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Transforming growth factor-β superfamily ligand trap ACE-536 corrects anemia by promoting late-stage erythropoiesis

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Rajasekhar N V S Suragani1, Samuel M Cadena1,3, Sharon M Cawley1,3, Dianne Sako1, Dianne Mitchell1, Robert Li1, Monique V Davies1, Mark J Alexander1, Matthew Devine1, Kenneth S Loveday1, Kathryn W Underwood1, Asya V Grinberg1, John D Quisel1, Rajesh Chopra2, R Scott Pearsall1, Jasbir Seehra1 & Ravindra Kumar1 Erythropoietin (EPO) stimulates proliferation of early-stage erythrocyte precursors and is widely used for the treatment of chronic anemia. However, several types of EPO-resistant anemia are characterized by defects in late-stage erythropoiesis1,2, which is EPO independent2,3. Here we investigated regulation of erythropoiesis using a ligand-trapping fusion protein (ACE-536) containing the extracellular domain of human activin receptor type IIB (ActRIIB) modified to reduce activin binding. ACE-536, or its mouse version RAP-536, produced rapid and robust increases in erythrocyte numbers in multiple species under basal conditions and reduced or prevented anemia in murine models. Unlike EPO, RAP-536 promoted maturation of latestage erythroid precursors in vivo. Cotreatment with ACE-536 and EPO produced a synergistic erythropoietic response. ACE-536 bound growth differentiation factor-11 (GDF11) and potently inhibited GDF11-mediated Smad2/3 signaling. GDF11 inhibited erythroid maturation in mice in vivo and ex vivo. Expression of GDF11 and ActRIIB in erythroid precursors decreased progressively with maturation, suggesting an inhibitory role for GDF11 in late-stage erythroid differentiation. RAP-536 treatment also reduced Smad2/3 activation, anemia, erythroid hyperplasia and ineffective erythropoiesis in a mouse model of myelodysplastic syndromes (MDS). These findings implicate transforming growth factor-b (TGF-b) superfamily signaling in erythroid maturation and identify ACE-536 as a new potential treatment for anemia, including that caused by ineffective erythropoiesis. Anemia is a common debilitating complication arising from diverse causes, including MDS, thalassemia, cancer chemotherapy, chronic kidney disease and hemorrhage4. Endogenous EPO and its cognate receptor are critical for the survival, proliferation and differentiation of erythroid progenitors during early-stage erythropoiesis5. Notably, erythroblast differentiation and maturation during late-stage

erythropoiesis is independent of EPO3,5. Anemia arising from late-stage defects is characterized by abortive maturation and ineffective erythropoiesis in which red blood cell (RBC) levels are deficient despite hypercellular bone marrow and elevated levels of endogenous EPO6,7. Ineffective erythropoiesis occurs in diseases such as MDS and β-thalassemia and often requires iron chelation therapy as an adjunct to chronic blood transfusion to alleviate potentially fatal iron overloading6,8,9. In patients with MDS, iron overloading due to transfusion dependence is also associated with decreased survival and increased progression to acute myeloid leukemia9. Members of the TGF-β superfamily, which include TGF-βs, activins, GDFs and bone morphogenetic proteins (BMPs)10, have been studied as potential regulators of erythropoiesis. Many studies have documented the effects of activin A and other superfamily members on erythroid precursor cells or cell lines11–16 and have implicated TGF-β superfamily signaling in other aspects of hematopoiesis16–18. However, there is a paucity of data regarding the role of these pathways in regulating erythropoiesis in vivo16–19. Notably, elevated levels of phosphorylated Smad2 and Smad3 (Smad2/3), key transcriptional mediators for multiple ligands in the TGF-β superfamily, have been reported in hematopoietic progenitors from patients with MDS20. Nonetheless, the role of TGF-β superfamily members in regulating erythroblast maturation during late-stage erythropoiesis remains unclear. Here we investigated how ActRIIB ligand traps affect the regulation of late-stage erythropoiesis. We used either ACE-536, a receptor fusion protein consisting of a modified human ActRIIB extracellular domain linked to the human IgG1 Fc domain, or RAP-536, a mouse version of ACE-536 with an identical ligandbinding domain. We introduced specific modifications in the wild-type ActRIIB extracellular domain: a single amino acid substitution (L79D), an N-terminal truncation of four amino acids and a C-terminal truncation of three amino acids. ACE-536 treatment increased RBC count, hemoglobin levels and hematocrit in a dose-dependent manner in mice, rats and monkeys under basal

1Acceleron

Pharma, Cambridge, Massachusetts, USA. 2Translational Development Department, Celgene, San Francisco, California, USA. 3These authors contributed equally to this work. Correspondence should be addressed to R.K. ([email protected]).

Received 29 October 2013; accepted 25 February 2014; published online 23 March 2014; doi:10.1038/nm.3512

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TBS RAP-536 Figure 1  ACE-536, a modified ActRIIB 4 100 100 104 10 fusion protein, increases RBC numbers, R1 80 80 103 103 * hemoglobin concentrations and hematocrit 60 60 R2 2 values by promoting maturation of late-stage BM 10 102 40 40 erythroblasts. (a) Differences in RBC numbers, 20 20 101 101 hemoglobin concentrations and hematocrit R3 0 0 100 100 values from vehicle (TBS, n = 8) after s.c. TBS ACE-536 TBS ACE-536 1 2 10 10 103 104 101 102 103 104 administration of ACE-536 to C57BL/6 mice CFU-E: spleen CFU-E: bone marrow 4 104 10 −1 twice weekly for 8 weeks at 0.1 mg kg 1,000 300 R1 (n = 9), 0.3 mg kg−1 (n = 7), 1 mg kg−1 (n = 8), 103 103 750 −1 −1 3 mg kg (n = 9) or 10 mg kg (n = 6). 2 2 200 R2 10 Spleen 10 * Absolute values for the highest dose are 500 101 101 shown in Supplementary Table 1. **P < 0.01, 100 250 R3 ***P < 0.001, as determined by one-way 100 100 0 0 analysis of variance (ANOVA) followed by 101 102 103 104 101 102 103 104 TBS ACE-536 TBS ACE-536 Ter-119 Dunnett’s post hoc analysis for multiple comparisons. (b) Differences in RBC numbers, Spleen Bone marrow 30 TBS hemoglobin concentrations and hematocrit 25 25 RAP-536 values from vehicle (TBS, n = 20) at day *** 20 20 29 after s.c. administration of ACE-536 to 15 *** 15 Sprague Dawley rats on days 1 and 15 (n = 20 10 10 per group). Absolute values are shown in 5 5 Supplementary Figure 1a. *P < 0.05, **P < 0.01, ** ***P < 0.001, as determined by ANOVA. 0 0 BasoE PolyE + OrthoE Late OrthoE BasoE PolyE + OrthoE Late OrthoE (c) Differences in RBC numbers, hemoglobin (R1) (R2) + Retic (R3) (R1) (R2) + Retic (R3) concentrations and hematocrit values from −1 −1 −1 baseline at day 13 after a single dose of ACE-536 to cynomolgus monkeys at 0.4 mg kg s.c. (n = 6), 2 mg kg s.c. (n = 6), 10 mg kg s.c. (n = 10), 30 mg kg−1 s.c. (n = 10) or 10 mg kg−1 intravenously (n = 10). Absolute values are shown in Supplementary Figure 1b. (d) Numbers of BFU-Es and CFU-Es obtained from bone marrow and spleen of C57BL/6 mice analyzed 48 h after administration of a single s.c. dose of ACE-536 (10 mg kg −1) or vehicle (n = 5 mice per group for spleen CFU-Es, otherwise n = 6 mice per group; all plated in duplicate). (e) Representative example of CD71 and Ter-119 flow cytometric profiles from the bone marrow (BM) and spleen of C57BL/6 mice analyzed at 72 h after s.c. treatment with vehicle (TBS) or RAP-536 (10 mg kg−1) (n = 5 per group). R1 corresponds to basophilic erythroblasts (BasoE), R2 corresponds to poly- and orthochromatophilic erythroblasts (PolyE + OrthoE) and R3 corresponds to late orthochromatophilic erythroblasts and reticulocytes (Late OrthoE + Retic). (f) Proportion of erythroid precursor populations as defined in e. Data in a–d and f are means ± s.e.m. Statistical significance between animals treated with vehicle and ACE-536 or RAP-536 is shown in c, d and f by *P < 0.05, **P < 0.01, ***P < 0.001, as determined by Student’s t-test.

