Tracking erythroid progenitor cells in times of need and times of plenty

Experimental Hematology 2015;-:-–-

Tracking erythroid progenitor cells in times of need and times of plenty Mark J. Koury Division of Hematology/Oncology, Vanderbilt University School of Medicine, Nashville, Tennessee (Received 11 August 2015; revised 22 September 2015; accepted 28 October 2015)

Red blood cell production rates increase rapidly following blood loss or hemolysis, but the expansion of erythropoiesis in these anemic states is tightly regulated such that rebound polycythemia does not occur. The erythroid cells that respond to erythropoietic stimulation or suppression are the progenitor stages of burst-forming units–erythroid (BFU-Es) and colony-forming units–erythroid (CFU-Es). Results from an early study of the changes in the size, location, and cell cycling status of BFU-E and CFU-E populations in mice under normal conditions, erythropoietic stimulation, and erythropoietic suppression are used as reference points to review subsequent developments related to erythroid progenitor populations and regulation of their size. The review concerns development of erythroid progenitor populations mainly in mice and humans, with a focus on the mechanisms related to the rapid but highly regulated expansion of erythropoiesis in spleens of erythropoietically stimulated mice. Current knowledge is used as a model of erythroid progenitor populations in mice under normal, erythropoietically suppressed, and erythropoietically stimulated conditions. Clinical applications of information learned from studies of erythropoietic expansion, in terms of current therapies for anemia, are reviewed. Copyright Ó 2015 ISEH - International Society for Experimental Hematology. Published by Elsevier Inc.

Erythrocytes (RBCs) carry oxygen from the lungs to the other tissues, enabling aerobic cellular respiration. Numbers of circulating RBCs in the blood have a narrow range, with daily turnover rates of about 1%, as the oldest RBCs are recognized as senescent and removed by macrophages, while a similar number of their replacements, the reticulocytes, enter the blood from the bone marrow. Normally, humans produce is about 2.0–2.5
1011 RBCs per day, but when RBCs are lost from bleeding or destroyed by hemolysis, the resultant anemia causes tissue hypoxia, leading to rapid expansion of marrow erythropoiesis. In mice, acute anemia increases erythropoiesis mainly in the spleen because of limited space for erythroid expansion in the marrow. This ‘‘stress’’ erythropoiesis in times of need for more RBCs is regulated largely by erythropoietin (EPO), the renal hormone that is produced in small amounts under normal conditions, but increases rapidly in response to kidney hypoxia. The erythropoietic response in times of need is highly regulated such that overshoot polycy-

Offprint requests to: Mark J. Koury, Division of Hematology/Oncology, Vanderbilt University School of Medicine, 777 Preston Research Building, Nashville, TN 37232-6307; E-mail: [email protected]

themia does not occur as recovery to baseline normal numbers of RBCs, a time of plenty, is achieved. In the rare cases of more than plenty RBCs, such as when an individual with polycythemia from acclimatization to high altitudes travels to sea level, erythropoiesis is suppressed until a smaller normal number of RBCs is achieved. Regulation of the erythropoietic process occurs mainly in the erythroid progenitor stages of differentiation. Examining the sizes and locations of these progenitor populations in times of need, times of plenty, and times of more than plenty, as reported nearly 40 years ago in Experimental Hematology [1], has advanced understanding of multiple factors that regulate RBC production and has helped in the development of clinical therapies for anemias. In 1977, Hiroshi Hara and Makio Ogawa published a study that examined the changes in erythroid progenitor cells, colony-forming units–erythroid (CFU-Es) and burstforming units–erythroid (BFU-Es), in mice under normal conditions, reduced erythropoietic demand, and increased erythropoietic stress [1]. This report, like several others that followed soon after the original descriptions of CFUEs [2] and BFU-Es [3] from Axelrad’s laboratory, indicated that erythropoietic stress increased CFU-E numbers modestly in the bone marrow and markedly in the spleen,

0301-472X/Copyright Ó 2015 ISEH - International Society for Experimental Hematology. Published by Elsevier Inc. http://dx.doi.org/10.1016/j.exphem.2015.10.007

2

M.J. Koury/ Experimental Hematology 2015;-:-–-

whereas BFU-E numbers decreased about twofold in marrow and increased twofold in the spleen. Suppression of erythropoietic demand had opposite effects on erythroid progenitor numbers in these two organs. The new information reported by Hara and Ogawa included: (i) BFU-Es normally found in the blood increased with erythropoietic stress and decreased with erythropoietic suppression. (ii) Despite the large differences in numbers of BFU-Es and CFU-Es in normal, erythropoietically stressed, or erythropoietically suppressed mice, the cell cycle status of BFUEs and CFU-Es was similar under all conditions in spleen and marrow [1]. These results indicated that BFU-Es migrate in the blood of mice from bone marrow to spleen, erythropoietic stress increases and erythropoietic suppression decreases BFU-E migration, and marked expansion of CFU-Es during erythropoietic stress involves more than simply an increased rate of erythroid cell proliferation. This review discusses some aspects of erythroid progenitor populations and their changes in response to erythropoietic demand that have been published since Hara and Ogawa’s publication and that may help explain their original results and provide insights into the development of new approaches in treating anemia.

Erythropoietic progenitor cells In the few years before Hara and Ogawa’s publication, knowledge about erythropoiesis had advanced rapidly following the characterization of two basic erythroid progenitor stages that precede microscopically identifiable erythroblasts, the CFU-E and the more immature BFU-E. These murine erythroid progenitors were detected by their growth, in semisolid or viscous medium containing EPO, into colonies of 8 to 64 morphologically identifiable erythroblasts after 2 days for CFU-Es [2] or much larger bursts or multiple colonies of erythroblasts after 8–14 days for BFU-Es [3]. Over the next few years, multiple studies by Eaves and colleagues characterized mouse and human CFU-Es, BFU-Es, and mature BFU-Es, intermediates with properties between those of the immature BFU-Es and CFU-Es [4]. Most of the properties of mouse and human erythroid progenitors were similar except for a relatively prolonged period of development in vitro for colonies (7–9 days) and immature bursts (17–20 days) of humans compared with those of mice [4]. Within a few years of their description, immature BFUEs were found to have a close relationship with the megakaryocytic lineage, as 40% of BFU-Es displayed both megakaryocytic and erythroid potential [5]. Subsequent studies reported that later-stage erythroid progenitors, including those in spleens of mice with phenylhydrazine (PHZ)-induced hemolytic anemia [6] and those isolated from human blood [7], retain megakaryocytic potential that is either lost or retained depending on their exposure in vitro to erythropoietic or thrombopoietic conditions.

