FGF-23 Is a Negative Regulator of Prenatal and Postnatal Erythropoiesis^*

Animals

Heterozygous Fgf-23 and 1α(OH)ase mice (both in C57BL/6 background) were interbred at 6–12 weeks to attain wild-type, Fgf-23^−/−, Fgf-23^−/−/1α(OH)ase^−/−, and 1α(OH)ase^−/− animals. Heterozygous 1α(OH)ase^−/− mice were generously provided by Dr. René St-Arnaud (Genetics Unit of Shriners Hospital, Montreal, Canada). All mice were housed in New York University College of Dentistry Animal Facility, kept on a light/dark (12h/12h) cycle at 23 °C, and received food (standard lab chow) and water ad libitum. Genomic DNA was obtained from tail snips, and routine PCR was used to identify the genotypes as previously described (20, 21). All animal studies were approved by the institutional animal care and use committee at New York University.

RNA Isolation

Total RNA was extracted from crushed whole tibiae, bone marrow, and kidney from 6-week-old mice using TRIzol reagent (Ambion; Invitrogen) according to the manufacturer's protocol (Molecular Research Center Inc., Cincinnati, OH). Synthesis of cDNA was performed using a high capacity cDNA reverse transcription kit as described by the manufacturer (Applied Biosystems, Foster City, CA). cDNA was amplified by quantitative real time PCR using the PerfecCTa® SYBR® Green SuperMix (Quanta Biosciences, Gaithersburg, MD). The following primers were used: Fgf-23, 5′-ACT TGG CCT TTA TTA GCC GGG TCT-3′ and 5′-AGA TGG CCT CCT CCC TGT GTT CAA-3′; FGFR1, 5′-TGA TGG GAG AGT CCG ATA GAG-3′ and 5′-CCT GAA GAC TGC TGG AGT TAA T-3′; FGFR2, 5′-CCA GCA CTG GAG CCT TAT TAT-3′ and 5′-GTG GTT GAT GGA CCC GTA TT-3′; FGFR3, 5′-CTA AAT GCC TCC CAC GAA GAT-5′ and 5′-CTG AGG ATG GAG CAT CTG TTA C-3′; FGFR4, 5′-CCT GAG GCC AGA TAC ACA GAT A-3′ and 5′-GGA TGA CTT GCC GAT GAT ACA C-3′; klotho, 5′-AAA TGG CTG GTT TGT CTG GGG-3′ and 5′-TAT GCC ACT CGA AAC CGT CCA-3′; erythropoietin (Epo), 5′-TCT ACG TAG CCT CAC TTC ACT-3′ and 5′-ACC CGG AAG AGC TTG CAG AAA-3′; hypoxia-induced factor 1α (HIF-1α), 5′-TCT CGG CGA AGC AAA GAG TCT-3′ and 5′-TAG ACC ACC GGC ATC CAG AAG-3′; hypoxia-induced factor 2-α (HIF-2α), 5′-GGG AAC ACT ACA CCC AGT GC-3′ and 5′-TCT TCA AGG GAT TCT CCA AGG-3′; transferrin, 5′-CCC TCT GTG ACC TGT GTA TTG-3′ and 5-CTT TCT CAA CGA GAC ACC TGA A-3′; transferrin receptor, 5′-TCC TGT CGC CCT ATG TAT CT-3′ and 5′-CGA AGC TTC AAG TTC TCC ACT A-3′; glucose transporter-1 (glut-1), 5′-CCC AGG TGT TTG GCT TAG A-3′ and 5′-CAG AAG GGC AAC AGG ATA CA-3′; phosphoglycerate kinase-1 (Pgk-1), 5′-CAC AGA AGG CTG GTG GAT AT-3′ and 5′-CTT TAG CGC CTC CCA AGA TAG-3′; and hypoxanthine-guanine phosphoribosyl transferase (HPRT-housekeeping gene), 5′-AAG CCT AAG ATG AGC GCA AG-3′ and 5′-TTA CTA GGC AGA TGG CCA CA-3′. Forty cycles (95 °C for 15 s; 60 °C for 30 s; and 72 °C for 30 s) were run on a Mastercycle Realplex² (Eppendorf, Hamburg, Germany), and reactions were analyzed. Melting curve and gel analyses (sizing and sequencing) verified single products of the appropriate base pair size.

