Chapter 32: Erythropoiesis

Erythrocytes evolved largely for the purpose of transporting oxygen to tissues. Thus, the size of the red cell mass and the rate of red cell production must be closely related to supply and demand for oxygen in the tissues. Toward the end of the 19th century, French mountaineers and physiologists established that a low tissue tension of oxygen stimulates red cell production.¹ In 1906, Paul Carnot, a professor at the Sorbonne, and Mademoiselle DeFlandre, his associate, suggested that hypoxia generates a humoral factor capable of stimulating red cell production.² Based on questionable experimental data, an influential biochemist Friederich Miescher³ erroneously proposed that marrow hypoxia directly stimulates red cell production. Finally, in 1950, in an ingenious study on parabiotic rats, Kurt Reissmann⁴ provided evidence for the existence of an indirect humoral mechanism. This work, and work of Erslev and colleagues^5,6 who demonstrated that the plasma from anemic rabbits and primates contains an erythrocyte-stimulating factor, provided a strong basis for an existence of the factor appropriately named erythropoietin (EPO). In 1957, Jacobson and coworkers⁷ reported that EPO was produced by the kidney, a finding that raised the possibility that EPO isolated in adequate amounts might be of therapeutic benefit to uremic patients. After EPO cloning and production of recombinant EPO in therapeutic quantities, EPO has proved to have not only indications for therapy of anemia, but also has extraerythroid effects that are yet to be fully elucidated, such as its effect on cell growth. Widespread use of EPO for the treatment of anemia has surpassed original expectations.

HEMOGLOBIN AND RED CELLS

Hemoglobin is present in the most primitive animal forms, such as Paramecium and Tetrahymena. Some crustaceans, such as Daphnia, are capable of developing an oxygen transport system without circulating red cells.⁸ One interesting exception is an Antarctic ice fish (Chaenocephalus aceratus) lacking hemoglobin.⁹ These ice fish compensate for the absence of hemoglobin by their unusual nitric oxide metabolism.^10,11,12 They have very large hearts and unusually large diameter capillaries. This permits a large volume of blood to circulate at high flow rate and at low vascular pressure because of decreased peripheral resistance. This permits their survival in the very high oxygen content of Antarctic waters.¹²

An erythroid cell that can synthesize, carry, and protect hemoglobin from oxidation was found only with the development of a circulatory system. Circulating nucleated erythrocytes first appear in the worms of the phylum Nemertina and in the sessile marine creatures of the phylum Phoronida. Erythropoiesis in these primitive invertebrates takes place near or on the peritoneal surface, derived from endothelial cells.¹³ Nonnucleated red cells are observed for the first time in the more advanced phylum Annelida. However, the evolutionary advantage derived from enucleation appears to be slight. Nucleated red cells are observed in more advanced animals, such as reptiles and birds.¹⁴ All mammalian erythrocytes are nonnucleated and in most species are disc shaped, but are oval in some species.¹⁵ Enucleation decreases the workload of heart as it reduces one third of the cell weight.

In nonmammalian species, the spleen is the fundamental erythropoietic organ. However, in some fish, the kidneys also are involved in red cell production.^16,17 In vertebrates, an evolutionary shift occurred from the spleen to the liver and from the liver to the bones cavities.¹⁸ The homeostatic regulation of blood or hemoglobin production has been studied in Daphnia,⁸ where a balance exists between oxygen need and hemoglobin production. In higher animals, this relationship is maintained by adjusting red cell production. Studies of birds,¹⁹ fish,²⁰ and mammals²¹ indicate that red cell production is controlled by EPO, which is capable of adjusting red cell production to the demands for oxygen in the tissues. EPO of mammals has considerable biologic similarity and genetic homology.²²

EMBRYONIC AND FETAL ERYTHROPOIESIS

The environment within the bone apparently is optimal for cellular proliferation and maturation. However, bone cavities do not develop until the fifth fetal month. Other, presumably less favorable, sites are responsible for red cell production during early embryonic life (Chap. 7). In the human, large nucleated blood cells are first formed in the yolk sac,²³ and some enucleate.²⁴ They cluster in blood islands that become enveloped by endothelial cells forming the vascular plexus of the yolk sac.²⁵ This is referred to as primitive erythropoiesis, and is contrasted with definitive erythropoiesis, which occurs in the fetal liver and in the marrow. During the second gestational month, erythropoiesis moves to fetal liver, wherein smaller, but still macrocytic, nonnucleated cells are produced.^26,27 At birth, the hepatic phase of blood cell production ceases, and erythropoiesis moves to the marrow (Chaps. 7 and 48 provide details of developmental switching of embryonic, fetal, and adult globin expression).

