- 1Pediatric Oncology Branch, Center for Cancer Research, National Cancer Institute, National Institutes of Health (NIH), Bethesda, MD, United States
- 2Institut de Génétique Moléculaire de Montpellier, University of Montpellier, Centre National de la Recherche Scientifique (CNRS), Montpellier, France
- 3Laboratory of Excellence GR-Ex, Paris, France
- 4Molecular and Clinical Nutrition Section, Intramural Research Program, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, MD, United States
Hematopoiesis, a process that results in the differentiation of all blood lineages, is essential throughout life. The production of 1x1012 blood cells per day, including 200x109 erythrocytes, is highly dependent on nutrient consumption. Notably though, the relative requirements for micronutrients during the perinatal period, a critical developmental window for immune cell and erythrocyte differentiation, have not been extensively studied. More specifically, the impact of the vitamin C/ascorbate micronutrient on perinatal as compared to adult hematopoiesis has been difficult to assess in animal models. Even though humans cannot synthesize ascorbate, due to a pseudogenization of the L-gulono-γ-lactone oxidase (GULO) gene, its generation from glucose is an ancestral mammalian trait. Taking advantage of a Gulo-/- mouse model, we show that ascorbic acid deficiency profoundly impacts perinatal hematopoiesis, resulting in a hypocellular bone marrow (BM) with a significant reduction in hematopoietic stem cells, multipotent progenitors, and hematopoietic progenitors. Furthermore, myeloid progenitors exhibited differential sensitivity to vitamin C levels; common myeloid progenitors and megakaryocyte-erythrocyte progenitors were markedly reduced in Gulo-/- pups following vitamin C depletion in the dams, whereas granulocyte-myeloid progenitors were spared, and their frequency was even augmented. Notably, hematopoietic cell subsets were rescued by vitamin C repletion. Consistent with these data, peripheral myeloid cells were maintained in ascorbate-deficient Gulo-/- pups while other lineage-committed hematopoietic cells were decreased. A reduction in B cell numbers was associated with a significantly reduced humoral immune response in ascorbate-depleted Gulo-/- pups but not adult mice. Erythropoiesis was particularly sensitive to vitamin C deprivation during both the perinatal and adult periods, with ascorbate-deficient Gulo-/- pups as well as adult mice exhibiting compensatory splenic differentiation. Furthermore, in the pathological context of hemolytic anemia, vitamin C-deficient adult Gulo-/- mice were not able to sufficiently increase their erythropoietic activity, resulting in a sustained anemia. Thus, vitamin C plays a pivotal role in the maintenance and differentiation of hematopoietic progenitors during the neonatal period and is required throughout life to sustain erythroid differentiation under stress conditions.
Introduction
Vitamin C (ascorbic acid, ascorbate), is an essential micronutrient to humans, possessing antioxidant activities. Vitamin C has been described as a regulator of numerous physiological processes including neuronal development (1), collagen synthesis (2), immune cell differentiation and function (3), response to infection (4), and most recently, in anti-cancer therapies (5, 6). Mechanistically, vitamin C serves as a cofactor for Jumonji-C domain-containing histone demethylases (JHDMs) and the ten-eleven translocation (TET) dioxygenases (7), altering the epigenome of the cell (7). An abnormal epigenetic remodeling leads to a dysregulated differentiation of stem cells, leukemic stem cells, as well as lymphocytes, amongst others (8–15). Moreover, vitamin C can impact cell physiology by scavenging reactive oxygen species (16). In the context of erythroid differentiation, oxidized ascorbate increases mitochondrial superoxide and exacerbates abnormal erythroblast differentiation whereas vitamin C, scavenging reactive oxygen species and reprogramming mitochondrial metabolism, rescues erythropoiesis (17). Thus, vitamin C regulates a myriad of cell processes through a wide diversity of mechanisms.
Of the >4,000 species of mammals, only higher primates, guinea pigs, and fruit bats have lost the ability to synthesize vitamin C from glucose, due to inactivation of L-gulono-γ-lactone oxidase (GULO), the enzyme that catalyzes the terminal step of L-ascorbic acid biosynthesis (18). Humans and guinea pigs independently lost the ability to synthesize ascorbate 40-50 and 20-25 million years ago, respectively, and the loss in bats was more recent (19–22). These data, together with the strong pseudogenization of the Gulo gene in higher primate and guinea pig genomes, strongly suggest that there was a selective pressure against vitamin C synthesis (22). From a dietary perspective, this pseudogenization has resulted in a requirement that these species consume diets that are rich in vitamin C, with severe vitamin C deficiency resulting in scurvy (16, 22). Nevertheless, prior to the dramatic onset of scurvy, vitamin C deficiency can negatively impact a myriad of other cellular processes. Based on our recent finding that vitamin C accelerates ex vivo human erythroid differentiation (17), it is of much interest to evaluate the impact of vitamin C levels on in vivo hematopoiesis, with a specific focus on erythroid differentiation.
To elucidate the role of vitamin C in regulating physiological in vivo hematopoiesis, we took advantage of the Gulo-/- mouse model which is dependent on dietary vitamin C for long-term survival (23, 24). Furthermore, as hematopoiesis is a dynamic process that changes as a function of ontogeny, we evaluated this process during the immediate postnatal period as well as in adults. During mouse development, hematopoiesis starts in the yolk sac and aorta-gonad-mesonephros region post conception, then moves to the placenta, fetal liver and spleen, before being sustained in the bone marrow (BM) from day 17.5 dpc (days post coitum) throughout the life of the animal (25–28). There are significant differences between fetal liver compared to postnatal BM HSCs, with the former showing a greater repopulating capacity than BM-HSCs (29–31). Moreover, BM hematopoietic stem cells (HSCs) from juvenile mice (<3-4 weeks of age) exhibit a much higher proliferative capacity than HSCs from adult mice (32). Notably though, differences in the metabolic regulation of BM HSCs in neonates and adults have not been extensively evaluated. Here, we find that ascorbic acid deficiency has a significantly greater impact on multiple hematopoietic differentiation pathways during the immediate postnatal period than in adults. Dramatic decreases in HSCs as well as multipotent progenitors (MPPs) and hematopoietic progenitors (HPCs) were associated with reduced numbers of common myeloid progenitors (CMPs) and megakaryocyte-erythrocyte progenitors (MEPs). Our data also show important differences in the sensitivity of progenitors to ascorbate levels–GMPs and mature myeloid cells were not affected while B and erythroid lineage cells were markedly reduced. Furthermore, while anemia resulting from vitamin C deprivation during the adult period was associated with a compensatory splenic erythropoiesis, these mice were unable to mount an efficient response to a hemolytic anemia. Thus, our data unveil a crucial role for vitamin C metabolism in supporting hematopoiesis during the postnatal period and highlight the importance of ascorbate in promoting erythropoiesis under stress conditions.
Results
Ascorbate availability regulates the fate of BM hematopoietic progenitor populations in neonatal and adult GULO-/- mice
Here, we studied the impact of ascorbate deficiency on hematopoiesis, evaluating potential differences during the neonatal and adult periods. To deplete mice of ascorbate during the neonatal period, pregnant Gulo−/− dams were first supplemented with an intermediate dose of vitamin C (0.5g/L) in their drinking water as compared to the standard high dose of 1.0g/L, and then immediately postpartum, vitamin C was decreased to 0.1g/L in the lactating dams (Figure 1A). The lowest vitamin C dose of 0.1g/L was not used during pregnancy as this level of deprivation resulted in a high mortality of pups. To evaluate the impact of ascorbate deficiency in the adult period, Gulo−/− mice were maintained in high dose ascorbate until 3-4 weeks of age and then deprived of vitamin C (supplementation decreased to 0.1g/L) for 14 days (Figure 1B). At the high dose of of vitamin C, plasma vitamin C levels were similar to those reported in WT mice (Figure 1C), ranging from 50-90μM (33, 34). The 2 week time period was selected because vitamin C levels in plasma and organs are significantly decreased by 1 week after withdrawal (34, 35), allowing evaluation of hematopoiesis during the neonatal period. Indeed, following vitamin C deprivation, ascorbate levels dropped markedly, from 33±11 and 51±13 μM in Gulo−/− pups and adults, to 2.2±0.4 and 1.9±0.5 μM, respectively (p<0.01, Figure 1C). Notably, while ascorbate levels were lower in Gulo−/− pups than adult mice receiving high doses of vitamin C, ascorbate deficiency during the neonatal period resulted in a generalized reduction in bone marrow cellularity in neonatal Gulo−/− mice, decreasing from 40.1x106±3.7x106 to 18.6x106±1.5x106 (p<0.0001) and this reduction was associated with a >2-fold reduction in both lineage-negative (Lin-) and Lin-Sca1+c-kit+ (LSK) hematopoietic progenitors (p<0.05, Figure 1D). This was not the case in ascorbate-deprived adult Gulo−/− mice where neither BM cellularity nor the numbers of LSK progenitors was altered (Figure 1D). These data suggested that ascorbate depletion has a more profound impact on the fate of BM progenitors in neonatal than adult mice.