conditions (Fig. 1a–c and Supplementary Fig. 1). Although a fusion protein containing the wild-type ActRIIB extracellular domain has been shown to increase muscle mass21, we observed no effects on body mass or on other blood cell counts in ACE-536–treated mice (Supplementary Fig. 1 and Supplementary Table 1). ACE-536 administration in mice led to elevation of RBC parameters that occurred within 3 d of a single dose and lasted for at least 30 d after dosing for 1 week (Supplementary Fig. 2a–c). A time-course analysis of reticulocyte production in rats following a single subcutaneous (s.c.) dose of RAP-536 also demonstrated that absolute reticulocyte numbers in peripheral blood increased to an average of twice the baseline levels during the 6–24 h interval after treatment and then declined to baseline levels coincident with rising RBC numbers (Supplementary Fig. 2d,e). RBC lifespan in mice was unaltered either during or after prolonged administration of RAP-536 (Supplementary Fig. 2f,g). These results indicate that ACE-536 promotes RBC formation under basal physiologic conditions. We next investigated potential therapeutic effects of ACE-536 and RAP-536 in murine models of acute or chronic anemia. Compared to vehicle (Tris-buffered saline (TBS)), RAP-536 treatment led to a more

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rapid recovery of RBC indices in both a rat model of acute anemia caused by blood loss (Supplementary Fig. 3a) and also a rat model of chemotherapy-induced anemia using carboplatin administration (Supplementary Fig. 3b). ACE-536 treatment significantly (P < 0.001) increased RBC indices in a mouse model of chronic kidney disease (Supplementary Fig. 3c,d). When the study was terminated, partially nephrectomized mice had reduced kidney weights and elevated serum concentrations of creatinine and urea compared with mice given sham surgery (Supplementary Fig. 3e–g), indicating persistence of renal insufficiency after nephrectomy. As expected, ACE-536 treatment did not affect kidney size or function (Supplementary Fig. 3e–g). To identify specific erythropoietic precursors affected by ACE-536, we assessed numbers of early-stage progenitors using erythroid burstforming unit (BFU-E) and erythroid colony-forming unit (CFU-E) assays, as well as of maturing erythroblasts, in both bone marrow and spleen of mice. By 48 h after treatment, ACE-536 treatment led to reduced BFU-Es and CFU-Es in bone marrow compared to vehicle (TBS) (P < 0.05) (Fig. 1d) and was associated with a similar trend in the spleen (P = 0.08). These effects of ACE-536 on erythroid progenitors contrast with the well-characterized proliferative effects of EPO 5.

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Figure 2  ACE-536 promotes maturation of erythroblasts independently Hemoglobin of EPO and acts synergistically with EPO to increase RBC production. (a) Differences in RBC numbers, hemoglobin concentrations and hematocrit values from vehicle determined at day 4 in C57BL/6 mice BFU-E CFU-E ProE BasoE PolyE OrthoE Retic RBC treated with ACE-536 (10 mg kg−1, intraperitoneal (i.p.) injection, day 0), neutralizing antibody against EPO (EPO mAb, 5.5 mg kg −1, i.p., days 0 ACE-536 responsive and 2) or the two in combination (n = 5 per group). *P < 0.05, **P < 0.01, EPO dependent ***P < 0.001 versus vehicle, as determined by Student’s t-test. (b) Renal levels of Epo mRNA in mice from a. *P < 0.05, **P < 0.01 versus vehicle; #P < 0.05 versus EPO mAb, as determined by Student’s t-test. (c) Differences in RBC numbers, hemoglobin concentrations and hematocrit values from vehicle determined at 72 h after separate or combination treatment with ACE-536 (10 mg kg−1, s.c.) and EPO (1,800 units per kg body weight (units kg −1), s.c.) in C57BL/6 mice (n = 4 per group). ***P < 0.001, ACE-536 + EPO versus vehicle. (d) Number of splenic erythroid precursor populations determined by flow cytometric analysis (as in Fig. 1) analyzed at 72 h after treatment of C57BL/6 mice (n = 4 per group) with EPO alone (1,800 units kg −1) or ACE-536 + EPO (10 mg kg−1). **P < 0.01, ***P < 0.001, ACE-536 + EPO versus EPO alone, as determined by Student’s t-test. Data in a–d are means ± s.e.m. (e) Comparison of the erythropoietic stages dependent on EPO with the stages exhibiting responsiveness to ACE-536. EPO stimulates proliferation and differentiation of early erythroid progenitors, and ACE-536 promotes erythrocyte maturation.

However, by day 7 of ACE-536 treatment (10 mg per kg body weight (mg kg−1), s.c., twice weekly), there was a trend toward increased BFU-E and CFU-E numbers (Supplementary Fig. 4). Differentiation profiling of terminal erythroid precursors from bone marrow and spleen, as determined by flow cytometric analysis with fluorophore-conjugated antibodies against CD71 and Ter-119 (refs. 22,23) (Fig. 1e), revealed that RAP-536 promoted the maturation of developing erythroblasts (Fig. 1e,f). Seventy-two hours after treatment with RAP-536, the number of basophilic erythroblasts was decreased and the numbers of poly- and orthochromatophilic erythroblasts and reticulocytes was concurrently increased in bone marrow and spleen as compared with cell numbers in vehicle-treated mice (Fig. 1e,f). For this analysis, R1, R2 and R3 cell populations were measured in bone marrow and spleen from mice treated with RAP-536 or TBS (R1 corresponds to basophilic erythroblasts, R2 to poly- and orthochromatophilic erythroblasts and R3 to late orthochromatophilic erythroblasts and reticulocytes). We confirmed the identities of each cell population by cytospin morphologic analysis (Supplementary Fig. 4e). We further confirmed the ability of RAP-536 to promote erythroblast maturation by analyzing its effects on Ter-119+ erythroid precursors from bone marrow and spleen that had been sorted using forward scatter (FSC, a measure of cell size) in combination with cell surface expression of either CD71 (ref. 24) (Supplementary Fig. 5) or CD44 (ref. 25) (Supplementary Fig. 6). These data indicate that compared to vehicle, at 72 h after treatment with RAP-536 there were reduced numbers of basophilic erythroblasts in bone marrow and spleen and increased numbers of late-stage (mature) erythroblasts and reticulocytes. Treatment of mice with RAP-536 did not affect apoptosis of Ter-119high erythroid precursors in bone marrow and spleen (Supplementary Fig. 7a) but did increase spleen weight by approximately 40% (Supplementary Fig. 7b,c), consistent with enhanced

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extramedullary erythropoiesis, as described earlier24. We did not detect a significant effect of RAP-536 on iron parameters such as serum iron, total iron binding capacity or iron saturation at week 1 or week 3 after treatment (Supplementary Fig. 7d–f). Similarly, we found no effect of RAP-536 on expression of Hamp (hepcidin) mRNA or Bmp6 mRNA in liver (Supplementary Fig. 7g,h), providing evidence that ACE-536 and RAP-536 do not act by altering iron homeostasis in vivo26. These findings indicate that endogenous TGF-β superfamily ligands sequestered by ACE-536 negatively regulate late-stage erythroid differentiation. To determine whether ACE-536–induced erythropoiesis is EPO dependent, we treated mice with ACE-536 or a neutralizing monoclonal antibody against EPO (EPO mAb). By day 4, ACE-536 treatment increased RBC number, hemoglobin levels and hematocrit (P < 0.001), whereas EPO mAb treatment decreased these parameters as expected (P < 0.001) (Fig. 2a). Notably, ACE-536 treatment completely rescued RBC parameters from the effects of EPO neutralization and even elevated RBC numbers significantly compared to vehicle (P < 0.05; Fig. 2a). These results demonstrate that the stimulatory effect of ACE-536 on RBC parameters is largely EPO independent at day 4. However, rescue of RBC parameters from the effects of EPO neutralization by ACE-536 was attenuated at day 7 (Supplementary Fig. 8a) compared to day 4, indicating that although accelerated maturation of late-stage erythroid precursors by ACE-536 is EPO independent, EPO is necessary for the continued supply of earlystage progenitors. Accordingly, Epo transcript levels in mice cotreated with ACE-536 and EPO mAb were increased at day 7 (Supplementary Fig. 8b) compared to day 4 (Fig. 2b). Cotreatment of mice with ACE-536 and EPO resulted in robust increases in RBC parameters at 72 h that were greater than the sum of the agents’ separate effects (Fig. 2c). Flow cytometric analysis of

