Abstract
Neudesin, originally identified as a neurotrophic factor, has primarily been studied for its neural functions despite its widespread expression. Using 8-week-old neudesin knockout mice, we elucidated the role of neudesin in the spleen. The absence of neudesin caused mild splenomegaly, shortened lifespan of circulating erythrocytes, and abnormal recovery from phenylhydrazine-induced acute anemia. Blood cross-transfusion and splenectomy experiments revealed that the shortened lifespan of erythrocytes was attributable to splenic impairment. Further analysis revealed increased erythrophagocytosis and decreased iron stores in the splenic red pulp, which was linked to the upregulation of Fcγ receptors and iron-recycling genes in neudesin-deficient macrophages. In vitro analysis confirmed that neudesin suppressed erythrophagocytosis and expression of Fcγ receptors through ERK1/2 activation in heme-stimulated macrophages. Finally, we observed that 24-week-old neudesin knockout mice exhibited severe symptoms of anemia. Collectively, our results suggest that neudesin regulates the function of red pulp macrophages and contributes to erythrocyte and iron homeostasis.
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Introduction
After development in the bone marrow, red blood cells (RBCs) enter the circulatory system and transport oxygen throughout the body. The structural flexibility and deformability of the RBC membrane are critical for transport through narrow blood capillaries, but they gradually decrease with age owing to physical and oxidative stresses1,2. Therefore, the body employs a mechanism to remove aged RBCs from circulation. After circulating for approximately 120 days in humans and 30 days in mice, aged RBCs are captured mainly by red pulp macrophages (RPMs) in the red pulp of the spleen. These macrophages digest aged RBCs and release heme-associated iron, which is recycled into new hemoglobin during erythropoiesis in the bone marrow3.
Several biological factors are implicated in the removal of aged RBCs. So-called “eat me” signals emerge on the surface of aged RBCs, triggering their clearance1. First, oxidative stress induces clustering of membrane protein band3, an abundant anion-transporter protein of RBCs1. The cluster is detected by naturally occurring anti-band3 autoantibodies, and complement activation through the classical pathway further enhances the opsonization of aged RBCs. This complex is recognized by Fcγ receptors (FcγRs) and complement receptor CR3 (Mac-1), which comprises CD11b and CD18 on the cell membrane of macrophages and induces erythrophagocytosis. As the second “eat me” signal, phosphatidylserine (PS) is exposed on the outer surface of RBCs2. Similar to other cell types, PS is located on the inner leaflet of healthy RBCs; however, under oxidative stress, osmotic shock, or other forms of cell stress, PS is exposed and accumulated in the outer leaflet of RBCs. Macrophages recognize PS-presenting RBCs with PS receptors, including Stabilin-2, Tim-1, and Tim-44,5. TAM receptors (Tyro3, Axl, Mer) also recognize PS through mediation by the bridging molecule Gas-61. The recognition of PS exposure by these receptors leads to phagocytosis of aged RBCs. Meanwhile, a “don’t eat me” signal functions as a suppressor of erythrophagocytosis. CD47 expressed on the surface of healthy RBCs binds to signal-regulatory protein alpha (SIRPα) present on macrophages, inhibiting erythrophagocytosis6. However, oxidative stress leads to a conformational change in the CD47 molecule, thereby switching on the “eat me” signal and inducing phagocytic signals through SIRPα, resulting in the clearance of aged RBCs7.
We previously identified and isolated a heme-binding secreted protein, neudesin, which is expressed in several tissues with the highest abundance in the central nervous system, with in vitro neurotrophic activity via the activation of intracellular signaling pathways8,9. Neudesin contains a heme/steroid-binding domain highly similar to cytochrome b5, which maintains the reduced state of hemoglobin in erythrocytes, and its neurotrophic activity is dependent on heme binding to this domain10. Although its receptor has not yet been identified, it is expected to be a G protein-coupled receptor (GPCR) owing to the inhibition of neudesin activity by inhibitors of the G protein-coupled signaling pathway8,9,11. In the adult mouse brain, neudesin is involved in appetite, anxiety behavior, and contextual fear memory12,13,14. Furthermore, neudesin has been increasingly studied in non-neural tissues. To date, our group demonstrated the functions of neudesin in adipose tissues15,16, and another group reported that neudesin regulates bovine adipogenesis and myogenesis17. Pathophysiologically, neudesin is also involved in tumorigenesis18,19,20.
In addition to its expression in neural and adipose tissues, neudesin mRNA is expressed in the spleen, although its splenic effects remain to be elucidated. In this study, we demonstrated the function of neudesin in the spleen. Eight-week-old neudesin knockout (KO) mice displayed abnormal RBC accumulation in the spleen. Circulating erythrocytes from neudesin KO mice showed enhanced resistance against hemolytic conditions in vitro as well as a shortened lifespan in vivo. These erythrocyte phenotypes were largely attributable to the hyperactivation of erythrophagocytosis by RPMs lacking neudesin, presumably owing to upregulation of FcγRs on RPMs of neudesin KO mice. Neudesin deficiency also led to impaired iron homeostasis in the spleen and incomplete recovery from drug-induced acute anemia. After maturation, 24-week-old neudesin KO mice exhibited normocytic anemia associated. Collectively, neudesin contributes to the homeostasis of erythrocytes and iron by regulating macrophage functions.
