the DM was selected as the culture condition to mimic fasting in vitro throughout the study. Forskolin under these culture conditions markedly enhanced the IR-induced killing of the cells from approximately 60 % to more than 90 %, in contrast to the nearly 30 % inhibition of IR-induced cell death obtained by forskolin in CM (Fig. 1B).

Figure 1.

Figure 1

Figure 1

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Starvation of ALL cells converts cAMP signaling into an enhancer of DNA damage-induced killing of the cells.A, REH cells (0.4 × 106 cells/ml) were incubated under various culture conditions to mimic fasting by inducing starvation of the cells, either by adding or removing fetal bovine serum (FBS) and/or glucose (Glc) from the culture medium, in the presence or absence of forskolin (Forsk, 70 μM) and /or irradiation (IR, 10 Gy). The cells were treated with forskolin for 45 min prior to IR, and the percentages of dead cells were monitored by flow cytometry of cells stained with propidium iodide (PI) 24 hours after IR. B, REH cells (0.4 × 106 cells/ml) were incubated in complete medium (CM, normal RPMI medium with 10 % FBS) or depleted medium (DM, glucose free RPMI without supplementation of FBS) in the presence or absence of forskolin (Forsk, 70 μM) and/or IR, 10 Gy as in panel A, and the percentages of dead cells were monitored by flow cytometry of cells stained with PI 24 hours after IR. C, REH cells (0.4 × 106 cells/ml) were cultured in CM or DM in the presence or absence of forskolin (Forsk, 70 μM) and /or IR, 10 Gy as in panel A. The percentages of dead cells were monitored by flow cytometry of cells stained with PI at 1, 2 or 4 hours after IR. D, REH cells (0.4 × 106 cells/ml) were incubated in CM or DM. The cells were treated with forskolin (Forsk, 70 μM) for 45 min prior to treatment with doxorubicin (Doxo, 300 nM). Flow cytometry analysis was performed 24 hours after the addition of doxorubicin and shown as the percentages of dead cells based on low forward scatter (FSC) and high side scatter. E, NALM-6 cells (0.4 × 106 cells/ml) and (F) 697 cells (0.4 × 106 cells/ml) were cultured in CM or DM in the presence or absence of forskolin (Forsk, 70 μM) and /or IR, 2.5 Gy. G and H, leukemic cells (0.5 × 106 cells/ml) freshly isolated from bone marrow aspirates of 2 patients diagnosed with ALL (ALL#82 and ALL#101) were cultured in SFEM medium. After 72 hours, the cells were incubated in CM or DM in the presence or absence of forskolin (Forsk, 70 μM) and/or IR, 5 Gy. The cells were treated with forskolin for 45 min prior to IR or doxorubicin, and the percentages of dead cells were monitored by flow cytometry of cells stained with PI 24 hours after IR. The data represent the mean ± SEM, n = 5 (panel A), n = 7 (panel B), n= 6 (panel C), n= 4 (panel D), n = 3 (panel E), n = 3 (panel F). ∗P < 0.05 (paired t-test). In all panels, ‘control’ represents cells cultured in the relevant medium alone. The content and abbreviations of the various culture media: 1) CM = RPMI + FBS, denotes normal RPMI supplemented with 10 % FBS 2) RMPI - FBS, denotes normal RPMI not supplemented with FBS 3) RPMI (- Glc) + FBS, denotes glucose-free RPMI supplemented with 10 % FBS 4) DM = RPMI (- Glc) - FBS, denotes glucose-free RPMI not supplemented with FBS 5) DM + Glc denotes glucose-free RPMI without FBS, but supplemented with 5 mM glucose 6) Earle′s Balanced Salt Solution. CM, complete medium; DM, depleted medium; PI, propidium iodide.

Having demonstrated that cAMP signaling enhanced the IR-induced cell death in DM measured after 24 hours (Fig. 1B), we next sought to determine how early the effect of starvation could be observed. To this end, the cells were exposed to IR in the presence or absence of forskolin in either CM or DM and then harvested at various time points after IR (1, 2 or 4 hours, respectively). As shown in Figure 1C, 10 Gy IR did not induce significant cell death as measured by PI staining at any of these time points. However, after culturing the cells in DM for 2 hours, forskolin alone reduced the viability of the cells, and together with IR enhanced the cell death to nearly 40 %. By 4 hours, a synergistic effect of combining forskolin and IR was observed. Thus, forskolin enhanced the IR-induced killing of the cells from less than 10 % to nearly 70 %, with forskolin alone killing approximately 35 % of the cells.

Although IR is commonly used for treating metastases of ALL in the brain and testis (35), other DNA-damaging agents including doxorubicin (Doxo) are frequently used in standard primary care therapy of ALL. Doxo was therefore used in a similar setup as described in Figure 1B. Due to the interference of Doxo with analyses of PI-stained cells, flow cytometry-based changes in the cells’ forward scatter (FSC, estimating cell size) and side scatter (SSC, estimating cell granularity) were used as a measure of cell death. As shown in Figure 1D, culturing the cells with the combination of forskolin and Doxo in DM enhanced the cell death (lower FSC and higher SSC) from approximately 35 % to 58 %. This supports the notion that starvation-mimicking fasting of the ALL cells reverses the cAMP-mediated protection and thereby enhances DNA damage-induced killing of the cells.

To investigate whether a similar effect could be observed in other ALL-derived cell lines, we included NALM-6 and 697 cells. cAMP signaling under starving conditions enhanced the IR-induced cell death both in NALM-6 (Fig. 1E) and 697 (Fig. 1F) cells.

To examine potential clinical relevance, primary leukemic cells isolated from bone marrow aspirates of two patients (ALL#82 and ALL#101) diagnosed with pediatric ALL (see Table 1 for patient characteristics) were utilized. The viability of the patient-derived ALL cells was generally low, due to both the handling of the bone marrow aspirates and the purification protocol itself, and was approximately 40% for ALL#82 and 60 % for ALL#101. The data in Figure 1, G and H are therefore presented as changes in IR-specific cell death and show that culturing the primary ALL cells in DM reverses the protective effects of forskolin on IR-induced cell death after 24 hours in the same manner as for REH cells. Thus, in DM, the IR-induced cell death was enhanced from approximately 22 % to 65 % in cells from ALL patient #82, and from less than 10 % to approximately 28 % in cells from ALL patient #101.