Figure 1 Ascorbate deficiency in neonatal GULO-/- mice results in a significant reduction in bone marrow hematopoietic progenitors. (A) Schema illustrating the protocol used to generate ascorbate-deficient GULO-/- pups. GULO-/- pregnant dams were supplemented with vitamin C (VitC) in the drinking water, either at 1g/L for the high dose group (upper panel) and 0.5g/L for the intermediate dose group (lower panel). Following the birth of pups, dams were either maintained at the high dose of vitamin C (1g/L) or the dose was decreased to 0.1g/L in the low dose group. (B) Schema illustrating the protocol used to generate adult GULO-/- mice (6-7 weeks) with ascorbate deficiency, by decreasing vitamin C in the drinking water of adult mice (4-5 weeks of age) from 1.0g/L to 0.1g/L for 14 days. (C) Plasma vitamin C concentrations were measured by HPLC in pups and adult mice maintained on high or low dose vitamin C as detailed in panels (A, B) Means ± SEM are presented (n=10-15 mice per group). (D) Total bone marrow (BM) cells, lineage- hematopoietic progenitors (Lin-), and Lin-Sca1+c-kit+ (LSK) progenitors were quantified in pups (left) and adult (right) GULO-/- mice supplemented with 1g/L of vitamin C (GULO-/-VitChigh, blue) or 0.1g/L of vitamin C (GULO-/-VitClow, orange). Representative contour plots showing the c-kit/Sca1 profiles in Lin- cells are presented and means ± SEM are indicated (n=9-12 mice per group). Statistical differences were evaluated by an unpaired two-tailed t-test. *p<0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001; ns, not significant.
To determine whether the decreased BM cellularity in Gulo−/− neonatal mice was specifically due to ascorbate depletion, 14 day old ascorbate-deficient Gulo−/− pups were either maintained on low vitamin C supplementation or switched to high vitamin C supplementation for an additional 14 days (Gulo−/− VitClow-rescue, Figure 2A). The 14 day rescue period was sufficient to restore ascorbate levels to baseline, increasing from 11±5 to 71±5 μM (p<0.0001, Figure 2B) and notably, Lin-, LSK, and LK (Lin-Sca1-c-kit+) progenitors were all significantly augmented, to levels similar to those detected in Gulo−/− VitChigh mice (Figure 2C). As the CD150 and CD48 SLAM markers can subdivide LSK into HSCs with long term reconstitution activity (CD150+CD48−LSK), multipotent progenitors (MPPs, CD150-CD48−LSK) and heterogeneous restricted progenitors (HPCs, CD150-/+CD48+LSK) (36–38), we evaluated their profiles in Gulo−/− VitChigh, Gulo−/− VitClow, and Gulo−/− VitClow-rescue mice. Importantly, the numbers of BM HSCs, MPPs, and HPCs were all significantly reduced in Gulo−/− VitClow mice as compared to Gulo−/− VitChigh mice (p<0.01, Figure 2D). Ascorbate rescue resulted in a trend towards increased HSCs and MPPs, and HPCs were markedly augmented, increasing from 16.0x103±1.0x103 to 34.6x103±4.2x103 (p<0.01, Figure 2D). This enhanced level of hematopoiesis correlated with a decreased percentage of BM HSC as well as an increased percentage of HPC following ascorbate rescue (p<0.05, Figure 2D). These data, together with an impressive increase in BM cellularity following vitamin C rescue, from 2.3x107±0.2x107 to 4.7x107±0.4x107 (p<0.001, Figure 2E), strongly suggest that adequate vitamin C levels are a prerequisite for HSC differentiation and the expansion of lineage committed hematopoietic cells.
Figure 2 Ascorbate rescue results in the increased generation of BM hematopoietic progenitors in GULO-/- pups. (A) Schema illustrating the rescue of ascorbate-deficient neonatal GULO-/- mice. Following the birth of pups, dams were either maintained at a high (1g/L, top) or low (0.1g/L, middle) dose of vitamin C for 28 days. For rescue experiments, GULO-/- pups maintained on low dose VitC (0.1g/L) were switched to a high dose (1g/L) at 14 days of life and mice were evaluated at day 28 (VitClow-rescue, bottom). (B) Plasma VitC concentrations were measured in all 3 groups at day 28 and means ± SEM are presented (n=4-5 per group). (C) Representative c-Kit/Sca1 plots of Lin- cells are shown (left) and absolute numbers ± SEM of Lin-, LSK, and LK progenitors are presented (n=5 mice per group). (D) Gating strategy used to define HSC (CD150+CD48-LSK), MPP (CD150-CD48-LSK) and HPC (CD150+/-CD48+LSK) populations (36) together with quantifications and percentages of each subset ± SEM are presented (n=5 mice per group). (E) Mean BM cellularity ± SEM in each condition (n=5 per group). Statistical analyses were performed using a one-way ANOVA (Tukey’s test). *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001; ns, not significant.
Myeloid cell progenitors are differentially impacted by ascorbate deficiency
HSCs undergo commitment to multipotent common myeloid progenitors (CMPs, CD34+CD16/32-LK) cells which can then differentiate into granulocyte-monocyte progenitors (GMPs, CD34+CD16/32+LK) or megakaryocyte-erythrocyte progenitors (MEPs, CD34-CD16/32-LK) (39–41). In accord with the HPC data (Figure 2E), the numbers of CMPs were reduced in ascorbate-deficient mice and increased to baseline following ascorbate rescue (Figure 3A). Interestingly, the total numbers of GMPs were not decreased in ascorbate-deficient conditions and indeed, GMP percentages were actually increased (p<0.001, Figure 3A). In contrast, MEPs were extremely sensitive to ascorbate levels, with a significantly lower level of MEP in Gulo−/− VitClow mice, and a rescue in these levels following ascorbate supplementation (p<0.001, Figure 3A). These results reveal differences in the sensitivity of myeloid progenitors to ascorbate levels. Furthermore, the finding that mature CD11b+ myeloid cell numbers were similar in Gulo−/− VitChigh, Gulo−/− VitClow, and Gulo−/− VitClow-rescue mice suggests that this lineage is not markedly impacted by BM ascorbate concentrations (Figure 3B). These data are consistent with a significantly higher frequency of CD11b+ BM myeloid cells under conditions of ascorbate deficiency and a decreased frequency upon ascorbate rescue (Figure 3B).
Figure 3 Differential impact of ascorbate deprivation on myeloid progenitor subsets. (A Gating strategy used to define CMP (CD34+CD16/32-LK), GMP (CD34+CD16/32+LK), and MEP (CD34-CD16/32-LK) subsets are shown together with representative contour plots. Quantifications and percentages of each subset ± SEM in 28 day old GULO-/- mice maintained under VitC-high, VitC-low, and rescue conditions are presented (n=5 mice per group). (B) CD11b+ myeloid cells were evaluated in BM of the indicated groups by flow cytometry and representative contour plots are shown (left). Total numbers as well as percentages of CD11b+ cells are presented as means ± SEM (n=5 mice per group). (C) Splenic cellularity ± SEM in each condition is shown (n=5 mice per group). (D) CD11b+ myeloid cells in spleen were evaluated as in panel (B). Means ± SEM are shown (n=5 mice per group). Statistical differences were analyzed using a one-way ANOVA (Tukey’s test). *p < 0.05; **p < 0.01; ***p < 0.001; ns, not significant.
The decrease in BM hematopoietic progenitors was coupled to a significantly reduced splenic cellularity in Gulo−/− VitClow as compared to Gulo−/− VitChigh mice, and numbers trended upwards following rescue (Figure 3C). However, consistent with the lack of impact of ascorbate deficiency on BM myeloid cells, the numbers of CD11b+ splenocytes were not altered by either ascorbate deprivation or rescue and again, the frequency of CD11b+ splenocytes was increased in ascorbate-deprived Gulo−/− mice (p<0.01, Figure 3D). The ensemble of these data reveals the importance of ascorbate-coupled regulation in the fate of early BM hematopoietic progenitors. Additionally, given the short half-life of myeloid cells (42), our data indicate that myeloid cell differentiation continues under conditions of ascorbate acid deficiency.
Ascorbate deficiency results in defective B lymphocyte differentiation with attenuated immunoglobulin production during the neonatal period
The hypocellular BM and spleen in ascorbate-deficient Gulo−/− mice (Figures 2E, 3C), together with a significant increase in the frequency of myeloid cells (Figures 3B, D), suggested that ascorbate levels might impact lymphoid lineage differentiation. We therefore evaluated both T and B lymphocyte differentiation in Gulo−/− mice as a function of their ascorbate status. Interestingly, ex vivo T cell differentiation (43) as well as the differentiation of regulatory and IL-17-secreting T cell subsets have been shown to be sensitive to ascorbic acid levels, via the vitamin C-regulated activity of Ten-eleven translocation (TET) enzyme dioxygenases (8, 9, 12, 14, 15). Importantly though, thymocyte differentiation in 14-day old Gulo−/− pups was globally intact, with similar absolute numbers of thymocytes as well as the different thymocyte sub-populations; double negative (DN), double positive (DP), and more mature single positive CD4 (SP4), single positive CD8 (SP8), and γδ thymocyte subsets (Figures S1A, B). Only the numbers of CD8+ intermediate single positive (ISP) thymocytes, representing a stage of TCR β-selection rearrangement wherein there is a proliferative burst (44–48), were lower in ascorbate-deficient as compared to ascorbate-sufficient Gulo−/− pups (Figure S1B). Indeed, in agreement with previous literature (10, 34), the numbers of splenic CD4+ and CD8+ T cells (Figure S1C) as well as subsets of naïve, central memory and effector T cells (data not shown) were similar in Gulo−/− VitClow and Gulo−/− VitChigh pups as well as adult mice. However, an extended 28 day ascorbate deprivation starting at birth led to a significant decrease in the total number of splenic CD3+ T cells (Figure S1D). Moreover, it is interesting to note that this did not reflect a global decrease in T lymphocytes as CD8+, but not CD4+, T cells were significantly reduced following ascorbate deprivation and were rescued by ascorbate supplementation (Figure S1D). Thus, long-term ascorbate deprivation during the neonatal period negatively impacts peripheral CD8+ T cell maintenance and/or homeostatic proliferation.