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Figure 3  GDF11-ActRIIB-Smad2/3 signaling in late-stage erythroid differentiation. (a) Representative inhibition curves for ACE-536 in combination with 5 100 15 10 250 * GDF11 (top) or GDF8 (bottom) based on two 4 200 14 independent experiments in an A204 cell–based 80 3 150 * * * assay measuring expression of a CAGA12-luciferase 13 9 2 100 reporter construct. Background represents results 60 12 1 50 obtained in the absence of ligand. (b) Representative 11 8 0 40 0 examples of a single experiment in which spleen sections from C57BL/6 mice (n = 5) with phlebotomyinduced anemia were immunostained for GDF11 (top) or Ter-119 (bottom) with hematoxylin counterstain. Arrowheads denote red pulp. Scale bar, 200 µm. (c) mRNA levels of the indicated genes in immature (Ter-119 −) and mature (Ter-119+) precursor cells in vivo, as determined by quantitative RT-PCR (qRT-PCR). Gata2 mRNA was used as a confirmatory marker for immature erythroid precursors, and hemoglobin β-subunit (Hbb) mRNA (Ter-119+/Ter-119− cells = 16.8) (data not shown) was used as a marker for Ter-119 + erythroid precursors. (d) Messenger RNA levels in synchronously differentiating Ter-119 − erythroid precursors ex vivo determined by qRT-PCR. Data in c and d are means ± s.e.m. from three independent experiments. (e) Levels of phosphorylated Smad2 (p-Smad2) as determined by enzyme-linked immunosorbent assay in fetal liver erythroid precursors treated ex vivo for 1 or 3 h with GDF11 (10 or 50 ng ml−1) in the absence or presence of ACE-536 (1 µg ml−1). Data are means of two independent experiments. C, untreated control. (f) Levels of phosphorylated Smad2 as determined by western blotting of total spleen extracts from EPO-pretreated mice treated with RAP-536 (10 mg kg −1, i.p.) or vehicle (TBS) for 6 h (n = 2 mice per group). (g) Left, representative examples of CD71 and Ter-119 flow cytometric profiles from spleens of C57BL/6 mice at 72 h after i.p. treatment with vehicle (TBS), EPO (1,800 IU kg −1, single dose) or EPO + GDF11 (1 mg kg−1, daily) (n = 3 mice per group). Right, proportion of erythroid precursor populations in regions R2 and R3. ##P < 0.01, versus TBS; **P < 0.01, GDF11 + EPO versus EPO alone, as determined by Student’s t-test. (h) RBC indices (left) and spleen weights (right) in C57BL/6 mice treated with GDF11 (0.1 mg kg−1, i.p., n = 4) or vehicle (PBS, n = 5) daily for 11 d. *P < 0.05 versus vehicle, as determined by Student’s t-test. Hgb, hemoglobin. (i) Left, serum GDF11 concentrations in subjects with MDS (n = 8, 67–91 years of age) versus age-matched normal individuals (n = 5). Right, Gdf11 mRNA levels in bone marrow and spleen from a NUP98-HOXD13 mouse model of MDS normalized to the levels in wild-type (WT) mice (n = 3 per group). *P < 0.05 versus wild-type mice, as determined by Student’s t-test. Data in c–e and g–i are means ± s.e.m. RBCs (1 × 106 cells µl–1)

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splenic erythroblasts at 72 h after treatment revealed that cotreatment with ACE-536 and EPO significantly increased maturation of basophilic erythroblasts compared to EPO alone, as shown by a shift in cells to more mature stages (Fig. 2d). The greater-thanadditive effect of combining ACE-536 and EPO treatment is probably due to the increased availability of early progenitors induced by EPO. Consistent with the concept that RAP-536 promotes erythroid precursor maturation, EPO-pretreated mice treated with RAP-536 exhibited fewer erythroid precursors (CD71highTer-119high) in S phase

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(Supplementary Fig. 8c) and more in the G0/G1 phase of the cell cycle (Supplementary Fig. 8d) compared to vehicle. Together, these results demonstrate that ACE-536 increases RBC formation by stimulating precursor maturation (Fig. 2e). We next explored signaling pathways responsible for the action of ACE-536. As determined under cell-free conditions by surface plasmon resonance, ACE-536 bound preferentially to GDF11, GDF8 and activin B (Smad2/3-activating ligands), as well as to BMP10 and BMP6 (ligands that activate Smad1, Smad5 and Smad8 (Smad1/5/8)),

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Figure 4  ACE-536 inhibits Smad2/3 Late stage: 12 months old Early stage: 6 months old activation, mitigates ineffective 13 9 14 9 erythropoiesis and ameliorates anemia in a * ** 8 8 * * NUP98-HOXD13 mouse model of MDS at 12 11 ### multiple stages of disease severity. (a) RBC ### 7 7 10 numbers and hemoglobin concentrations 6 6 9 (top) and morphological enumeration of 8 5 5 hematopoietic precursors in bone marrow 7 4 6 4 (bottom) in vehicle-treated wild-type (WT) WT MDS MDS WT MDS MDS MDS MDS MDS MDS mice (n = 5), vehicle-treated MDS mice + + + + + + + + + + TBS TBS RAP-536 TBS TBS RAP-536 TBS RAP-536 TBS RAP-536 (n = 5) and MDS mice (n = 6) treated with −1 RAP-536 (10 mg kg , twice weekly) for 8 weeks ending at approximately 6 months of 26% 24% 32% 43% age (early stage). *P < 0.05, **P < 0.01, versus 48% 52% 56% 74% vehicle-treated MDS mice; ###P < 0.001 68% 75% versus wild-type mice, as determined by Student’s t-test. (b) RBC numbers and WT + TBS MDS + TBS MDS + RAP-536 MDS + TBS MDS + RAP-536 hemoglobin concentrations (top) and morphological enumeration of hematopoietic Erythroid precursors Nonerythroid cells precursors in bone marrow (bottom) in MDS mice treated with RAP-536 (10 mg kg−1, WT + TBS MDS + TBS MDS + RAP-536 twice weekly, n = 5) or vehicle (n = 4) for 7 weeks ending at approximately 12 months of age (late stage). *P < 0.05 versus vehicle-treated MDS mice, as determined by Student’s t-test. Data in a and b are means ± s.e.m. (c) Representative phosphorylated Smad2/3 immunostaining (brown) in hemotoxylin-counterstained spleen sections from a vehicle-treated wild-type mouse, a vehicle-treated MDS mouse and an MDS mouse treated with RAP-536 (10 mg kg −1, twice weekly) for 7 weeks ending at 12 months of age. Scale bars, 100 µm and 10 µm (inset).