Results
Neudesin-deficient mice have more erythrocytes in the spleen and abnormalities in circulating erythrocytes
In addition to the brain and white adipose tissue where neudesin’s function has been previously reported, neudesin is also expressed in the spleen (Fig. 1a). However, its function in the spleen is not clear. Therefore, we first examined the spleens of 8-week-old WT and neudesin KO mice. The spleens of neudesin KO mice were modestly larger and slightly (but significantly) heavier than those of WT mice (Fig. 1b, c), suggesting that neudesin influenced spleen contents. Therefore, we investigated which type of cells contributed to mild splenomegaly in neudesin KO mice. Flow cytometric analysis showed that the spleens of neudesin KO mice contained approximately 1.5-fold more TER119+ RBCs compared with those of WT mice (Fig. 1d). The number of CD45+ white blood cells in the spleens was comparable between WT and neudesin KO mice. These data suggest that neudesin is involved in RBC homeostasis, as in the case of cytochrome b5 reductase, which has a heme-binding domain similar to the heme/steroid-binding domain of neudesin10. To assess this possibility, peripheral blood was hematologically assessed, revealing almost normal RBCs except for a slight reduction in mean corpuscular hemoglobin (MCH) and an increase in platelet count in neudesin KO mice (Table 1). Additionally, the RBCs of neudesin KO mice were significantly more resistant to hypertonic and sugar-depleted hemolytic conditions (Fig. 1e). These data indicated that the RBCs of neudesin KO mice were more deformable. Because the deformability of erythrocytes is used as an indicator of cellular senescence1, circulating erythrocytes in neudesin KO mice may be a younger population than those in WT mice. We further measured the ratios of PS exposure in erythrocytes and naturally occurring antibody-bound erythrocytes, which revealed no significant difference between WT and neudesin KO mice (Fig. 1f, g). These results showed that there was little change in the percentage of aged erythrocytes between WT and neudesin KO mice.
a Relative neudesin mRNA levels in tissues of 8-week-old C57BL/6 male mice (n = 5) were quantified by RT-qPCR. WAT, white adipose tissue. BM, bone marrow. b A representative picture of spleens from 8-week-old wildtype (WT) and neudesin knockout (KO) male mice. c Spleens of neudesin KO mice (n = 15, male) were significantly heavier than those of 8-week-old WT male mice (n = 15, P = 0.0393 by unpaired t test). d Splenocyte suspensions from 8-week-old WT (n = 16) and neudesin KO male mice (n = 15) were assessed by flow cytometric analysis using anti-TER119 and anti-CD45 mAbs, and cell numbers of TER119+ red blood cells and CD45+ white blood cells in the spleen were counted. P = 0.0098, and 0.1862, respectively, by unpaired t test. e Hemolysis ratios were measured after incubating blood cells from WT and KO male mice for 16 h in Ringer’s solution (left), hypertonic solution (middle), or sugar-depleted solution (right). P = 0.8905, 0.0059, and <0.0001, respectively, by unpaired t-test (n = 8 for each group). Exposure of phosphatidylserine on the outer plasma membrane of erythrocytes was measured by annexin V binding (f) (n = 15 for each group, male, P = 0.2840), and naturally occurring antibody-bound erythrocytes were detected with anti-mouse IgG and IgM antibodies (g) (WT: n = 15, KO: n = 16, male, P = 0.3183).
Erythrocytes in neudesin KO mice have a shortened lifespan owing to impairment of the spleen rather than of erythrocytes
Our data suggest that neudesin is implicated in the senescence of RBCs. Accordingly, we examined the lifespan of RBCs in WT and neudesin KO mice using direct in vivo biotinylation of RBCs. Sulfo-NHS-biotin was injected intravenously to label RBCs, and the lifespan of RBCs was determined by flow cytometric analysis of peripheral biotinylated RBCs (Fig. 2a, b). The half-life of labeled erythrocytes in WT mice was approximately 17.2 ± 3.0 days; in contrast, that in neudesin KO mice decreased to about 13.3 ± 1.7 days (Fig. 2c).
a Scheme of experiments to measure erythrocyte lifespan in vivo using intravenous administration of sulfo-NHS-biotin at 8 weeks of age. RBCs were collected from tail veins on days 3, 10, 17, and 24 after administration. Biotinylated RBCs were detected using fluorescently conjugated streptavidin. b More than 99% of RBCs were biotinylated on day 3 after administration (3 d; gray histogram in the left panel). The right panel shows representative histograms of WT (white) and neudesin KO (gray) on day 24 after administration (24 d). c The percentage of biotinylated RBCs from WT and neudesin KO male mice was tracked by flow cytometry. WT: n = 12, KO: n = 10. P = 0.0010. Blood samples were collected from WT (n ≧ 7) and neudesin KO (n ≧ 9) male mice on days 0, 4, and 8 after intraperitoneal administration of phenylhydrazine at 8 weeks of age. RBC counts (d) (P = 0.4199), hemoglobin concentrations (Hb) (e) (P = 0.0019), and hematocrit levels (Ht) (f) (P = 0.1057) were measured. g Relative neudesin mRNA levels in spleens of WT male mice (n = 3) were quantified by RT-qPCR on days 0, 2, 4, and 6 after intraperitoneal administration of phenylhydrazine. h The scheme of experiments to measure the lifespan of RBCs transfused from WT or neudesin KO male mice into WT or neudesin KO male mice at 8 weeks of age. RBCs were collected on days 1, 8, 15, and 22 after transfusion. i The percentage of survival of transfused RBCs was tracked by flow cytometry. WT RBCs transfused into WT mice: n = 10, WT RBCs into KO mice: n = 6, KO RBCs into WT mice: n = 5, KO RBCs into KO mice: n = 6. (WT RBCs > KO mice vs. WT RBCs > WT mice: P = 0.0260, KO RBCs > WT mice vs. WT RBCs > WT mice: P > 0.9999, KO RBCs > KO mice vs. WT RBCs > WT mice: P < 0.0001, KO RBCs > WT mice vs. WT RBCs > KO mice: P = 0.3728, KO RBCs > KO mice vs. WT RBCs > KO mice: P = 0.9843, KO RBCs > KO mice vs. KO RBCs > WT mice: P = 0.0077). j The scheme of experiments to measure the lifespan of erythrocytes 4 weeks after splenectomy or sham-splenectomy. RBCs were collected on days 3, 10, 17, and 24 after administration. k The percentage of biotinylated RBCs from splenectomized and sham-splenectomized male mice was tracked by flow cytometry. WT sham: n = 4, KO sham: n = 4, WT splenectomy: n = 6, KO splenectomy: n = 6. (KO sham vs. WT sham: P = 0.0087, WT splenectomy vs. WT sham: P = > 0.9999, KO splenectomy vs. WT sham: P = > 0.9999, WT splenectomy vs. KO sham: P = 0.0117, KO splenectomy vs. KO sham: P = 0.0016, KO splenectomy vs. WT splenectomy: P = > 0.9999) Data are shown as means ± SD of two or more experiments. *P < 0.05 and **P < 0.01 by a 2-way analysis of variance followed by Sidak’s post-test.