Blood Collection and Hematologic Analysis

Peripheral blood was collected posteuthanasia from 6-week-old mice by cardiac puncture into EDTA-coated tubes to prevent clotting. Blood samples were then shipped overnight to Cornell University Veterinary Clinic for automated complete blood count.

Fetal Liver Collection

Time breeding of Fgf-23^+/− mice was carried out. Pregnant females were sacrificed at E15.5, and embryos were obtained by C-section. Genomic DNA was obtained from tail snips, and routine PCR was used to identify the genotypes of the embryos as previously described (20, 21). Fetal livers were isolated and fetal liver cell suspensions were prepared from WT and Fgf-23^−/− mice.

Isolation and Assessment of Blood, Bone Marrow, Spleen, and Liver Cells by Flow Cytometry

Bone marrow was isolated from dissected tibiae and femora from 6-week-old mice by flushing in Iscove's modified Dulbecco's medium (IMDM) (Sigma-Aldrich) supplemented with 20% fetal bovine serum (20% IMDM) through a 26-gauge Becton Dickinson needle. Marrow cells were dispersed by manual agitation and then filtered to remove foreign particles. Spleens from 6-week-old mice and fetal livers from E15.5 embryos were surgically removed and homogenized into a cell suspension in 20% IMDM. Flow cytometry analysis for peripheral blood, bone marrow, spleen, and fetal liver cells were carried out in a BD FACSort^TM flow cytometer equipped with 488 argon lasers (BD Biosciences, San Jose, CA). For immunostaining, cells were washed and resuspended in 1× PBS containing 0.1% BSA. Mouse Fc receptor was blocked prior to staining using CD16/32 antibody to reduce nonspecific binding. After the addition of antibodies, cells were incubated for 40 min on ice; for peripheral blood, red blood cells were further lysed using BD FACS lysing solution (BD Biosciences). Labeled cells were then washed with 1× PBS and analyzed by flow cytometry. Appropriate isotype controls were kept for each set. Forward and side scatter patterns were gated, excluding the debris. A total of 20,000 events were collected and analyzed using FlowJo software version 7.6.5. Erythroid lineage was assessed using Ter119 APC/CD71 PE markers combined with the forward scatter properties (26). Hematopoietic stem/progenitor cells were differentiated using SLAM markers (CD150 PE/CD48 APC), Sca-1 FITC (Ly6A-E), c-Kit Percp Cy5.5 (CD117), CD90 PE (Thy-1), and APC-tagged lineage mixture comprising of antibodies against CD3, B220 (CD45R), Ly6G and Ly6C (Gr-1), CD11b (Mac-1), and TER119. c-Kit⁺Sca1⁺ cells were gated on lineage negative fraction to analyze LSK (Lin⁻c-Kit⁺Sca1⁺). The LSK cells were then analyzed on a Thy-1^low gate to obtain KTLS population (LSK Thy^low). CD45.1 PE and CD45.2 FITC antibodies were used to differentiate donor and recipient populations after transplantation. All antibodies except SLAM markers were purchased from BD Pharmingen. SLAM markers CD150 and CD48 were purchased from e-Biosciences (San Diego, CA). Cell apoptosis was assessed using the TACS annexin V kit (Trevigen Inc., Gaithersburg, MD).

Methylcellulose Cultures

Cell suspensions were prepared from 6-week-old bone marrow, spleen, and fetal liver (E15.5) in 20% IMDM. Aliquots were then plated in a methylcellulose medium supplemented with recombinant cytokines for colony assays of murine cells (Methocult M3434; StemCell Technologies, Vancouver, Canada). Cytokines included: IL-3 (4 ng/10 μl), stem cell factor (20 ng/10 μl), granulocyte macrophage-colony stimulating factor (2 ng/10 μl), and erythropoietin (2 units/10 μl). Fetal liver and bone marrow cells were plated at a concentration of 1 × 10⁴, whereas spleen cells were plated at a concentration of 1 × 10⁵ cells/35-mm tissue culture dish. Cells were incubated in a humidified chamber at 37 °C with 5% CO₂. Three different types of colonies were scored 8 and 12 days postculture including: burst-forming unit of erythroid cells (BFU-E) and the most primitive colony-forming units of granulocyte-erythrocyte-macrophage-megakaryocyte cells (CFU-GEMM).