During the neonatal period, the volume of available marrow space is almost the same as the total volume of hematopoietic cells and marrow vasculature.²⁸ This process continues for a few years until the growth of bones and bone cavities exceeds the growth of hematopoietic mass. However, whenever the demand on erythropoiesis increases (blood loss, hypoxia, ineffective erythropoiesis, or hemolysis), the lack of reserve space in neonates and small children reactivates extramedullary erythropoiesis in the liver and spleen.²⁹ In adults, expansion of marrow space continues, and the amount of fatty tissue gradually increases in all bone cavities. Because of the abundant marrow space, compensatory reactivation of extramedullary sites rarely occurs in later life. Extramedullary hematopoiesis during adult years indicates pathologic rather than compensatory blood formation, such as seen in primary myelofibrosis (Chap. 86) wherein the stem cells have abnormal interaction with the extracellular matrix.³⁰ During fetal life, EPO production is primarily hepatic.³¹ At birth, a gradual switch to renal production of EPO occurs. In the adult, the kidney is responsible for approximately 85 percent of total production.^32,33

PROGENITOR CELLS

Our ability to evaluate early erythropoiesis rests on functional assays of hematopoietic progenitors. The developmentally earliest progenitor committed to the erythroid lineage is the burst-forming unit–erythroid (BFU-E). It was initially termed a burst because it contains cells still capable of migration. These cells form smaller clusters around a larger central colony, giving the appearance of a sunburst with satellite colonies (Fig. 32–1). However, all the cells in the colony and its satellites are derived from a single BFU-E and, thus, are clonal. BFU-E takes longer than more mature erythroid progenitors to form a colony of erythroblasts (10 to 14 days) and form a large colony approximately 2000 to 3000 cells. BFU-E express low levels of EPO receptors (EPORs). BFU-E mature into colony-forming unit–erythroid (CFU-E), the more mature erythroid progenitor. A CFU-E is identified through more differentiated erythroid progenitor that in vitro form smaller colonies (50 to 200 cells) that mature in 3 to 5 days with EPOR density and EPO dependency increase gradually as progenitor cells mature, culminating at the level of the CFU-E.^34,35 BFU-E and CFU-E cannot be identified by microscopy (Chap. 31), but they can be studied in vitro by their ability to generate microscopically recognizable hemoglobinized precursors (i.e., erythroblasts) by so-called clonogenic assays on semisolid media.

PRECURSOR CELLS

In contrast, cells that constitute the latter stages of erythropoiesis can be identified by light microscopy (Chap. 31). The earliest morphologically recognizable erythroid precursor in the adult marrow is the pronormoblast. Pronormoblasts are conspicuously large and they have large uncondensed nuclei and deep basophilic cytoplasm as a result of the presence of numerous RNA-containing polyribosomes. Pronormoblast has a volume of 900 fL, 10 times the volume of the mature red blood cell. With each successive division, the precursor cells give rise to daughter cells of half their volume. Furthermore, with each division there is an increase in hemoglobin synthesis and condensation of nucleus. Thus, when the pronormoblasts divide to become basophilic normoblasts, the daughter cells have less blue cytoplasm because of hemoglobin synthesis and also a greater condensation of the nucleus. When the basophilic normoblasts divide further, they give rise to mature cells with more cytoplasmic hemoglobin that is stainable with both acid and basic dyes resulting in muddy-colored cytoplasm. These cells are called polychromatophilic normoblasts. The offspring of polychromatophilic normoblasts are called orthochromic normoblasts. Their nuclear chromatin is completely condensed and cytoplasm is pink from complete hemoglobinization. These cells do not divide further. Following extrusion of the nucleus, the enucleated cells derived from orthochromic orthochromatic erythroblasts are termed reticulocytes, named after the cytoplasmic remnants of the endoplasmic reticulum and the persistence of a few mitochondria and strings of ribosomes seen when stained with supravital dyes. Reticulocytes remain in the marrow for 48 to 72 hours before being released to the blood. The reticulocytes have an irregular polylobated shape and various membrane-bound organelles.³⁶ In the blood, immature erythrocytes (reticulocytes) undergo further maturation with the removal of vestiges of organelles and reconditioning of the membrane to become mature red blood cells with the morphology of a biconcave disk.³⁷