These bipotent megakaryocyte–erythroid progenitors (MEPs) differentiate along the erythroid lineage when increased expression of transcription factors that induce erythroid differentiation, including GATA1 [8], KLF1 [9,10], and LMO2 [10], are accompanied by decreased expression of transcription factors that induce megakaryocytic differentiation, including KLF1 antagonism of FLI1 [9] and MYB-induced miR486-3p suppression of MAF translation [11]. Thus, erythroid progenitor cells were defined as a continuum beginning with immature BFUEs, which can be restricted to erythroid differentiation despite their megakaryocytic potential, and ending with CFU-Es. The immediate progeny of the CFU-Es, the proerythroblasts (ProEBs), are the first morphologically recognized stage of erythroid differentiation. Based on their appearance on Giemsa staining during terminal differentiation, ProEBs pass through basophilic, polychromatophilic, and orthochromatic erythroblast stages as they accumulate hemoglobin before enucleating to form reticulocytes. Erythroblasts in the terminal erythroid stages, termed erythroid precursor cells, are now often identified by flow cytometry-based surface expression of specific proteins, glycoproteins, and/or cell size [12–14]. However, the transition between the operationally defined CFU-E and microscopically identified ProEB stages has been uncertain. Overlap in the CFU-E and ProEB populations is possible, as some ProEBs may complete three or four generations of cell divisions, giving rise to a colony of 8–16 cells in the CFU-E assay. Therefore, in some situations, these two closely related and potentially overlapping populations will be designated as CFU-E/ProEBs.

Changes in erythroid progenitor populations with varying erythropoietic demand More than a decade before the descriptions of the erythroid progenitor cells, Erslev noted that mitotic divisions in the erythroid precursors and the ratios of ProEBs to later stages of precursors were the same under conditions of suppressed and stimulated erythropoiesis [15]. He concluded that erythropoiesis was controlled by either the rate of hematopoietic stem cell (HSC) differentiation into ProEBs or the rate of ProEB replication [15]. Although bleeding has been reported to increase proliferation and self-renewal of phenotypically sorted HSCs [16], and EPO administration has been reported to influence differentiation of multipotent HSC progeny toward the erythroid lineage [17], the vast majority of studies have focused on the effects of erythropoietic stress or suppression on the erythroid progenitor cells. The importance of EPO in vivo as the principal hypoxiainducible stimulator of erythropoietic responses was well recognized before identification of the CFU-E and the BFU-E [18]. Administration of an EPO preparation to

M.J. Koury/ Experimental Hematology 2015;-:-–-

mice increased erythroblast populations in the marrow, especially in the spleen, but no change in the radiolabeled mitoses in the populations could be detected, and a slightly shortened S phase calculated by double radiolabeling could not account for large increases in the erythroblast populations [19]. The requirement for EPO in the media of CFU-E and BFU-E assays, the restriction of the effects of EPO to hematopoietic cells of the erythroid lineage, and the absence of an effect of high-EPO states on the proliferation of a specific stage of erythroblast differentiation indicated that the target cell populations for EPO’s regulation of erythropoiesis were very likely the BFU-Es and/or CFU-Es. Therefore, several investigators examined the effects of increased or decreased EPO levels in vivo on the numbers, location, and proliferation status of BFU-E and CFU-E populations [1,3,20–23]. Marrow and spleen were hematopoietic sites examined in these investigations, but Hara and Ogawa, who had established that BFU-Es, but not CFU-Es, circulated in the blood [24], also examined BFU-Es in the blood [1]. About the same time as Hara and Ogawa’s publication, the purification of EPO was reported [25], but several more years were required before the cloning of EPO and availability of large quantities of purified EPO. Therefore, in addition to using partially purified EPO preparations, investigators frequently increased endogenous EPO by inducing hemolytic anemia or blood-loss anemia. They suppressed endogenous EPO by hypertransfusing erythrocytes or creating a posthypoxia condition. Hara and Ogawa had previously found that PHZ-induced hemolytic anemia in mice increased CFU-Es modestly in the marrow and markedly in the spleen, whereas BFU-Es decreased modestly in the marrow and increased modestly in the spleen [24]. Importantly, they reported that the splenic increases in BFU-Es were preceded by increases in the normally low levels of circulating BFU-Es [24]. However, PHZ causes direct oxidant stress in erythroid progenitors, as well as inducing hemolytic anemia. Thus, confirmation of increased circulating BFU-Es with administration of an EPO preparation or with blood loss provided evidence that the increase in splenic BFU-Es and the maintenance of elevated splenic CFU-Es at later times in erythropoietically stressed mice were potentially related to an influx of migrating BFU-Es, which likely originated in the marrow, where BFU-E numbers declined [1]. Hypertransfusioninduced polycythemia had the opposite effects of acute anemia on BFU-E and CFU-E populations in marrow, spleen, and blood, indicating that EPO very likely mediated these changes [1]. Decreases in marrow BFU-Es during stimulated erythropoiesis and increases in marrow BFU-Es during suppressed erythropoiesis were about twofold compared with the normal baseline numbers. However, the changes in CFU-Es were severalfold in the marrow and more than an order of magnitude in the spleen. Other early investigations reported similar patterns in the BFU-

3

E and/or CFU-E populations in murine marrow and spleen with increased or decreased erythropoiesis [3,20–23]. The prominent role of the spleen in mice during erythropoietic stress is due to the rapid increase in CFU-Es and their progeny, but Hara and Ogawa’s demonstration that circulating BFU-E numbers increased with erythropoietic stress provided a mechanism for sustaining the expanded erythropoiesis.

Mobility, proliferation, and differentiation of BFU-Es The relationship between circulating BFU-Es, their homing to the hematopoietic tissues, and their subsequent differentiation to the CFU-E stage in hematopoietic tissues remains relatively unexamined compared with the later differentiation stages of CFU-Es through reticulocytes. The prominence of splenic stress erythropoiesis in mice has facilitated investigation into the response to erythropoietic stimulation, as compared with studies with humans in which the marrow is the site of both normal and stress erythropoiesis. In the same year as Hara and Ogawa’s article, BFU-Es were reported to circulate in human blood [26,27]. The circulating BFU-Es lodge in the marrow through an interaction of p67 non-integrin laminin-binding protein expressed on their surface with non-integrin laminins expressed by the bone marrow stroma [28]. Table 1 lists growth factors and adhesion proteins that play a role in localization, proliferation, and differentiation of erythroid progenitor and precursor cells during normal and stress erythropoiesis. Stem cell factor (SCF) expression on the surface of stromal cells plays a key role in the lodging of circulating BFU-E in both marrow and spleen following PHZ-induce anemia [29]. KIT, the surface transmembrane receptor for SCF, has intrinsic kinase activity, which, on binding of SCF (in either the fixed stromal form or soluble form), leads to activation of several intracellular pathways that regulate survival, proliferation, and differentiation of both BFU-Es and CFU-Es. These pathways include those involving RAS, Raf-1, mitogen-associated protein kinase (RAS-Raf-MAPK); phosphatidylinositol-3 kinase, protein kinase B (PI3K-AKT); and phospholipase Cg, protein kinase C, inositol trisphosphate (PLC-PKC-IP3) [30]. The importance of integrins containing the b1 component for interactions of BFU-Es with their microenvironment was illustrated in b1-integrin knockout mice by decreases in splenic BFU-E populations, with essentially no increase in response to erythropoietic stress [31]. During their differentiation into CFU-Es, the interactions of BFU-Es with the hematopoietic microenvironment are not well characterized, but several regulatory factors are known. These regulatory factors and the erythroid progenitors that they affect are illustrated in Figure 1 for mice in states of normal, suppressed, and stimulated erythropoiesis. Eaves and her colleagues helped identify factors in BFU-E development under normal and

M.J. Koury/ Experimental Hematology 2015;-:-–-

4

Table 1. Growth factors and adhesion proteins that regulate normal and stress erythropoiesis Growth factor/adhesion protein