B lymphocyte differentiation during the neonatal period was acutely sensitive to vitamin C levels as CD19+ BM B cells were significantly reduced by a 14-17 day ascorbate deprivation (p<0.01, Figure 4A). However, BM B cells in adult Gulo−/− mice were not impacted by vitamin C depletion and splenic B cell numbers remained high (Figures 4A and S2). While the numbers of c-kit+ pro-B progenitors in ascorbate-deprived neonatal mice trended lower in Gulo−/− VitClow than Gulo−/− VitChigh mice, these levels did not reach significance (Figure 4A). Notably though, an extended 28 day vitamin C deprivation period during the neonatal period resulted in a massive loss in pro-B cells in the BM, decreasing from 9.8x105±1.1x105 to 1.7x105±0.4x105 (p<0.0001, Figure 4B), and this was associated with an almost complete loss in CD19+ B cells (11.2x106±1.0x106 to 1.2x106±0.2x106, p<0.0001 Figure 4B). The reduction in B cell differentiation was directly due to low ascorbate levels as a 14 day rescue with high vitamin C supplementation restored pro-B as well as mature B cells in the BM (Figure 4B). Furthermore, the reduction in splenic B cells was alleviated by vitamin C supplementation (Figure 4C). Thus, ascorbate deficiency results in a dramatic decrease in B cell differentiation.
Figure 4 Ascorbate deprivation during the neonatal period results in a significant loss in B cell lymphopoiesis and reduced immunoglobulin production. (A) B cell differentiation was evaluated in the BM of pups (left) and adult (right) GULO-/- mice supplemented with 1g/L of vitamin C (GULO-/-VitChigh) or 0.1g/L of vitamin C (GULO-/-VitClow). Total numbers of pro-B cells, evaluated as CD19+c-kit+CD25-, and CD19+ B cells were quantified and means ± SEM are shown (n=10-12 mice per group). (B) Pro-B and B lymphocytes were evaluated at day 28 in BM of GULO-/-VitChigh, GULO-/-VitClow, and GULO-/-VitClow-rescue mice. Representative contour plots as well as absolute numbers of pro-B and B cells are shown (n=5 mice per group). (C) Representative contour plots of CD19+ splenic B cells in 28 day old GULO-/- mice maintained under VitC-high, VitC-low, and rescue conditions together with absolute numbers of splenic B cells (n=5 mice per group). (D) Serum immunoglobulins (IgM, IgG and IgA) were monitored in neonatal (18 day old, left) and adult (right) GULO-/- mice supplemented with high or low vitamin C by ELISA. Means ± SEM (μg/ml) are presented (n= 5-8 mice per group). Statistical analyses were performed using an unpaired two-tailed t-test in panels (A, D); and a one-way ANOVA (Tukey’s test) in panels B-C. *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.001; ns, not significant.
The important impact of vitamin C supplementation on B cell differentiation during the neonatal period raised the possibility that immunoglobulin production might be impacted by plasma/BM ascorbate levels. Indeed, basal IgM and IgG levels were significantly lower in ascorbate-deficient as compared to ascorbate-sufficient Gulo−/− pups (p<0.05-0.01, Figure 4D) whereas they were not altered in Gulo−/− adult mice (Figure 4D). Ascorbic acid supplementation has been shown to positively regulate plasma cell differentiation in adult Gulo−/− mice (49) and the data presented here add to our understanding of vitamin C metabolism in B lymphocytes. During the first two weeks of postnatal life, a period that is characterized by an impaired ability to mount a humoral response (50), ascorbic acid levels play a critical role in B cell function, regulating immunoglobulin production.
BM and splenic erythroid lineage cells are negatively impacted by ascorbic acid deprivation during the neonatal period
Based on the hypocellularity of the BM of ascorbate-deficient neonatal Gulo−/− mice and the dramatic decrease in MEPs (Figure 3A), we quantified Ter119+ erythroid cells–a lineage that comprises a large proportion of BM hematopoietic cells (51). In agreement with the BM hypocellularity in ascorbate-deficient Gulo−/− pups, the absolute number of Ter119+ BM cells was significantly reduced following ascorbate deprivation; 8.4x106±0.8x106 and 22.4x106±2.4x106 erythroid cells in ascorbate-deficient and sufficient pups, respectively (p<0.001, Figure 5A). In contrast, the numbers of Ter119+BM cells in adult Gulo−/−VitClow mice revealed no significant differences with Gulo−/−VitChigh mice (Figure 5A), in accord with their normal BM cellularity (Figure 1D). Finally, vitamin C supplementation at 14 days of life resulted in a complete restoration of Ter119+ BM cell numbers by day 28 (p<0.01, Figure 5B). Interestingly though, the percentages of Ter119+ erythroid cells in the spleen were not significantly augmented, suggesting differences in the regulation of erythroid differentiation in these organs (Figure 5C).
Figure 5 Ascorbate deprivation leads to decreased erythroid cells in GULO-/- pups. (A) Ter119+ erythroid BM cells were quantified in pups (left) and adult (right) GULO-/- mice supplemented with 1g/L of vitamin C (GULO-/-VitChigh) or 0.1g/L of vitamin C (GULO-/-VitClow; n=10-12 mice per group). Representative contour plots are shown and quantification of Ter119+ cells are presented as means ± SEM. (B) Ter119+ BM cells were evaluated in 28 day old GULO-/- mice maintained under VitC-high, VitC-low, and rescue conditions. The percentages and absolute numbers of Ter119+ cells are shown (n=5 per group). (C) Splenic Ter119+ erythroid cells were evaluated as in panel (B) and means ± SEM are presented (n=5 mice per group). Statistical differences were analyzed using an unpaired two-tailed t-test in panel (A); and a one-way ANOVA (Tukey’s test) in panels (B, C). *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001; ns, not significant.
Unlike humans, mice exhibit erythropoietic activity in the spleen under stress conditions. It was therefore of interest to determine whether splenic erythropoietic activity is regulated by ascorbate levels. In accord with the data in 28 day old Gulo−/− VitClow mice–showing decreased numbers of splenic Ter119+ cells (Figure 5C), erythroid cell numbers were also reduced at day 14 of life (p<0.05). However, Ter119+ cells were not impacted following ascorbic acid deprivation during the adult period (Figure 6A). To assess whether vitamin C deprivation resulted in a compensatory increase in splenic erythropoiesis or alternatively, a block in erythroid differentiation, we evaluated the relative levels of early and late erythroblasts in pups and adults. Erythropoiesis is maintained in the mouse spleen during the neonatal period, with erythroid precursors detected until 6-7 weeks of age (51–53) and indeed, we found that early erythroblasts, characterized as Ter119+CD44hiFSChi, were detected in spleens of Gulo−/− VitChigh mice during the neonatal period but were almost undetectable in adults (20.4±3.6% vs 4.5±0.8%, Figure 6B). Importantly though, the percentages of early erythroblasts in both Gulo−/− VitClow pups and adults was markedly increased as compared to ascorbate-replete animals (p<0.01, Figure 6B). The continued presence of early erythroblasts in Gulo−/− VitClow adult mice (23.4±5.5%) reveals a secondary erythropoietic activity that was associated with a significantly increased spleen area in adult Gulo−/− mice (p<0.05, Figure 6C). It was though insufficient as the hematocrit in adult Gulo−/− mice was still reduced compared to ascorbate-sufficient mice (p<0.01, Figure 6D). Collectively, these data demonstrate the critical role of ascorbate metabolism in supporting splenic erythroid differentiation, with insufficient compensation in low ascorbate conditions.
Figure 6 Ascorbate deficiency is associated with splenic erythropoiesis in GULO-/- mice. (A) Ter119+ cells in the spleen were assessed in GULO-/- pups and adult mice in the indicated conditions. Representative contour plots and quantifications are presented. (B) Within the splenic Ter119+ subset, early (pro, basophilic and polychromatic) erythroblasts were distinguished from late (orthochromatic) erythroblasts, reticulocytes, and RBCs as a function of their CD44/FSC profiles. Representative CD44/FSC contour plots and quantifications in individual mice are presented (n=8-12 mice per group). (C) Spleen sizes in adult GULO-/-VitChigh and GULO-/-VitClow mice were quantified as a function of their area (cm2) and means ± SEM (left) as well as representative images are shown (right). (D) Hematocrits in GULO-/-VitChigh and GULO-/-VitClow mice were evaluated, and percentages are shown (n=5-6 mice with two replicates). Statistical analyses were performed using an unpaired two-tailed t-test in panels (A, C, D) and a two-way ANOVA (Tukey’s test) in panel (B). *p < 0.05; **p < 0.01; ***p < 0.001; ns, not significant.