Table 1.

Characteristics of patient-derived ALL cells

ALL#82 ALL#101
Age: years (y), months (m) 5y 11m 9y 4 m
Sex Female Female
Bone marrow infiltration at diagnosis (% CD19+/CD10+) 92% 83%
Cytogenetics ETV6-RUNX1 positive No cytogenetic aberrations, no high-risk markers detected
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The starvation-induced killing of ALL cells is via apoptosis and involves the production of ROS

Having established that starvation turns cAMP from an inhibitor to an enhancer of IR- and Doxo-induced cell death, the next step was to verify that the mode of cell death was apoptosis. Two commonly used methods for identifying and analyzing apoptosis were used; the assessment of the loss of mitochondrial membrane potential and the analysis of Annexin V binding to phosphatidylserine-exposing cells (36, 37). The loss of mitochondrial membrane potential was assessed by staining the cells with the tetramethylrhodamine methyl ester (TMRM) dye (36), and the TMRM intensity was measured by flow cytometry. Cells with low TMRM intensity exhibit reduced mitochondrial membrane potential, indicative of apoptotic cells (Fig. 2A). As shown in Figure 2B, the percentage of IR-treated cells with low TMRM intensity was enhanced by forskolin in cells cultured in DM. Moreover, forskolin increased the percentage of Annexin V-positive cells after IR-treatment in DM from 40 % to 82 %, while reducing the percentage of Annexin V-positive cells by 30 % when combined with IR in CM (Fig. 2C). In addition, inhibition of caspases by Z-VAD significantly reduces the cell death from nearly 80 % to 35 % when ALL cells were treated with forskolin and IR for 24 hours in DM (Fig. 2D). Together, these data suggest that starvation enhances DNA damage-induced apoptosis of ALL cells by reversing the inhibitory effects of cAMP signaling.

Figure 2.

Figure 2

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Starvation enhances the DNA damage-induced killing of ALL cells by apoptosis and involves the production of ROS.A and C, REH cells (0.4 × 106 cells/ml) were cultured in CM or DM in the presence or absence of forskolin (Forsk, 70 μM) and /or IR, 10 Gy. The cells were treated with forskolin for 45 min prior to IR. (A and B) The loss of mitochondrial membrane potential was assessed by flow cytometry of cells stained with tetramethylrhodamine methyl ester (TMRM) 24 hours after IR. A, the gating strategy, with TMRM-low cells representing the apoptotic cell population in one representative experiment, Control in CM = unstimulated cells cultures in CM and IR + Forsk in DM = IR plus forskolin-treated cells cultured in DM. B, the quantification of the TMRM experiments. The data represent the mean ± SEM, n = 3, ∗P < 0.05 (paired t test). C, the percentages of apoptotic cells (Annexin V+ cells) were assessed by flow cytometry of cells stained with Annexin V and 7-AAD, 24 hours after IR. The data represent the mean ± SEM, n = 4 ∗P < 0.05 (paired t-test). D, REH cells (0.4 x 106 cells/ml) were incubated in DM in the presence or absence of the caspase inhibitor Z-VAD (100 μM) for 1 hour, followed by treatment with or without forskolin (Forsk, 70 μM) for 45 min prior to IR, 10 Gy. The percentages of dead cells were monitored by flow cytometry of cells stained with PI, 24 hours after IR. The data represent the mean ± SEM, n = 4, ∗P < 0.05 (paired t-test). E, REH cells (0.4 × 106 cells/ml) were cultured in CM or DM in the presence or absence of forskolin (Forsk, 70 μM) and /or IR, 10 Gy. The cells were treated with forskolin for 45 min prior to IR, and total reactive oxygen species (ROS) levels were monitored by flow cytometry of cells stained with CellROX green 1 hour after IR. The data represent the mean ± SEM, n = 6, ∗P < 0.05 (paired t-test). F, REH cells (0.4 x 106 cells/ml) were cultured in DM in the presence or absence of 30 mM NAC for 30 min, followed by incubation with or without forskolin (Forsk, 70 μM) for 45 min prior to IR, 10 Gy. Total ROS levels were analyzed as in panel A. The data represent the mean ± SEM, n = 3, ∗P < 0.05 (paired t-test). G, REH cells (0.4 × 106 cells/ml) were cultured in DM in the presence or absence of 30 mM of NAC for 30 min before incubating the cells with or without forskolin (Forsk, 70 μM) for 45 min prior to IR, 10 Gy. The percentages of dead cells were monitored by flow cytometry of cells stained with PI, 24 hours after IR. The data represent the mean ± SEM, n = 5, ∗P < 0.05 (paired t-test). In all panels, ‘control’ represents cells cultured in the relevant medium alone. CM, complete medium; DM, depleted medium; ROS, reactive oxygen species; TMRM, tetramethylrhodamine methyl ester.

The next aim was to reveal the mechanisms by which forskolin enhances DNA damage-induced apoptosis in starving ALL cells. Starvation is known to enhance ROS production, and hence, to oxidative stress resulting in cell death (38, 39, 40, 41, 42, 43). Accordingly, ROS levels were assessed by flow cytometry analyses of ALL cells stained with the CellROX green reagent. Starvation alone enhanced the ROS levels nearly 7-fold (Fig. 2E). Neither forskolin nor IR alone further enhanced the ROS levels significantly, whereas forskolin added prior to IR further enhanced the IR-specific ROS levels more than 4-fold in DM compared to the levels in forskolin plus IR-treated cells in CM. In line with the marginal effects on cell death of depleting the cells of either only glucose (medium 2) or only serum (medium 3) (Fig. 1A), the ROS levels were barely changed when culturing the cells under these conditions as compared to culturing the cells in CM (Fig. S1). The ROS scavenger N-acetyl cysteine (NAC) reduced the levels of ROS in starving cells (cells cultured in DM) as well as the ROS levels in starving cells treated with the combination of forskolin and IR (Fig. 2F). Notably, when measuring the cell death 24 hours after IR in DM, the ROS scavengers NAC reduced the IR-induced cell death in the presence of forskolin from nearly 90 % to 38 % (Fig. 2G). Taken together, these data indicate that culturing the ALL cells in starvation medium enables forskolin to enhance the ROS levels in IR-treated cells, which in turn promotes the DNA damage-induced cell death.