Source

Erythroid progenitor/ precursor

SCF (KIT ligand)

Hematopoietic stromal cells

BFU-E and CFU-E

Non-integrin laminins

Hematopoietic stromal cells Hematopoietic stromal cells Hematopoietic stromal cells Plasma (adrenal cortex) Plasma (renal cortex)

BFU-E

b1-integrins and binding partners Hedgehog and BMP4 Glucocorticoid hormones EPO

FAS ligand

Erythroblasts

Gas6

Erythroblasts and hematopoietic stromal cells EBI central macrophages EBI central macrophages EBI central macrophages EBI central macrophages EBI central macrophages Hematopoietic stromal cells

VCAM-1 av-Integrins Erythroblast-macrophage protein CD169 (Siglec1) CD163 TGF-b superfamily ligands

BFU-E BFU-E BFU-E and CFU-E CFU-E, ProEB, early BasoEB CFU-E, ProEB, early BasoEB ProEBs and differentiating EBs CFU-E, ProEBs, and differentiating EBs CFU-E, ProEBs, and differentiating EBs CFU-E, ProEBs, and differentiating EBs CFU-E, ProEBs, and differentiating EBs CFU-E, ProEBs, and differentiating EBs Differentiating erythroblasts

erythropoietically stressed conditions using W/Wv mice, which are chronically anemic and have deficiencies in Kit, the SCF receptor, and f/f mice, which are impaired in responding to erythropoietic stress and have a deficiency of Smad5, a signaling pathway protein for bone morphogenetic proteins [20,32]. Numbers of bone marrow and spleen erythroid progenitors from immature BFU-E through CFUE stages are decreased in W/Wv mice and normal in f/f mice [32]. SCF was found to be required for erythroid differentiation from the BFU-E through CFU-E stages in both mice [29] and humans [33]. After EPO administration to irradiated or hypertransfused f/f mice, splenic generation of transient splenic erythropoietic foci in vivo was absent, and CFU-E numbers were diminished, suggesting deficiency in a pre-CFU-E stage of erythroid progenitor that expanded during erythropoietic stress [20]. In erythropoietically stimulated mice, a population of splenic erythroid progenitors in a stage prior to CFU-Es and, corresponding to the population of BFU-Es absent in f/f mice, has been characterized [34]. These progenitors, termed stress BFU-Es, required several factors in addition to SCF for rapid growth and differentiation into CFU-Es and mature erythroblasts over a period of 5 to 7 days. After recovery from acute

Erythroid cell receptor/ signaling KIT/PI3K-AKT RAS-Raf-MAPK PLC-PKC-IP3 p67 Non-integrin- binding protein b1-Integrins and binding partners BMP4 receptor/Smad5

Effect on erythroid cells Lodgment in hematopoietic tissue; Survival and proliferation Lodgment in hematopoietic tissue Proliferation during stress erythropoiesis Proliferation during stress erythropoiesis

Glucocorticoid receptor/PPAR-a EPO-R-JAK2/Stat5 RAS-Raf-MAPK PI3K-AKT FAS-Caspases

Proliferation during stress erythropoiesis

Survival inhibition and differentiation

TAM receptors (Axl)/ GATA1

Survival and differentiation during stress erythropoiesis

a4b1-Integrin

Adherence to EBI central macrophages

ICAM-4

Adherence to EBI central macrophages

Erythroblast-macrophage protein Sialylated glycoproteins

Adherence to EBI central macrophages Adherence to EBI central macrophages

Unknown

Adherence to EBI central macrophages

TGF-b superfamily receptors/Smad2,3

Differentiation inhibition

Survival and differentiation

anemia, stress BFU-E populations in spleens of mice require repletion from a source in the marrow [34]. Bone morphogenetic protein 4 (BMP4) is the principal growth factor required by stress BFU-Es and provided by the splenic hematopoietic stroma in mice [34] and, presumably, by the marrow in humans. In addition to BMP4 and SCF, growth differentiation factor 15 (GDF15) and a hedgehog protein appear to be required for a maximal response in the development of stress BFU-Es in both mice and humans [35]. A mechanism for the expansion without differentiation of BFU-Es of PHZ-induced anemic mice [36] and BFU-Es of human marrow [37] is based on the synthetic glucocorticoid hormone dexamethasone, which increases self-renewal of these progenitors. Glucocorticoids, which are increased in PHZ-induced anemia, activate their nuclear receptors, which act in an apparent complex with peroxisome proliferator-activated receptor a (PPAR-a) to control transcription of genes involved in the expansion of BFU-Es [38]. During this expansion of BFU-E numbers, their differentiation is inhibited posttranscriptionally by the induction of ZFP36L2, an RNA-binding protein that binds specific mRNAs involved in erythroid differentiation [36].

M.J. Koury/ Experimental Hematology 2015;-:-–-

5

A Normal erythropoiesis in marrow

------------------------------

SCF ---------------------------------------------

Normal EPO --------------------

B Suppressed erythropoiesis in marrow

------------------------------

SCF -------------------------------------- ↓ EPO (↑ Fas, ↑ Fas ligand) -----------

C Stimulated erythropoiesis in spleen

---------------------------

SCF ------------------------

-------- ↑ Glucocorticoids + Hedgehog + BMP4 ------------- ↑ EPO (↓ Fas, ↓ Fas ligand, ↑ Gas6 ) -------

MEP

Megakaryocte progenitor

BFU-E

CFU-E

Erythroblast

Macrophage

Figure 1. Model of erythroid progenitors in mice under three erythropoietic conditions. (A) Marrow during normal erythropoiesis. (B) Marrow during erythropoiesis suppressed by hypertransfusion. (C) Spleen during erythropoiesis stimulated by blood loss. Under normal or suppressed conditions, human marrow events are similar to those of mice, whereas under stimulated erythropoiesis conditions, the events illustrated for the murine spleen occur in the human marrow. Time periods below each section depict when specific systemic cytokines, such as EPO or glucocorticoids, or locally produced cytokines, such as SCF, Fas ligand, Gas6, BMP4, and Hedgehog, are required. Under normal conditions (A), formation of EBIs (niches of terminal erythroid differentiation) occurs when macrophages interact with CFU-Es or with late-stage BFU-Es that rapidly become CFU-Es. Suppression of erythropoiesis (B) inhibits formation of new EBIs and reduces CFU-Es and their erythroblast progeny in EBIs. In erythropoietically stimulated mice (C), BFU-Es expand in the spleen before EBI formation, which is increased as BFU-Es and their CFU-E progeny accumulate. With erythropoietic stimulation (C), expanded CFU-Es and their erythroblast progeny reside in the EBIs. The expansion of EBI numbers during erythropoietic stimulation may be much greater than depicted in (C), as space limitations in the figure prevent us from illustrating these frequently exponential increases. During and after erythropoietic stress, BFU-Es migrate from the marrow to the mouse spleen (not shown).