The number of erythroid precursor cells determines to a great extent the number of red cells produced. The proerythroblasts also contain EPORs that, in the presence of higher than normal levels of EPO, may accelerate their entry into their first mitotic division. This process may lead to a shortened marrow transit time of erythroblasts³⁸ and result in release of still immature erythrocytes (polychromatophilic macrocytes), so-called stress reticulocytes (Fig. 32–2).³⁹ Creation of a normal sized and shaped red cells, devoid of organelles, is the end result of an orderly transformation of a proerythroblast with a large nucleus and a volume of approximately 900 fL to a hemoglobinized anucleate disc-shaped cell with a volume of approximately 90 fL. Although cytoplasmic maturation is continuous, the interposed mitotic divisions cause a stepwise reduction in cytoplasmic and nuclear volumes, enabling recognition of proerythroblasts, erythroblasts, and polychromatophilic macrocytes (reticulocytes) with light microscopy (Chap. 31). Direct measurements of the number of marrow erythroblasts and reticulocytes have shown approximately 50 erythroblasts and approximately 124 reticulocytes for each proerythroblast (Table 32–1).^40,41 This distribution conforms to the number of cells in a theoretic erythroid pyramid (Table 32–1, Fig. 32–3). In the pyramid, each erythroblast undergoes five mitotic divisions over 5 days before the orthochromatic erythroblast loses its nucleus and as an immature erythrocyte enters a 2- to 3-day period of maturation before its release from the marrow. The size and shape of these erythroid pyramids undoubtedly vary, but such variations play a role in the physiologic control of red cell production. When production is suppressed, as in low EPO as seen in anemia of chronic renal disease, the distribution of erythroblasts appears normal, with no morphologic or ferrokinetic evidence of ineffective erythropoiesis but the number of erythroid progenitors is decreased.³⁸ When production is increased, as in severe hemolytic anemia, the pyramid of erythroid precursors also appear normal, with no evidence of additional mitotic divisions but the number of erythroid progenitors is increased. Consequently, the rate of red cell production largely depends on the number of erythroid progenitors formed.

As the erythroblast matures, its synthetic activities increase rapidly, producing all proteins characteristic of mature red blood cells, particularly globin. Eventually 95 percent of all protein in the red cell is hemoglobin, almost all hemoglobin A (α₂β₂) in adults, with only small amounts of hemoglobin F (α₂γ₂) and hemoglobin A₂ (α₂δ₂). Hemoglobin F is unequally distributed and is present only in some erythrocytes, designated as F cells (Chaps. 48 and 49).

EPOR density declines sharply on early erythroblasts, and EPORs are absent from the more mature erythroblast forms while the number of receptors for transferrin increases, reflecting the increased demands for iron for heme synthesis.

ERYTHROBLAST ENUCLEATION

The microenvironment may be important for proliferation and maturation of erythroblasts. However, in situ secreted or circulating growth factors and cytokines appear to be less important for precursor cells than for progenitor cells. Intercellular adhesion molecules secure the structural integrity of the marrow, and fibronectin is of special importance for erythroblasts.⁴² Loss of fibronectin receptors heralds the translocation of polychromatophilic macrocytes (reticulocytes) into blood, but some newly emerging erythrocytes remain sticky even after release and are temporarily sequestered by the spleen (Chap. 6). Because erythroid colonies developed in vitro consist principally of nucleated red cells, enucleation may primarily be induced by marrow stromal cells (Chaps. 5 and 31).

The extrusion of the spent pyknotic nuclei at terminal erythroid maturation is unique to mammals. This process results in the formation of pliable biconcave disc from rigid spheroidal cells. Enucleation decreases workload of the heart as it reduces one-third of the erythrocyte weight. The retinoblastoma protein and its effector, E2f-2, are critical for erythroid cells to exit the cell cycle for enucleation to take place.⁴³ During terminal differentiation, the plasma membrane forms an envelope around the nucleus, followed by the formation of the contractile actin ring on the plasma membrane to move the nucleus for disposal.⁴⁴ Rac guanosine triphosphatases (GTPases) and their effector mDia2 are required for the contractile actin ring movement in the release of the nucleus.⁴⁵ After separation from the cell, the expelled nucleus displays phosphatidylserine and is recognized and engulfed by macrophages.⁴⁶ Emp, an erythroblast–macrophage protein initially identified as a mediator of erythroblast–macrophage interactions, also plays a role in the enucleation.⁴⁷ Emp associates with F-actin, an interaction important for the normal distribution of F-actin in both erythroblasts and macrophages. Emp null mice do not extrude their nuclei from erythroid cells. Thus, Emp appears to be required for erythroblast enucleation.⁴⁸

Microscopic determination of marrow cellularity and proportion of erythroblasts permits semiquantitative evaluation of erythropoiesis. However, the presence of ineffective erythropoiesis in disease states, such as iron deficiency, anemia of chronic disease, megaloblastic anemias, and thalassemias, makes the morphologic approach misleading (Chaps. 37, 41, 42, and 48). Red cell production can be accurately estimated by ferrokinetic studies using ⁵⁹Fe. Similarly, the amount of the final product of erythropoiesis, the red cell mass, can also be accurately measured. Unfortunately, the ever-increasing regulation of even minute amounts of radioisotopes used in vivo makes these methods available in only a few specialized centers.