ROS is involved in starvation-induced autophagy promoted by cAMP-signaling in the presence of DNA-damaging agents

Autophagy is generally regarded as a cellular survival mechanism to cope with stressful conditions such as starvation or DNA damage, and it is one of the processes known to link ROS and viability (44, 45, 46). In line with this, we previously showed that cAMP-mediated inhibition of DNA damage-induced apoptosis in ALL cells was accompanied by enhanced ROS-mediated autophagy (9). Here however, we observed that when ALL cells are cultured in DM, cAMP-mediated enhancement of IR-induced ROS levels leads to cell death. This may suggest that ROS produced by starving cells leads to a process known as cell death with autophagy (47). In this process, starving cells respond to stress by increasing ROS and autophagy levels in an attempt to survive, but the cells ultimately fail to adapt and thereby undergo apoptosis despite the induced autophagy. We therefore tested if cAMP signaling in starving cells would enhance the IR-induced levels of ROS-mediated autophagy.

Autophagy was analyzed by two established methods; Western blot analysis to detect the lipidated form of LC3 (LC3-II) (48), and by measuring CYTO-ID intensity using flow cytometry (49). The CYTO-ID dye stains autophagic vesicles, and our previous work has shown that CYTO-ID staining yields results comparable to LC3-II detection (9, 10). Measurements of LC3-II were performed in the presence or absence of the lysosomal inhibitor bafilomycin A1, which induces accumulation of autophagosomes and correlates with an increased rate of autophagic flux (50). As shown (Fig. 3A, left panel), culturing the ALL cells in DM, by itself enhanced the levels of LC3-II. When quantifying these effects (Fig. 3, right panel), it was shown that forskolin cultured in DM further enhanced the level of LC3-II, and more than doubled the levels of LC3-II in the IR-treated cells. The ability of cAMP signaling to enhance IR-induced autophagy was confirmed by staining the cells with CYTO-ID (Fig. 3B). Knockdown of the autophagy-promoting gene ULK1 by siRNA reduced the CYTO-ID staining of the cells (Fig. 4B), and the knockdown of ULK1 protein was verified by western blot analyses (Fig. S2). NAC reduced the elevated ROS levels by nearly 30 % (Fig. 3C), demonstrating the involvement of ROS in the forskolin-mediated enhancement of IR-induced autophagy. In support of our hypothesis to assess whether starvation-induced autophagy contributes to cell death in ALL cells, we inhibited autophagy by knocking down ULK1 (Fig. 3D), or by using the autophagy inhibitor 3-MA (Fig. 3E). Neither approach reduced the cell death observed in cells treated with IR and forskolin.

Figure 3.

Figure 3

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ROS is involved in starvation-induced autophagy promoted by cAMP signaling in the presence of DNA damaging agents.A, REH cells (0.8 × 106 cells/ml) were cultured in CM or DM in the presence or absence of forskolin (Forsk, 70 μM) for 45 min prior to IR, 7 Gy. The cells were harvested 24 hours after IR, and Baf1 (5 nM) was added for the last 4 hours. Total cell lysates were subjected to immunoblot analyses with antibodies against LC3. Antibodies against vinculin were used as a control of equal loading. Left panel shows one representative Western blot of four independent experiments. Right panel shows a quantified overview of LC3-II signal intensity relative to the vinculin signal. The data represent the mean ± SEM, n = 4, ∗P < 0.05 (paired t-test). B, REH cells (0.4 × 106 cells/ml) were transfected with control siRNA or with siRNA against ULK1 as described in Materials and Methods. After 4 hours, cells were cultured in DM for 45 prior to IR, 5 Gy. For monitoring of autophagy, the cells were subjected to CYTO-ID staining 24 hours after IR, and the mean fluorescence intensity (MFI) was analyzed by flow cytometry. The data represent the mean ± SEM, n = 4, ∗P < 0.05 (paired t-test). C, REH cells (0.4 × 106 cells/ml) were cultured in DM in the presence or absence of 10 mM NAC for 30 min, followed by incubation with forskolin (Forsk, 70 μM) for 45 min prior to IR, 10 Gy. The cells were subjected to CYTO-ID staining 24 hours after IR, and MFI was analyzed by flow cytometry. The data represent the mean ± SEM, n = 4, ∗P < 0.05 (paired t-test). D, REH cells (0.4 × 106 cells/ml) were transfected with control siRNA or with siRNA against ULK1 and treated with or without forskolin (Forsk, 70 μM) and /or IR, 5 Gy as in panel B. The percentages of dead cells were monitored by flow cytometry of cells stained with PI 24 hours after IR. The data represent the mean ± SEM, n = 3, ∗P < 0.05 (paired t-test). E, REH cells (0.4 × 106 cells/ml) were incubated in DM in the presence or absence of the autophagy inhibitor 3-MA (500 μM) for 1 hour, followed by treatment with or without forskolin (Forsk, 70 μM) for 45 min prior to IR, 10 Gy. The percentages of dead cells were monitored by flow cytometry of cells stained with PI 24 hours after IR. The data represent the mean ± SEM, n = 4, ∗P < 0.05 (paired t-test). In all panels, ‘control’ represents cells cultured in the relevant medium alone. CM, complete medium; DM, depleted medium; MFI, mean fluorescence intensity.

Figure 4.