CFU-Es reside in erythroblastic islands As BFU-Es differentiate and reach the CFU-E stage of differentiation, they become associated with the structural niche of terminal mammalian erythropoiesis, the erythro-

blastic island (EBI). An erythroblastic island consists of a central macrophage surrounded by erythroid cells that vary in differentiation stage from the CFU-E through the nascent reticulocyte [39,40]. Analysis of the erythroid

6

M.J. Koury/ Experimental Hematology 2015;-:-–-

progenitors in EBIs of spleens of mice with PHZ-induced anemia revealed that CFU-Es were located in EBIs, but extremely few BFU-Es were found in EBIs as compared with their prevalence in the entire spleen [41]. Mice treated with liposomal clodronate to destroy macrophages and phlebotomy to induce anemia had about 50% survival of splenic BFU-Es, but less than 3% survival of splenic CFU-Es, when compared with phlebotomized controls that were not treated with liposomal clodronate [42]. Chronic deficiency of macrophages in mice caused decreased marrow erythropoiesis, but only slight or no anemia because of a concomitant decline in erythrocyte removal [43,44]. However, splenic erythropoiesis was diminished in response to PHZ-induced hemolysis [43] or phlebotomy [44] in these mice, confirming that functioning macrophages are required to respond to erythropoietic stress [42]. Hypertransfused rats have decreased numbers of marrow EBIs, and the EBIs that are present have decreased numbers of erythroblasts, which are immature compared with erythroblasts in EBIs of normal controls [45]. Loss of EBIs with hypertransfusion is consistent with CFU-Es being part of EBIs and with their numbers declining in response to decreased EPO, as reported by Hara and Ogawa and others. Therefore, BFU-Es may initiate EBI formation by associating with central macrophages, but they appear to differentiate rapidly into CFU-Es. On the other hand, the loss of all but 3% of splenic CFU-Es in PHZ-treated mice that have been depleted of macrophages by liposomal clodronate indicates that few CFU-Es exist outside of EBIs, and if CFU-Es initiate EBI formation by associating with a central macrophage, they appear to do so very rapidly after reaching the CFU-E stage. Figure 1 illustrates both late BFU-E and early CFU-E stages as initiators of normal EBI formation. All stages of erythroid differentiation from ProEBs to orthochromatic erythroblasts are found in EBIs of normal rat marrow [45]. Possible sources of this heterogeneity in erythroid stages in an EBI include: (i) EBI initiation with one macrophage interacting with cells of different stages of maturation, e.g., a BFU-E and a CFU-E, as illustrated in Figure 1C; (ii) EBI initiation by one BFU-E or CFU-E with one descendant CFU-E/ProEB self-renewing and another differentiating; and (iii) later addition of a CFU-E or a BFU-E to a previously established EBI. The absence of circulating CFU-Es, even during periods of erythropoietic stress, as reported by Hara and Ogawa [1,24], may be related to their rapid incorporation into and retention within the EBIs by the network of multiple adhesion proteins and their binding partners that mediate interactions between erythroid cells and central macrophages in EBIs [39,40]. The pairs of adhesion proteins and their respective binding partners include: (i) erythroid a4b1-integrin and macrophage vascular cell adhesion molecule (VCAM-1); (ii) erythroid intercellular adhesion

molecule 4 (ICAM-4) and macrophage integrins with av components; (iii) erythroblast-macrophage protein (EMP) in a homophilic interaction on both erythroid cells and macrophages; (iv) erythroid sialylated glycoproteins and macrophage CD169/Siglec 1; and (v) an unknown erythroid cell protein and macrophage hemoglobinhaptoglobin receptor (CD163). Thus, CFU-Es and ProEBs do not mobilize and circulate in the blood, even during increased erythropoietic demand, indicating that they are stably associated with macrophages in EBIs. However, some adaptations in response to erythropoietic stimulation may facilitate rapid expansion of the numbers of EBIs. In EBIs reconstituted in vitro, some CFU-Es/ProEBs lose adherence to central macrophages [46,47], and in vivo, these progenitors may be mobile in splenic or marrow hematopoietic tissue without entering circulating blood. Within the spleen or marrow, the mobile CFU-Es/ProEBs can potentially interact with macrophages to initiate new EBIs. Macrophage numbers may also be expanded, as newly differentiated macrophages with phenotypes of central macrophages are generated from monocyticmacrophage precursor cells. Indeed, newly generated macrophages are required for the erythropoietic response to large-scale, PHZ-induced hemolysis [48].

CFU-E dependence on EPO and the prevention of apoptosis With tritiated thymidine suicide analyses of cell cycle status of the splenic and marrow erythroid progenitors, both Hara and Ogawa [1] and Iscove [21], who published his findings at the same time, found no effects of erythropoietic status or site of erythropoiesis. They found that about 30%–40% of BFU-Es and about 60%–75% of CFU-Es were in the DNA synthesis phase of the cell cycle, regardless of whether the progenitors were from the marrow or spleen and whether they were from stimulated, normal, or suppressed mice [1,21]. These results indicated that EPO was not a mitogen for BFU-Es or CFU-Es. The large and prompt increases in CFU-E numbers following erythropoietic stimulation led to the notion that an erythroid progenitor population that immediately preceded the CFU-E stage was regulated by EPO. These results were later clarified when it became apparent that EPO’s major action was on the maintenance of viability of CFU-Es and ProEBs through the prevention of apoptosis [49,50]. Because CFU-Es are in active cell cycle as assessed by the tritiated thymidine results, their enhanced survival is accompanied by rapid increases in their numbers. Subsequent results with knockout of the genes for EPO and its receptor in mice confirmed the onset of EPO dependence at the CFU-E stage of differentiation [51]. In both mice and humans, the differentiation stages with the highest expression of EPO receptor and accompanying dependence on EPO for survival extend from CFU-Es/ProEBs into the early

M.J. Koury/ Experimental Hematology 2015;-:-–-

basophilic erythroblast stage [49,52–54]. The transmembrane EPO receptor (EPO-R) undergoes a conformational change after binding EPO that leads to signaling through its chaperone protein Janus kinase-2 (JAK2), which associates with the cytoplasmic portion of EPO-R. Multiple pathways mediate EPO-R-JAK2 signaling, including signal transduction and activation of transcription 5 (STAT5), RAS-Raf-MAPK, and PI3K-AKT [18]. Further examination of the CFU-E/ProEB populations indicated that despite all members of the population having a similar differentiation potential, the range of EPO concentrations required to prevent apoptosis varied widely [55]. This heterogeneity in EPO responsiveness among CFU-Es/ProEBs provided an explanation for the graded responsiveness to EPO in vivo and in vitro, but the source of susceptibility to apoptosis was not apparent until the role of FAS (CD95), a membrane protein of the tumor necrosis factor (TNF) receptor family that activates caspases that induce apoptosis, was defined in erythroid cells. Human CFU-Es/ProEBs display high levels of FAS, which induces apoptosis when they bind FAS ligand (FL) in the presence of low levels of EPO, but not when EPO levels are increased [56]. Importantly, the predominant source of FL for human CFU-Es/ProEBs is mature erythroblasts [56]. Thus, the close spatial arrangement in the EBIs of the maturing erythroblasts and CFU-Es/ProEBs provides a localized feedback mechanism for apoptosis in the EPO-dependent stages of CFU-Es/ProEBs that is removed by increased EPO levels. A similar Fas/FL negative feedback loop has been described in mice, but the EPOdependent CFU-Es/ProEBs not only express the most Fas, they also produce the most FL in the EBI [57]. Furthermore, EPO reduces expression of both Fas and FL in these EPO-dependent cells [57], such that this autoregulation feedback allows a low steady-state erythropoiesis in the mouse spleen with the ability to rapidly expand during periods of EPO-mediated erythropoietic stress [58]. Simulated erythropoietic recoveries after blood loss anemia based on FL produced by the EPO-dependent cells in mouse EBIs and by mature erythroblasts in human EBIs are consistent with the relatively more rapid recovery from acute blood loss observed in mice compared with humans [59]. A potential mechanism based on autocrine or paracrine stimulation within the EBIs has been proposed for the secreted factor, growth arrest-specific 6 (Gas6), which is induced in erythroid progenitor cells by EPO and enhances their survival response to EPO [60]. Mice deficient in Gas6 have impaired recovery from PHZ-induced anemia, as do mice with absence in erythroid cells of Axl, one of three TAM tyrosine kinase receptors for Gas6 [60,61]. Mice deficient in Axl have impaired expression of EPO receptors and the erythroid transcription factor GATA1 in EPO-dependent erythroblast stages [61], providing a mechanism for Gas6 enhancement of EPO-dependent survival.