Chapters 7, 30, 47, and 48 discuss developmental control of erythropoiesis, differential use of enzyme and globin genes, and the crucial differences between embryonic yolk sac and fetal/adult definite erythropoiesis. This chapter focuses mainly on adult erythropoiesis.

Erythropoiesis is a tightly regulated system, but the details are still not fully elucidated. Much remains to be learned from uncovering the molecular basis of many congenital and acquired mutations that disrupt the control of erythropoiesis. Two competing hypothesis have been proposed to explain the differentiation of the hematopoietic progenitors cells toward erythroid lineage.

DETERMINISTIC MODEL OF LINEAGE COMMITMENT

According to the deterministic model of lineage commitment, specific extracellular signals, such as cytokines, play an instructive role in lineage specification.

Multipotential progenitors (Chap. 18) and erythroid unipotential progenitors, the BFU-E, require stem cell factor, interleukin-3, granulocyte-macrophage colony-stimulating factor, and/or thrombopoietin for growth and survival (Fig. 32–4).

STOCHASTIC MODEL OF ERYTHROID DIFFERENTIATION

In contrast, the stochastic model proposes that spontaneous formation of a set of transcription factors, independently of extrinsic signals, mediates lineage commitment. These transcription factors activate a unique set of genes for a particular lineage and repress the action of alternative transcription factors and cytokines play only a permissive role. Most of evidence based on gene targeting studies and in vitro culture studies support the stochastic model of differentiation. Several transcription factors, such as GATA-1, FOG1, erythroid Kruppel-like factor (EKLF), PU.1, and SCL/TAL1 (stem cell leukemia/T-cell acute lymphoblastic leukemia 1 factor) have been characterized that are involved in erythroid differentiation.

The GATA family of zinc-finger transcription factors was first identified as the nuclear factors that bind to the GATA sequence in the enhancer region of the globin genes.^49,50 GATA-1 protein is expressed during erythroid differentiation, with highest expression in CFU-Es and pronormoblasts. GATA-1 promotes erythroid differentiation by activating several erythroid-specific genes and represses transcription of Kit receptor and GATA-2. GATA-1 deficient mice die at embryonic day 10.5 with severe anemia from maturation arrest at the stage of pronormoblasts.⁵¹ In vitro, GATA-1 null the embryonic stem cells fail to mature beyond pronormoblast and undergo apoptosis. GATA-1 and its cofactor CBP are essential for the formation of an erythroid-specific histone acetylation pattern of histones at the active globin genes and the β-globin locus control region.⁵² GATA-1, along with EPO, induces expression of the antiapoptotic protein Bcl-x_L⁵³ and interacts with multiple proteins, including FOG-1 and PU.1,⁵⁴ and FOG-1 acts as a cofactor for GATA-1.⁵⁵ GATA-1 interaction with PU.1 appears to counteract erythropoiesis by inducing differentiation of pluripotent stem cell to myeloid and B lymphopoiesis and inhibition of erythropoiesis.^54,56,57 Whereas PU.1 absence appears to be required for completion of terminal erythroid differentiation, low levels of PU.1 expression are essential for fetal erythropoiesis and for proper augmentation of adult erythropoiesis at times of stress.⁵⁸

Friends of GATA

FOG-1, a member of friend of GATA family of zinc finger proteins, act as cofactors for GATA-1. It was first identified in a yeast two-hybrid screen for GATA-1 interacting proteins.⁵⁵ It binds to the amino zinc finger of GATA-1. FOG-1−/− mice die during embryonic days 10.5 to 11.5 from severe anemia with arrest in erythroid maturation at a stage similar to that observed in the GATA-1− mice.⁵⁹ FOG-1 physically interacts with GATA-1 to augment or inhibit its transcriptional activity depending on the promoter context.