Serum FGF-23 and Epo Measurements

Serum FGF-23 and erythropoietin (Epo) concentrations were measured in samples from 6-week-old mice using the mouse FGF-23 C-terminal ELISA kit (Immutopics International, San Clemente, CA) and the Quantikine rat/mouse Epo immunoassay kit (R&D Systems, Minneapolis, MN), according to the manufacturer's protocols.

FGF-23 Protein Injections

Wild-type C57BL/6 mice (6–8 weeks old) or 1α(OH)ase^−/− and WT littermates were given a single injection intraperitoneally of recombinant human FGF-23 protein (5 μg) (R&D Systems) or vehicle (PBS) and analyzed 24 h later for hematopoietic cellularity in peripheral blood, bone marrow, and spleens as well as for hematology, as described above.

Oxygen Treatment

Fgf-23^−/− mice and WT littermates were placed in a sealed Plexiglass chamber where oxygen was administered through a tube at a rate of 4 liters/min for 1 h. The room and chamber temperatures were maintained at ∼20 °C throughout the experiment. Mice were not anesthetized or restrained, and food and water were available ad libitum during the entire treatment.

Adhesion Assay

Bone marrow cell adherence was determined using 96-well plates coated with 5 μg of fibronectin (Sigma) overnight. Whole bone marrow cells from 6-week-old WT and Fgf-23^−/− mice were plated at a density of 10⁵ cells in 100 μl of 2% IMDM, seeded in triplicate, and incubated for 40 min at 37 °C. Cells were then fixed in 4% paraformaldehyde (Sigma), stained using 0.25% crystal violet (Sigma), and lysed with 0.1% Triton X-100 (Sigma) in 1× PBS. The plates were read at an absorbance of 550 nm, where readings represented that higher optical density values corresponded to a higher adhesion.

In Vitro Transmigration Assay

Chemotaxis toward stromal-derived factor 1 (SDF-1α) was assessed using a dual chamber Transwell with a pore size insert of 8 μm. 10⁵ whole bone marrow cells from 6-week-old WT and Fgf-23^−/− mice in 100 μl of 2% IMDM were seeded in triplicate to the upper chamber of 24-well plates containing inserts (BD Falcon). 100 ng of SDF-1α (Peprotech, Rocky Hill, NJ) was added to the lower chamber containing 600 μl of medium. Bone marrow cells were incubated for 3 h at 37 °C, 5% C0₂ in the presence or absence (to detect spontaneous migration) of SDF-1α. Afterward, the nonmigrated cells remaining in the Transwells and the migrated fraction were both collected and counted. Migrated cells were expressed as a percentage of migrated cells per total cells seeded.

In Vivo Homing

Homing experiments were performed using tail vein injection of 2.5 × 10⁶ whole bone marrow cells from 6-week-old wild-type or Fgf-23^−/− (CD45.2; Ly5.2) mice into 8-week-old myeloablated B6.SJL (CD45.1; Ly5.1) recipient mice. Myeloablation was achieved by a lethal dose of irradiation (900 rads). Recipient mice were sacrificed 24 h post-transplantation, and peripheral blood, spleen, and bone marrow cells were collected. Flow cytometry was performed on cell suspensions from each tissue collected and analyzed for the presence of CD45.2⁺ donor cells to determine the numbers of homed cells to the BM in comparison with spleen and peripheral blood. A minimum of 50,000 events were acquired in a BD FACSort ^TM flow cytometer equipped with 488 argon lasers (BD Biosciences).