7

Expansion of CFU-Es and ProEBs Although prevention of apoptosis by EPO provides a mechanism for the rapid increases in CFU-Es in erythropoietically stimulated mice and prompt decreases in erythropoietically suppressed mice, the higher proliferation rate of CFU-Es compared with BFU-Es regardless of erythropoietic status appears to be related to the microenvironment provided by EBIs and other growth factors produced at more distant sites. Human CFU-Es/ProEBs require EPO to survive, but their proliferation in vitro depends on insulin-like growth factor 1 (IGF-1) and SCF [62], with SCF not only supporting most of the proliferation, but also preventing further differentiation [63]. The studies describing dexamethasone induction of increased self-renewal and inhibited differentiation in BFU-Es did not find the effects in CFU-Es [36–38]. However, other studies reported that dexamethasone acts in concert with SCF and EPO in vitro to greatly expand (up to one millionfold) human progenitors at the CFU-E/ProEB stage without inducing further differentiation [64,65]. Indeed, human ProEBs derived from peripheral blood and treated with both EPO and dexamethasone exhibited direct interference of EPO receptor signaling by activated glucocorticoid receptors, which inhibited further differentiation but did not inhibit EPO’s anti-apoptotic effects [66]. In vivo studies in mice with hypofunctional nuclear glucocorticoid receptors revealed normal, steady-state erythropoiesis, but when challenged by PHZ-induced anemia or hypoxia, these mice had no CFU-E increases in the spleen or erythrocyte responses in the blood compared with wild-type mice [67]. Studies with mice deficient in Raf-1 kinase and/or Fas are consistent with glucocorticoids supporting proliferation without differentiation through prolonged Raf-1 expression while suppressing Fas expression [68]. Thus, steady-state erythropoiesis in the marrow, which requires EPO and SCF, is not dependent on glucocorticoid hormone, but the generation of CFU-E from BFU-E and expansion of CFU-E/ProEBs in the spleen during stress erythropoiesis require glucocorticoid hormone in a manner similar to its requirement for BMP4. That glucocorticoid hormones can induce greater expansion of erythroid progenitor populations does not explain the relatively high rate of mitosis in CFU-Es compared with BFU-Es. Stimulation of proliferation by direct contact with central macrophages or high concentrations of growth factors secreted in the immediate area surrounding the central macrophage may be increasing CFU-E/ProEB proliferation. In this regard, EBIs formed in vitro from mice [47] and humans [44] have revealed that cell cycle duration is decreased in CFU-Es/ProEBs when they are in direct contact with central macrophages. When CD169þ macrophages, like central macrophages in EBIs, are stimulated by dexamethasone in vitro, attached ProEBs are induced to proliferate, indicating that glucocorticoids may induce late erythroid progenitor cell proliferation indirectly

8

M.J. Koury/ Experimental Hematology 2015;-:-–-

through effects on central macrophages in addition to direct effects on erythroid progenitor self-renewal [69]. Among the potential secreted central macrophage products that may accumulate in the area of the EBI and stimulate CFU-E/ProEB proliferation is BMP4, which is produced by and accumulates around splenic macrophages following EPO administration in mice [70]. Survival and differentiation of erythroblasts The differentiating erythroblasts are the progeny of CFUEs/ProEBs that also reside in EBIs but do not have a direct EPO requirement for survival. However, erythroblasts have large intracellular accumulations of BCL-XL, the antiapoptotic member of the BCL-2 family, which is induced by EPO in the preceding CFU-E/ProEB stages [71,72], whereas the pro-apoptotic protein BIM, another BCL-2 family member, is decreased [72]. Pharmacologic studies with molecules designed to trap members of the transforming growth factor b (TGF-b) superfamily have indicated that in normal steady-state erythropoiesis and erythropoiesis during various anemias, erythroblast differentiation is negatively regulated by members of this family, which includes TGF-b, activins, BMPs, and growth differentiation factors (GDFs) [73]. This negative regulation of differentiation appears to begin in the latter half of the EPOdependent stages, that is, the pro-erythroblasts and early basophilic erythroblasts, and persists through the enucleating orthochromatic stage [73]. Applying knowledge of erythropoietic stimulation to treatment of clinical anemias Despite limitations in the investigation of erythropoietic regulation in humans, several clinical applications related to basic laboratory research into erythropoietic regulation have been achieved since the early studies by Hara and Ogawa and their contemporaries. These applications have been developed to treat chronic underproduction anemias. The cloning of EPO and large-scale production of recombinant human EPO (rhEPO) provided a therapeutic agent that is used routinely to treat the anemia of renal failure [74]. Because renal disease causes an EPO deficiency state, the correction of anemia by rhEPO administration in these patients has been very effective [74]. Prolonging the in vivo half-life of rhEPO severalfold through bioengineered increases in glycosylation [75] (Darbepoetin) or by polyethylene glycol conjugation of rhEPO [76] (continuous erythropoietin receptor activator [CERA]) allowed administration to be spaced at intervals of multiple weeks. However, as recovery of circulating RBCs approached near-normal levels in renal patients treated with rhEPO or the long-acting stimulators of EPO receptors, an increase in thrombotic cardiovascular events became evident [77]. In addition to increased thrombotic vascular disease, concerns about enhanced tumor growth in patients