Figure 4

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Starvation-induced killing of ALL cells involves ROS-mediated activation of p38 MAPK.A, REH cells (0.8 × 106 cells/ml) were cultured in CM or DM in the presence or absence of forskolin (Forsk, 70 μM) for 45 min prior to IR, 7 Gy. The cells were harvested 1 hour after IR, and total cell lysates were subjected to immunoblot analyses with antibodies against p38 MAPK (p38), phosphorylated p38 MAPK (pp38). Antibodies against vinculin were used as a control of equal loading. Left panel (upper) shows one representative Western blot of six independent experiments. Right panel shows a quantified overview of pp38 signal intensity relative to the vinculin signal. The data represent the mean ± SEM, n = 6, ∗P < 0.05 (paired t-test). Left panel (lower) shows a representative Western blot from one of one experiment using antibodies against p38 and vinculin. B, REH cells (0.8 × 106 cells/ml) were cultured in DM in the presence or absence of 40 μM of the p38 MAPK inhibitor (SB 202190) for 30 min, followed by incubation with or without forskolin (Forsk, 70 μM) for 45 min prior to IR, 7 Gy. The cells were harvested 1 hour after IR, and total cell lysates were subjected to immunoblot analyses with antibodies against p38, pp38 and vinculin. Left panel (upper) shows one representative Western blot of four independent experiments. Right panel shows a quantification of the pp38 signal intensity relative to vinculin signal. The data represent the mean ± SEM, n = 3, ∗P < 0.05 (paired t-test). Left panel (lower) shows a representative Western blot from one of one experiment using antibodies against p38 and vinculin. C, REH cells (0.8 × 106 cells/ml) were cultured in DM in the presence or absence of 10 mM NAC for 30 min, followed by incubation with or without forskolin (Forsk, 70 μM) for 45 min prior to IR, 7 Gy. The cells were harvested 1 hour after IR, and total cell lysates were subjected to immunoblot analyses with antibodies against pp38 and vinculin. Left panel shows one representative Western blot of three independent experiments. Right panel shows quantification of the pp38 signal intensity relative to the vinculin signal. The data represent the mean ± SEM, n = 4, ∗P < 0.05 (paired t-test). In all panels, ‘control’ represents cells cultured in the relevant medium alone. CM, complete medium; DM, depleted medium.

Starvation enhances cAMP/DNA damage-induced p38 MAPK activation via ROS

The next objective was to elucidate the mechanistic link between the enhanced ROS levels and the induced apoptosis in starving ALL cells treated with forskolin and IR. ROS is reported to activate mitogen-activated kinases such as p38 MAPK (51, 52, 53), which in turn can induce apoptosis (54, 55, 56, 57). Accordingly, the activation of p38 MAPK was assessed by Western blot analyses of the phosphorylated form of the kinase. Culturing the cells in DM significantly enhanced the effect of forskolin on IR-induced pp38 MAPK protein levels as examined 1 hour after exposure to IR (Fig. 4A). Neither forskolin nor IR alone significantly enhanced the kinase activity compared to the activities noted in cells cultured in CM, but kinetic experiments revealed that the effect of forskolin on IR-induced phosphorylation of p38 MAPK declined after 3 hours (Fig. S3). The specificity of the p38 MAPK activation assay was verified by the fact that the elevated level of pp38 MAPK in the forskolin/IR-treated cells was significantly reduced by the p38 kinase inhibitor SB202190 (Fig. 4B). The involvement of ROS in the forskolin/IR-mediated activation of p38 MAPK was further demonstrated by pre-treating the cells with the ROS scavenger NAC, which reduced the pp38 MAPK levels in IR-treated cells co-stimulated with forskolin for 1 hour in DM (Fig. 4C).

Apoptosis induced by forskolin and IR in starving ALL cells involves the activation of p38 MAPK

Activation of p38 MAPK has in other cell systems been shown to link elevated ROS levels to apoptosis (54, 56, 58, 59). Accordingly, we addressed the involvement of p38 MAPK in the forskolin plus IR-mediated killing of ALL cells cultured in DM. The p38 MAPK inhibitor SB202190 significantly reduced the death of forskolin plus IR-treated cells by 30 %, as assessed both by PI- (Fig. 5A) and TMRM-staining (Fig. 5B).

Figure 5.

Figure 5

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Apoptosis induced by forskolin and IR in starving ALL cells involves the activation of p38 MAPK.A and D, REH cells (0.4 × 106 cells/ml) were cultured in DM in the presence or absence of 10 μM of the p38 MAPK inhibitor SB 202190 for 30 min, followed by incubation with forskolin (Forsk, 70 μM) for 45 min prior to IR, 10 Gy. A, the percentages of dead cells were monitored by flow cytometry of cells stained with PI 2 hours after IR. The data represent the mean ± SEM, n = 4, ∗P < 0.05 (paired t-test). B, the loss of mitochondrial membrane potential was assessed by flow cytometry of cells stained with TMRM 2 hours after IR and shown as percentage of TMRM-low cells, representing apoptotic cells. The data represent the mean ± SEM, n = 3, ∗P < 0.05 (paired t-test). C, total ROS levels were analyzed by staining the cells with CellROX green 1 hour after IR, and the MFI was analyzed by flow cytometry. The data represent the mean ± SEM, n = 4, ∗P < 0.05 (paired t-test). D, the cells were subjected to CYTO-ID staining for detection of autophagy 24 hours after IR, and the MFI was analyzed by flow cytometry. The data represent the mean ± SEM, n = 3, ∗P < 0.05 (paired t-test). E, model explaining how DNA damage-induced killing of ALL cells is enhanced under starving culture conditions in the presence of cAMP signaling. Our model suggests that starvation promotes cAMP-mediated (via forskolin, Forsk) enhancement of DNA damage-induced (via IR) killing of ALL cells by enhancing the ROS levels. Increased ROS levels activate p38 MAPK, which subsequently leads to apoptosis. Additionally, ROS is involved in starvation-induced autophagy, which is prompted by cAMP signaling in the presence of a DNA damaging agent. Notably, activation of p38 MAPK appears to function both upstream and downstream of ROS. According to our model, the cells die with – but not due to the high levels of autophagy. In all panels, ‘control’ represents cells cultured in the relevant medium alone. DM, depleted medium; MFI, mean fluorescence intensity.