In Vitro Ter119+ Cell Isolation

Bone marrow was isolated from dissected tibiae and femora from 6-week-old mice by flushing in IMDM (Sigma-Aldrich) supplemented with 20% fetal bovine serum (20% IMDM) through a 26-gauge Becton Dickinson needle. Marrow cells were dispersed by manual agitation and then filtered to remove foreign particles. These cells were sorted using Ter119-conjugated magnetic beads (Miltenyi Biotec) and stained with CD71^PE and Ter119^FITC antibodies (BD Biosciences) to obtain early to late stage erythroblast populations using flow cytometry. Purified Ter119+ cells were seeded in triplicate in 24-well plates for 24 h at a cell density of 10⁵ cells/well and then treated for 4 h with or without 5 ng/ml of recombinant FGF-23 (R&D Systems).

Statistics

Statistical significance was evaluated by Student's t test for comparison between two groups or by one-way analysis of variance followed by Tukey's test for multiple group comparisons. All analyses were performed using GraphPad Prism 4.0, and all values were expressed at means ± S.E. p values less than 0.05 were considered significant.

Expression of Fgf-23 and Its Signaling Components in Erythroid Cells

High expression of Fgf-23 in bone has been reported by several groups, confirming that bone is the principal source of Fgf-23 production (20, 27, 28). However, expression of Fgf-23 in specific bone marrow cells remains unknown. Here, we determined mRNA expression of Fgf-23 and several fgf-23 signaling components (klotho and FGFR1–4) in isolated BM erythroid cells (Ter119⁺) of adult WT mice. Real time quantitative RT-PCR revealed that WT Ter119+ erythroid cells highly express Fgf-23, klotho, and FGFR1, 2, and 4 but showed minimal FGFR3 expression (Fig. 1). These data suggest that erythroid cells are capable of undergoing active Fgf-23 signaling.

Analysis of the Fgf-23^−/− Mice Hematologic Characteristics

To assess the impact of FGF-23 on erythropoiesis, we performed complete blood count analysis in peripheral blood (PB) of 6-week-old WT, heterozygous (Fgf-23^+/−), and Fgf-23 null (Fgf-23^−/−) mice. Fgf-23^−/− mice were found to exhibit significantly elevated erythrocyte (RBC) numbers in comparison with WT and Fgf-23^+/− mice (Fig. 2A). In addition, a significant increase in RBC distribution width (Fig. 2B) and a marked reduction in the red blood cell indices, mean cell volume, and mean corpuscular hemoglobin were detected in Fgf-23^−/− mice (Fig. 2, C and D), characteristic of hypochromic microcytic anemia. No differences were found in any of the examined hematologic parameters between WT and Fgf-23^+/− mice.

Fgf-23 Deficiency Results in Aberrant Erythropoiesis

Because of the significant increase in circulating red blood cell numbers, we further investigated by flow cytometry the effect of Fgf-23 deficiency in erythrocyte differentiation in PB and BM as the origin of postnatal hematopoiesis. Based on the known role of osteoblasts in the regulation of hematopoiesis (22, 23, 29) and published data showing severe reduction in bone mineral density and osteoblast numbers in Fgf-23^−/− mice (14, 20, 21, 30), we expected to find severely altered erythropoiesis in the BM of Fgf-23^−/− mice.