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within a tenfold range of Kd values (Supplementary Tables 2 and 3). Additional analysis using a cell-based reporter gene assay revealed that ACE-536 potently inhibited Smad2/3 signaling induced by GDF11 and GDF8, but not signaling induced by other ligands tested, including activins and TGF-β1 (Fig. 3a, Supplementary Fig. 9a–d and Supplementary Table 4). Previous studies have implicated activin A in erythropoiesis. We therefore investigated the possibility that the effects of ACE-536 on RBC production depend on intact signaling by activin A, activin C or activin E, perhaps as part of a compensatory response to direct effects on other ligand pathways. The erythropoietic response to RAP-536 was unaltered by pretreatment of wild-type mice with antibody against activin A or in mice lacking inhibin C or inhibin E (Supplementary Fig. 10a–c), confirming that the response occurs independently of these ligands. Despite high-affinity binding to BMP6 and BMP10 under cell-free conditions (Supplementary Table 2), ACE-536 did not inhibit Smad1/5/8 signaling induced by these ligands in cellbased assays (Supplementary Fig. 9e–g), suggesting that ACE-536 could not compete with endogenous receptors for these ligands. We conducted additional experiments to investigate the GDF11ActRIIB-Smad2/3 pathway as a potential regulator of erythropoiesis. In spleen sections from wild-type mice, GDF11 immunostaining was prominent in the erythropoietic niche (red pulp area), as demarcated by Ter-119 immunostaining (Fig. 3b). In erythroid precursors from mouse fetal liver, Gdf11, Acvr2b and certain other members of the TGF-β superfamily were expressed at elevated levels in immature (Ter-119−) compared to mature (Ter-119+) erythroid precursors, whereas Gdf8 mRNA was not detectable at either stage (Fig. 3c and Supplementary Table 5). During synchronous differentiation of erythroid precursors isolated from mouse fetal liver ex vivo, the levels of Gdf11 mRNA and (Acvr2b) mRNA were highest at the erythroid progenitor stage and then declined progressively over 48 h (Fig. 3d). Treatment of erythroid precursors from mouse fetal

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liver with GDF11 ex vivo increased Smad2/3 phosphorylation, and cotreatment with ACE-536 blunted this increase by more than half (Fig. 3e). Similarly, RAP-536 treatment in vivo inhibited Smad2/3 phosphorylation in mouse spleen (Fig. 3f) but did not affect Smad 1/5/8 phosphorylation (data not shown). Together, these results indicate that erythroid precursors downregulate GDF11 and its cognate receptor ActRIIB progressively with maturation and respond to exogenous GDF11 treatment with Smad2/3 activation, raising the possibility of cell-autonomous signaling during terminal erythroid differentiation. These results also show that ACE-536 or RAP-536 is able to inhibit this GDF11-induced signaling. To better understand the role of GDF11 signaling in erythro­poiesis, we treated early-stage Ter-119− cells with GDF11 and found increased expression of multiple TGF-β superfamily target genes involved in cellular proliferation, such as Myc, Id1, Atf4 and Pdgfa (Supplementary Table 6). Consistent with this observation, cotreatment with EPO and GDF11 in vivo over a 72-h period increased the numbers of CD71highTer-119low (R2) immature erythroid precursors (P < 0.01) and concurrently reduced the numbers of CD71highTer-119high (R3) erythroid precursors (P < 0.01) in bone marrow and spleen, as compared to EPO treatment alone (Fig. 3g). This result indicates that GDF11 negatively regulates late-stage erythroid differentiation. Similarly, chronic administration of GDF11 to wild-type mice produced mild anemia with reduced RBC indices and caused concomitant increases in spleen weight (~25%) and in the number of erythroid precursors (~37%, data not shown), indicative of erythroid hyperplasia and ineffective erythropoiesis (Fig. 3h). In addition, we used flow cytometric analysis to monitor the differentiation of mouse splenic erythroid progenitors ex vivo in the presence or absence of exogenous GDF11. Compared to vehicle, GDF11 reduced the number of mature erythroblasts (Supplementary Fig. 11), indicating that GDF11 inhibits terminal erythroid differentiation. Unlike treatment in vivo, GDF11

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letters treatment ex vivo did not alter the number of CD71highTer-119low cells compared to vehicle (Supplementary Fig. 11), suggesting that cells whose maturation is blocked by GDF11 do not survive under ex vivo conditions. Notably, concurrent treatment with ACE-536 attenuated these inhibitory effects of GDF11, as indicated by increased numbers of mature erythroblasts (Supplementary Fig. 11). These findings indicate that the inhibitory effect of GDF11 on terminal erythroid differentiation results at least in part from a direct action on erythroid precursors. In EPO-treated mice, expression of ActRIIB mRNA in spleen was increased significantly (P < 0.05) at 24 h but then reduced signi­ ficantly (P < 0.05) at 48 h after treatment, as compared to vehicle (Supplementary Fig. 12a). Expression of GDF11 mRNA followed a similar, albeit nonsignificant, trend (Supplementary Fig. 12a). The levels of GDF11 mRNA and ActRIIB mRNA were also concurrently reduced in mice at 48 h following acute blood loss compared to vehicle-treated mice (Supplementary Fig. 12b). Together, these results support a model in which key components of the GDF11-Smad2/3 pathway in erythroid progenitors are transiently upregulated upon acute EPO stimulation to promote erythroid expansion and then coordinately downregulated to allow precursors to undergo maturation in response to an acute demand for RBCs. The ability of RAP536 to enhance the acute response to EPO (Fig. 2c) or to accelerate recovery from acute blood loss (Supplementary Fig. 3a) is consistent with this model. Embryonic loss-of-function mutations in GDF11 alter patterning of the axial skeleton and produce abnormalities of the palate and kidney27. GDF11 has also been implicated in negative regulation of neurogenesis28 and retinal development29, but no previous studies of GDF11 have reported changes in erythropoiesis. In agreement with results presented here, GDF11 has recently been observed in bone marrow and spleen, with high expression measured in mouse spleen30. On the basis of our findings, we investigated circulating GDF11 concentrations in a limited sample of humans with the heterogeneous disorder MDS and found these subjects to have elevated levels in comparison with age-matched control subjects (Fig. 3i), consistent with the erythroid hyperplasia and ineffective erythropoiesis that characterizes this syndrome. We therefore investigated the effects of RAP-536 in the NUP98HOXD13 mouse model of MDS, which is characterized by abortive precursor maturation and ineffective hematopoiesis31. Untreated MDS mice displayed elevated expression of GDF11 mRNA in bone marrow and particularly in spleen compared to wild-type mice (Fig. 3i). As expected, 6-month-old MDS mice developed severe anemia compared to wild-type mice (Fig. 4a). Bone marrow analyses revealed increased numbers of erythroid precursors (Fig. 4a) and a lower myeloid/ erythroid (M/E) ratio (Supplementary Fig. 13d) in MDS mice compared to wild-type mice, indicative of ineffective erythropoiesis. In 6-month-old MDS mice, RAP-536 treatment significantly increased RBC count (+16.9%, P < 0.01) and hemoglobin level (+12.5%, P < 0.05) (Fig. 4a), reduced erythroid precursor cell count in bone marrow (Fig. 4a, P < 0.05) and normalized the M/E ratio to that of wild-type mice (Fig. 4a and Supplementary Fig. 13d). Similarly, in 12-monthold MDS mice, RAP-536 treatment significantly increased RBC count (+18.3%, P < 0.05) and hemoglobin level (+13.0%, P < 0.05), reduced erythroid precursor cell count (P < 0.01) and improved the M:E ratio, as compared to vehicle (Fig. 4b and Supplementary Fig. 13e), without affecting apoptosis (Supplementary Fig. 13a). RAP-536 treatment did not affect the absolute number of myeloid precursors (data not shown). Flow cytometry confirmed that RAP-536 treatment reduced