This demonstrated a functional link between p38 MAPK, cAMP-mediated ROS levels and DNA damaged-induced apoptosis in starving ALL cells, but the mechanism underlying ROS generation was still not established. It should be noted, that p38 MAPK in other cell types is not only activated by ROS, but can also induce ROS levels (58). We therefore analyzed ROS levels in cells cultured in CM or DM in the presence or absence of SB202190, and we showed that inhibition of p38 MAPK completely abrogated the ROS induction one hour after IR exposure in cells cultured in DM in the presence of forskolin (Fig. 5C). Similarly, SB202190 also prevented the induction of autophagy, as measured by CYTO-ID staining 24 hours post-IR (Fig. 5D). This suggests that p38 MAPK activation may function both upstream and downstream of ROS production, as well as upstream of cAMP-induced autophagy in starving IR-treated ALL cells (Fig. 5E).

Intermittent fasting sensitizes ALL cells to DNA-damaging treatment in vivo

Depletion of glucose and serum in vitro appeared to reverse the cAMP-induced protection against IR-induced cell death, and the next aim was therefore to test the effect of starvation in an in vivo animal model of ALL. To this end, we examined the impact of intermittent fasting on the ability of IR to reduce the progression of leukemia in a previously described (17) xenograft model of human ALL. Bone marrow transplantation of human ALL cells in immune deficient mice is a well-established model for studying leukemia progression (60). In this model, the ALL cells in the bone marrow are continuously exposed to cAMP driven by PGE2 produced by stromal cells (12), thereby mimicking the forskolin-supplemented conditions used in vitro. Using the same experimental setup and conditions for establishing the xenograft ALL model in SCID mice, the animals were subjected to intermittent fasting or maintained on a normal diet for approximately 25 days. Leukemia progression was monitored using in vivo imaging by detecting the firefly luciferase (fLuc) activity from the transduced ALL cells. The effect of whole-body IR (3.5 Gy) on leukemia progression was monitored in both fasting and non-fasting mice. As shown in Figure 6, the leukemia progressed gradually over more than 3 weeks, and the experiment was terminated based on pre-determined endpoints and clinical signs of illness. Intermittent fasting and suboptimal doses of IR both reduced the leukemic cell growth when applied separately, and when combined nearly completely suppressed the leukemia progression. Together with the results from the in vitro starvation cultures of ALL cells, our data support the notion that intermittent fasting enhances IR-induced responses in ALL cells.

Figure 6.

Figure 6

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Intermittent fasting enhances the effect of IR on ALL in vivo. ALL Xenograft SCID mice (5 mice per treatment group) were subjected to intermittent fasting (as described in Materials and Methods) or fed a normal diet. The mice were treated with or without suboptimal doses of whole-body IR totaling 3.5 Gy (1 Gy on the 9th day and 2.5 Gy on the 12th day post i.t. injection) as described in the Experimental Procedures section. Development of leukemia was monitored by noninvasive in vivo imaging of luminescence from the leukemic cells of the xenograft mice. Upper panel, luminescence images of xenograft REH mice taken at the experimental endpoint (23 days after intratibial injection). Lower panel, Xenograft luciferase activity [photons per second, (p/s)] over time. Ctrl = mice subjected to a normal diet; Fasting = mice subjected to intermittent fasting; IR = mice treated with whole body; IR + fasting = mice subjected to intermittent fasting and treated with IR. Each data point represents the mean ± SEM signal intensity of the 5 xenograft mice in each treatment group. ∗P < .05 (unpaired Student t-test).

Discussion

Strategies to improve the therapeutic index of cytotoxic DNA damaging treatments are needed to minimize the devastating long-term effects of current multimodal chemotherapy protocols for pediatric ALL. Previous work from our group has shown that cAMP signaling protects ALL cells from cell death induced by IR and DNA damaging chemotherapy in vitro (7, 8, 14, 16), and drives leukemia progression in a xenograft mouse model (17). Growing evidence suggests that intermittent fasting can improve the therapeutic index of cancer therapies by sensitizing cancer cells to chemotherapy (61, 62, 63). In the present study, we have explored how nutrient deprivation of leukemic cells may affect the sensitivity to DNA-damaging treatment, both in vitro and in vivo, through modulation of the cAMP signaling pathway.

In vitro, starvation was mimicked by culturing ALL cells under commonly used starvation conditions involving culture media depleted of serum and/or glucose (29, 30, 31, 32). Among the tested combinations, only complete depletion of both glucose and serum (i.e., depleted medium, DM) effectively abolished the inhibiting effects of cAMP signaling on IR-induced apoptosis. Under these culture conditions, cAMP signaling instead enhanced the IR-mediated killing of ALL-derived cell lines as well as primary leukemic blasts from the bone marrow of pediatric ALL patients. In these experiments, IR served as a convenient model for DNA damage in vitro. However, as the clinical relevance of IR is limited to ALL cases with metastasis to brain or testes (35), we confirmed that cAMP signaling under starvation conditions also enhanced the killing of ALL cells treated with doxorubicin (Doxo). With Doxo being a chemotherapeutic agent widely used in clinical protocols (64), our results indicated a broader relevance.

In our previous research on ALL cells cultured under normal culture conditions, we showed that cAMP-mediated inhibition of IR-induced apoptosis involves prominent inhibition of p53 accumulation (7). This led us to hypothesize that under starving culture conditions, the shift in cAMP-signaling towards enhanced IR-induced apoptosis would be accompanied by enhanced levels of p53. However, this proved not to be the case. As shown in Figure S4, p53 levels were substantially reduced in starved cells regardless of cAMP activation, suggesting that apoptosis in this context occurs via p53-independent mechanisms. The noted p53 reductions could be the result of the G1 cell cycle arrest observed in depleted cell cultures (Fig. S5). Similar p53-independent apoptosis with downregulation of p53 protein has been linked to G1 arrest in both MCF-7 breast cancer cells and HCT116 human colorectal carcinoma cells (65, 66).