We confirmed the increase in total RBC numbers by analysis of erythroid cells and determined the maturation stage of erythroblasts (Ter119⁺) by the loss of CD71 expression, as reported by Asari et al. (31). A representative dot plot illustrating different erythroid populations is shown in Fig. 3A. Our results show a significant increase in the differentiation of immature (pro-E; Ter119^+med CD71^+hi), as well as mature erythroid cells (Ter119hi⁺) in PB (Fig. 3, B and C) and BM (Fig. 3, D and E) of Fgf-23^−/− mice. In addition, the capacity of Fgf-23-deficient HSCs to generate erythroid colonies (BFU-E) in vitro was considerably higher compared with WT BM cells (Fig. 4A), suggesting that loss of Fgf-23 results in increased erythroid progenitor cell activity. Furthermore, we examined whether the increase in erythrocyte numbers is due to increased Epo being released by the kidneys. Our data show that circulating Epo levels were elevated in Fgf-23^−/− mice compared with WT littermates (Fig. 4B). In addition, we found that Epo mRNA expression was significantly increased in bone marrow, liver, and kidney of Fgf-23-deficient mice (Fig. 4C) because of induced HIF signaling, as determined by significant up-regulation of the hypoxia-inducible transcription factors HIF-1α and HIF-2α mRNA expression in all three tissues examined (Fig. 4, D and E). However, HIF-1α and HIF-2α mRNA expression was suppressed in Fgf-23^−/− bone (Fig. 4, D and E), leading to significantly reduced Epo expression in bone (Fig. 4C, inset) both of which are not surprising because fgf-23-deficient mice have severely decreased osteoblast numbers. Recent reports have indicated that osteoblasts are a source of local Epo production in the bone, which would account for our observed decrease in bone Epo and HIF expression (32). Additionally, mRNA expression of hypoxia-responsive genes including transferrin, transferrin receptor, glucose transporter-1, and phosphoglycerate kinase-1 was significantly elevated both in liver and bone marrow of Fgf-23^−/− mice (Fig. 5, A–H). These findings suggest that a lack of Fgf-23 in mice results in increased erythrocyte production in the bone marrow and release into the circulation by altering the BM environment and rendering it hypoxic. Hypoxia in turn induces HIF signaling, which activates local and systemic Epo production that stimulates erythropoiesis in Fgf-23 mutant mice. To further address the role of hypoxia in mediating the observed erythroid cell changes, we treated Fgf-23^−/− mice and WT littermates with 100% oxygen for 1 h and measured several erythroid cell parameters. We have found that oxygen treatment significantly reduced serum Epo levels in Fgf-23^−/− mice and returned them to control levels (Fig. 4F). Furthermore, after only 1 h of oxygen treatment, mRNA expression of renal and BM HIF-1α was significantly decreased in Fgf-23^−/− mice compared with untreated Fgf-23^−/− mice (Fig. 4, G and I). Renal Epo mRNA was also found significantly decreased with oxygen treatment in Fgf-23^−/− mice (Fig. 4H). Importantly, BM Ter119+ erythroid populations were also rescued in Fgf-23^−/− mice treated with oxygen compared with untreated mice (Fig. 4J). These data further support our hypothesis that a lack of Fgf-23 stimulates a hypoxic BM environment, which activates Epo-induced erythropoiesis in our Fgf-23 mutant mice.

Fgf-23 Deficiency Alters Early Hematopoietic Progenitors

Because our data suggest that Fgf-23 results in aberrant erythropoiesis, we investigated whether these changes also occur in early hematopoietic progenitors. The frequency of CD150⁺CD48⁻ (SLAM) cells, enriched for HSCs, was 2-fold higher in Fgf-23^−/− mice compared with WT littermates, both in peripheral blood (2.3 ± 0.3 and 1.2 ± 0.3, respectively; Fig. 6A) and bone marrow (0.9 ± 0.1 and 0.41 ± 0.1, respectively; Fig. 6B), indicating that a loss of Fgf-23 results in increased HSC production. Similarly, other HSC markers, LSK (Lin⁻Sca-1⁺c-Kit⁺) and KTLS (c-Kit⁺Thy-1⁺Lin⁻Sca-1⁺), were also markedly elevated in PB and BM of Fgf-23^−/− mice compared with WT littermates (data not shown). We have also found that the increase in HSCs coincided with a significant decrease in HSC apoptosis in Fgf-23^−/− BM, as assessed by a combination of SLAM and annexin V staining (Fig. 6C). To determine whether the perturbations in the bone marrow and peripheral blood arise at the progenitor level, we characterized the capacity of hematopoietic progenitor cells from WT and Fgf-23^−/− mice for myeloid differentiation within their local BM environment. As assessed by colony-forming assay, the number of multilineage colony-forming cells (CFU-GEMM) per femur was significantly increased in Fgf-23^−/− bone marrow compared with WT littermates (Fig. 6D). Similarly, the number of total colonies formed in Fgf-23^−/− BM was also increased (data not shown). Therefore, our results demonstrate not only that the frequency of HSCs was drastically elevated in peripheral blood and bone marrow of Fgf-23^−/− mice, but also that this increase correlated with a significant increase in the in vitro proliferative capacity and absolute numbers of hematopoietic progenitor cells from Fgf-23^−/− mice, providing evidence for increased function and activity of Fgf-23-deficient HSCs.