GATA-2 was initially cloned as a GATA motif-binding factor is present in all erythroid cells; and, targeted deletion of GATA-2 resulted in embryonic lethality at day 10.5 from ablation of blood cell development.⁶⁰ GATA-1 and GATA-2 directly regulate GATA-2 transcription in a reciprocal fashion during erythroid differentiation.^61,62 GATA-2 autoregulates its transcription by binding to its own regulatory elements in the promoter region. This autoregulation is abolished by the displacement of GATA-2 by GATA-1 (GATA-2/GATA-1 switch), an interaction facilitated by FOG-1.⁶³ Chromatin immunoprecipitation studies indicate that FOG-1 facilitates occupancy by GATA-1 at selected cis-regulatory chromatin elements. Double knockout of GATA-1 and GATA-2 results in embryonic lethality with complete absence of primitive erythropoiesis.⁶⁴ The severity of this phenotype compared to either single GATA-1 or GATA-2 knockout suggests overlapping functions of these two transcription factors in primitive erythropoiesis.

Kruppel-like Factor

EKLF is a zinc finger protein identified by subtractive hybridization of the mRNA of erythroid cells with common messages in a myeloid cell line.⁶⁵ It interacts with CACCC sequence in the β-globin promoter, where it modifies chromatin structure permitting β-globin gene transcription. EKLF-deficient mice die at embryonic day 14.5 to 15 from severe anemia from defective definitive erythropoiesis.⁶⁶ There is a marked decrease in β-globin mRNA and protein levels in EKLF-deficient erythroid cells. Large amounts of iron accumulate in the reticuloendothelial system of EKLF-deficient mice, consistent with an ineffective erythropoiesis.

STEM CELL LEUKEMIA/T-CELL ACUTE LYMPHOBLASTIC LEUKEMIA 1

SCL/TAL1 is a member of basic helix-loop-helix transcription factors essential for maturation of the erythroid and megakaryocytic lineages.⁶⁷ Knockout of SCL/TAL1 leads to failure of hematopoiesis.⁶⁸ Selective rescue of SCL/TAL1 null embryonic stem cells under the control of stem cell enhancer revealed differentiation blocks in erythroid and megakaryocytic maturation.⁶⁹ Conditional knockout studies have revealed that erythroid and megakaryocytic precursors do not develop in the marrow of mice upon deletion of SCL/TAL1.⁷⁰ Heterodimerization of SCL with other transcription factors, such as E2A, is a prerequisite for its functions.⁷¹

BCL11A, a transcription factor initially identified in lymphoid cells, regulates erythroid differentiation, especially in switching from fetal to adult hemoglobin.⁷¹ Fetal hemoglobin (HbF) levels decline after birth and are then replaced by adult hemoglobin A. The molecular mechanisms responsible for this switch are not completely known. Genome-wide association findings have provided a major breakthrough in understanding this phenomenon.⁷² There is an inverse correlation between BCL11A and HbF expression in erythroid cells. BCL11A occupies several discrete sites in the β-globin gene cluster and likely plays an important role in hemoglobin switching during erythroid differentiation.

GROWTH ARREST-SPECIFIC 6 PROTEIN

Growth arrest-specific 6 (Gas6) protein is a secreted vitamin K–dependent protein that interacts with cell membranes and leads to intracellular signaling (via its receptor tyrosine kinases). Gas6 receptors are expressed in hematopoietic tissue, megakaryocytes, myelomonocytic precursors, and marrow stromal cells. Gas6 amplifies the erythropoietic response to EPO using a mouse model of Gas6 knockout.⁷³ Gas6 is known to downregulate the expression of inflammatory cytokines such as tumor necrosis factor-α by macrophages.⁷⁴

Figure 32-5 outlines the interrogation of the molecular mechanisms that regulate lineage-specific differentiation and commitment reveals the existence of separate megakaryocytic/erythroid progenitors versus both myeloid and lymphoid lineages.⁷⁵

ERYTHROPOIETIN, OXYGEN SENSING, AND HYPOXIA-INDUCIBLE FACTOR

Erythropoietin

The principal hormone regulating erythropoiesis is EPO, which is produced principally in the kidney.⁷ Erythroid progenitors express their own EPO.⁷⁶ Different levels of kidney-produced EPO are optimal for various stages of erythroid maturation.⁷⁷ Purification of EPO provided a partial protein sequence that led to cloning of the gene and permitted mass production of the recombinant protein.⁷⁸ EPO and its recombinant form are heavily glycosylated α-globulins with a molecular mass of 34,000 daltons and a specific activity of approximately 200,000 IU/mg.^79,80 Sixty percent of the molecular weight of the recombinant protein is contributed by amino acids; the remaining 40 percent is composed of carbohydrate. Using molecular probes for EPO, mRNA enabled the localization of the synthesis of EPO to renal cortical interstitial cells^81,82 of endothelial or fibroblastic lineage. The cells appear to function in an all-or-none fashion, with the overall production of mRNA dependent on the number of cells activated.⁸³