Because neudesin KO mice had a shortened lifespan of erythrocytes under normal conditions, we compared erythrocytes from WT and neudesin KO mice during acute anemia induced by phenylhydrazine injection. Phenylhydrazine injection decreased RBC counts and hematocrit levels, which recovered 4 and 8 days after injection in WT neudesin KO and mice, respectively (Fig. 2d, f). Neudesin KO mice showed impaired hemoglobin concentrations after injection (Fig. 2e). Neudesin mRNA levels were barely detected in the bone marrow but remarkably increased in the spleens of WT mice after induction of acute anemia (Fig. 2g). These data suggest that neudesin is necessary for iron recycling in acute anemic conditions.
To further investigate the cause of the shortened lifespan of RBCs in neudesin KO mice, we performed a blood cross-transfusion experiment. Biotinylated WT or neudesin KO RBCs were transfused into WT or neudesin KO mice (Fig. 2h). Consequently, mice in both neudesin KO groups demonstrated accelerated clearance of biotinylated RBCs compared with that of WT mice transfused with the same RBCs (Fig. 2i). These results suggest that some environments surrounding erythrocytes rather than the properties of the erythrocytes themselves cause the shortened lifespan of RBCs in neudesin KO mice.
The splenomegaly phenotype in neudesin KO mice implies that impaired spleen function leads to a shortened erythrocyte lifespan in neudesin KO mice. Thus, we examined the effect of splenectomy on erythrocyte lifespan (Fig. 2j). In splenectomized neudesin KO mice, the lifespan of erythrocytes was recovered equally to that of splenectomized WT mice, while the erythrocyte lifespan of sham KO mice remained shorter than that of sham WT mice (Fig. 2k). These data indicate that the shortened lifespan of RBCs in neudesin KO mice might be caused by increased removal of aged RBCs in the spleen.
Enhanced erythrophagocytosis and decreased iron accumulation by RPM lacking neudesin
The spleen is known to recognize and remove old, malformed, and damaged erythrocytes. RPMs are the primary phagocytes that engulf these erythrocytes in the spleen. To investigate whether neudesin was involved in the phagocytic function of RPM, we first examined RPM populations in neudesin KO mice. The ratios, absolute cell numbers, and distributions of RPMs showed little difference between WT and neudesin KO mice (Fig. 3a–c). Next, to investigate the erythrophagocytic activity of RPMs, erythrocytes were labeled ex vivo with PKH26 fluorescent dye and traced in WT and neudesin KO mice (Fig. 3d–f). Three days after PKH26+ RBCs were administered, about 50% of WT RPMs had engulfed PKH26+ RBCs (Fig. 3e, f). In neudesin KO mice, the ratio of PKH+ RPMs increased significantly to about 60% (Fig. 3f). Considering that neudesin mRNA was relatively abundant in F4/80+ RPMs (Fig. 3g), it can be assumed that neudesin suppresses phagocytic activity in an autocrine/paracrine manner. In addition to erythrophagocytosis, RPMs were involved in the accumulation and distribution of iron. We examined the involvement of neudesin in iron homeostasis. Perl’s Prussian staining, which specifically stains iron, revealed decreased iron accumulation in neudesin KO RPMs (Fig. 3h). Quantitative determinations of the non-heme iron contents in spleens, livers, kidneys, and blood serum revealed that splenic iron content reduced by ~40% in neudesin KO mice (Fig. 3i, j).
a Splenocytes from 8-week-old WT and neudesin KO male mice were stained with anti-F4/80 and anti-CD11b mAbs. Representative flow cytometry plots show populations of red pulp macrophages (RPM). b Chart showing the number of RPMs from 8-week-old WT and neudesin KO male mice (WT: n = 15, KO: n = 15, P = 0.6308). c Autofluorescence of spleen sections from 8-week-old WT and neudesin KO male mice were observed to visualize RPMs. Dotted lines indicate boundaries between red pulp (rp) and white pulp (wp). d Scheme of experiments to measure erythrophagocytosis in vivo using transfusion of PKH26-conjugated RBCs into 8-week-old WT and KO male mice. On day 3 after transfusion, splenocytes were collected and analyzed by flow cytometry to detect PKH26+ RPMs. e Representative histograms of RPMs from 8-week-old WT (white) and KO (gray) male mice. f The percentage of PKH+ RPMs from 8-week-old WT and neudesin KO male mice tracked by flow cytometry. *P < 0.05 by unpaired t-test (WT: n = 15, KO: n = 15, P = 0.0469). g Splenocytes from 8-weeks old male mice were sorted with lineage-specific antibodies, and relative neudesin mRNA expression levels were quantified by RT-qPCR (n = 5). h Spleen sections from 8-week-old WT and neudesin KO male mice were stained with Perl’s Prussian blue to visualize the iron deposits. Dotted lines indicate boundaries between red pulp (rp) and white pulp (wp). i Non-heme iron content was quantified in spleens, livers, and kidneys from 8-week-old WT and neudesin KO male mice. Each symbol represents an individual mouse. Spleen: P = 0.0383, liver: P = 0.0737, and kidney: P = 0.1793 by unpaired t-test (n = 8). j The graph shows serum iron levels of 8-week-old WT and neudesin KO male mice. P = 0.1924 by unpaired t-test (n = 6) Each symbol represents an individual mouse. Data are shown as means ± SD of three or more experiments.