Starvation conditions, like deprivation of serum and glucose, are known to enhance intracellular ROS levels, contributing to oxidative stress and consequently to cell death (38, 39, 40). Previous studies have shown that cAMP signaling enhances the DNA damage-induced intracellular levels of ROS in ALL cells (9). In the present study, we found that starvation itself enhanced the ROS levels, which were further elevated by forskolin and IR. This was verified using the ROS scavenger NAC. Based on these results, we investigated the mechanisms whereby elevated ROS enhanced IR-mediated cell death. The stress-activated MAPK p38 is a known mediator of ROS- and DNA damage-induced apoptosis (51, 59). In accordance with other studies (67, 68, 69), we showed that cAMP signaling under starvation conditions activated p38 MAPK. The p38 MAPK inhibitor SB202190 reduced both p38 MAPK activation and apoptosis in IR/forskolin-treated cells, supporting a functional link between these events. Moreover, as the ROS scavenger NAC attenuated the p38 MAPK activation, and SB202190 reduced the ROS levels, this suggests a bidirectional feedback loop between ROS and p38 MAPK. Such a dual role of p38 MAPK in regulation of ROS levels has been reported in other models, including hypoxia-induced injury in PC12 cells (58).

p38 MAPK is also implicated in autophagy, a process often upregulated under metabolic stress (70, 71). While autophagy generally promotes survival under stress, excessive or dysregulated autophagy is also known to contribute to cell death (72, 73). Previous work showed that cAMP-mediated enhancement of ROS levels promotes survival of ALL cells exposed to DNA-damaging agents under normal conditions (9). In contrast, the current findings demonstrate markedly increased autophagy in starved cells, particularly in IR plus forskolin-treated cells. Moreover, inhibition of p38 MAPK with SB202190 prevented autophagy and simultaneously reduced apoptosis, and NAC lowered autophagy levels, suggesting an interplay between ROS, p38 MAPK, autophagy, and apoptosis in starving ALL cells.

Importantly, blocking autophagy by knocking down ULK1 or with the autophagy inhibitor 3-MA did not reduce the cell death induced by IR and forskolin, indicating that autophagy is not required for apoptosis in this setting. Rather, these data are consistent with the concept of cell death with autophagy, a phenomenon where autophagy is activated as a cellular stress response but fails to induce protection. This situation has been reported in several contexts where autophagy is induced as an attempt to restore homeostasis, yet the overwhelming stress or damage exceeds the cell’s protective capacity (74). Thus, our results support a model where cAMP signaling enhances ROS-mediated activation of p38 MAPK, which in turn promotes apoptosis in a context of elevated but non-protective autophagy. This distinction is important and underlines the complexity in the manner that autophagy regulates apoptotic cell death (75).

To study the relevance of our in vitro observations in an in vivo model, we explored the impact of intermittent fasting on DNA-damaging treatment of ALL in a xenograft model of human ALL in SCID mice. The results revealed that suboptimal IR doses nearly abolished leukemia progression in fasting mice, indicating that fasting enhances the efficacy of DNA-damaging therapies in vivo, consistent with the in vitro data. Using a similar xenograft model, leukemia progression in mice on a normal diet was previously shown to be driven by cAMP signaling provided by PGE2 from bone marrow stromal cells (17). Under these conditions, blocking PGE2 production by the cyclooxygenase-inhibitor indomethacin reduced the leukemia progression (17). Since fasting may increase cAMP levels via compounds like for instance glucagon (18, 19), the beneficial effects of intermittent on IR-mediated inhibition of leukemia progression could seem as contradictory to the previous results from mice fed on a regular diet. However, consistent with the in vitro results, cAMP signaling in the context of fasting appears to shift from a protective to a pro-apoptotic role.

Although glucagon receptor expression has been reported on ALL cells (20, 21), the metabolic implications of glucagon in ALL remain poorly understood. To examine shifts in glycolytic metabolism under starvation-mimicking conditions, oxygen consumption rates (OCR) and extracellular acidification rates (ECAR) were measured using a Seahorse XFe 24 analyzer. In line with previous reports by Buss et al. (76), IR-treatment resulted in increased ECAR and OCR under both normal and starving conditions (Fig. S6). However, we did not find changes in metabolism that could explain the shifted cAMP response in ALL cells cultured in DM.

To summarize, the present study demonstrates that intermittent fasting enhances the anti-leukemic effects of DNA-damaging treatment in vivo. Supported by the in vitro data, our results imply that cAMP signaling under starvation conditions augments the DNA damage-mediated killing of ALL cells by promoting ROS-dependent activation of p38 MAPK (summarized in Fig. 5E). These data suggest that fasting may allow for lower doses of chemotherapy, potentially reducing side effects and long-term complications of the treatments. Although the present study aligns with growing literature on the beneficial effects of fasting in hematological malignancies (12, 24, 26, 27, 28, 77), clinical translation should proceed with great caution. Pediatric ALL patients frequently experience hypoglycemia (78), and the effects of intermittent fasting on human cancer prognosis still remain elusive due to lack of high-quality randomized clinical trials. Fasting interventions could be considered as an adjunct therapy in selected patient subgroups, like for instance the high percentage of obese pediatric ALL patients (79, 80) who are at increased risk of treatment failure.