Anemia-induced erythropoiesis is dependent on ascorbic acid metabolism
The data presented above demonstrated the importance of ascorbic acid for erythroid differentiation under physiological conditions and revealed the potential of splenic erythropoietic activity in adult Gulo−/− mice to compensate for ineffective erythropoiesis under conditions where ascorbic acid levels were limiting. It was therefore of interest to determine the role of ascorbic acid in responding to a pathological situation of hemolytic anemia. We therefore subjected adult Gulo−/−mice to a chemically induced hemolytic anemia by injecting phenylhydrazine (PHZ) over 5 days as shown in Figure 7A. Serum vitamin C levels were not altered by PHZ, remaining significantly reduced in the low-dose group (p<0.001, Figure 7B). While vitamin C deprivation was associated with a basal decrease in weight (34) (Figure 7C), PHZ treatment only impacted weight loss in this group (decrease of 15.1±0.3%, p<0.01, Figure 7C). Importantly, these Gulo−/− VitClow mice did not exhibit an increase in hematocrit following PHZ-induced anemia. Hematocrits initially fell sharply in both groups at day 2, to 23.4±1.5% (Gulo−/− VitCHigh) and 18.1±2.8% (Gulo−/− VitCLow), as previously described (54). However, by day 5, the high dose vitamin C group recovered to 40.9±1.9% (p<0.0001) while the low dose group remained unchanged (Figure 7D). The percentage of Ter119+ BM erythroid cells was not increased following PHZ treatment of the vitamin C-high Gulo−/−mice group, but they were decreased in the vitamin C-low group, resulting in a 3-fold lower numbers of Ter119+ BM erythroid cells (p<0.001, Figure 7E). Furthermore, in the vitamin C-high Gulo−/−mice group, there were compensatory changes in erythroblast differentiation. Proerythroblasts, basophilic erythroblasts, polychromatic erythroblasts, orthochromatic erythroblasts/reticulocytes, and RBCs can be distinguished by their CD44/FSC profiles (55, 56) and we detected augmented BM polychromatic and orthochromatic erythroblasts following PHZ treatment (p<0.05-0.0001, Figure 7F). Importantly though, hemolytic anemia in vitamin C-low Gulo−/−mice did not result in significant changes in any of these BM erythroid subsets (Figure 7F).
Figure 7 Hemolytic anemia results in changes in bone marrow erythroid differentiation in ascorbate-sufficient but not ascorbate-deficient GULO-/- adults. (A) Schema illustrating the experimental protocol used to induce hemolytic anemia in adult GULO-/- mice by phenylhydrazine (PHZ). PHZ was administered IP at a dose of 40mg/kg on day 0 and at 20mg/kg on days 1 and 3, and then mice were euthanized at day 5. (B) Plasma vitamin C concentrations were measured by HPLC in control and PHZ-treated GULO-/-VitChigh and GULO-/-VitClow mice at day 5. Means ± SEM are presented (n=5-6 per group). (C) Weights of control and PHZ-treated GULO-/-VitChigh and GULO-/-VitClow mice were measured at day 5 post-treatment. Means ± SEM are shown (n=5-6 per group). (D) Hematocrits were monitored in PHZ-treated GULO-/- mice on days 2 and 5 and means ± SEM are presented (n=5 per group). (E) The impact of PHZ-induced hemolytic anemia on Ter119+ BM cells was evaluated at day 5 in the indicated conditions. Representative contour plots, percentages, and absolute numbers of Ter119+ cells are presented. (F) The differentiation state of Ter119+ erythroblasts was evaluated as a function of CD44/FSC profiles, distinguishing proerythroblasts (Pro), basophilic erythroblasts (Baso), polychromatic erythroblasts (Poly), orthochromatic erythroblasts (Ortho), reticulocytes, and RBCs as indicated (left). The percentages of cells in each stage of erythroid differentiation were quantified in the indicated conditions at day 5 of PHZ-induced anemia (right). Statistical analyses were performed using an unpaired two-tailed t-test in panels (B–E) and a two-way ANOVA (Tukey’s test) in panel (F). *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001; ns, not significant.
As discussed above, mice generally respond to anemia by activating a stress erythropoiesis in the spleen (57–59). We therefore evaluated erythropoietic activity in the spleens of anemic Gulo−/−mice. An increased response of the vitamin C-high as compared to vitamin C-low group was demonstrated by the significantly increased spleen size in the former (p<0.01, Figure 8A). This was associated with a dramatic increase in the frequency of splenic Ter119+ erythroid cells in the vitamin C high group (p<0.001, Figure 8B). Most strikingly, vitamin C-high Gulo−/−mice responded to hemolytic anemia with increases in splenic basophilic, polychromatic, and orthochromatic erythroblast/reticulocyte subsets whereas the vitamin C-low group only exhibited increases in the basophilic erythroblast subset (Figure 8C). Thus, under conditions of hemolytic stress, vitamin C-deficient mice are not able to massively increase their erythropoietic activity, either in the BM nor spleen, resulting in an inability to overcome severe anemia.
Figure 8 Impaired stress erythropoiesis in ascorbate-deficient GULO-/- mice in response to acute hemolytic anemia. (A) Spleen area (cm2) in PHZ-treated GULO-/- mice was measured at day 5 and means ± SEM (left) as well as representative images (right) are shown. (B) The impact of PHZ-induced hemolytic anemia on the percentages and absolute numbers of splenic Ter119+ cells in GULO-/-VitChigh and GULO-/-VitClow mice are shown at day 5. Representative Ter119/SSC contour plots, percentages, and absolute numbers of Ter119+ cells are presented (n=5-6 mice per group). (C) Changes in splenic proerythroblast, basophilic, polychromatic, orthochromatic erythroblast, and reticulocyte subsets as well as RBCs were monitored in PHZ-treated GULO-/-VitChigh and GULO-/-VitClow mice as described in (Figure 7F). Quantifications of erythroid precursors as well as representative CD44/FSC contour plots are shown. Statistical analyses were performed using an unpaired two-tailed t-test in panels (A, B) and a two-way ANOVA (Tukey’s test) in panel (C). *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001; ns, not significant.
Discussion
Hematopoiesis is a dynamic process that allows all blood lineage cells to be maintained throughout life. This process relies on the continual presence of self-renewing, multipotent HSCs but it is notable that the perinatal period is a critical developmental window for immune cell and erythrocyte generation (60–62). Nonetheless, the differential roles of metabolic parameters in regulating perinatal versus adult hematopoiesis have not been extensively addressed. Using a murine GULO-deficient mouse model, we show that vitamin C deficiency during the perinatal but not adult period results in a hypocellular BM. These data suggested that perinatal hematopoiesis exhibits a significantly greater dependence on the vitamin C micronutrient than hematopoiesis occurring during the adult period and indeed, HSC, MPP, and HPC progenitors were dramatically reduced in BM of vitamin C-deficient Gulo-/- pups. Furthermore, our data show a differential impact of vitamin C on different hematopoietic lineages; neither BM nor splenic myeloid cells were reduced, even upon extended vitamin C deprivation, whereas B lymphoid and erythroid cells were markedly decreased under conditions of vitamin C deficiency. Thus, our data reveal a pivotal role for vitamin C in HSC maintenance during the postnatal period and point to differences in the sensitivity of different hematopoietic lineages to ascorbate metabolism.
One major difficulty in elucidating the mechanism(s) via which vitamin C exerts its actions on human hematopoietic differentiation is due to the lack of appropriate animal models. Aside from primates, who lost the ability to synthesize vitamin C at the split between the Strepsirrhini and Haplorrhini primate suborders (63, 64), guinea pigs and bats are the only two other species that have independently lost the ability to synthesize vitamin C due to mutations in the Gulo gene (20, 21, 65). As such, rat and murine models with mutations in the Gulo gene have been important for evaluations of physiological processes that are modulated by vitamin C levels (23, 66, 67). Notably though, these animal models are not flawless because they lack compensatory mechanisms that have evolved in higher primates. Additionally, while humans develop scurvy when plasma vitamin C levels are <10µM, mice can tolerate much lower plasma vitamin C levels (<1µM), highlighting important physiological differences between species (6, 68, 69).
Higher primates have likely been subjected to severe, but as yet unknown, environmental pressures that resulted in mutations in the GULO gene as well as in the urate oxidase (UOX) gene (70). In the 21st century, the loss of the enzymes encoded by these genes increases the risk of Homo Sapiens to develop multiple diseases, including hypertension, cardiovascular disease, and infection, amongst others (70). That being said, humans appear to have evolved mechanisms to potentially overcome some of the drawbacks associated with significantly reduced levels of vitamin C. While the liver produces ascorbate at the rate of 200 mg/kg/day in vitamin C-producing mammals (71, 72), humans survive on massively lower levels, with FDA recommendation of 1-1.5 mg/kg/day (73). A further argument that strongly suggests that humans have evolved compensatory mechanisms for vitamin C loss is the finding that mice and rats with an inactivated Gulo gene require significantly higher doses of vitamin C than humans, ranging from 80-300 mg/kg/day (23, 66, 67). As fruit bats also consume more massive levels of vitamin C, reaching 250 mg/kg/day (74), it is tempting to hypothesize that they have not developed compensatory changes. Importantly, two known compensatory changes in humans are both associated with erythrocytes, the most abundant cell in the human body. First, human, but not mouse, erythrocytes contain an ascorbate-reducible b-type cytochrome, likely contributing to their ability to reduce the one electron-oxidized form of ascorbate, monodehydroascorbate. Moreover, a b-type cytochrome is also highly present on erythrocytes in guinea pigs, another mammal that lost the ability to produce vitamin C (75). Second, the postnatal erythroid expression of GLUT1, promoting the transport of glucose as well as dehydroascorbic acid (DHA)–the two-electron oxidized form of vitamin C (76–78), is a specific feature of those mammals that have lost the ability to synthesize ascorbate from glucose (57, 79). Indeed, human red blood cells harbor greater than 200,000 molecules of GLUT1 per cell (80, 81), allowing the shuttling of DHA across the membrane followed by its rapid reduction to ascorbate [reviewed by (82)]. Nonetheless, Gulo-/- mice with low ascorbate levels exhibit changes in multiple metabolites (83) and like humans, they develop vascular disease as demonstrated by aortic wall damage (23). Together, these data support the use of Gulo-/- mice to evaluate the impact of vitamin C deficiency on hematopoiesis.