Migration and Homing of Fgf-23-deficient BM Cells

Elevated HSCs in the bone marrow could be due to alterations in HSC homing or retention of the cells in the correct microenvironment and/or defects in their migratory function. To address this, we assessed surface expression of molecules required for HSC homing/migration such as the chemokine SDF-1α/CXCL-12. Serum SDF-1α levels were considerably elevated in Fgf-23^−/− mice compared with in WT mice (Fig. 7A). It has been shown that elevation of SDF-1 levels in peripheral blood results in hematopoietic progenitor and stem cell mobilization to the peripheral circulation (33). Our data also showed that BM cells from Fgf-23-deficient mice exhibited no change in chemotaxis toward an SDF-1α gradient compared with WT BM cells (Fig. 7B).

We further carried out in vivo homing experiments and transplanted WT and Fgf-23^−/− bone marrow cells (CD45.2⁺) into lethally irradiated B6.SJL WT (CD45.1⁺) recipient mice. Studies have shown that lethal irradiation creates a noncompetitive host HSC pool that can be easily replaced by donor HSCs (34, 35). Homing efficiency was assessed 24 h post-transplantation in the bone marrow, peripheral blood, and spleen of the recipient mice. We found a significant increase in the number of Fgf-23^−/− cells seeding the host bone marrow (Fig. 7, D and F) compared with transplanted WT BM cells. Furthermore, these experiments showed a marked reduction in Fgf-23^−/− BM cell localization in peripheral blood (Fig. 7, D–E). However, localization of Fgf-23^−/− and WT transplanted BM cells was similar in the spleens of irradiated recipient mice (Fig. 7, D and G). Finally, we assessed the ability of BM cells to adhere to fibronectin, a substance found in the bone microenvironment. We detected no significant differences between WT and Fgf-23^−/− BM cell adherence capabilities, indicating that Fgf-23^−/− BM cells are as capable as WT BM cells of migrating from the BM to the peripheral blood (Fig. 7C). Collectively, our results confirm that loss of Fgf-23 results in increased production of HSCs in the bone marrow and no evidence of a defect in their migratory function or in physical association with the regulatory components of the microenvironment niche.

Assessment of Spleen Cellularity

To assess whether loss of Fgf-23 results in hematopoietic abnormalities in hematopoietic organs other than the bone marrow, we investigated the cellular composition of the spleens of Fgf-23^−/− mice. Hematopoietic differentiation of homogenized spleen cells showed that the frequency of both immature and mature erythroid cells was significantly lower in Fgf-23^−/− mice (Fig. 8, A and B). These findings were confirmed by CFU assays showing a marked reduction in the number of erythroid colonies (BFU-E; Fig. 8C), as well as the primitive granulocyte-erythrocyte-macrophage-megakaryocyte (CFU-GEMM) progenitors and, subsequently, the number of total colonies formed (data not shown) in the spleens of Fgf-23^−/− mice compared with in WT mice, ruling out the possibility that extramedullary erythropoiesis is the cause of the elevated erythroid populations in Fgf-23^−/− mice. Furthermore, the frequency of HSC-enriched CD150⁺CD48⁻ (SLAM) and their colony forming ability were greatly higher in Fgf-23^−/− than in WT spleens (Fig. 8, D and E). It is possible that increased HSC frequency in our Fgf-23-deficient mice results in increased differentiation into the lymphoid lineage, specifically T-lymphocytes, because of heighten immune response. This may occur at the expense of the myeloid lineage, explaining the decrease in splenic erythroid populations. Further investigation of the different lymphoid populations is required to properly understand this phenomenon.