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erythroid hyperplasia in MDS mice at both ages (Supplementary Fig. 13b,c). A time-course analysis in MDS mice treated with RAP-536 for 7 months showed a sustained elevation in RBC numbers for the duration of the study (Supplementary Fig. 13f). Finally, spleen sections from untreated MDS mice displayed increased phosphorylated Smad2/3 immunostaining compared to wild-type mice, and RAP-536 treatment blunted this increase (Fig. 4c), suggesting that RAP-536 can mitigate aberrant Smad2/3 signaling in MDS mice. Together, these results indicate that inhibition of Smad2/3 signaling with RAP-536 ameliorates anemia, erythroid hyperplasia and ineffective erythropoiesis in MDS mice regardless of disease severity. Our findings, obtained with a modified ActRIIB ligand trap, reveal a role for TGF-β superfamily ligands in regulating terminal erythroid differentiation by a Smad2/3-dependent process. Whereas Smad1/5/8-dependent signaling mediated by BMP4 has previously been implicated in the regulation of stress erythropoiesis32, our results implicate GDF11-ActRIIB-Smad2/3–dependent signaling as a key regulatory mechanism in proliferating erythroid precursors that controls their late-stage maturation under both steady-state and stress conditions. It remains to be determined whether additional TGF-β superfamily members involved in Smad2/3 signaling also participate in this process. Although our findings raise the possibility of GDF11-Smad2/3 signaling in a cell-autonomous mechanism, paracrine signaling by accessory cells in the erythropoietic niche cannot be excluded33. TGF-β superfamily signaling pathways may represent new EPO-independent therapeutic targets that are particularly relevant to the treatment of anemia in patients with diseases characterized by ineffective erythropoiesis. ACE-536 therefore provides an opportunity to treat anemia associated with disorders, such as MDS and β-thalassemia, characterized by ineffective erythropoiesis with high EPO levels, an accumulation of immature erythroid precursors and decreased maturation of these cells. The related fusion protein sotatercept consists of the unmodified extracellular domain of wild-type human ActRIIA receptor attached to the Fc portion of human IgG1 and shares with ACE-536 the ability to bind GDF11 and increase erythrocytic parameters. Unlike ACE-536, sotatercept binds activins with high affinity. ACE-536 and sotatercept34 have completed phase 1 clinical studies in healthy volunteers, and phase 2 studies are ongoing with these agents for anemia in patients with MDS or thalassemia. Methods Methods and any associated references are available in the online version of the paper. Note: Any Supplementary Information and Source Data files are available in the online version of the paper. Acknowledgments We thank J. Knopf for critical review and T. Ahern for editorial contributions. We acknowledge the Cell Biology, Protein Purification and Preclinical Pharmacology groups at Acceleron Pharma for their contributions in support of this work. We also thank G. Paradis and A. Parmelee for their assistance with flow cytometry. NUP98-HOXD13 MDS mice and age-matched FVB wild-type mice were obtained from P. Aplan’s laboratory at the US National Institutes of Health. Inhbctm1Zuk and Inhbctm2Zuk mice were obtained from M. Matzuk, Baylor College of Medicine. AUTHOR CONTRIBUTIONS R.N.V.S.S., S.M. Cadena, S.M. Cawley, J.D.Q., R.C., R.S.P., J.S. and R.K. planned and designed the experiments. R.N.V.S.S., S.M. Cadena, S.M. Cawley, D.S. and R.L. conducted the experiments. R.N.V.S.S., S.M. Cadena, S.M. Cawley, D.S., D.M., R.L., M.V.D., M.D., K.S.L., K.W.U. and A.V.G. collected and interpreted data. R.N.V.S.S. and M.J.A. drafted and revised the manuscript.

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letters COMPETING FINANCIAL INTERESTS The authors declare competing financial interests: details are available in the online version of the paper.

1. Bowen, D. What is ineffective erythropoiesis in myelodysplastic syndromes? Leuk. Lymphoma 18, 243–247 (1995). 2. Libani, I.V. et al. Decreased differentiation of erythroid cells exacerbates ineffective erythropoiesis in β-thalassemia. Blood 112, 875–885 (2008). 3. Eshghi, S., Vogelezang, M.G., Hynes, R.O., Griffith, L.G. & Lodish, H.F. α4β1 integrin and erythropoietin mediate temporally distinct steps in erythropoiesis: integrins in red cell development. J. Cell Biol. 177, 871–880 (2007). 4. Smith, R.E. Jr. The clinical and economic burden of anemia. Am. J. Manag. Care 16 (suppl.), S59–S66 (2010). 5. Hattangadi, S.M., Wong, P., Zhang, L., Flygare, J. & Lodish, H.F. From stem cell to red cell: regulation of erythropoiesis at multiple levels by multiple proteins, RNAs, and chromatin modifications. Blood 118, 6258–6268 (2011). 6. Tanno, T. & Miller, J.L. Iron loading and overloading due to ineffective erythropoiesis. Adv. Hematol. 2010, 358283 (2010). 7. Ginzburg, Y. & Rivella, S. β-thalassemia: a model for elucidating the dynamic regulation of ineffective erythropoiesis and iron metabolism. Blood 118, 4321–4330 (2011). 8. Cao, A., Moi, P. & Galanello, R. Recent advances in β-thalassemias. Pediatr. Rep. 3, e17 (2011). 9. Jabbour, E., Kantarjian, H.M., Koller, C. & Taher, A. Red blood cell transfusions and iron overload in the treatment of patients with myelodysplastic syndromes. Cancer 112, 1089–1095 (2008). 10. Rider, C.C. & Mulloy, B. Bone morphogenetic protein and growth differentiation factor cytokine families and their protein antagonists. Biochem. J. 429, 1–12 (2010). 11. Yu, J. et al. Importance of FSH-releasing protein and inhibin in erythrodifferentiation. Nature 330, 765–767 (1987). 12. Broxmeyer, H.E. et al. Selective and indirect modulation of human multipotential and erythroid hematopoietic progenitor cell proliferation by recombinant human activin and inhibin. Proc. Natl. Acad. Sci. USA 85, 9052–9056 (1988). 13. Shiozaki, M., Kosaka, M. & Eto, Y. Activin A: a commitment factor in erythroid differentiation. Biochem. Biophys. Res. Commun. 242, 631–635 (1998). 14. Maguer-Satta, V. et al. Regulation of human erythropoiesis by activin A, BMP2, and BMP4, members of the TGFβ family. Exp. Cell Res. 282, 110–120 (2003). 15. Akel, S., Petrow-Sadowski, C., Laughlin, M.J. & Ruscetti, F.W. Neutralization of autocrine transforming growth factor-β in human cord blood CD34+CD38−Lin− cells promotes stem-cell-factor-mediated erythropoietin-independent early erythroid progenitor development and reduces terminal differentiation. Stem Cells 21, 557–567 (2003).

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16. Söderberg, S.S., Karlsson, G. & Karlsson, S. Complex and context dependent regulation of hematopoiesis by TGF-β superfamily signaling. Ann. NY Acad. Sci. 1176, 55–69 (2009). 17. Shav-Tal, Y. & Zipori, D. The role of activin A in regulation of hemopoiesis. Stem Cells 20, 493–500 (2002). 18. Blank, U. & Karlsson, S. The role of Smad signaling in hematopoiesis and translational hematology. Leukemia 25, 1379–1388 (2011). 19. Shiozaki, M. et al. Evidence for the participation of endogenous activin A/erythroid differentiation factor in the regulation of erythropoiesis. Proc. Natl. Acad. Sci. USA 89, 1553–1556 (1992). 20. Zhou, L. et al. Inhibition of the TGF-β receptor I kinase promotes hematopoiesis in MDS. Blood 112, 3434–3443 (2008). 21. Cadena, S.M. et al. Administration of a soluble activin type IIB receptor promotes skeletal muscle growth independent of fiber type. J. Appl. Physiol. 109, 635–642 (2010). 22. Suragani, R.N. et al. Heme-regulated eIF2α kinase activated Atf4 signaling pathway in oxidative stress and erythropoiesis. Blood 119, 5276–5284 (2012). 23. Socolovsky, M. et al. Ineffective erythropoiesis in Stat5a−/−5b−/− mice due to decreased survival of early erythroblasts. Blood 98, 3261–3273 (2001). 24. Liu, Y. et al. Suppression of Fas-FasL coexpression by erythropoietin mediates erythroblast expansion during the erythropoietic stress response in vivo. Blood 108, 123–133 (2006). 25. Chen, K. et al. Resolving the distinct stages in erythroid differentiation based on dynamic changes in membrane protein expression during erythropoiesis. Proc. Natl. Acad. Sci. USA 106, 17413–17418 (2009). 26. Camaschella, C. BMP6 orchestrates iron metabolism. Nat. Genet. 41, 386–388 (2009). 27. McPherron, A.C., Lawler, A.M. & Lee, S.J. Regulation of anterior/posterior patterning of the axial skeleton by growth/differentiation factor 11. Nat. Genet. 22, 260–264 (1999). 28. Wu, H.H. et al. Autoregulation of neurogenesis by GDF11. Neuron 37, 197–207 (2003). 29. Kim, J. et al. GDF11 controls the timing of progenitor cell competence in developing retina. Science 308, 1927–1930 (2005). 30. Loffredo, F.S. et al. Growth differentiation factor 11 is a circulating factor that reverses age-related cardiac hypertrophy. Cell 153, 828–839 (2013). 31. Lin, Y.W., Slape, C., Zhang, Z. & Aplan, P.D. NUP98-HOXD13 transgenic mice develop a highly penetrant, severe myelodysplastic syndrome that progresses to acute leukemia. Blood 106, 287–295 (2005). 32. Paulson, R.F., Shi, L. & Wu, D.C. Stress erythropoiesis: new signals and new stress progenitor cells. Curr. Opin. Hematol. 18, 139–145 (2011). 33. Iancu-Rubin, C. et al. Stromal cell-mediated inhibition of erythropoiesis can be attenuated by Sotatercept (ACE-011), an activin receptor type II ligand trap. Exp. Hematol. 41, 155–166.e17 (2013). 34. Sherman, M.L. et al. Multiple-dose, safety, pharmacokinetic, and pharmacodynamic study of sotatercept (ActRIIA-IgG1), a novel erythropoietic agent, in healthy postmenopausal women. J. Clin. Pharmacol. 53, 1121–1130 (2013).