with malignancies who received rhEPO to hasten recovery from chemotherapy-induced anemias led to more restricted use of rhEPO and long-acting stimulators of EPO receptors [78]. A polyethylene glycol-conjugated synthetic peptide with an amino acid sequence that is unrelated to EPO, but activates EPO receptors (peginesatide), was effective in treating patients who had developed antibodies to rhEPO that caused clinical pure red cell aplasia after exposure to rhEPO [79]. However, a subsequent increased incidence of acute lethal cardiovascular events associated with peginesatide led to discontinuation of its clinical use. These clinical problems associated with rhEPO and closely related medications that directly replicate its action have resulted in development of other agents that can increase erythropoiesis in underproduction anemias. These other erythropoietic agents have actions at various stages of erythroid progenitor differentiation. In aplastic anemia, an immune-mediated marrow failure process markedly decreases numbers of multilineage progenitors and lineagespecific progenitors, including the erythroid progenitors. In those patients with aplastic anemia who do not respond to immunosuppressive therapy, the thrombopoietin receptor agonist eltrombopag can increase erythropoiesis alone or, more commonly, in concert with other hematopoietic lineages [80]. In these cases, the increase in hematopoietic progenitor cells resulting from eltrombopag activity leads to increased supplies of BFU-E and their progeny. For those disorders that specifically affect the erythroid lineage progenitors, glucocorticoids have long been the standard therapy for the inherited disease Diamond–Blackfan anemia (DBA), with the large majority of patients having initial responses, but variable rates of relapse [81]. The role of mutations in genes related to production and function of ribosomes, as well as the role of p53 activity in the loss of BFU-Es and CFU-Es in DBA, has indicated that the ability of corticosteroids to improve erythropoiesis is more likely related to their ability to expand erythroid progenitor populations than to their immunosuppressive actions [82,83]. The ability of PPAR-a agonists to enhance erythropoiesis in vivo and in vitro when corticosteroid levels are elevated indicates that these agonists may have the potential to be effective in DBA patients who experience corticosteroid toxicity or resistance [38]. During the EPO-dependent period, one approach to the thrombotic and cardiovascular problems associated with rhEPO and related activators of EPO-Rs is to increase endogenous EPO production through administration of stabilizers of hypoxia-inducible factor 2a (HIF-2a), the transcription factor responsible for EPO transcription [84]. Roxadustat, an inhibitor of the prolyl hydroxylases that target HIF-2a for proteasomal degradation, has been found, in anemic renal failure patients, to increase circulating EPO concentrations in a controlled manner while improving iron availability for erythropoiesis [85]. A second approach to improving the response to EPO is to combine it with

M.J. Koury/ Experimental Hematology 2015;-:-–-

another agent that acts by a separate mechanism, but enhances EPO responses. The TGF-b ligand trap agents based on activin type II receptor A (sotatercept) [86] and activin type II receptor B (ACE-536) [87] increase red cell production in healthy individuals. In murine models of two human anemias with ineffective erythropoiesis, myelodysplasia [73] and b-thalassemia [88], the murine forms of sotatercept and ACE-536 improved erythropoiesis by enhancing erythroblast differentiation. The spleens of these mice had increased expression of the TGF-b superfamily member GDF-11, which signals through the Smad 2/3 pathways and inhibits erythroblast differentiation [73,88]. The increased GDF-11 and other members of the TGF-b superfamily are neutralized by TGF-b ligand trap agents. Although these agents facilitated differentiation of erythroblasts in the post EPO-dependent cells, they also increased FAS and FAS ligand expression so that numbers of proerythroblasts were decreased, such that the improved erythropoiesis was transient [88]. However, when combined with EPO, which decreases erythroid FAS and FAS ligand expression [57], the TGF-b ligand trap agents produced a sustained increase in erythropoiesis [88].

Conclusions Anemia from blood loss with recovery to baseline hemoglobin levels is a relatively common event. The recovery process after blood loss must first upregulate erythropoiesis promptly, but then smoothly downregulate it to prevent rebound polycythemia. The original notion of EPO’s induction of mitosis in a specific erythroid population in this recovery process has evolved into concepts of finely regulated apoptosis in erythroid progenitors that can be inhibited or induced with erythropoietic stimulation or suppression, respectively, and erythroid progenitor fates that balance limited self-renewal with differentiation such that progenitor expansion accompanies increased erythropoietic demand. Stress erythropoiesis occurring mainly in the spleen of mice provides an ideal and easily accessed site for examination of the regulation of erythropoiesis in a time of need. As in 1977, experiments in mice continue to be important because the micro-environment provided by the EBI and role of the central macrophage in the viability and proliferation of CFU-Es and later stages of differentiation are much more difficult to study in humans. However, the results of the experiments in mice should continue to provide important information that can be translated into the development of treatments for various types of anemias.

References 1. Hara H, Ogawa M. Erythropoietic precursors in mice under erythropoietic stimulation and suppression. Exp Hematol. 1977;5:141–148.

9

2. Stephenson JR, Axelrad AA, McLeod DL, Shreeve MM. Induction of colonies of hemoglobin-synthesizing cells by erythropoietin in vitro. Proc Natl Acad Sci USA. 1971;68:1542–1546. 3. Axelrad A, McLeod D, Shreeve M, Heath D. Properties of Cells that Produce Erythropoietic Colonies In Vitro. In: Robinson W, ed. Hemopoiesis in Culture. Bethesda, MD: U.S. Government Printing Office; 1974. p. 226–237. 4. Eaves CJ, Humphries RK, Eaves AC. In Vitro Characterization of Erythroid Precursor Cells and Erythropoietic Differentiation. In: Stamatoyannopoulos G, Nienhuis AW, eds. Cellular and Molecular Regulation of Hemogolobin Switching. New York: Grune and Stratton; 1979. p. 251–273. 5. McLeod DL, Shreeve MM, Axelrad AA. Chromosome marker evidence for the bipotentiality of BFU-E. Blood. 1980;56:318–322. 6. Vannucchi AM, Paoletti F, Linari S, et al. Identification and characterization of a bipotent (erythroid and megakaryocytic) cell precursor from the spleen of phenylhydrazine-treated mice. Blood. 2000;95:2559–2568. 7. Goldfarb AN, Wong D, Racke FK. Induction of megakaryocytic differentiation in primary human erythroblasts: a physiological basis for leukemic lineage plasticity. Am J Pathol. 2001;158:1191–1198. 8. Mancini E, Sanjuan-Pla A, Luciani L, et al. FOG-1 and GATA-1 act sequentially to specify definitive megakaryocytic and erythroid progenitors. EMBO J. 2012;31:351–365. 9. Siatecka M, Bieker JJ. The multifunctional role of EKLF/KLF1 during erythropoiesis. Blood. 2011;118:2044–2054. 10. Bianchi E, Zini R, Salati S, et al. c-myb supports erythropoiesis through the transactivation of KLF1 and LMO2 expression. Blood. 2010;116:e99–e110. 11. Bianchi E, Bulgarelli J, Ruberti S, et al. MYB controls erythroid versus megakaryocyte lineage fate decision through the miR-4863p-mediated downregulation of MAF. Cell Death Differ. 2015;22: 1906–1921. 12. Socolovsky M, Nam H, Fleming MD, Haase VH, Brugnara C, Lodish HF. Ineffective erythropoiesis in Stat5a(/)5b(/) mice due to decreased survival of early erythroblasts. Blood. 2001;98:3261–3273. 13. Chen K, Liu J, Heck S, Chasis JA, An X, Mohandas N. Resolving the distinct stages in erythroid differentiation based on dynamic changes in membrane protein expression during erythropoiesis. Proc Natl Acad Sci USA. 2009;106:17413–17418. 14. Hu J, Liu J, Xue F, et al. Isolation and functional characterization of human erythroblasts at distinct stages: implications for understanding of normal and disordered erythropoiesis in vivo. Blood. 2013;121: 3246–3253. 15. Erslev A. Hematology: control of red cell production. Annu Rev Med. 1960;11:315–332. 16. Cheshier SH, Prohaska SS, Weissman IL. The effect of bleeding on hematopoietic stem cell cycling and self-renewal. Stem Cells Dev. 2007;16:707–717. 17. Grover A, Mancini E, Moore S, et al. Erythropoietin guides multipotent hematopoietic progenitor cells toward an erythroid fate. J Exp Med. 2014;211:181–188. 18. Koury MJ. Erythropoietin: the story of hypoxia and a finely regulated hematopoietic hormone. Exp Hematol. 2005;33:1263–1270. 19. Papayannopoulou T, Finch CA. On the in vivo action of erythropoietin: a quantitative analysis. J Clin Invest. 1972;51:1179–1185. 20. Gregory CJ, Tepperman AD, McCulloch EA, Till JE. Erythropoietic progenitors capable of colony formation in culture: response of normal and genetically anemic W-W-V mice to manipulations of the erythron. J Cell Physiol. 1974;84:1–12. 21. Iscove NN. The role of erythropoietin in regulation of population size and cell cycling of early and late erythroid precursors in mouse bone marrow. Cell Tissue Kinet. 1977;10:323–334. 22. Pannacciulli IM, Massa GG, Saviane AG, Ghio RL, Bianchi GL, Bogliolo GV. Effect of bleeding on in vivo in vitro colony-forming hemopoietic cells. Acta Haematol. 1977;58:27–33.