FGF-23 Affects Hematopoiesis Independent of Changes in the Bone Marrow Environment

Previous studies have shown that Fgf-23 is expressed in fetal liver, heart, and somites of mice at E11.5 days postcoitum (20). Examining prenatal erythropoiesis allows us to investigate the effects of Fgf-23 prior to inflammatory disease onset commonly observed in these mice (i.e., emphysema and renal insufficiency). To determine whether the hematopoietic changes in Fgf-23^−/− mice are specific to the bone marrow environment or due to aberrant production or function of HSCs, we examined whether hematopoietic stem cell populations in Fgf-23^−/− mice are affected before their translocation from fetal liver to the bone marrow.

We evaluated fetal liver cell populations at embryonic day E15.5, and we found that Fgf-23 deletion results in a significant increase in mature erythrocyte populations (Fig. 9A). Colony-forming assays also showed that the high frequency of erythroid cells in Fgf-23^−/− fetal livers correlated with an actual increase in functional erythroid progenitors and more BFU-E colonies from Fgf-23^−/− fetal liver cells (Fig. 9B). Interestingly, similar to the BM data, the frequency of HSC-enriched CD150⁺CD48⁻ (SLAM) cells was considerably higher in Fgf-23^−/− than in WT fetal livers (Fig. 9, C and D). LSK (Lin⁻Sca-1⁺c-Kit⁺) and KTLS (c-Kit⁺Thy-1⁺Lin⁻Sca-1⁺) cells were also found to be greatly increased in Fgf-23^−/− fetal livers (Fig. 9, E–G). Taken together, our data suggest that Fgf-23 regulates hematopoiesis independently of changes in the bone environment.

Effect of FGF-23 Administration on Erythropoiesis

To further investigate the mechanism by which Fgf-23 affects erythropoiesis, we injected intraperitoneally recombinant human FGF-23 protein (or vehicle (PBS)) into wild-type C57BL/6 mice and evaluated in vivo changes in hematopoiesis. Serum FGF-23 levels were markedly elevated after injection of FGF-23, confirming successful delivery of the protein (Fig. 10A). Our data show that exogenous administration of one single dose of FGF-23 protein in WT mice leading to high levels of FGF-23 results in erythropoietic changes opposite to the ones observed in mice lacking Fgf-23. Specifically, we detected a substantial reduction in erythropoiesis in all tissues examined (PB, BM, and spleen) in mice injected with FGF-23 (Fig. 10, C, D, F–H, and K–L). This correlated with an actual decrease in functional erythroid cells, which generated considerably fewer erythroid colonies in the BM (Fig. 10I). We also examined whether the decrease in erythrocyte numbers were due to a decrease in erythropoietin. Our data show that circulating Epo levels were in fact significantly reduced in mice injected with FGF-23 compared with WT littermates (Fig. 10B). These data further emphasize the role of Fgf-23 in regulating Epo-mediated changes in erythropoiesis. Furthermore, the frequency of HSC-enriched cells in peripheral blood (Fig. 10E), bone marrow (Fig. 10J), and spleen (Fig. 10M) was significantly reduced in mice injected with FGF-23. The changes induced by high FGF-23 levels were observed within 24 h of FGF-23 administration, indicating its potency.

To further determine the significant role of Fgf-23 in regulating erythropoiesis, we isolated and treated BM-derived Ter119⁺ cells from WT and Fgf-23^−/− littermate mice and treated the cells with FGF-23 protein or vehicle (PBS) for 4 h. We found that untreated Fgf-23^−/− Ter119⁺ cells showed significantly elevated mature erythroid cells (Ter119hi⁺) compared with untreated WT controls, whereas WT cells treated with FGF-23 protein resulted in reduced Ter119hi⁺ cell frequency (Fig. 11A), similar to our in vivo data. Importantly, FGF-23 treatment of Fgf-23-deficient erythroid cells rescued erythropoiesis and brought Ter119hi⁺ cell frequency to control levels (Fig. 11A). Furthermore, FGF-23 treatment resulted in a significant decrease in Epo and HIF mRNA expression in Fgf-23^−/− cells and similar to untreated WT levels (Fig. 11, C and D). Our data also show that Fgf-23 acts on the Epo-R. Loss of Fgf-23 results in high expression of the Epo-R in isolated Ter119⁺ cells and FGF-23 treatment restores the frequency of Epo-R positive cells to normal control levels (Fig. 11B). Taken together, our data strongly suggest that Fgf-23 plays an important role in the regulation of erythropoiesis through the HIF/Epo pathway.