Experimental procedures

Reagents and antibodies

Forskolin (#F6886), doxorubicin (#D1515), PI (#P4170), glucose (#G7021), 3-methyladenine 3-MA (M9281) and Z-VAD-FMK (#627610) were obtained from Sigma-Aldrich. CYTO-ID Autophagy detection kit ver. 2.0 (ENZ-KIT175) was purchased from Enzo Life Sciences. N-acetyl-L-cysteine (NAC) (#A9165), CellROX Green Oxidative Stress Reagent (C10444) and MitoProbe™ TMRM Assay Kit (#M20036) and eBioscience™ Annexin V Apoptosis Detection Kits (# 88-8006-74) were purchased from Thermo Fisher Scientific. The phospho-p38 MAPK (pp38) rabbit monoclonal antibody (#4511), the p38 MAPK rabbit polyclonal antibody (#9212), the ULK1 (A705, #4776) rabbit polyclonal antibody and the LC3B (# 2775) rabbit polyclonal antibody were obtained from Cell Signaling Technology. SB 202190, the p38 MAPK inhibitor and the vinculin mouse monoclonal antibody (#V9131) were from Sigma-Aldrich, the p53 (DO-1, #SC-126) mouse monoclonal antibody and the β-actin mouse monoclonal antibody (C4 #sc-47778) were purchased from Santa Cruz Biotechnology.

Cell culturing and primary cell isolation

The B-cell precursor acute lymphoblastic leukemia cell lines REH (81), NALM-6 (82), and 697 (83) were maintained as previously described (27) at a density between 2 × 105 and 1 × 106 cells/ml in RPMI 1640 supplemented with 10% heat inactivated FBS, 125 U/ml penicillin, and 125 μg/ml streptomycin as previously described (10).

To verify the physiological relevance of the data we obtained on ALL cell lines, key experiments were repeated on primary leukemic blasts derived from pediatric patients newly diagnosed with ALL. Because of the limited number of cells obtained from the patient-derived bone marrow aspirates, only selected experiments were performed. Primary ALL cells were isolated from bone marrow aspirates of newly diagnosed ALL patients in line with previously established procedure (7). The expression of CD19 and CD10 on isolated primary ALL cells were determined by flow cytometry analyses with antibodies directed against CD19 (MACS Miltenyi Biotec #130-091-328) and CD10 (BioLegend #312218) as previously described (7). Collection of bone marrow aspirates from children diagnosed with ALL was performed after obtaining informed consent by parents through the Norwegian Childhood Cancer Biobank (REK 2016/943) (84), in accordance with the Declaration of Helsinki. Sample collection was approved by the Regional Ethics Committee of Norway region South-East C (REK 2014/883).

DNA damage induction by IR or doxorubicin

DNA damage was induced in the ALL cells by either exposing the cells to IR using a X-Strahl RS320 X-ray irradiator at a rate of 3.9 Gy/min or by treating the cells with doxorubicin (Sigma-Aldrich). Both DNA-damaging agents resulted in the similar levels of cell death, as measured by flow cytometry analyses of FSC and SSC. Due to the interference of doxorubicin with analyses of cell death by PI staining and autophagy by CYTO-ID-staining, IR was generally used as the source to introduce DNA damage in the ALL cells.

Western blot analysis

Cells were harvested and lysed in radioimmunoprecipitation (RIPA) buffer as previously described (85). The protein concentration in the lysates was measured using the Pierce BCA Protein Assay Kit (Thermo Fisher Scientific). Equal amounts of protein were loaded on a 4% to 20% Criterion TGX Precast Midi Protein Gel (Bio-Rad, Hercules, CA). Proteins were transferred to Trans-Blot Turbo Midi PVDF Transfer Pack (Bio-Rad) by using the Trans-Blot Turbo Transfer System and detected using standard immunoblotting techniques. Proteins were visualized using SuperSignal™ West Dura Extended Duration Substrate kit (Thermo Fisher Scientific) according to the manufacturer’s instructions. Images were captured using ChemiDoc Imaging System (Bio-Rad) and intensity of protein bands was quantified by using the Image Lab 6.1 software (Bio-Rad, https://www.bio-rad.com/en-no/product/image-lab-software?ID=KRE6P5E8Z). Western immunoblot analyses were performed as previously detailed (17).

Flow cytometry for analyses of cell death, ROS levels and autophagy

All flow cytometry analyses were performed on a NovoCyte (Acea Biosciences Inc.) equipped with three lasers (488, 405, 640 nm) and 13 detection channels. For analyzing the fraction of dead cells in the cell cultures, cells were incubated with PI (20 mg/mL) at 4 °C for 10 minutes. PI was excited at 488 nm and detected in the 572/28 nm channel. Analyses of apoptotic cells were performed by measuring changes in the mitochondrial membrane potential by using the MitoProbe™ tetramethylrhodamine methyl ester (TMRM) Assay Kit according to the manufacturer’s protocol (Thermo Fisher Scientific Cat #M20036). TMRM was excited by the 488 nm laser and detected in the 585/40 nm channel. Apoptosis was also measured with the eBioscience™ Annexin V Apoptosis Detection Kit measuring the staining of annexin V and 7-AAD according to the manufacturer’s protocol (Thermo Fisher Scientific Cat# 88-8006-74). Annexin V was excited at 405 nm, and detected in the 445/45 nm channel, whereas 7-AAD was excited by the 488 nm laser and detected in the 675/30 nm channel. Staining of autophagosomes was performed by using the CYTO-ID Autophagy detection kit (Enzo Life Sciences) according to manufacturer’s protocol, and the dye was excited by the 488 nm laser and detected in the 530/30 nm channel. Levels of reactive oxygen species (ROS) were measured by using the CellROX Green Oxidative Stress Reagent (Thermo Fisher Scientific). The cells were incubated with the CellROX reagent (5 mmol/L) for 30 minutes at 37 °C, and the dye was excited by the 488 nm laser and detected in the 530/30 nm channel.

Knockdown of proteins by transfection with small-interfering RNA oligonucleotides

REH cells (4 × 106) were transfected with small interfering RNA (siRNA) using a nucleofector device (Amaxa Biosciences) and the Nucleofector® Kit R (Lonza) according to the manufacturer’s instructions and using the program G-009 as previously described (10). For knockdown of ULK1 we used 1.6 μM of ULK1 siRNA (L-005049-00-0010). Equimolar concentration of a non-targeting siRNA (D-001810- 01-05) was used as control. All siRNAs were obtained from Dharmacon. After transfection, the cells were incubated for 4 hours before further treatments were initiated.