FcγR upregulation on the surface of RPM in neudesin KO mice
Erythrophagocytosis is triggered by several types of phagocytic receptors on the surface of RPMs1. We, therefore searched for receptors that mediate accelerated erythrophagocytosis in neudesin KO mice. The quantification of receptors by flow cytometry revealed the upregulation of FcγRs (FcγR1, FcγR2/3, and FcγR4) on the surface of RPMs in neudesin KO mice, whereas the expression of receptors recognizing PS (Stabilin-2, Tim-4, and Axl) remained unchanged (Fig. 4a, b). Furthermore, the levels of CD163 molecules, which are erythroblast adhesion receptors21, also increased in the neudesin KO RPMs. To investigate whether the upregulation of FcγRs and CD163 is induced at the transcriptional level in neudesin KO RPMs, RPMs from WT and neudesin KO mice were isolated with anti-F4/80 microbeads, and Adgre1-coding F4/80 molecules, Fcgr1, Fcgr2b, Fcgr3, Fcgr4, and CD163 mRNAs were quantified (Fig. 4c). In neudesin KO RPMs, Fcgr4 and CD163 mRNAs were significantly enhanced compared with those in WT RPMs. The expression levels of Fcgr1, Fcgr2b, and Fcgr3 genes tended to increase in neudesin KO RPMs without significance. Hmox1 and Slc40a1, which code for heme oxygenase 1 and iron transporter Ferroportin-1, respectively, were also upregulated in neudesin KO RPMs, suggesting the possibility of enhanced heme catabolism and export after digestion of RBCs (Fig. 4d). Spic, which controls the development of red pulp macrophages22, was also increased in neudesin KO RPMs (Fig. 4d).
a The mean fluorescence intensity (MFI) of surface receptors on the gated red pulp macrophages (RPM) from 8-week-old WT and neudesin KO male mice was measured by flow cytometry (n ≧ 6, F4/80: P = 0.9388, CD11b: P = 0.9629, VCAM1: P = 0.6770, Stabilin-2: P = 0.1617, Tim-4: P = 0.7969, Axl: P = 0.6595, FcγR1: P = 0.0370, FcγR2/3: P = 0.0356, FcγR4: P < 0.0001, SIRPα: P = 0.6267, CD163: P = 0.0064, and CD206: P = 0.3749 by unpaired t test) b Representative histograms of RPMs treated with anti-FcγR1 (left panel) and anti-CD163 (right panel) from 8-week-old WT (white) and neudesin KO (gray) male mice. c RPMs from 8-week-old WT and KO male mice were isolated with magnetic bead-conjugated anti-F4/80 antibody, and relative mRNA expression levels of genes encoding macrophage receptors were quantified by RT-qPCR (n = 5 each groups, Adgre1: P = 0.3676, Fcgr1: P = 0.2295, Fcgr2b: P = 0.0848, Fcgr3: P = 0.1061, Fcgr4: P = 0.0396, and Cd163: P = 0.0421 by unpaired t-test). d Relative mRNA expression levels of genes regulating iron recycling were quantified by RT-qPCR with isolated RPMs from 8-week-old WT and KO male mice (n = 5 each groups, Hmox1: P = 0.0465, Slc40a1: P = 0.0260, and Spic: P = 0.014 by unpaired t-test). Data are shown as the mean ± SD of four or more experiments. Each symbol represents an individual mouse. *P < 0.05 and **P < 0.01, and ***P < 0.001 by unpaired t-test.
Neudesin suppresses erythrophagocytosis and FcγRs expression through ERK activation in bone marrow-derived macrophages
To examine whether the suppressive effects of neudesin on phagocytosis are restricted during erythrophagocytosis by RPMs, the phagocytic activities of bone marrow-derived macrophages (BMDMs) were determined (Fig. 5a–c). To induce RPM-like cells, we used granulocyte-macrophage colony-stimulating factor (GM-CSF) and hemin-stimulated BMDMs because these cells have properties similar to those of RPMs23,24,25. Bone marrow cells from WT and neudesin KO mice were cultured in vitro in the presence of macrophage colony-stimulating (M-CSF) factor following additional stimulation with GM-CSF and hemin. The differentiation efficiency of neudesin KO bone marrow cells into F4/80+ macrophages was comparable to that of WT cells (Fig. 5b). Consistent with the in vivo results, neudesin KO BMDMs engulfed PKH26+ RBCs more frequently than did WT BMDMs (Fig. 5c). However, WT and neudesin KO BMDMs phagocytosed apoptotic thymocytes and killed E. coli to a similar extent (Fig. 5c). Treatment with recombinant neudesin protein inhibited the enhanced erythrophagocytosis of neudesin KO BMDMs (Fig. 5d). These data suggest that neudesin negatively regulates phagocytosis with specificity toward erythrocytes. The slight upregulation of FcγR1 and Fcγ2/3 were also observed on the surface of neudesin KO BMDMs, and suppressed by the addition of recombinant neudesin protein into the culture medium (Fig. 5e). To investigate the intracellular signaling pathway mediating the function of neudesin in BMDMs, phosphorylation of ERK1/2, AKT, and CREB, which is promoted by neudesin in neural precursor cells9, were analyzed by western blotting. Phosphorylation of ERK1/2 was decreased in neudesin KO BMDMs, and upregulated by recombinant neudesin (Fig. 5f, g). AKT and CREB pathways were unchanged between WT and neudesin KO BMDMs and after treatment with recombinant neudesin protein. The MEK inhibitor U0126 enhanced erythrophagocytosis and surface expression of FcγR1 in WT BMDMs (Fig. 5h, i), suggesting that neudesin exerts the inhibitory effect on BMDMs through activation of ERK1/2.