The experiments presented here highlight the critical need for adequate serum vitamin C levels in hematopoiesis in the Gulo-/- mouse model, especially during the postnatal period. Previous research elegantly showed that vitamin C deprivation in adult mice results in an increased frequency of BM HSCs in the absence of any changes in the frequency of MPPs, HPCs, CMPs, or MEPs (10). In our studies, the marked BM hypocellularity in Gulo-/- pups, secondary to vitamin C deprivation in the lactating dams, was associated with a marked decrease in HSC, MPP, and HPC numbers but with no changes in the frequency of these cells. These data highlight differences in the impact of ascorbate depletion during the neonatal period as compared to the adult period. It should also be noted that ascorbate depletion was responsible for the loss of these progenitors during the neonatal period as progenitor numbers were rescued by a 2 week repletion with vitamin C. Our data also highlight differences in the dependence of more committed progenitors to ascorbate levels; while CMP and MEP cells were markedly reduced in Gulo-/- pups following vitamin C depletion in the dams, GMPs were maintained and their frequency was even augmented. This was associated with a maintenance of BM and splenic CD11b+ myeloid cells whereas Ter119+ erythroid cells were markedly reduced. While the mechanisms responsible for differential vitamin C dependence of GMPs and MEPs remains to be determined, it is important to note that vitamin C serves as a cofactor for the dioxygenase activity of Tet2 (84) and the effect of vitamin C on HSCs has been shown to be mediated, at least in part, by Tet2 (10, 11). Thus, it will be of much interest to evaluate how the kinetics of ascorbate depletion during the neonatal and adult periods regulates Tet2 activity in progenitor subsets. Our data strongly suggest that multiple hematopoietic processes, starting with the differentiation and maintenance of hematopoietic stem cells, are significantly more sensitive to vitamin C levels during the neonatal than the adult period.
As regards the impact of vitamin C deprivation on immune responsiveness, a large number of studies suggest that vitamin C supplementation may alleviate or prevent infections (4). Data emerging from the Covid pandemic has even pointed to a role for vitamin C in improving immune function in patients infected with SARS-CoV-2 (85, 86). Furthermore, a specific impact of vitamin C on immunity during the newborn period is suggested by an exciting recent study performed in the Central-African Republic; offspring born to vitamin C-deficient mothers had low serum vitamin C levels and they were at a significantly higher risk of being infected with an enteric virus (87). In our study, vitamin C deficiency resulted in a significant reduction in B lymphocytes in pups and an almost complete loss in the generation of B cells following 4 weeks of vitamin C deprivation. Furthermore, low numbers of peripheral B cells in Gulo-/- pups was associated with significantly reduced serum IgM/IgG levels. Taken together, it would be of much interest to determine whether humoral and cellular immunity are negatively impacted in infants, as well as adults, with hypovitaminosis-C.
Within the context of human erythrocyte differentiation, ascorbate has been found to play a role in red cell function. Serum ascorbate levels are inversely correlated with the osmotic fragility of red cells and moreover, in patients with diabetes, lower erythrocyte ascorbate levels are associated with increased RBC rigidity (69). Similarly, in patients with end-stage renal disease, low serum vitamin C levels correlate with hypochromic red cells whereas patients with high vitamin C levels harbor an increased fraction of red cells with normal hemoglobin content (88). These correlative studies are in agreement with ex vivo erythropoiesis assays showing that vitamin C promotes the erythroid differentiation of human hematopoietic progenitors (17). Furthermore, while there is a crosstalk between Tet2 and ascorbate, the mechanism is related to the protection of erythroblasts from oxidative stress (17, 89), in a manner that appears to be independent of Tet2’s dioxygenase activity (90, 91). The data presented here reveal the importance of serum ascorbate levels in supporting in vivo erythroid differentiation, with low levels of ascorbate severely attenuating the ability of vitamin C-deficient mice to respond to anemia.
Vitamin C supplementation has been evaluated as a means of improving erythroid differentiation in hemodialysis patients as well as in patients with anemia of chronic disease, and importantly, it has shown potential clinical benefit in some patients (92–95). However, the impact of vitamin C on erythropoiesis does not appear to be homogeneous; it did not improve hemoglobin recovery in patients with iron-deficiency anemia (96) and in patients with sickle cell disease (SCD, the combined treatment of vitamin C and vitamin E increased markers of hemolysis (97). Interestingly though, a study of SCD patients in Saudi Arabia revealed significantly lower plasma vitamin C levels compared to healthy controls (98). Together, these studies highlight the paucity of information regarding the mechanisms via which vitamin C regulates ineffective erythropoiesis. Our identification of an abnormal stress erythropoiesis in vitamin C-deficient Gulo-/- mice underscores the importance of developing new murine models for studying human erythroid pathologies, especially erythroid disorders that manifest during the newborn period.
Methods and materials
GULO-/- Mice
GULO-/- mice (23) were maintained and bred under specific pathogen-free conditions in the NIDDK animal facility (NIH, Bethesda, MD). Mice were fed ad libitum on a regular chow diet (NIH-07) on a 12h light/12h dark cycle and supplemented with ascorbic acid (Sigma-Aldrich, St. Louis, MO) in the drinking water. Water was changed every 2 days. In experiments performed on 14-17 day old GULO-/- pups, dams were either maintained on 1g/L of ascorbic acid pre- and post-delivery (high vitamin C group) or started on 0.5g/l during pregnancy and then shifted to 0.1g/L post-delivery (low-vitamin C group, Figure 1A). For experiments performed on adult GULO-/- mice (6-7 weeks of age), all mice were maintained on high vitamin C supplementation (1g/L in drinking water) until 14 days prior to experimentation when vitamin C levels were either maintained or decreased to 0.1g/L (Figure 1B). For rescue experiments, 14 day old GULO-/- pups (and dams) in the low vitamin C group described above were shifted to high dose ascorbic acid (1g/L) for 14 days, until 28 days of age (referred to as VitClow-rescue, Figure 2A).
Animal care and experiments were performed in accordance with National Institutes of Health (NIH) guidelines and all experiments were approved by the Animal Care and Use Committee of the NIDDK.
Induction of anemia
Haemolytic anaemia was induced in adult GULO-/- mice (Figure 5A) maintained on either high (1g/L) or low (0.1g/L) vitamin C supplementation. Anemia was induced by phenylhydrazine hydrochloride (PHZ, Sigma-Aldrich) diluted in PBS. Mice were injected intraperitoneally at days 0, 1, and 3, with doses of 40mg/kg, 20mg/kg, and 20mg/kg, respectively. Haematocrits were measured in retroorbital samples on days 2 and 5. Mice were euthanized on day 5 for analyses.
Vitamin C and haematocrit measurements
Whole blood was collected by retro-orbital bleeds using heparinized capillary tubes and haematocrits were evaluated. For vitamin C measurements, plasma was further processed for HPLC analyses as described previously (99). Briefly, whole blood was centrifuged at 250g for 5 min at 4C. One volume of plasma was mixed with four volumes of cold methanol/1mM EDTA and centrifuged at >14000g for 10 min at 4C. The supernatant was then transferred to a fresh tube and either analyzed by HPLC or stored at -80C. Ascorbic acid was analyzed by HPLC coupled with electrochemical detection (100).
Thymocyte, splenocyte and BM preparations
Murine thymus, spleen, and bone marrow were collected after euthanizing the mice. BM was flushed using a 25gauge needle into 2% fetal bovine serum (FBS) PBS. Single cell thymocyte and splenocyte suspensions were generated by physical disruption of tissue and filtration through 70µm nylon cell strainers. Cells were counted and further processed for flow cytometry.
Flow cytometry
Single cell suspensions were stained with three separate antibody cocktails for analyses of BM, spleen, and thymus. BM subsets were identified using the following conjugated mAbs: Ter119, CD25 (Invitrogen, Thermo Fisher Scientific, Waltham, MA), c-kit, Sca1, CD19 and CD44 (BD Biosciences, San Jose, CA). Lineage-differentiated BM cells were eliminated using a dump consisting of mAbs against CD11b, Gr1 and CD3. HSCs, MPPs, and HPCs were identified within the lineage- Sca1+c-kit+ (LSK) lineage as a function of CD150 and CD48 profiles while CMP, GMP, and MEP were evaluated on lineage-c-kit+ (LK) progenitors as a function of CD34 and CD16/32 expression (36, 38). For thymus, cells were stained with the following directly conjugated mAbs: CD3, c-kit, CD4, CD8, CD44, TCRγδ (BD Biosciences) and CD25 (Invitrogen). Non-T lineage cells were excluded using a dump consisting of mAbs against CD19, Gr1, CD11b. Splenic subsets were distinguished using the following directly conjugated mAbs: Ter119 (Invitrogen), CD44, CD3, CD4, CD8, CD62L, TCRγδ, CD19 and CD11b (BD Biosciences). Stained cells were analyzed by flow cytometry using the LSR II-Fortessa (BD Biosciences). All data analyses were performed using Diva (BD Biosciences), and FlowJo Mac v.10.6.2 software (Tree Star).