10

M.J. Koury/ Experimental Hematology 2015;-:-–-

23. Peschle C, Magli MC, Cillo C, et al. Kinetics of erythroid and myeloid stem cells in post-hypoxia polycythaemia. Br J Haematol. 1977;37:345–352. 24. Hara H, Ogawa M. Erthropoietic precursors in mice with phenylhydrazine-induced anemia. Am J Hematol. 1976;1:453–458. 25. Miyake T, Kung CK, Goldwasser E. Purification of human erythropoietin. J Biol Chem. 1977;252:5558–5564. 26. Ogawa M, Grush OC, O’Dell RF, Hara H, MacEachern MD. Circulating erythropoietic precursors assessed in culture: characterization in normal men and patients with hemoglobinopathies. Blood. 1977; 50:1081–1092. 27. Clarke BJ, Housman D. Characterization of an erythroid precursor cell of high proliferative capacity in normal human peripheral blood. Proc Natl Acad Sci USA. 1977;74:1105–1109. 28. Bonig H, Chang KH, Nakamoto B, Papayannopoulou T. The p67 laminin receptor identifies human erythroid progenitor and precursor cells and is functionally important for their bone marrow lodgment. Blood. 2006;108:1230–1233. 29. Broudy VC, Lin NL, Priestley GV, Nocka K, Wolf NS. Interaction of stem cell factor and its receptor c-kit mediates lodgment and acute expansion of hematopoietic cells in the murine spleen. Blood. 1996;88:75–81. 30. Munugalavadla V, Kapur R. Role of c-Kit and erythropoietin receptor in erythropoiesis. Crit Rev Oncol Hematol. 2005;54:63–75. 31. Ulyanova T, Jiang Y, Padilla S, Nakamoto B, Papayannopoulou T. Combinatorial and distinct roles of alpha(5) and alpha(4) integrins in stress erythropoiesis in mice. Blood. 2011;117:975–985. 32. Gregory CJ, Eaves AC. Three stages of erythropoietic progenitor cell differentiation distinguished by a number of physical and biologic properties. Blood. 1978;51:527–537. 33. Dai CH, Krantz SB, Zsebo KM. Human burst-forming unitserythroid need direct interaction with stem cell factor for further development. Blood. 1991;78:2493–2497. 34. Paulson RF, Shi L, Wu DC. Stress erythropoiesis: new signals and new stress progenitor cells. Curr Opin Hematol. 2011;18:139–145. 35. Xiang J, Wu DC, Chen Y, Paulson RF. In vitro culture of stress erythroid progenitors identifies distinct progenitor populations and analogous human progenitors. Blood. 2015;125:1803–1812. 36. Zhang L, Prak L, Rayon-Estrada V, et al. ZFP36L2 is required for self-renewal of early burst-forming unit erythroid progenitors. Nature. 2013;499:92–96. 37. Narla A, Dutt S, McAuley JR, et al. Dexamethasone and lenalidomide have distinct functional effects on erythropoiesis. Blood. 2011;118:2296–2304. 38. Lee HY, Gao X, Barrasa MI, et al. PPAR-alpha and glucocorticoid receptor synergize to promote erythroid progenitor self-renewal. Nature. 2015;522:474–477. 39. Chasis JA, Mohandas N. Erythroblastic islands: niches for erythropoiesis. Blood. 2008;112:470–478. 40. Manwani D, Bieker JJ. The erythroblastic island. Curr Top Dev Biol. 2008;82:23–53. 41. Sadahira Y, Mori M. Role of the macrophage in erythropoiesis. Pathol Int. 1999;49:841–848. 42. Sadahira Y, Yasuda T, Yoshino T, et al. Impaired splenic erythropoiesis in phlebotomized mice injected with CL2MDP-liposome: an experimental model for studying the role of stromal macrophages in erythropoiesis. J Leukoc Biol. 2000;68:464–470. 43. Chow A, Huggins M, Ahmed J, et al. CD169(þ) macrophages provide a niche promoting erythropoiesis under homeostasis and stress. Nat Med. 2013;19:429–436. 44. Ramos P, Casu C, Gardenghi S, et al. Macrophages support pathological erythropoiesis in polycythemia vera and beta-thalassemia. Nat Med. 2013;19:437–445. 45. Mohandas N, Prenant M. Three-dimensional model of bone marrow. Blood. 1978;51:633–643.