a Scheme of experiments to measure phagocytosis of bone marrow-derived macrophages (BMDMs) in vitro. At 1 h after adding fluorescent baits, cells were collected and analyzed by flow cytometry to detect phagocytosis by BMDMs (n = 9). b Percentages of induced F4/80+ macrophages from WT and neudesin KO bone marrow cells (n = 9, P = 0.5997 by unpaired t-test). c Percentage of phagocytosis by BMDMs against RBCs, apoptotic thymocytes, and killed E. coli. *P < 0.05 by unpaired t-test (n ≧ 8, RBCs: P = 0.0164, apoptotic thymocytes: P = 0.2423, and E. coli: P = 0.3560 by unpaired t-test). d Pre-treatment with recombinant neudesin (100 ng/mL) for 2 days into the culture medium inhibited enhanced erythrophagocytosis by neudesin KO BMDMs (n = 6 each group, Control:WT vs. Control:KO: P = 0.0231, Control:WT vs. + neudesin:WT: P = 0.9998, Control:WT vs. + neudesin:KO: P = 0.9994, Control:KO vs. + neudesin:WT: P = 0.0192, Control:KO vs. + neudesin:KO: P = 0.0297, and + neudesin:WT vs. + neudesin:KO: P = 0.997 by a 2-way analysis of variance followed by Sidak’s post-test). e Mean fluorescence intensity (MFI) of surface macrophage receptors on BMDMs induced from WT and neudesin KO bone marrow cells were measured by flow cytometry. The effects of pre-treatment with recombinant neudesin (100 ng/mL) were also investigated (n = 6 each group, Fcgr1; Control:WT vs. Control:KO: P = 0.0001, Control:WT vs. + neudesin:WT: P = 0.371, Control:WT vs. + neudesin:KO: P = 0.972, Control:KO vs. + neudesin:WT: P < 0.0001, Control:KO vs. + neudesin:KO: P = 0.0003, and + neudesin:WT vs. + neudesin:KO: P = 0.19, Fcgr2/3; Control:WT vs. Control:KO: P = 0.0234, Control:WT vs. + neudesin:WT: P < 0.0001, Control:WT vs. + neudesin:KO: P = 0.0014, Control:KO vs. + neudesin:WT: P < 0.0001, Control:KO vs. + neudesin:KO: P < 0.0001, and + neudesin:WT vs. + neudesin:KO: P = 0.1137, and Fcgr4; Control:WT vs. Control:KO: P = 0.9866, Control:WT vs. + neudesin:WT: P = 0.3411, Control:WT vs. + neudesin:KO: P = 0.9993, Control:KO vs. + neudesin:WT: P = 0.2027, Control:KO vs. + neudesin:KO: P = 0.9966, and + neudesin:WT vs. + neudesin:KO: P = 0.2835 by a 2-way analysis of variance followed by Sidak’s post-test). f Cell lysates from WT and neudesin KO BMDMs were extracted 30 min after incubation with recombinant neudesin (100 ng/mL). Phosphorylated ERK1/2, ERK1/2, phosphorylated AKT, AKT, phosphorylated CREB, CREB, and β-Actin were detected by western blotting. g Relative expression of p-ERK/ERK, p-AKT/AKT, and p-CREB/CREB was determined (n = 4, p-ERK/ERK Control: P = 0.0435, p-ERK/ERK +neudesin: P = 0.3870, p-AKT/AKT Control: P = 0.8851, p-AKT/ AKT +neudesin: P = 0.3878, p-CREB/CREB Control: P = 0.6681, p-CREB/CREB +neudesin: P = 0.8863 by a 2-way analysis of variance followed by Sidak’s post-test)). h Pre-treatment with MEK inhibitor U0126 (100 nM) for 2 days into the culture medium enhanced erythrophagocytosis by WT, but not neudesin KO BMDMs (n = 6 each group, Control:WT vs. Control:KO: P = 0.0457, Control:WT vs. + U0216:WT: P = 0.0238, Control:WT vs. + U0216:KO: P = 0.0336, Control:KO vs. + U0216:WT: P = 0.9894, Control:KO vs. + U0216:KO: P = 0.9988, + U0216:WT vs. + U0216:KO: P = 0.9984 by a 2-way analysis of variance followed by Sidak’s post-test). i Pre-treatment with MEK inhibitor U0126 (100 nM) for 2 days into the culture medium enhanced surface expression of FcγR1 on WT, but not neudesin KO BMDMs. (n = 6 each group, Control:WT vs. Control:KO: P = 0.0019, Control:WT vs. + U0216:WT: P = 0.0034, Control:WT vs. + U0216:KO: P = 0.0004, Control:KO vs. + U0216:WT: P > 0.9999, Control:KO vs. + U0216:KO: P = 0.9897, + U0216:WT vs. + U0216:KO: P = 0.9446 by a 2-way analysis of variance followed by Sidak’s post-test)*P < 0.05, and ***P < 0.001 by a 2-way analysis of variance followed by Sidak’s post-test. Each symbol represents an individual mouse. Data are shown as means ± SD of three or more experiments.