ELISA
The total serum IgM, IgG and IgA protein levels were measured by ELISA (Thermo-Fisher Scientific). Plates were coated with respective purified and pre-titrated monoclonal antibodies and developed with pre-titrated HRP-conjugated anti-mouse polyclonal antibodies specific for each mouse Ig isotype. Isotype controls were serially diluted and used to generate a standard curve. The serum samples were diluted, added to the plate as technical triplicates and their Ig concentrations were determined using the SoftMax-Pro software (Molecular Devices, LLC, San Jose, CA).
Statistical analyses
Data were analysed using GraphPad Prism software version 9.3.1 (Graph Pad Prism, La Jolla, CA). p-values were calculated using either unpaired two-tailed t-tests, one-way ANOVA (Tukey’s multiple comparison test) or two-way ANOVA (Tukey’s multiple comparison test), as specified. p-values are indicated in individual figure legends.
Data availability statement
The raw data supporting the conclusions of this article will be made available by the authors, without undue reservation.
Ethics statement
This study was reviewed and approved by the Animal Care Committee of the NIDDK, National Institutes of Health (NIH).
Author contributions
IP, MP, AM, VZ, ML, PCV, and NT conceived the study. IP, MP, AM, JM, PGM, SK, VZ, ML, PCV, and NT were involved in study design. IP, MP, AM, JM, VZ, and PCV performed experiments. VZ, ML, PCV, and NT supervised the study. All authors participated in data analysis and discussions. IP, MP, PCV, and NT wrote the manuscript and all authors critically reviewed the manuscript. All authors contributed to the article and approved the submitted version.
Funding
IP, AM, and PGM were funded by fellowships from the French Labex EpiGenMed, the French Ministry of Health, and the Clarin-COFUND EU Program. IP and AM were also funded by the intramural NIH research program and PGM by NIH DK32094. This work was supported by generous funding from the FRM, ARC, the French national (ANR) research grants and the French laboratory consortiums (Labex) EpiGenMed and GR-EX. ML and PCV are supported by the NIDDK (ZIA DK054506 23) and NT by NCI intramural NIH research programs (ZIA BC 011924 and ZIA BC 011923).
Acknowledgments
We thank all members of our lab for their scientific critique and support and are grateful to the NIH animal facility staff for their expert assistance. The schematics were created with BioRender.com.
Conflict of interest
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
Publisher’s note
All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article, or claim that may be made by its manufacturer, is not guaranteed or endorsed by the publisher.
Supplementary material
The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fimmu.2022.898827/full#supplementary-material
References
1. Figueroa-Mendez R, Rivas-Arancibia S. Vitamin c in health and disease: Its role in the metabolism of cells and redox state in the brain. Front Physiol (2015) 6:397. doi: 10.3389/fphys.2015.00397
2. Murad S, Grove D, Lindberg KA, Reynolds G, Sivarajah A, Pinnell SR. Regulation of collagen synthesis by ascorbic acid. Proc Natl Acad Sci U.S.A. (1981) 78:2879–82. doi: 10.1073/pnas.78.5.2879
3. Ang A, Pullar JM, Currie MJ, Vissers MCM. Vitamin c and immune cell function in inflammation and cancer. Biochem Soc Trans (2018) 46:1147–59. doi: 10.1042/BST20180169
5. Padayatty SJ, Levine M. Vitamin c: the known and the unknown and goldilocks. Oral Dis (2016) 22:463–93. doi: 10.1111/odi.12446
6. Shenoy N, Creagan E, Witzig T, Levine M. Ascorbic acid in cancer treatment: Let the phoenix fly. Cancer Cell (2018) 34:700–6. doi: 10.1016/j.ccell.2018.07.014
7. Lee Chong T, Ahearn EL, Cimmino L. Reprogramming the epigenome with vitamin c. Front Cell Dev Biol (2019) 7:128. doi: 10.3389/fcell.2019.00128
8. Sasidharan Nair V, Song MH, Oh KI. Vitamin c facilitates demethylation of the Foxp3 enhancer in a tet-dependent manner. J Immunol (2016) 196:2119–31. doi: 10.4049/jimmunol.1502352
9. Yue X, Trifari S, Aijo T, Tsagaratou A, Pastor WA, Zepeda-Martinez JA, et al. Control of Foxp3 stability through modulation of TET activity. J Exp Med (2016) 213:377–97. doi: 10.1084/jem.20151438
10. Agathocleous M, Meacham CE, Burgess RJ, Piskounova E, Zhao Z, Crane GM, et al. Ascorbate regulates haematopoietic stem cell function and leukaemogenesis. Nature (2017) 549:476–81. doi: 10.1038/nature23876
11. Cimmino L, Dolgalev I, Wang Y, Yoshimi A, Martin GH, Wang J, et al. Restoration of TET2 function blocks aberrant self-renewal and leukemia progression. Cell (2017) 170:1079–1095 e1020. doi: 10.1016/j.cell.2017.07.032
12. Song MH, Nair VS, Oh KI. Vitamin c enhances the expression of IL17 in a Jmjd2-dependent manner. BMB Rep (2017) 50:49–54. doi: 10.5483/BMBRep.2017.50.1.193
13. Mingay M, Chaturvedi A, Bilenky M, Cao Q, Jackson L, Hui T, et al. Vitamin c-induced epigenomic remodelling in IDH1 mutant acute myeloid leukaemia. Leukemia (2018) 32:11–20. doi: 10.1038/leu.2017.171
14. Oyarce K, Campos-Mora M, Gajardo-Carrasco T, Pino-Lagos K. Vitamin c fosters the In vivo differentiation of peripheral CD4(+) Foxp3(-) T cells into CD4(+) Foxp3(+) regulatory T cells but impairs their ability to prolong skin allograft survival. Front Immunol (2018) 9:112. doi: 10.3389/fimmu.2018.00112
15. Iamsawat S, Tian L, Daenthanasanmak A, Wu Y, Nguyen HD, Bastian D, et al. Vitamin c stabilizes CD8+ iTregs and enhances their therapeutic potential in controlling murine GVHD and leukemia relapse. Blood Adv (2019) 3:4187–201. doi: 10.1182/bloodadvances.2019000531
16. Zhitkovich A. Nuclear and cytoplasmic functions of vitamin c. Chem Res Toxicol (2020) 33:2515–26. doi: 10.1021/acs.chemrestox.0c00348
17. Gonzalez-Menendez P, Romano M, Yan H, Deshmukh R, Papoin J, Oburoglu L, et al. An IDH1-vitamin c crosstalk drives human erythroid development by inhibiting pro-oxidant mitochondrial metabolism. Cell Rep (2021) 34:108723. doi: 10.1016/j.celrep.2021.108723
18. Burns JJ. Missing step in man, monkey and guinea pig required for the biosynthesis of l-ascorbic acid. Nature (1957) 180:553. doi: 10.1038/180553a0
19. Birney EC, Jenness R, Ayaz KM. Inability of bats to synthesise l-ascorbic acid. Nature (1976) 260:626–8. doi: 10.1038/260626a0
20. Nishikimi M, Kawai T, Yagi K. Guinea Pigs possess a highly mutated gene for l-gulono-gamma-lactone oxidase, the key enzyme for l-ascorbic acid biosynthesis missing in this species. J Biol Chem (1992) 267:21967–72. doi: 10.1016/S0021-9258(19)36707-9
21. Nishikimi M, Fukuyama R, Minoshima S, Shimizu N, Yagi K. Cloning and chromosomal mapping of the human nonfunctional gene for l-gulono-gamma-lactone oxidase, the enzyme for l-ascorbic acid biosynthesis missing in man. J Biol Chem (1994) 269:13685–8. doi: 10.1016/S0021-9258(17)36884-9
22. Drouin G, Godin JR, Page B. The genetics of vitamin c loss in vertebrates. Curr Genomics (2011) 12:371–8. doi: 10.2174/138920211796429736
23. Maeda N, Hagihara H, Nakata Y, Hiller S, Wilder J, Reddick R. Aortic wall damage in mice unable to synthesize ascorbic acid. Proc Natl Acad Sci U.S.A. (2000) 97:841–6. doi: 10.1073/pnas.97.2.841
24. Chen Y, Curran CP, Nebert DW, Patel KV, Williams MT, Vorhees CV. Effect of vitamin c deficiency during postnatal development on adult behavior: functional phenotype of gulo-/- knockout mice. Genes Brain Behav (2012) 11:269–77. doi: 10.1111/j.1601-183X.2011.00762.x
25. Godin I, Cumano A. Of birds and mice: hematopoietic stem cell development. Int J Dev Biol (2005) 49:251–7. doi: 10.1387/ijdb.041945ig
26. Yamane T. Mouse yolk sac hematopoiesis. Front Cell Dev Biol (2018) 6:80. doi: 10.3389/fcell.2018.00080
27. Palis J. Primitive and definitive erythropoiesis in mammals. Front Physiol (2014) 5:3. doi: 10.3389/fphys.2014.00003
28. Kikuchi K, Kondo M. Developmental switch of mouse hematopoietic stem cells from fetal to adult type occurs in bone marrow after birth. Proc Natl Acad Sci U.S.A. (2006) 103:17852–7. doi: 10.1073/pnas.0603368103
29. Morrison SJ, Wandycz AM, Akashi K, Globerson A, Weissman IL. The aging of hematopoietic stem cells. Nat Med (1996) 2:1011–6. doi: 10.1038/nm0996-1011
30. Hackney JA, Charbord P, Brunk BP, Stoeckert CJ, Lemischka IR, Moore KA. A molecular profile of a hematopoietic stem cell niche. Proc Natl Acad Sci U.S.A. (2002) 99:13061–6. doi: 10.1073/pnas.192124499
31. Heissig B, Ohki Y, Sato Y, Rafii S, Werb Z, Hattori K. A role for niches in hematopoietic cell development. Hematology (2005) 10:247–53. doi: 10.1080/10245330500067249