46. Spike BT, Dibling BC, Macleod KF. Hypoxic stress underlies defects in erythroblast islands in the Rb-null mouse. Blood. 2007;110: 2173–2181. 47. Rhodes MM, Kopsombut P, Bondurant MC, Price JO, Koury MJ. Adherence to macrophages in erythroblastic islands enhances erythroblast proliferation and increases erythrocyte production by a different mechanism than erythropoietin. Blood. 2008;111: 1700–1708. 48. Haldar M, Kohyama M, So AY, et al. Heme-mediated SPI-C induction promotes monocyte differentiation into iron-recycling macrophages. Cell. 2014;156:1223–1234. 49. Koury MJ, Bondurant MC. Maintenance by erythropoietin of viability and maturation of murine erythroid precursor cells. J Cell Physiol. 1988;137:65–74. 50. Koury MJ, Bondurant MC. Erythropoietin retards DNA breakdown and prevents programmed death in erythroid progenitor cells. Science. 1990;248:378–381. 51. Wu H, Liu X, Jaenisch R, Lodish HF. Generation of committed erythroid BFU-E and CFU-E progenitors does not require erythropoietin or the erythropoietin receptor. Cell. 1995;83:59–67. 52. Wickrema A, Bondurant MC, Krantz SB. Abundance and stability of erythropoietin receptor mRNA in mouse erythroid progenitor cells. Blood. 1991;78:2269–2275. 53. Broudy VC, Lin N, Brice M, Nakamoto B, Papayannopoulou T. Erythropoietin receptor characteristics on primary human erythroid cells. Blood. 1991;77:2583–2590. 54. Wickrema A, Krantz SB, Winkelmann JC, Bondurant MC. Differentiation and erythropoietin receptor gene expression in human erythroid progenitor cells. Blood. 1992;80:1940–1949. 55. Kelley LL, Koury MJ, Bondurant MC, Koury ST, Sawyer ST, Wickrema A. Survival or death of individual proerythroblasts results from differing erythropoietin sensitivities: a mechanism for controlled rates of erythrocyte production. Blood. 1993;82: 2340–2352. 56. De Maria R, Testa U, Luchetti L, et al. Apoptotic role of Fas/Fas ligand system in the regulation of erythropoiesis. Blood. 1999;93: 796–803. 57. Liu Y, Pop R, Sadegh C, Brugnara C, Haase VH, Socolovsky M. Suppression of Fas-FasL coexpression by erythropoietin mediates erythroblast expansion during the erythropoietic stress response in vivo. Blood. 2006;108:123–133. 58. Koulnis M, Liu Y, Hallstrom K, Socolovsky M. Negative autoregulation by Fas stabilizes adult erythropoiesis and accelerates its stress response. PLoS One. 2011;6:e21192. 59. Eymard N, Bessonov N, Gandrillon O, Koury MJ, Volpert V. The role of spatial organization of cells in erythropoiesis. J Math Biol. 2015; 70:71–97. 60. Angelillo-Scherrer A, Burnier L, Lambrechts D, et al. Role of Gas6 in erythropoiesis and anemia in mice. J Clin Invest. 2008; 118:583–596. 61. Tang H, Chen S, Wang H, Wu H, Lu Q, Han D. TAM receptors and the regulation of erythropoiesis in mice. Haematologica. 2009;94: 326–334. 62. Muta K, Krantz SB, Bondurant MC, Wickrema A. Distinct roles of erythropoietin, insulin-like growth factor I, and stem cell factor in the development of erythroid progenitor cells. J Clin Invest. 1994; 94:34–43. 63. Muta K, Krantz SB, Bondurant MC, Dai CH. Stem cell factor retards differentiation of normal human erythroid progenitor cells while stimulating proliferation. Blood. 1995;86:572–580. 64. Panzenbock B, Bartunek P, Mapara MY, Zenke M. Growth and differentiation of human stem cell factor/erythropoietin-dependent erythroid progenitor cells in vitro. Blood. 1998;92:3658–3668. 65. Von Lindern M, Zauner W, Mellitzer G, et al. The glucocorticoid receptor cooperates with the erythropoietin receptor and c-Kit to

M.J. Koury/ Experimental Hematology 2015;-:-–-

66.

67. 68.

69.

70.

71.

72.

73.

74. 75. 76.

77.

enhance and sustain proliferation of erythroid progenitors in vitro. Blood. 1999;94:550–559. Stellacci E, Di Noia A, Di Baldassarre A, Migliaccio G, Battistini A, Migliaccio AR. Interaction between the glucocorticoid and erythropoietin receptors in human erythroid cells. Exp Hematol. 2009;37: 559–572. Bauer A, Tronche F, Wessely O, et al. The glucocorticoid receptor is required for stress erythropoiesis. Genes Dev. 1999;13:2996–3002. Rubiolo C, Piazzolla D, Meissl K, et al. A balance between Raf-1 and Fas expression sets the pace of erythroid differentiation. Blood. 2006; 108:152–159. Falchi M, Varricchio L, Martelli F, et al. Dexamethasone targeted directly to macrophages induces macrophage niches that promote erythroid expansion. Haematologica. 2015;100:178–187. Millot S, Andrieu V, Letteron P, et al. Erythropoietin stimulates spleen BMP4-dependent stress erythropoiesis and partially corrects anemia in a mouse model of generalized inflammation. Blood. 2010;116:6072–6081. Rhodes MM, Kopsombut P, Bondurant MC, Price JO, Koury MJ. Bcl-x(L) prevents apoptosis of late-stage erythroblasts but does not mediate the antiapoptotic effect of erythropoietin. Blood. 2005;106: 1857–1863. Koulnis M, Porpiglia E, Porpiglia PA, et al. Contrasting dynamic responses in vivo of the Bcl-xL and Bim erythropoietic survival pathways. Blood. 2012;119:1228–1239. Suragani RN, Cadena SM, Cawley SM, et al. Transforming growth factor-beta superfamily ligand trap ACE-536 corrects anemia by promoting late-stage erythropoiesis. Nat Med. 2014;20:408–414. Erslev AJ. Erythropoietin. N Engl J Med. 1991;324:1339–1344. Koury MJ. Sugar coating extends half-lives and improves effectiveness of cytokine hormones. Trends Biotechnol. 2003;21:462–464. Locatelli F, Reigner B. C.E.R.A.: pharmacodynamics, pharmacokinetics and efficacy in patients with chronic kidney disease. Expert Opin Investig Drugs. 2007;16:1649–1661. Horl WH. Anaemia management and mortality risk in chronic kidney disease. Nat Rev Nephrol. 2013;9:291–301.

11

78. Hedley BD, Allan AL, Xenocostas A. The role of erythropoietin and erythropoiesis-stimulating agents in tumor progression. Clin Cancer Res. 2011;17:6373–6380. 79. Macdougall IC, Rossert J, Casadevall N, et al. A peptide-based erythropoietin-receptor agonist for pure red-cell aplasia. N Engl J Med. 2009;361:1848–1855. 80. Desmond R, Townsley DM, Dumitriu B, et al. Eltrombopag restores trilineage hematopoiesis in refractory severe aplastic anemia that can be sustained on discontinuation of drug. Blood. 2014;123: 1818–1825. 81. Vlachos A, Ball S, Dahl N, et al. Diagnosing and treating Diamond Blackfan anaemia: results of an international clinical consensus conference. Br J Haematol. 2008;142:859–876. 82. Horos R, von Lindern M. Molecular mechanisms of pathology and treatment in diamond blackfan anaemia. Br J Haematol. 2012;159: 514–527. 83. Varricchio L, Migliaccio AR. The role of glucocorticoid receptor (GR) polymorphisms in human erythropoiesis. Am J Blood Res. 2014;4:53–72. 84. Koury MJ, Haase VH. Anaemia in kidney disease: harnessing hypoxia responses for therapy. Nat Rev Nephrol. 2015;11:394–410. 85. Besarab A, Provenzano R, Hertel J, et al. Randomized placebocontrolled dose-ranging and pharmacodynamics study of roxadustat (FG-4592) to treat anemia in nondialysis-dependent chronic kidney disease (NDD-CKD) patients. Nephrol Dial Transplant. 2015;30: 1665–1673. 86. Ruckle J, Jacobs M, Kramer W, et al. Single-dose, randomized, double-blind, placebo-controlled study of ACE-011 (ActRIIAIgG1) in postmenopausal women. J Bone Miner Res. 2009;24: 744–752. 87. Attie KM, Allison MJ, McClure T, et al. A phase 1 study of ACE536, a regulator of erythroid differentiation, in healthy volunteers. Am J Hematol. 2014;89:766–770. 88. Dussiot M, Maciel TT, Fricot A, et al. An activin receptor IIA ligand trap corrects ineffective erythropoiesis in beta-thalassemia. Nat Med. 2014;20:398–407.