Certain 5′ sequences located 6000 to 12,000 bp upstream also affect EPO gene transcription.⁸⁴ These sequences are not hypoxia sensitive but appear necessary for tissue and cellular specificity.⁸⁴ Hepatic production is contributed primarily by hepatocytes but is a much less important source than is the kidney.⁸⁵ During fetal life, however, hepatic EPO production is of major importance for red cell production (Chap. 7).^86,87 EPO production is regulated exclusively at the level of its transcription by hypoxia at the transcription level. The transcriptional activation of the EPO gene is controlled by a specific sequence located in the 3′ flanking region termed hypoxia-responsive element.^88,89,90 The core of the enhancer is constituted by the sequence CACGTGCT and mutations in this core sequence abolish hypoxia responsiveness.

EPO is not stored; it is secreted immediately.^81,82,83 Circulating recombinant EPO and presumably native EPO have a half-life (T_1/2) of 4 to 12 hours, with a volume of distribution slightly larger than that of the plasma volume.⁹¹ EPO is degraded after it binds to EPOR (see “Erythropoietin Receptor” below).⁹²

Erythropoietin Receptor

Interaction of EPO with its receptor EPOR results in (1) stimulation of erythroid cell division, (2) erythroid differentiation by induction of erythroid-specific protein expression, and (3) prevention of erythroid progenitor apoptosis.⁹³ Earlier models of this interaction were based on the ligand (EPO)-induced homodimerization of EPOR. In reality, EPOR is a preformed homodimer that undergoes a major conformational change upon binding,⁹⁴ which initiates the EPO-specific erythroid signal transduction cascade (Fig. 32–6). The cytoplasmic portion of EPOR contains a positive regulatory domain that interacts with Janus kinase 2 (JAK2).⁹⁵ Immediately after EPO binding, JAK2 cross-phosphorylates the EPOR itself, and other proteins such as STAT5 (signal transducer and activator of transcription 5), thus initiating a cascade of erythroid-specific signaling.⁹⁶ JAK2/STAT5 signaling plays an essential role in EPO–EPOR–mediated regulation of erythropoiesis (see Fig. 32–6).⁹⁷ Deficiency of EPO–EPOR is lethal by abrogating fetal liver erythropoiesis (but not the “primitive” yolk sac erythropoiesis). However, in these EPO or EPOR knockout mice, differentiation of pluripotential stem cells to BFU-E occurs, but not the subsequent erythroid differentiation. This occurrence demonstrates the crucial role of EPO in terminal erythroid maturation.^35,98,99 The C-terminal cytoplasmic portion of EPOR also possesses a domain essential for prevention of apoptosis (see Fig. 32–6) by inducing expression of Bcl-x_L via phosphoinositide 3′-kinase (PI3K).⁵³ However, the cytoplasmic portion of EPOR also contains a negative regulatory domain¹⁰⁰ that interacts with hematopoietic cell phosphatase (HCP, also known as SHP1) and down-modulates signal transduction.¹⁰¹ Once recruited by EPOR tyrosine (Y)429, HCP attaches to the cytoplasmic EPOR domain and dephosphorylates JAK2. Inactivation of the HCP binding site leads to prolonged phosphorylation of JAK2/STAT5.^101,102 CIS3 (also known as SOCS3), another negative regulator of erythropoiesis, binds to the cytoplasmic portion of the EPOR Y401 and suppresses EPO-dependent JAK2/STAT5 signaling.^103,104 Thus, deletion of the distal C-terminal cytoplasmic portion of EPOR results in a truncated EPOR, abolishes negative regulatory elements, and results in increased proliferation of erythroid progenitor cells. Gain-of-function mutations resulting from deletion of the negative regulatory domain of the EPOR gene (Chap. 57) have been demonstrated in a small proportion of individuals with primary familial and congenital polycythemia, but are rarely found in erythroleukemia¹⁰⁵; however, the rearranged EPOR has also been identified in a subtype of high-risk B-progenitor acute lymphoblastic leukemia.¹⁰⁶

Because the activation signal after EPO binding to its receptor is rapidly downregulated and EPO briskly disappears after binding to EPOR, EPO–EPOR internalization is one mechanism of downregulation of EPO signaling.⁹² After EPO binds to the receptor, EPO–EPOR complexes are ubiquinated, rapidly internalized, and targeted for degradation. This process involves two proteolytic systems, the proteosomes that remove part of the intracellular domain of EPOR at the cell surface and the lysosomes that degrade the EPO–EPOR complex in the cytoplasm.¹⁰⁷