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Materials. ACE-536 consists of a modified human ActRIIB extracellular domain (residues 24–131) linked to the human IgG1 Fc domain (including the hinge, CH2 and CH3 regions) and was generated as described previously35. Specific modifications in the wild-type ActRIIB extracellular domain are a single amino acid substitution (L79D), an N-terminal truncation of four amino acids and a C-terminal truncation of three amino acids. RAP-536 consists of the same modified human ActRIIB extracellular domain linked to the mouse IgG2a Fc domain and was generated similarly. RAP-536 will be provided for research proposals of scientific merit directed towards the validation and potential extension of the current studies under a mutually agreeable material transfer agreement, subject to the availability of the materials. Wild-type animals. Experimental procedures conducted at Acceleron were performed according to protocols approved by the Acceleron Pharma Institutional Animal Care and Use Committee. Male C57BL/6 mice (2–6 months of age) were obtained from Taconic Laboratories, and timed-pregnant mice serving as the source for fetuses at embryonic (E) days 12.5–14.5 were obtained from Harlan Laboratories. Sprague Dawley rats (males >250 g) were obtained from Harlan Laboratories except for rats (males and females >250 g) used in a 1-month toxicology study performed at Charles River Laboratories. Cynomolgus monkeys (Macaca fascicularis, males and females 2–5 years of age) used in a 1-month toxicology study were obtained from Charles River Laboratories. Administration of ACE-536. The rat and monkey studies presented in Figure 1 were performed and analyzed at Ricerca and Charles River Laboratories, respectively. Experimental procedures conducted at these facilities were performed according to protocols approved by the Institutional Animal Care and Use Committee for Ricera and the Institutional Animal Care and Use Committee for Charles River Laboratories, Preclinical Services, Reno, Nevada. All other studies were performed at Acceleron. Mice, rats and monkeys were treated with ACE-536, EPO (Henry Schein, Melville, NY; cat. #028881), neutralizing antibody against EPO (R&D Systems, Minneapolis, MN; cat. #MAB959; 5.5 mg kg−1), GDF11 (generated and purified in-house) or vehicle (PBS or Tris-buffered saline) by either i.p. or s.c. injection as indicated. Body weights were measured with an electronic scale at the time of dosing. At study termination, mice were euthanized by CO2 asphyxiation, and bone marrow, spleens, kidneys, whole blood and serum were collected. Hematologic studies. The submandibular vein of mice was pierced by a sterile lancet, and 100 µl of blood was collected in an EDTA-coated microtainer tube. Terminal blood samples were taken from euthanized mice via cardiac puncture. Complete blood counts were determined using a VetScan HM2 Hematology System (Abaxis) or an Advia 120 Automated Hematology Analyzer (Bayer). Blood samples from rats in the 1-month toxicology study were collected into EDTA-containing tubes via the retro-orbital plexus under CO2/O2 anesthesia. Blood samples from cynomolgus monkeys were collected by venipuncture into tubes containing EDTA. Serum samples were analyzed using automated hematology analyzers. Reticulocyte counts in peripheral blood were determined by staining with thiazole orange and flow cytometry. Anemia models. For anemia induced by blood loss, Sprague Dawley rats were anesthetized by isoflurane, and 20% of total blood volume as determined by body weight36 was removed via cannulated jugular vein and replaced with PBS, leading to a 15% drop in RBC count. For chronic renal insufficiency induced by partial nephrectomy, at 10 weeks of age, male C57BL/6 mice were partially (five-sixths) nephrectomized or received a sham operation at Taconic Laboratories. Mice arrived at Acceleron 6 d later and were allowed to acclimate to the animal facility for 3 d before baseline blood sampling via the submandibular vein (7 d before dosing). Metal wound clips were removed at the time of the first dose, and mice were assessed for proper wound healing. For chemotherapy-induced anemia, Sprague Dawley rats were anesthetized by isoflurane and received a single injection of PBS or carboplatin (75 mg kg−1) via cannulated jugular vein on day 0; they were then treated with ACE-536 or vehicle (TBS) beginning on day 11.

doi:10.1038/nm.3512

Mouse model of myelodysplastic syndromes. NUP98-HOXD13 MDS mice31 and age-matched FVB wild-type mice were obtained from P. Aplan’s laboratory at the US National Institutes of Health and maintained at Taconic. Experiments with these mice were approved by the Acceleron Pharma Institutional Animal Care and Use Committee. Disease severity increases with age, eventually progressing to acute myeloid leukemia in the majority of mice, and the mice have a mean lifespan of approximately 14 months. Starting at approximately 4 months of age, mice exhibit anemia, leukopenia, ineffective erythropoiesis and bone marrow that is normocellular to hypercellular31. To monitor the effects of chronic administration, MDS mice were treated with RAP-536 (10 mg kg−1, s.c., twice weekly) or vehicle beginning at 4 months of age and continuing for 7 months, and blood samples (50 µL) were collected at baseline and monthly thereafter for complete blood count analysis. Activin-null mice. Mice with homozygous loss-of-function mutations in Inhbc (Inhbctm1Zuk)37, which encodes activin C, and Inhbe (Inhbctm2Zuk)37, which encodes activin E, were obtained from M. Matzuk, Baylor College of Medicine. Age-matched wild-type mice were used as controls. Mice at 6–11 weeks of age were treated with RAP-536 (10 mg kg−1, s.c.) twice weekly for 1 week. Blood samples collected after twice weekly treatment for 1 week were subjected to complete blood count analysis. Neutralization of activin A. Neutralizing antibody against human activin A was generated in-house, validated by surface plasmon resonance and cell-based inhibition assays (data not shown) and administered to mice at a dose of 10 mg kg−1. The elimination half-life of the antibody was determined in vivo to set the dosing frequency (data not shown). Male C57BL/6 mice at 6–8 weeks of age were pretreated s.c. with antibody against activin A (10 mg kg−1) or vehicle (PBS) three times weekly for 1 week. Vehicle-pretreated mice were then treated with RAP-536 (10 mg kg−1, s.c.) twice weekly for 1 week, and antibody-pretreated mice were continued on antibody treatment (10 mg kg−1, three times weekly, s.c.) for 1 more week or treated for 1 more week with antibody (10 mg kg−1, three times weekly, s.c.) and RAP-536 (10 mg kg−1, twice weekly, s.c.) in combination. Blood samples obtained at study termination were subjected to complete blood count analysis. Erythroid burst-forming units and colony-forming units. Male mice 6–8 weeks of age were administered ACE-536 (10 mg kg−1, s.c) or vehicle (TBS). On day 2, mice were killed by CO2 asphyxiation and peripheral blood and the spleen, tibias and femurs were collected. Peripheral blood was used for complete blood counts. Spleen cells and bone marrow cells were isolated and plated in methylcellulose-based media to assess clonogenic progenitors of the erythroid lineage. For assessment of CFU-Es, bone marrow and spleen cells were cultured at 2.5 × 104 and 2.3 × 104 cells/ml, respectively, in methylcellulose medium containing rmEPO (Methocult M3334, Stem Cell Technologies), and colonies were counted independently by four investigators on day 2. For assessment of BFU-Es, bone marrow and spleen cells were cultured at 5.0 × 103 and 2.3 × 104 cells/ml, respectively, in methylcellulose medium containing rmSCF, rmIL-3, rmIL-6 and rmEPO (Methocult M3434, Stem Cell Technologies), and colonies were counted similarly on day 12. Profiling of erythroid differentiation. Erythroid differentiation was monitored with mouse-specific antibodies (BD Biosciences) against CD71 (transferrin receptor) (PE-conjugated, cat. #553267, 1:100 dilution; or FITCconjugated, cat. #553266, 1:200 dilution), CD44 antigen (APC-conjugated, cat. 559250, 1:200 dilution) and Ter-119 (glycophorin-A–associated protein) (PE-conjugated, cat. #553673, 1:100 dilution; or APC-conjugated, cat. #557909, 1:100 dilution) and analyzed by flow cytometry as described previously22,24,25. Briefly, 1 × 106 cells from bone marrow or spleen were suspended in Iscove’s medium containing 5% FBS and incubated with FITC- or APC-labeled antibody against CD71 and CD44 and PE-labeled antibody against Ter-119 (BD Biosciences) for 20 min at 37 °C before blocking nonspecific sites with mouse Fc blocker (BD Biosciences). Cells were washed twice with PBS containing 5% FBS, and samples were analyzed on a flow cytometer (BD Biosciences FACSCalibur). Dead cells in the samples were excluded from analysis by counterstaining with propidium iodide (BD Biosciences). The percentages