Lentivirus production in HEK293T cells

Lentiviral vectors containing genes coding for fLuc and enhanced green fluorescent protein (EGFP) were produced by transfecting HEK293T cells with pMD2.G envelope plasmid, pCMVΔ8.91 packaging plasmid, and pSLIEW transfer plasmid as described previously (60).

Lentiviral transduction of REH cells

REH cells were lentivirally transduced with a lentiviral vector containing genes encoding fLuc and EGFP, as previously described (17). In brief, cells (5 × 105 cells/well) in culture medium were seeded in 48-well plates. Lentiviral concentrates were added to the cells, and spinfection was performed by centrifugation of the plates at 900g for 50 min at 34 °C. The plates were then incubated at 37 °C and 5% CO2 in a humidified atmosphere for 2 days before removing the viral particles by two repeated washes at 300g for 10 min at 4 °C. A small aliquot was taken from the cells to analyze the amount of EGFP+ cells by flow cytometry, and the remaining cells were subjected to intratibial injection into SCID mice.

Xenograft model of ALL in SCID mice

Xenografts of ALL in mice were established in 6- to 8-weeks old SCID mice. The mice (The Jackson Laboratory) were intratibially (i.t.) injected with lentivirally-transduced ALL cells (2 × 105 cells) expressing EGFP and fLuc (9, 17, 86).The proximal end of the tibia was exposed as the knee was kept in a flexed position. A 23 G needle was used to drill a hole into the tibia before injecting the ALL cells (40 μl per animal) using a 31 G insulin syringe. The procedure was performed under deep isoflurane anesthesia and the animals received local and general analgesics both during the procedure and a maintenance dose 6 to 8 hours later. One day after successful engraftment of ALL cells, the animals were subjected to either intermittent fasting or normal diets for approximately 25 days. The fasting regimen consisted of 7 cycles of intermittent fasting: one day with sequential 6 hours of feeding (from 8:30 am to 2:30 pm) and 18 hours of fasting, followed by one day of normal feeding. The mice were treated with or without a suboptimal dose of whole-body IR, totaling 3.5 Gy (1 Gy in the first week and 2.5 Gy in the second week), and the progression of leukemia was followed by in vivo imaging detecting the fLuc activity from the leukemic cells by using an In Vivo Imaging System (IVIS) Spectrum CT instrument from PerkinElmer. Animals with high leukemia burden or those that displayed symptoms of suffering (according to predetermined criteria) were humanely euthanized. Animals were kept under appropriate housing conditions with water ad libitum. Experiments were approved by the Norwegian Food Safety Authority (FOTS ID 29918).

Extracellular flux analyses of glycolytic metabolism

Glycolytic metabolism was measured using the Seahorse Extracellular Flux 24 well system (Seahorse XFe 24) using the glycolytic rate assay test as described (87). Briefly, cells were incubated in CM or DM for 6 hours prior to resuspension in Seahorse glycolysis rate assay medium (RPMI pH 7.4, 10 mM glucose, 2 mM L-glutamine, 1 mM sodium pyruvate, 1 mM HEPES buffer, no sodium bicarbonate or phenol red). The cells were then seeded into Cell-Tak-coated (Corning) Seahorse culture plates at a density of 8 x 105 cells per well and incubated in a non-CO2 incubator for 1 hour. Basal measurements of ECAR and OCR were measured in 5 cycles (3 minute measurement, 2 minute waiting) followed by injection of rotenone antimycin A (final concentration 0.5 μM). After 5 measurement cycles, 2-deoxy-D-glucose (2-DG) (final concentration 50 mM) was injected, followed by 5 measurement cycles. Results were analyzed as previously described (88).

Statistical analyses

Statistical analyses were performed by using the GraphPad Prism10 software (https://www.graphpad.com/updates/prism-10-4-1-release-notes). The paired t-test was used to investigate for statistically significant differences unless otherwise stated. The graphs are presented as mean values from at least three independent experiments as indicated in the figure legends, with error bars showing the SEM. Each bar displays the individual data points.

Data availability

All data relevant to this study are included within this article.

Supporting information

This article contains supporting information (1).

Conflict of interest

The authors declare that they have no conflicts of interest with the contents of this article.

Acknowledgments

The authors acknowledge the support of Nina-Cathrin Robinson at Oslo University Hospital in obtaining the bone marrow aspirates of newly diagnosed ALL patients. The authors thank Olaf Heidenreich (Newcastle University) for the kind gift of the pSLIEW vector.

Author contributions

A. Y., N. R., S. B., M. M. B., M. C. M. K., B. S. S., J. A. W., S. N., and H. K. B. writing–review and editing; A, Y. and H. K. B. writing–original draft; A. Y. visualization; A. Y. and H. K. B. validation; A. Y. software; A. Y., N. R., S. B., J. A. W., and H. K. B. methodology; A. Y., N. R., S. B., M. M. B., M. C. M. K., J. A. W., and S. N. investigation; A. Y., N. R., S. B., and J. A. W. formal analysis; A. Y. and S. B. data curation; A. Y., N. B., B. S. S., S. N. and H. K. B. conceptualization; M. M. B., M. C. M. K., and H. K. B. resources; H. K. B. supervision; H. K. B. project administration; H. K. B. funding acquisition.

Funding and additional information

This work was supported by the University of Oslo, the Norwegian Childhood Cancer Society, the UNIFOR FRIMED research fund, the Blix Family Legacy, and the Anders Jahre Research foundation. These organizations are public or nonprofit organizations that support science in general; they had no role in gathering, analyzing, or interpreting the data.

Reviewed by members of the JBC Editorial Board. Edited by Patrick J. O'Brien

Supporting information

Supporting information
mmc1.pdf (347.1KB, pdf)

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Supplementary Materials

Supporting information
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Data Availability Statement

All data relevant to this study are included within this article.

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