Mature neudesin-deficient mice exhibit normocytic anemia
To investigate whether impaired iron recycling at a young age affects the later metabolism of erythrocytes, we examined peripheral RBCs and splenocytes of mature WT and neudesin KO mice at 24 weeks of age (Fig. 6). Mature neudesin KO mice exhibited significantly decreased RBC counts, hemoglobin concentrations, and hematocrit levels compared with WT mice, whereas mean corpuscular volume (MCV), MCH, and mean corpuscular hemoglobin concentration (MCHC) were similar (Fig. 6a). Serum concentrations of erythropoietin and bilirubin were unaltered in 24-week-old WT and neudesin KO mice (Fig. 6c). The increase in spleen weight and the enhanced expression of FcγR4 on RPMs were preserved in 24-week-old neudesin KO mice (Fig. 6d, g). However, the expression of FcγR1 increased compared with that at 8 weeks of age irrespective of the genotype, with little difference observed between 24-week-old WT and KO mice (Fig. 6g). These data suggest that abnormal iron recycling in youth develops into severe anemia with aging in neudesin KO mice.
a, b Blood cell counts of 24-week-old WT and neudesin KO male mice. RBC, red blood cell count (WT: n = 16, KO: n = 15, P = 0.0193); Hb, hemoglobin (WT: n = 13, KO: n = 12, P = 0.0055); Ht, hematocrit (WT: n = 13, KO: n = 12, P = 0.0016); MCV, mean corpuscular volume (WT: n = 13, KO: n = 12, P = 0.6175); MCH, mean corpuscular hemoglobin (WT: n = 13, KO: n = 12, P = 0.5218); MCHC, mean corpuscular hemoglobin concentration (WT: n = 13, KO: n = 12, P = 0.4683); WBC, white blood cell count (WT: n = 6, KO: n = 5, P = 0.4938); PLT, platelet count (WT: n = 13, KO: n = 12, P = 0.3715). c Serum concentration of erythropoietin (Epo, WT: n = 12, KO: n = 13, P = 0.2198) and total bilirubin (WT: n = 11, KO: n = 9, P = 0.6419) from 24-week-old WT and neudesin KO male mice. d Spleens of neudesin KO male mice were significantly heavier than those of 24-weeks-old WT male mice (WT: n = 13, KO: n = 12, P = 0.0494). e Splenocyte suspensions from 24-week-old WT and neudesin KO male mice were assessed by flow cytometric analysis, and cell numbers of TER119+ red blood cells (WT: n = 13, KO: n = 12, P = 0.2580) and CD45+ white blood cells (WT: n = 13, KO: n = 12, P = 0.3981) in the spleen were determined. f Chart showing the number of RPMs from 24-week-old WT and neudesin KO male mice (WT: n = 13, KO: n = 12, P = 0.4752). g Mean fluorescence intensity (MFI) of surface macrophage receptors on RPMs from 24-week-old WT and neudesin KO male mice was measured by flow cytometry (WT: n = 13, KO: n = 12, FcγR1: P = 0.6545, FcγR2/3: P = 0.3510, FcγR4: P = 0.0027). *P < 0.05, **P < 0.05, and ***P < 0.001 by unpaired t-test. Each symbol represents an individual mouse. Data are shown as means ± SD of two or more experiments.
Discussion
Neudesin is a secretory heme-binding protein that was initially identified as a neurotrophic factor and was thereafter considered as a regulator of obesity development and energy expenditure11. In this study, we additionally find that neudesin plays a role in erythrocyte homeostasis by regulating splenic erythrophagocytosis.
Neudesin KO mice (8 weeks old) displayed mild splenomegaly with increased accumulation of RBCs without obvious anemia or polycythemia, and their peripheral RBCs were relatively tolerant to osmotic stress and sugar depletion. These data implied that senescent RBCs were removed at an earlier aging stage owing to accelerated turnover of RBCs in the spleens of neudesin KO mice. Concordantly, in vivo biotinylation assays demonstrated the shortened lifespan of RBCs, and cross-transfusion data excluded the possibility of faster RBC aging in KO mice. Splenectomy experiments revealed that the shortened lifespan of RBCs was caused by abnormalities in the spleen of neudesin KO mice. Unexpectedly, the lifespan of RBCs in splenectomized WT mice was similar to that of sham WT mice, even though the spleen is mainly responsible for removing aged RBCs. When stressed RBCs are transfused, macrophages transiently accumulate in the liver and remove stressed red blood cells as needed26. A similar compensatory phenomenon may be occurring in the livers of splenectomized mice.
Nevertheless, despite increased RBC clearance, we could not detect the compensatory enhancement of erythropoiesis in the bone marrow or spleen of neudesin KO mice. The ratios of TER119+CD71+ erythroblasts in the bone marrow and spleen and the concentrations of serum erythropoietin were unaltered in WT and neudesin KO mice (Supplementary Fig. 1). Extramedullary hematopoiesis might occur independently of erythropoietin in non-splenic tissues in neudesin KO mice.
It is considered that the phagocytic ability of RPMs was increased mainly by upregulated surface expression of FcγRs in neudesin KO mice. CD163 molecules, which are endocytic receptors for hemoglobin-haptoglobin complexes, were also upregulated in neudesin KO RPMs. Previously, CD163 has been shown to mediate cell-cell interactions between macrophages and developing erythroblasts in erythroblastic islands and function as an innate immune sensor for bacteria21,27. It is possible that CD163 is also involved in the adhesion with RBCs and/or recognition of senescent RBCs by RPMs, and that upregulation of CD163 on neudesin KO RPM might contribute to the accumulation of RBCs and upregulation of erythrophagocytosis in the spleen.