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of CD71/CD44- and Ter-119-positive cells were gated and data recorded. Differential sorting of erythroblasts was carried out on a MoFlo cell sorter at the Tufts Laser Cytometry and Core Facility at the Tufts University School of Medicine. Cytospin and benzidine Giemsa staining of cells for morphological analysis was carried out as described previously22. Morphological images were obtained with a Nikon Eclipse 80i microscope fitted with a 100× oil-immersion objective and processed with NIS-Elements imaging software (Nikon). Isolation of erythroid precursors. Mouse fetal liver cells consisting almost exclusively of erythroid precursors were isolated from E12.5–E14.5 C57BL/6 embryos as described22. In brief, cells were sorted to obtain immature (Ter-119−) and mature (Ter-119+) erythroid precursor populations by labeling with a multilineage biotinylated antibody cocktail (BD Biosciences) followed by biotinylated antibody against Ter-119 (BD Biosciences; cat. #553672, 1:200 dilution) according to a two-step purification procedure with an Easy Sep magnetic separation kit (Stem Cell Technologies, cat. #18556). Splenocytes were obtained from wild-type mice treated with EPO (2,000 units kg −1) for 24 h. Mature RBCs in the cell suspension were lysed with RBC cell lysis buffer (Sigma, R7757) according to the manufacturer’s instructions. The remaining cells were cultured in serum-free SP34 expansion medium supplemented with EPO (2.5 U/ml), mouse stem cell factor (100 ng/ml), insulin-like growth factor-1 (40 ng/ml), dexamethasone (1 µM), β-estradiol (1 µM), holotransferrin (200 µg/ml), 1% BSA and β-mercaptoethanol (100 µM) with partial medium changes as described previously38. After 5 d in culture, expanded Ter-119− cells were separated using a Ter-119 separation kit, transferred into serum-free SP-34 suboptimal differentiation medium supplemented with EPO (2 U/ml), holotransferrin (100 µg/ml) and β-mercaptoethanol (100 µM), and seeded at a cell density of 3.5 × 105 cells/ml in fibronectin-coated culture dishes (BD Biosciences). Cells were treated with GDF11 (25 ng/ml) for 48 h with or without ACE-536 (25 µg/ml). Determination of ligand binding affinities and inhibition. The ligand binding profile of ACE-536 was determined by surface plasmon resonance as described35. Potency of ligand inhbition by ACE-536 was determined in gene reporter assays. The effects of ACE-536 on ligand signaling through Smad2/3 were tested with A204 cells transfected with a pGL3-CAGA12luciferase reporter construct generated in-house as described35. Similarly, A549 cells transfected with this reporter construct were used to evaluate effects of ACE-536 on TGF-β1 signaling39. The effects of ACE-536 on ligand signaling through Smad1/5/8 were tested with HepG2 (ref. 40) or T98G (ref. 41) cells transfected with a BMP response element (BRE4)-luciferase reporter construct generated in-house as described35. Quantitation of Smad2/3 protein. C57BL/6 mice were treated with EPO (1,800 units kg−1, i.p.) to initiate erythroid expansion as described24 and treated 40 h later with RAP-536 (10 mg kg−1, i.p.) or vehicle (TBS). Mice were euthanized 6 h after treatment with RAP-536 or vehicle (n = 2 per group), and splenic cell extracts were prepared with radioimmunoprecipitation assay lysis buffer (Boston Bioproducts, DT-115D) containing 1× phosphatase inhibitor and protease inhibitors (Sigma, P-8340). Cell extracts (100 µg) were separated by 12% SDS-PAGE and analyzed by western blotting as described22. In brief, membranes were first probed with antibody against phosphorylated Smad2/3 (Santa Cruz Biotechnology, sc-11769-R, 1:750 dilution), then stripped with Restore buffer (Pierce) and reprobed with antibody against Smad2 (Cell Signaling Technology, cat. #3103, 1:1,000 dilution) to control for total protein loading. Primary antibodies were detected with horseradish peroxidase– conjugated secondary antibody against rabbit IgG (Santa Cruz Biotechnology, sc-2030, 1:5,000 dilution) and chemiluminescence (Pierce). For ex vivo experiments, erythroid precursors isolated from mouse fetal liver were cultured overnight in serum-free SP34 medium (Invitrogen) with EPO (2 U/ml), insulin (10 µg/ml) and holotransferrin (200 µg/ml). Cells were treated with GDF11 for 1 h or 3 h with and without ACE-536 (1 µg/ml). Levels of Smad2 phosphorylation were determined using a Phospho-Smad2 (Ser465/467) Sandwich ELISA Kit (Cell Signaling Technology, cat. #7348) according to the manufacturer’s instructions.