32. Bowie MB, McKnight KD, Kent DG, McCaffrey L, Hoodless PA, Eaves CJ. Hematopoietic stem cells proliferate until after birth and show a reversible phase-specific engraftment defect. J Clin Invest (2006) 116:2808–16. doi: 10.1172/JCI28310
33. Iwama M, Amano A, Shimokado K, Maruyama N, Ishigami A. Ascorbic acid levels in various tissues, plasma and urine of mice during aging. J Nutr Sci Vitaminol (Tokyo) (2012) 58:169–74. doi: 10.3177/jnsv.58.169
34. Kim H, Bae S, Yu Y, Kim Y, Kim HR, Hwang YI, et al. The analysis of vitamin c concentration in organs of gulo(-/-) mice upon vitamin c withdrawal. Immune Netw (2012) 12:18–26. doi: 10.4110/in.2012.12.1.18
35. Vissers MC, Bozonet SM, Pearson JF, Braithwaite LJ. Dietary ascorbate intake affects steady state tissue concentrations in vitamin c-deficient mice: tissue deficiency after suboptimal intake and superior bioavailability from a food source (kiwifruit). Am J Clin Nutr (2011) 93:292–301. doi: 10.3945/ajcn.110.004853
36. Oguro H, Ding L, Morrison SJ. SLAM family markers resolve functionally distinct subpopulations of hematopoietic stem cells and multipotent progenitors. Cell Stem Cell (2013) 13:102–16. doi: 10.1016/j.stem.2013.05.014
37. Boyer SW, Schroeder AV, Smith-Berdan S, Forsberg EC. All hematopoietic cells develop from hematopoietic stem cells through Flk2/Flt3-positive progenitor cells. Cell Stem Cell (2011) 9:64–73. doi: 10.1016/j.stem.2011.04.021
38. Challen GA, Boles N, Lin KK, Goodell MA. Mouse hematopoietic stem cell identification and analysis. Cytometry A (2009) 75:14–24. doi: 10.1002/cyto.a.20674
39. Kondo M, Weissman IL, Akashi K. Identification of clonogenic common lymphoid progenitors in mouse bone marrow. Cell (1997) 91:661–72. doi: 10.1016/S0092-8674(00)80453-5
40. Akashi K, Traver D, Miyamoto T, Weissman IL. A clonogenic common myeloid progenitor that gives rise to all myeloid lineages. Nature (2000) 404:193–7. doi: 10.1038/35004599
41. Ema H, Morita Y, Suda T. Heterogeneity and hierarchy of hematopoietic stem cells. Exp Hematol (2014) 42:74–82 e72. doi: 10.1016/j.exphem.2013.11.004
42. Lebson L, Nash K, Kamath S, Herber D, Carty N, Lee DC, et al. Trafficking CD11b-positive blood cells deliver therapeutic genes to the brain of amyloid-depositing transgenic mice. J Neurosci (2010) 30:9651–8. doi: 10.1523/JNEUROSCI.0329-10.2010
43. Manning J, Mitchell B, Appadurai DA, Shakya A, Pierce LJ, Wang H, et al. Vitamin c promotes maturation of T-cells. Antioxid Redox Signal (2013) 19:2054–67. doi: 10.1089/ars.2012.4988
44. Rodewald HR, Fehling HJ. Molecular and cellular events in early thymocyte development. Adv Immunol (1998) 69:1–112. doi: 10.1016/S0065-2776(08)60606-9
45. Hoffman ES, Passoni L, Crompton T, Leu TM, Schatz DG, Koff A, et al. Productive T-cell receptor beta-chain gene rearrangement: coincident regulation of cell cycle and clonality during development in vivo. Genes Dev (1996) 10:948–62. doi: 10.1101/gad.10.8.948
46. Blom B, Spits H. Development of human lymphoid cells. Annu Rev Immunol (2006) 24:287–320. doi: 10.1146/annurev.immunol.24.021605.090612
47. Ciofani M, Zuniga-Pflucker JC. Notch promotes survival of pre-T cells at the beta-selection checkpoint by regulating cellular metabolism. Nat Immunol (2005) 6:881–8. doi: 10.1038/ni1234
48. Swainson L, Kinet S, Manel N, Battini JL, Sitbon M, Taylor N. Glucose transporter 1 expression identifies a population of cycling CD4+ CD8+ human thymocytes with high CXCR4-induced chemotaxis. Proc Natl Acad Sci U.S.A. (2005) 102:12867–72. doi: 10.1073/pnas.0503603102
49. Qi T, Sun M, Zhang C, Chen P, Xiao C, Chang X. Ascorbic acid promotes plasma cell differentiation through enhancing TET2/3-mediated DNA demethylation. Cell Rep (2020) 33:108452. doi: 10.1016/j.celrep.2020.108452
50. Astori M, Finke D, Karapetian O, Acha-Orbea H. Development of T-b cell collaboration in neonatal mice. Int Immunol (1999) 11:445–51. doi: 10.1093/intimm/11.3.445
51. Myneni VD, Szalayova I, Mezey E. Differences in steady-state erythropoiesis in different mouse bones and postnatal spleen. Front Cell Dev Biol (2021) 9:646646. doi: 10.3389/fcell.2021.646646
52. Barminko J, Reinholt B, Baron MH. Development and differentiation of the erythroid lineage in mammals. Dev Comp Immunol (2016) 58:18–29. doi: 10.1016/j.dci.2015.12.012
53. Chen L, Wang J, Liu J, Wang H, Hillyer CD, Blanc L, et al. Dynamic changes in murine erythropoiesis from birth to adulthood: implications for the study of murine models of anemia. Blood Adv (2021) 5:16–25. doi: 10.1182/bloodadvances.2020003632
54. Socolovsky M, Nam H, Fleming MD, Haase VH, Brugnara C, Lodish HF. Ineffective erythropoiesis in Stat5a(-/-)5b(-/-) mice due to decreased survival of early erythroblasts. Blood (2001) 98:3261–73. doi: 10.1182/blood.V98.12.3261
55. Chen K, Liu J, Heck S, Chasis JA, An X, Mohandas N. Resolving the distinct stages in erythroid differentiation based on dynamic changes in membrane protein expression during erythropoiesis. Proc Natl Acad Sci U.S.A. (2009) 106:17413–8. doi: 10.1073/pnas.0909296106
56. Liu J, Zhang J, Ginzburg Y, Li H, Xue F, De Franceschi L, et al. Quantitative analysis of murine terminal erythroid differentiation in vivo: novel method to study normal and disordered erythropoiesis. Blood (2013) 121:e43–49. doi: 10.1182/blood-2012-09-456079
57. Montel-Hagen A, Blanc L, Boyer-Clavel M, Jacquet C, Vidal M, Sitbon M, et al. The Glut1 and Glut4 glucose transporters are differentially expressed during perinatal and postnatal erythropoiesis. Blood (2008) 112:4729–38. doi: 10.1182/blood-2008-05-159269
58. Moreau R, Tshikudi Malu D, Dumais M, Dalko E, Gaudreault V, Romero H, et al. Alterations in bone and erythropoiesis in hemolytic anemia: Comparative study in bled, phenylhydrazine-treated and plasmodium-infected mice. PloS One (2012) 7:e46101. doi: 10.1371/journal.pone.0046101
59. Paulson RF, Hariharan S, Little JA. Stress erythropoiesis: definitions and models for its study. Exp Hematol (2020) 89:43–54 e42. doi: 10.1016/j.exphem.2020.07.011
60. Laky K, Dugan P, Frischmeyer-Guerrerio PA. Hematopoietic reconstitution of neonatal immunocompetent mice to study conditions with a perinatal window of susceptibility. Sci Rep (2018) 8:12254. doi: 10.1038/s41598-018-30767-1
61. Gao X, Xu C, Asada N, Frenette PS. The hematopoietic stem cell niche: from embryo to adult. Development (2018) 145. doi: 10.1242/dev.139691
62. Pucella JN, Upadhaya S, Reizis B. The source and dynamics of adult hematopoiesis: Insights from lineage tracing. Annu Rev Cell Dev Biol (2020) 36:529–50. doi: 10.1146/annurev-cellbio-020520-114601
63. Nakajima Y, Shantha TR, Bourne GH. Histochemical detection of l-gulonolactone: phenazine methosulfate oxidoreductase activity in several mammals with special reference to synthesis of vimin c in primates. Histochemie (1969) 18:293–301. doi: 10.1007/BF00279880
64. Pollock JI, Mullin RJ. Vitamin c biosynthesis in prosimians: evidence for the anthropoid affinity of tarsius. Am J Phys Anthropol (1987) 73:65–70. doi: 10.1002/ajpa.1330730106
65. Cui J, Yuan X, Wang L, Jones G, Zhang S. Recent loss of vitamin c biosynthesis ability in bats. PloS One (2011) 6:e27114. doi: 10.1371/journal.pone.0027114
66. Mizushima Y, Harauchi T, Yoshizaki T, Makino S. A rat mutant unable to synthesize vitamin c. Experientia (1984) 40:359–61. doi: 10.1007/BF01952551
67. Horio F, Ozaki K, Yoshida A, Makino S, Hayashi Y. Requirement for ascorbic acid in a rat mutant unable to synthesize ascorbic acid. J Nutr (1985) 115:1630–40. doi: 10.1093/jn/115.12.1630
68. Campbell EJ, Vissers MC, Bozonet S, Dyer A, Robinson BA, Dachs GU. Restoring physiological levels of ascorbate slows tumor growth and moderates HIF-1 pathway activity in gulo(-/-) mice. Cancer Med (2015) 4:303–14. doi: 10.1002/cam4.349
69. Tu H, Li H, Wang Y, Niyyati M, Wang Y, Leshin J, et al. Low red blood cell vitamin c concentrations induce red blood cell fragility: A link to diabetes Via glucose, glucose transporters, and dehydroascorbic acid. EBioMedicine (2015) 2:1735–50. doi: 10.1016/j.ebiom.2015.09.049
70. Johnson RJ, Gaucher EA, Sautin YY, Henderson GN, Angerhofer AJ, Benner SA. The planetary biology of ascorbate and uric acid and their relationship with the epidemic of obesity and cardiovascular disease. Med Hypotheses (2008) 71:22–31. doi: 10.1016/j.mehy.2008.01.017
71. Chatterjee IB. Evolution and the biosynthesis of ascorbic acid. Science (1973) 182:1271–2. doi: 10.1126/science.182.4118.1271