Another incompletely understood mechanism of erythropoiesis regulation is the presence of several EPOR isoforms, some of which may have an inhibitory function on erythropoiesis.^108,109,110

Nonerythroid Effect of Erythropoietin Signaling

Soon after the erythroid effects of recombinant EPO were described, nonerythroid effects were identified.¹¹¹ Some of these effects are beneficial, including roles in neural, cardiovascular, and retinal tissues, and in immune function and in tissue repair. It has been claimed that the hormone also exerts beneficial effects on athletic performance and improved neurocognition, but these are not convincingly substantiated. The effects of EPO in nonerythroid tissues are the result of EPO binding to EPOR, and, as in erythroid cells, the EPO–EPOR interaction initiates a signal transduction process that regulates the survival, growth and differentiation of the involved tissue.¹¹² EPO and EPOR play a physiologic role in many nonerythroid cells including endothelial cells,¹¹³ megakaryocytes, and cells of the brain, heart, uterus, breast, and testis. However, in some tissues (e.g., brain, heart, and kidney) the signaling mechanism may be different because EPO can interact with EPOR and CD131 heterodimers.^114,115

Detrimental EPO effects include a poorly understood increased cancer mortality,^95,116,117 increased blood pressure, and thrombosis.¹¹⁴

Hypoxia-Inducible Factors

Under normal conditions, EPO production is mediated by decreased oxygen saturation of hemoglobin, that is, hypoxemia.⁷⁷ Hypoxia is an important factor in development, energy metabolism, vasculogenesis, iron metabolism, tumor promotion and is the principal regulator of erythropoiesis. The response to hypoxia is controlled by transcriptional factors termed hypoxia-inducible factors (HIFs).^118,119 Adaptive physiologic responses to hypoxia serve to (1) increase O₂ delivery to cells, (2) allow cells to survive under reduced O₂ by activating glycolysis, and (3) reduce the formation of reactive oxygen species.¹²⁰ HIFs are heterodimeric transcription factors composed of a highly-regulated α subunit and a constitutively expressed β subunit that belongs to the basic helix-loop-helix containing the PER-ARNT-SIM (PAS)-domain family of transcription factors. The first HIF to be discovered, HIF-1, is induced in hypoxic cells and binds to a cis-acting nucleotide sequence of hypoxia-controlled genes referred to as the hypoxia-responsive element, first identified in the 3′-flanking region of the human EPO gene.¹²¹ Two other HIF homologues, HIF-2 and HIF-3, have been identified. HIF-2 has more limited tissue expression than HIF-1 but it is the principal regulator of EPO expression.^118,119 Many hypoxia-inducible genes are directly regulated by HIF-1. Approximately 3 percent of all genes expressed in endothelial tissue are HIF-1 regulated.¹²² The half-life of HIF-1α in the cell is minutes under normoxic conditions. HIF-1 and HIF-2 α subunits are rapidly degraded by the von Hippel-Lindau (VHL) protein–ubiquitin–proteasome pathway.¹²³ The targeting and subsequent polyubiquitination of HIF α subunits requires VHL, iron, O₂, and proline hydroxylase activity, and this complex constitutes the oxygen sensor (Fig. 32-7).^124,125

This degradation of HIFs α subunits is initiated by a posttranslational hydroxylation event at residue proline 564 (P564) that is mediated by one of several iron-containing proline hydroxylases (PHDs). The hydroxylation of HIFs α subunits facilitates binding to the VHL protein and subsequent ubiquitination and proteasomal degradation. Osteosarcoma protein 9 (OS-9) binds to both HIF-1α and PHD2 and is required for efficient prolyl hydroxylation.¹²⁶ Under hypoxic conditions, HIF-1 and HIF-2 α proteins are not degraded and are translocated to the cell nucleus where they dimerize with HIF-β to form the HIF heterodimer that activates transcription through binding to specific hypoxia-responsive elements on target genes. Another regulatory step involves O₂-dependent asparaginyl-hydroxylation of asparagine (N) 803 in HIF-1α that requires the enzyme HIF-3, also known as FIH-1 (factor inhibiting HIF-1). Hydroxylation of N803 during normoxia blocks the binding of transcription factors p300 and CBP to HIF-1, resulting in inhibition of HIF-1–mediated gene transcription. Under hypoxic conditions, HIF α subunits are not hydroxylated. The unmodified protein escapes VHL-binding, ubiquitination, and degradation (see Fig. 32–7). When N803 of HIF-1α is not asparaginyl-hydroxylated, p300 and CBP can bind to the HIF-1 heterodimer, allowing transcriptional activation of HIF-1 target genes.