The transcription factor Spi-C plays an essential role in heme-dependent differentiation from monocytes to RPMs3,22. Spic KO mice showed decreased erythrophagocytosis by splenic macrophages and increased splenic iron stores. In contrast, in addition to increased erythrophagocytosis (Fig. 3f) and decreased splenic iron store (Fig. 3h, i), neudesin KO mice showed upregulated expression of Spic mRNA in RPMs (Fig. 4d). It is possible that the upregulation of Spic in neudesin KO RPMs, which might enhance the properties of RPMs as iron-recycling macrophages, causes upregulation of erythrophagocytosis and abnormal iron homeostasis.
Neudesin is a secretory protein expressed in many tissues other than the spleen such as the nervous system and adipose tissues. We previously reported that neudesin functions as a negative regulator of sympathetic activity and affects heat production and fatty acid oxidation in brown adipose tissue and enhanced lipolysis in white adipose tissue16. Thus, there remains a possibility that the splenic phenotypes of neudesin KO mice are secondary effects from other tissues on the spleen and macrophages. However, as shown in Fig. 5e, BMDMs from neudesin KO, cultured independently of other tissues, exhibited upregulation of FcγRs, and recombinant neudesin protein counteracted the upregulation. Furthermore, to investigate whether neudesin proteins can act on the spleen in an endocrine manner from other tissues, we examined intravenous administration of recombinant neudesin proteins. A single dose or three consecutive days administration of neudesin proteins at least did not rescue the splenic phenotypes of neudesin KO such as increased splenic cell number and increased expressions of FcγRs on RPMs (Supplementary Figs. 2 and 3). These data suggest that, at least in the regulation of FcγRs, neudesin acts in an autocrine/paracrine manner within the spleen. Further studies using neudesin conditional KO mice will reveal more details about the functions of neudesin.
Our study demonstrated that neudesin, which has been identified as a heme-binding secretory factor, suppressed erythrophagocytosis of RPMs and BMDMs. The differences between WT and neudesin KO mice were small at 8 weeks of age under normal conditions, but the small differences led to delayed recovery from drug-induced anemia and subsequent anemia in mature adulthood. The receptor for neudesin and the physiological significance of its affinity for heme remain unrevealed. Future studies should focus on further characterizing neudesin.
Methods
Mice
All Animal studies were conducted in accordance with the International Guiding Principles for Biomedical Research Involving Animals and approved by the Animal Research Committee of Kobe Pharmaceutical University (approval number 2018-044, 2019-028, 2020-059, 2021-061, 2022-034, 2023-043), and the care of the animals was implemented following the Animal Research: Reporting of in Vivo Experiments guidelines. We have complied with all relevant ethical regulations for animal use. Neudesin knockout (KO) mice were reported previously16. Mice were housed in a temperature-controlled pathogen-free animal facility with a 12-h light and dark cycle with free access to water and food. For in vivo study, 8- and 24-week-old male WT and KO mice in the C57BL/6N background were used.
Quantitative RT-PCR
RNA was extracted using the Sepasol-RNA I Super G (Nacalai tesque, Kyoto, Japan). Up to 1 µg of RNA was used for reverse transcription into cDNA and Real-time quantitative RT-PCR was performed using the THUNDERBIRD Probe qPCR/RT Set (TOYOBO, Osaka, Japan). The 18 S ribosomal RNA level was used as an internal control. The PCR primers used are listed in Supplementary Table 1.
Flow cytometry
Splenocytes were prepared by applying pressure to the spleen using the head of a syringe and collected through a 70-μm cell strainer. Splenocytes and blood cells were incubated with fluorochrome-conjugated antibodies and other reagents, and immunofluorescent cells were analyzed and sorted using FACSAriaIII and FlowJo software (BD Biosciences, Franklin Lakes, NJ, USA). Antibodies and reagents used are listed in Supplementary Table 2, and gating strategies are shown in Supplementary Fig. 4.
Blood cell count
Peripheral blood samples were harvested in EDTA-coated microtubes (Health Wave Japan, Tokyo, Japan) and analyzed with an automatic blood cell counter PCE-310 (Erma Inc., Tokyo, Japan) using the mouse species program.
Hemolysis test
Erythrocytes (0.4% hematocrit) were incubated at 37 °C for 16 h in Ringer solution containing 125 mM NaCl, 5 mM KCl, 1 mM MgSO4, 32 mM HEPES, 5 mM glucose, and 1 mM CaCl2, pH = 7.4, 305 mOsm. Hypertonic solution was obtained by adding 400 mM sucrose to the Ringer solution. In studies on glucose depletion, glucose was removed from the Ringer solution. After incubation, the samples were centrifuged for 4 min at 400 × g, 4 °C, and the supernatants were harvested. To quantify hemolysis, the hemoglobin concentration of the supernatant was determined photometrically at 405 nm. The absorption of the supernatant of erythrocytes lysed in distilled water was defined as 100% hemolysis.
RBC biotinylation, transfusion, and splenectomy
In vivo biotinylation and transfusion of RBCs followed by flow cytometry were performed at 8 weeks of age. Splenectomy was performed at 4 weeks of age, followed by in vivo biotinylation at 8 weeks of age. RBCs were labeled in vivo by intravenous injection of 30 mg/kg sulfo-NHS-biotin (Thermo Fisher Scientific). Small samples of peripheral blood were collected from the tail vein 3 days after biotin labeling and stained using Alexa Fluor 594-conjugated streptavidin. Blood samples were analyzed at 7-day intervals to quantify biotin-labeled cells in the circulation. For transfusions of biotinylated RBC, donor mice received the biotinylating reagent. Two days after injection, mice were euthanized via isoflurane overdose and promptly exsanguinated by cardiocentesis. Whole blood was collected into sterile tubes in a 1:7 mixture with anti-coagulant citrate phosphate dextrose adenine solution. After washing with PBS, 100 μL of diluted blood (1 × 109 cells per mouse) was administered to the recipient mice via retro-orbital injection under anesthesia.