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Immunohistochemistry. Immunohistochemical staining was performed at the Histopathology and Tissue Shared Resource Facility at the Lombardi Comprehensive Cancer Center, Georgetown University. Briefly, 5-µm-thick sections from formalin-fixed, paraffin-embedded tissues were deparaffinized with xylenes and rehydrated through a graded-alcohol series. Heatinduced epitope retrieval was performed by immersing the tissue sections in 10 mM citrate buffer (pH 6.0) with 0.05% Tween at 98 °C for 20 min. Immunohistochemical staining was performed using a VectaStain Kit (Vector Labs) according to the manufacturer’s instructions. Briefly, slides were treated with 3% hydrogen peroxide, avidin and biotin blocking solution (Invitrogen) and 10% normal (goat) serum and exposed to primary antibodies against GDF11 (Abcam, ab124721, 1:60 dilution), Ter-119 (BD Biosciences, cat. #553669, 1:100 dilution) or p-Smad 2/3 (Abcam, ab52903, 1:20 dilution) overnight at 4 °C. Slides were exposed to biotin-conjugated anti-rabbit secondary antibodies (Vector Labs), Vectastain ABC reagent and DAB chromagen (Dako). Slides were counterstained with hematoxylin (Harris modified hematoxylin, Fisher), treated with 1% ammonium hydroxide, dehydrated and mounted with Acrymount. Alternate sections with primary antibody omitted were used as negative controls. Images were obtained with a Nikon Eclipse 80i microscope fitted with 10×, 20× or 100× oil-immersion objectives and processed with NIS-Elements imaging software (Nikon). Differential cell counts in bone marrow smears. Femurs were opened with a scalpel blade, and smears were prepared by gently wiping the marrow with a small brush moistened with PBS containing 5% FBS. Staining and evaluation of bone marrow smears and differential cell counts were carried out by RADIL Services (Columbia, MO). Expression profiling by PCR array. mRNA isolated from immature (Ter-119−) and mature (Ter-119+) erythroid precursors from fetal mouse liver was analyzed by expression profiling of 84 genes involved in TGF-β superfamily signaling (using the Mouse TGF-β/BMP Signaling Pathway PCR Array, SA Biosciences, PAMM-035Z), and 84 target genes responsive to TGF-β superfamily signaling (using the Mouse TGF-β Signaling Targets PCR Array, SA Biosciences, PAMM-235Z). Each 96-well array plate was assayed using 500 ng of input cDNA (5.2 ng of cDNA per well). Reactions and data analyses were carried out according to the manufacturer’s instructions. Data were normalized using Gapdh mRNA as control, and gene expression differences were determined by the 2−∆∆Ct method according to the manufacturer’s protocol. Quantitative real-time PCR. Kidney or liver tissue was disrupted by homogenization (Tissue Tearor, BioSpec Products) and total RNA was extracted with a Ribopure Kit (Ambion) according to the manufacturer’s instructions. Nucleotide concentration was determined with a NanoDrop 1000 spectrophotometer (Thermo Scientific). Real-time PCR reactions consisted of 100 ng total RNA and primers, Eukaryotic 18S rRNA Endogenous Control (Applied Biosystems), and the TaqMan RNA-to-CT 1-Step Kit (Applied Biosystems). Amplifications were performed in a 7300 Real-Time PCR System (Applied Biosystems). Taqman primers for EPO, hepcidin, BMP6, GDF11, ActRIIB, Gata-2 and β-globin were purchased from Life Technologies (Epo: Mm01202755_m1; Hamp: Mm00519025_m1; Bmp6: Mm01332882_m1; Gdf11: Mm01159973_m1; Acvr2b: Mm01348449_m1; Gata2: Mm00492301_ m1; Hbb: Mm00731743_m1). Measurement of creatinine and urea nitrogen. Creatinine and blood urea nitrogen were quantified using a VetScan HM2 Hematology System (Abaxis). Quantitation of red blood cell life span. RBC life span was determined as described42 with minor modifications. Circulating RBCs were biotinylated in vivo by five i.p. injections of 500 µl EZ-Link Sulfo-NHS-Biotin (Pierce Biotechnology) in sterile water (one injection of 8 mg/ml concentration on day −2 and four 4 mg/ml injections on day −1 spaced 2.5 h apart). On day 0, animals were randomly assigned to treatment with vehicle (TBS) or RAP-536 (10 mg kg−1, s.c., twice weekly; n = 5 per group). On day 0 and weekly

doi:10.1038/nm.3512

control subjects (n = 5) was obtained from Tissue Solutions Ltd. (Glasgow, Scotland) from hospitals in the United States according to IRB-approved protocols and with IRB-approved consent forms for molecular and genetic analysis. GDF11 levels were measured using an ELISA kit (MyBioSource Inc., San Diego, CA, MBS939778) according to the manufacturer’s instructions. Each sample was diluted 1:2 in sample diluent supplemented with protease inhibitor cocktail (Sigma-Aldrich).

Detection of apoptosis. Apoptosis in erythroid precursors was determined by flow cytometry using an apoptosis detection kit (BD Biosciences, cat. #556547). Erythroid precursors in bone marrow were stained with fluorophore (APC)-conjugated anti-Ter-119 antibodies and counter stained with annexin V–specific antibody conjugated to FITC (BD Biosciences, cat. #556547, 1:100 dilution) according to the manufacturer’s instructions. Late-stage apoptotic and necrotic cells in the samples were excluded from the analysis by counterstaining with propidium iodide. The percentages of annexin V+ and Ter-119+ cells were gated and the data were recorded.

Statistical analyses. Data are reported as means ± s.e.m. unless indicated otherwise. Non-normally distributed variables were log-transformed so that data were normally distributed before statistical analysis. Responses to multiple ACE-536 doses were compared by one-way ANOVA followed by Dunnett’s post hoc analysis for multiple comparisons. Rate of recovery from anemia induced by blood loss or chemotherapy was assessed by two-way ANOVA for repeated measures, comparing the interaction of the main effects of treatment and time, with post hoc analysis by Tukey’s honest significant difference test for multiple comparisons. Otherwise, data were analyzed by nonpaired Student’s t-test for between-group comparisons.

Quantitation of reticulocytes. Reticulocyte counts in peripheral blood from rats were measured by flow cytometry after staining with thiazole orange (Sigma Aldrich). Absolute counts of reticulocytes were determined using the following formula: absolute reticulocyte count = % reticulocytes × 10 −2 × RBC × 1012 l−1. Cell cycle analysis. Mice were injected i.p. with BrdU from a BD Biosciences Flow kit (cat. #552598) 1 h before they were killed. Erythroid precursors were obtained from bone marrow and spleen and incubated with antibodies against CD71 and Ter-119 as described43 with minor modifications. BrdU-labeled cells were detected with a fluorescently labeled antibody from the BD Biosciences Flow kit and stained with 7-AAD according to the manufacturer’s instructions. Cells were then analyzed by flow cytometry as described43. Administration and quantitation of GDF11. Purified recombinant mouse GDF11 was generated in-house and administered to wild-type mice (0.1 mg kg−1, i.p., daily) as described previously30 for 11 d. Serum from male and female subjects with MDS (n = 8, 67–91 years of age) and age-matched normal

35. Sako, D. et al. Characterization of the ligand binding functionality of the extracellular domain of activin receptor type IIb. J. Biol. Chem. 285, 21037–21048 (2010). 36. Lee, H.B. & Blaufox, M.D. Blood volume in the rat. J. Nucl. Med. 26, 72–76 (1985). 37. Lau, A.L., Kumar, T.R., Nishimori, K., Bonadio, J. & Matzuk, M.M. Activin βC and βE genes are not essential for mouse liver growth, differentiation, and regeneration. Mol. Cell. Biol. 20, 6127–6137 (2000). 38. Menon, M.P. et al. Signals for stress erythropoiesis are integrated via an erythropoietin receptor-phosphotyrosine-343-Stat5 axis. J. Clin. Invest. 116, 683–694 (2006). 39. Ignotz, R.A. & Honeyman, T. TGF-β signaling in A549 lung carcinoma cells: lipid second messengers. J. Cell. Biochem. 78, 588–594 (2000). 40. Scharpfenecker, M. et al. BMP-9 signals via ALK1 and inhibits bFGF-induced endothelial cell proliferation and VEGF-stimulated angiogenesis. J. Cell Sci. 120, 964–972 (2007). 41. Townson, S.A. et al. Specificity and structure of a high affinity activin receptor-like kinase 1 (ALK1) signaling complex. J. Biol. Chem. 287, 27313–27325 (2012). 42. Roy, C.N. et al. Hepcidin antimicrobial peptide transgenic mice exhibit features of the anemia of inflammation. Blood 109, 4038–4044 (2007). 43. Koulnis, M. et al. Identification and analysis of mouse erythroid progenitors using the CD71/TER119 flow-cytometric assay. J. Vis. Exp. 54, e2809 (2011).

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thereafter, 2.5 µl blood samples were collected via the tail vein in 500 µl of PBS containing 5% BSA. Cells were centrifuged at 300g for 5 min at 4 °C, and pellets were resuspended in 500 µl of PBS containing 5% BSA. AlexaFluor 488–conjugated streptavidin (Invitrogen, cat. #S-11223) was added to a final concentration of 1 mg/ml and incubated for 30 min at 4 °C. Cells were then centrifuged at 300g for 5 min at 4 °C. Pelleted cells were resuspended in 500 µl of PBS containing 5% BSA and analyzed by flow cytometry.

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