Genetic Ablation of Vitamin D in Fgf-23^−/− Mice

Fgf-23 is a known regulator of 1,25(OH)₂D₃ and phosphate homeostasis. Fgf-23^−/− mice exhibit highly elevated serum vitamin D and phosphate levels along with severe skeletal abnormalities and extraosseous calcifications (14, 20, 21, 30). Elimination of vitamin D in Fgf-23^−/− mice reversed the described abnormalities in these mice (21), indicating that vitamin D partly mediates the function of Fgf-23 to regulate phosphate homeostasis and bone mineralization. Therefore, we investigated whether the elevated levels of vitamin D could account for the elevated levels of erythropoiesis in Fgf-23^−/− mice. We generated mice deficient in both Fgf-23 and 1α-hydroxylase genes, the enzyme required to convert vitamin D to its active form, 1,25(OH)₂D₃, (Fgf-23^−/−/1α(OH)ase^−/−). Here, we compared and analyzed data obtained from wild-type, Fgf-23^−/−, Fgf-23^−/−/1α(OH)ase^−/−, and 1α(OH)ase^−/− animals.

Complete blood count analysis revealed that ablation of vitamin D from Fgf-23^−/− mice partially rescued the erythroid changes observed in our Fgf-23^−/− mice. Specifically, RBC numbers were significantly increased in Fgf-23^−/− (as described above), Fgf-23^−/−/1α(OH)ase^−/−, and 1α(OH)ase^−/− animals compared with WT littermates (Fig. 12A); however, RBC counts in both Fgf-23^−/−/1α(OH)ase^−/− and 1α(OH)ase^−/− were markedly lower than in Fgf-23 single knock-out mice (Fig. 12A).

Because of the significant increase in circulating red blood cell numbers in all three mutant genotypes (Fgf-23^−/−, Fgf-23^−/−/1α(OH)ase^−/−, and 1α(OH)ase^−/−), we investigated further the impact of vitamin D deficiency in erythroid cell differentiation in BM, by flow cytometry. Interestingly, both Fgf-23^−/− and Fgf-23^−/−/1α(OH)ase^−/− mice had significantly elevated immature (pro-E) and mature (Ery-C) erythroid cells (Fig. 12, B and C) compared with WT controls. Erythroid cell populations, however, in 1α(OH)ase^−/− mice did not differ from WT mice, but they were considerably decreased compared with Fgf-23^−/− and Fgf-23^−/−/1α(OH)ase^−/− mice (Fig. 12, B and C). Evaluation of early SLAM enriched HSC progenitors showed elevated frequency of HSCs in Fgf-23^−/− and Fgf-23^−/−/1α(OH)ase^−/− mice, but not in 1α(OH)ase^−/− mice compared with WT littermates (Fig. 12D). SLAM⁺ cells were, however, greatly decreased in 1α(OH)ase^−/− mice compared with Fgf-23^−/− mice.

To further address the interplay between Fgf-23 and vitamin D in regulating hematopoiesis, we treated 1α(OH)ase^−/− mice with exogenous FGF-23 protein. As we have previously shown, FGF-23 administration significantly reduced both erythroid cell and HSC frequencies in the BM of WT cells (Fig. 12, E and F). However, exogenous FGF-23 significantly reduced the frequency of both immature and mature erythroid cells in 1α(OH)ase^−/− mice (Fig. 12, E and F). In summary, these data suggest that elimination of vitamin D cannot rescue the hematopoietic changes observed in our Fgf-23^−/− mice, suggesting that these changes are independent of vitamin D, and thus Fgf-23 plays a vital role in regulating hematopoiesis and, particularly, erythropoiesis.