73. Council NR. Dietary reference intakes for vitamin c, vitamin e, selenium, and carotenoids. Washington (DC) (2000).
74. Milton K, Jenness R. Ascorbic acid content of neotropical plant parts available to wild monkeys and bats. Experientia (1987) 43:339–42. doi: 10.1007/BF01945577
75. Su D, May JM, Koury MJ, et al. Human erythrocyte membranes contain a cytochrome b561 that may be involved in extracellular ascorbate recycling. J Biol Chem (2006) 281:39852–9. doi: 10.1074/jbc.M606543200
76. Bianchi J, Rose RC. Glucose-independent transport of dehydroascorbic acid in human erythrocytes. Proc Soc Exp Biol Med (1986) 181:333–7. doi: 10.3181/00379727-181-42261
77. Vera JC, Rivas CI, Fischbarg J, Golde DW. Mammalian facilitative hexose transporters mediate the transport of dehydroascorbic acid. Nature (1993) 364:79–82. doi: 10.1038/364079a0
78. Rumsey SC, Kwon O, Xu GW, Burant CF, Simpson I, Levine M. Glucose transporter isoforms GLUT1 and GLUT3 transport dehydroascorbic acid. J Biol Chem (1997) 272:18982–9. doi: 10.1074/jbc.272.30.18982
79. Montel-Hagen A, Kinet S, Manel N, Mongellaz C, Prohaska R, Battini JL, et al. Erythrocyte Glut1 triggers dehydroascorbic acid uptake in mammals unable to synthesize vitamin c. Cell (2008) 132:1039–48. doi: 10.1016/j.cell.2008.01.042
80. Helgerson AL, Carruthers A. Equilibrium ligand binding to the human erythrocyte sugar transporter. evidence for two sugar-binding sites per carrier. J Biol Chem (1987) 262:5464–75. doi: 10.1016/S0021-9258(18)45595-0
81. Mueckler M. Facilitative glucose transporters. Eur J Biochem (1994) 219:713–25. doi: 10.1111/j.1432-1033.1994.tb18550.x
82. May JM, Qu Z, Morrow JD. Mechanisms of ascorbic acid recycling in human erythrocytes. Biochim Biophys Acta (2001) 1528:159–66. doi: 10.1016/S0304-4165(01)00188-X
83. Duggan GE, Joan Miller B, Jirik FR, Vogel HJ. Metabolic profiling of vitamin c deficiency in gulo-/- mice using proton NMR spectroscopy. J Biomol NMR (2011) 49:165–73. doi: 10.1007/s10858-011-9485-5
84. Inoue S, Lemonnier F, Mak TW. Roles of IDH1/2 and TET2 mutations in myeloid disorders. Int J Hematol (2016) 103:627–33. doi: 10.1007/s12185-016-1973-7
85. Cerullo G, Negro M, Parimbelli M, Pecoraro M, Perna S, Liguori G, et al. The long history of vitamin c: From prevention of the common cold to potential aid in the treatment of COVID-19. Front Immunol (2020) 11:574029. doi: 10.3389/fimmu.2020.574029
86. Hoang BX, Shaw G, Fang W, Han B. Possible application of high-dose vitamin c in the prevention and therapy of coronavirus infection. J Glob Antimicrob Resist (2020) 23:256–62. doi: 10.1016/j.jgar.2020.09.025
87. Moya-Alvarez V, Koyembi JJ, Kaye LM, Mbecko JR, Sanke-Waigana H, Djorie SG, et al. Vitamin c levels in a central-African mother-infant cohort: Does hypovitaminosis c increase the risk of enteric infections? Matern Child Nutr (2021) 17:e13215. doi: 10.1111/mcn.13215
88. Seibert E, Richter A, Kuhlmann MK, Wang S, Levin NW, Kotanko P, et al. Plasma vitamin c levels in ESRD patients and occurrence of hypochromic erythrocytes. Hemodial Int (2017) 21:250–5. doi: 10.1111/hdi.12467
89. Guo S, Jiang X, Wang Y, Chen L, Li H, Li X, et al. The protective role of TET2 in erythroid iron homeostasis against oxidative stress and erythropoiesis. Cell Signal (2017) 38:106–15. doi: 10.1016/j.cellsig.2017.07.002
90. Yan H, Wang Y, Qu X, Li J, Hale J, Huang Y, et al. Distinct roles for TET family proteins in regulating human erythropoiesis. Blood (2017) 129:2002–12. doi: 10.1182/blood-2016-08-736587
91. Qu X, Zhang S, Wang S, Wang Y, Li W, Huang Y, et al. TET2 deficiency leads to stem cell factor-dependent clonal expansion of dysfunctional erythroid progenitors. Blood (2018) 132:2406–17. doi: 10.1182/blood-2018-05-853291
92. Morales-Borges R, Rivera-Rodrígue KM. Effectiveness of vitamin c in the treatment of anemia in patients with chronic diseases: A case study (2020). Available at: http://www.bookpi.org/.
93. Taji Y, Morimoto T, Okada K, Fukuhara S, Fukui T, Kuwahara T. Effects of intravenous ascorbic acid on erythropoiesis and quality of life in unselected hemodialysis patients. J Nephrol (2004) 17:537–43.
94. Giancaspro V, Nuzziello M, Pallotta G, Sacchetti A, Petrarulo F. Intravenous ascorbic acid in hemodialysis patients with functional iron deficiency: a clinical trial. J Nephrol (2000) 13:444–9.
95. Sezer S, Ozdemir FN, Yakupoglu U, Arat Z, Turan M, Haberal M. Intravenous ascorbic acid administration for erythropoietin-hyporesponsive anemia in iron loaded hemodialysis patients. Artif Organs (2002) 26:366–70. doi: 10.1046/j.1525-1594.2002.06888.x
96. Li N, Zhao G, Wu W, Zhang M, Liu W, Chen Q, et al. The efficacy and safety of vitamin c for iron supplementation in adult patients with iron deficiency anemia: A randomized clinical trial. JAMA Netw Open (2020) 3:e2023644. doi: 10.1001/jamanetworkopen.2020.23644
97. Arruda MM, Mecabo G, Rodrigues CA, Matsuda SS, Rabelo IB, Figueiredo MS. Antioxidant vitamins c and e supplementation increases markers of haemolysis in sickle cell anaemia patients: a randomized, double-blind, placebo-controlled trial. Br J Haematol (2013) 160:688–700. doi: 10.1111/bjh.12185
98. Hasanato RM. Zinc and antioxidant vitamin deficiency in patients with severe sickle cell anemia. Ann Saudi Med (2006) 26:17–21. doi: 10.5144/0256-4947.2006.17
99. Li H, Tu H, Wang Y, Levine M. Vitamin c in mouse and human red blood cells: an HPLC assay. Anal Biochem (2012) 426:109–17. doi: 10.1016/j.ab.2012.04.014
Keywords: vitamin C, ascorbate, GULO, hematopoiesis, erythropoiesis, anemia, neonatal, development
Citation: Phadke I, Pouzolles M, Machado A, Moraly J, Gonzalez-Menendez P, Zimmermann VS, Kinet S, Levine M, Violet P-C and Taylor N (2022) Vitamin C deficiency reveals developmental differences between neonatal and adult hematopoiesis. Front. Immunol. 13:898827. doi: 10.3389/fimmu.2022.898827
Received: 17 March 2022; Accepted: 02 September 2022;
Published: 30 September 2022.
Edited by:
Nina Cabezas-Wallscheid, Max Planck Institute for Immunobiology and Epigenetics, GermanyReviewed by:
Roshana Thambyrajah, Mar Institute of Medical Research (IMIM), SpainMarie-Laure Arcangeli, Institut National de la Santé et de la Recherche Médicale (INSERM), France
Copyright © 2022 Phadke, Pouzolles, Machado, Moraly, Gonzalez-Menendez, Zimmermann, Kinet, Levine, Violet and Taylor. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.
*Correspondence: Mark Levine, markl@bdg8.niddk.nih.gov; Pierre-Christian Violet, pierre-christian.violet@nih.gov; Naomi Taylor, naomi.taylor@igmm.cnrs.fr