Acute anemia model with phenylhydrazine
Mice were intraperitoneally administered 2 mg/mouse phenylhydrazine (Sigma-Aldrich, St. Louis, MO, USA) dissolved in 200 μL PBS. At 2, 4, and 6 days after injection, peripheral blood was collected.
Histological analysis
Spleens were fixed in 4% paraformaldehyde in PBS for 24 h and then embedded in paraffin blocks. Paraffin sections of the spleen were prepared using a rotary microtome (RM2135, Leica, USA) and Prussian blue iron staining was performed. Iron Stain Kit (Prussian Blue Stain) (Abcam, ab150674) was used by following manufacturer’s procedure. Autofluorescence of RPM was observed using an IX70 inverted fluorescent microscope (Olympus, Tokyo, Japan).
In vivo and in vitro phagocytosis assays
RBCs were labeled with PKH26, washed three times with PBS, and resuspended at a density of 3 × 107 cells in 100 μL in PBS. Mice were injected through the retro-orbital venous sinus with PKH26-labeled RBCs. Splenocytes were collected 5 days after injection, and stained with fluorescence-conjugated anti-F4/80 and anti-CD11b mAb. RPMs that had ingested PKH26-labeled RBCs were detected by flow cytometry. For in vitro phagocytosis assays, PKH26-labeled RBCs, PKH26-labeled apoptotic thymocytes prepared by treatment with 10 μM dexamethasone at 37°C for 4 h after PKH26-labeling, or TRITC-labeled killed E. coli BioParticles (Sigma) were added to BMDMs at the cell number ratio of 10:1, 10:1, and 100:1, respectively. After incubation for 1 h, cells were collected, hemolyzed, and stained with fluorescence-conjugated anti-F4/80 mAb. F4/80+ BMDMs that had ingested fluorescent cells were detected by flow cytometry.
Tissue and serum iron levels
Serum and tissue iron concentrations were measured using a commercially available colorimetric iron quantification kit (BioAssay Systems, Hayward, CA, USA) according to the manufacturer’s instructions.
Isolation of RPMs
RPMs were isolated from 1 × 108 splenocytes by positive selection with anti-F4/80 microbeads, MACS columns, and MACS separators according to the manufacturer’s protocols (Miltenyi Biotech, Bergisch Gladbach, Germany).
Cell culture of BMDMs
Bone marrow cells (2 × 107 cells/mL) were cultured in complete RPMI (RPMI-1640 medium supplemented with 10% FBS, 0.03 mg/mL L-glutamine, 100 units/mL penicillin, and 100 mg/mL streptomycin) supplemented with murine M-CSF (25 ng/mL) for 7 days, followed by stimulation with complete RPMI supplemented with 20 ng/mL murine GM-CSF and 4 μM hemin for 2 days25.
Western blotting
BMDM were homogenized by sonication for 30 s in ice-cold RIPA buffer with a protease and phosphatase inhibitor (Nacalai tesque, Kyoto, Japan), and centrifuged for 10 min at 14,000 × g. In total, 5 μg of total protein from cell lysate was run on an 12.5% sodium dodecyl sulfate-polyacrylamide gel electrophoresis. After electrophoresis, proteins were transferred to a methanol-hydrated PVDF (polyvinylidene fluoride) membrane for 90 min and blocked with Blocking One (Nacalai tesque) for 30 min at RT. Membranes were probed overnight at 4 °C with a primary antibody. After overnight incubation, membranes were washed three times with PBST for 5 min and then incubated for 1 h at RT with an anti-rabbit IgG horseradish-peroxidase-conjugated secondary antibody. Membranes were then incubated in Chemi-Lumi One reagent (Nacalai tesque) at RT and then imaged using ImageQuant LAS 4000 (FUJIFILM, Tokyo, Japan). Protein loading was confirmed by probing for β-actin. The quantification of the western blots was performed by densitometry using ImageJ10. Expression levels were determined with ImageJ software. The antibodies used are listed in Supplementary Table 2.
Statistical analysis and reproducibility
Data were analyzed using Prism software (GraphPad Software). Data are expressed as the mean ± SD. Unpaired t test and 2-way analysis of variance followed by Sidak’s post-test were used as described in the figure legends. All p values are two-sided, and p values less than 0.05 were considered statistically significant. Sample sizes and number of replicates are described in the figure legends.
Reporting summary
Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.
Data availability
Source data for the graphs are available as Supplementary Data file.
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Acknowledgements
The authors would like to thank members of the Department of Microbial Chemistry for insightful discussions and support. We thank Editage (www.editage.com) for English language editing. This work was supported by a MEXT KAKENHI Grant-in-Aid for Scientific Research (C) Grant Number 20K07060 (to Y.N.) from the Ministry of Education, Culture, Sports, Science, and Technology (MEXT).
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Y.N., Y.M., T. Mukae, T. Mikami, R.S., and N.K. performed the experiments, Y.N. performed data analysis, Y.N. wrote the manuscript, and H.K., N.I., and M.K. supervised the study.
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Nakayama, Y., Masuda, Y., Mukae, T. et al. A secretory protein neudesin regulates splenic red pulp macrophages in erythrophagocytosis and iron recycling. Commun Biol 7, 129 (2024). https://doi.org/10.1038/s42003-024-05802-9
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DOI: https://doi.org/10.1038/s42003-024-05802-9
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