HIF-2 Transcription Factor

HIF-1α and HIF-2α exhibit a high degree sequence homology but have differing mRNA expression patterns: HIF-1α is expressed ubiquitously, whereas HIF-2α expression is restricted to certain tissues.^118,127 The kidney is the main site of EPO production (i.e., renal interstitial cells), and HIF-2 and, to a lesser degree, HIF-1 are the principal regulator of EPO transcription in the kidney.^118,127 In other tissues, such as brain¹²⁸ and liver⁸⁷ (which generates approximately 15 percent of circulating EPO), EPO gene transcription is HIF-2–dependent.¹²⁷ The discovery of an iron-responsive element in the 5′ untranslated region of HIF-2α reveals a novel regulatory link between iron availability and HIF-2α expression¹²⁹ that may also influence control of erythropoiesis. The importance of HIF-2α in regulation of EPO gene was demonstrated by a gain-of-function HIF-2α mutation causing erythrocytosis.¹³⁰

Hypoxia-Independent Regulation of Hypoxia-Inducible Factor

While O₂-dependent regulation of the HIF-1α subunit is mediated by prolyl hydroxylases, VHL protein, and the proteasomal complex, hypoxia-independent regulation of HIF-1α has been uncovered. This novel mechanism involves the receptor of activated protein kinase C (RACK1) as a HIF-1α–interacting protein that promotes prolyl hydroxylase/VHL-independent proteasomal degradation of HIF-1α. RACK1 competes with heat shock protein 90 (HSP90) for binding to the PAS-A domain of HIF-1α. HIF-1α degradation is abolished by loss-of-function RACK1. RACK1 binds to the proteasomal subunit, elongin-C, and promotes ubiquitination of HIF-1α (see Fig. 32–7). Therefore, RACK1 and HSP90 are the essential components of an O₂/PHD/VHL-independent mechanism for regulating HIF-1α.¹³¹

The rapid degradation of HIF is complex and tightly regulated, and mutations affecting the genes that encode the regulatory factors may underlie some of the unexplained congenital polycythemias.

This complex (Chaps. 34 and 57) constitutes the oxygen sensor (see Fig. 32–7).^124,125,132

INSULIN-LIKE GROWTH FACTOR-1, RENIN–ANGIOTENSIN SYSTEM, AND HEMATOPOIESIS

Although in vitro studies of erythropoiesis have provided crucial information about the regulation of erythropoiesis, many experiments were performed in the presence of serum and serum-component proteins capable of stimulating and inhibiting erythropoiesis.^133,134 Using serum-free conditions, insulin-like growth factor-1 (IGF-1) can partially substitute for EPO in BFU-E cultures. Furthermore, anephric, nonanemic patients with no detectable EPO have elevated levels of IGF-1.¹³⁵

NIX-DEPENDENT CLEARANCE OF MITOCHONDRIA BY AUTOPHAGY

During terminal erythroid differentiation, erythroid cells discard all their internal organelles including mitochondria. Autophagy has been suggested to play an important role in this process based on early morphologic studies.^136,137 This is confirmed by studies using chemical inhibitors of autophagy and small interfering ribonucleic acid (siRNA) knockdown of essential autophagy associated genes.^138,139 At the initial stage of autophagy, a double-membrane structure is formed to sequester cytoplasmic components in autophagosomes. Autophagosomes then fuse with lysosomes and become autophagolysosomes to degrade the sequestered components. Mice deficient in a bcl-2 family member, Nix, a protein that is expressed during erythropoiesis and regulates mitochondrial apoptosis (autophagy), display defects in the clearance of mitochondria. Interestingly, the formation of autophagosomes in reticulocytes is normal in the absence of Nix, suggesting that Nix is not required for the initiation of autophagy or the formation of autophagosomes. Instead, mitochondria remain clustered outside of autophagosomes in Nix−/− reticulocytes.¹³⁸ This indicates that Nix is required for the sequestration of mitochondria by autophagosomes. Another study using virally transformed Nix−/− erythroid cells also reported defective inclusion of mitochondria by autophagosomes.¹⁴⁰ Therefore, Nix deficiency leads to a specific defect in mitochondrial removal without causing a general block in autophagy or erythroid maturation. Nix−/− reticulocytes are defective in the loss of mitochondrial membrane potential during in vitro maturation.¹³⁸

DOWNREGULATION OF CELL SURFACE PROTEINS AND OTHER ORGANELLES

During erythroid maturation, reticulocytes release small vesicles containing cellular proteins. These vehicles are exosomes that are involved in the clearance of cell-surface proteins, such as acetyl cholinesterase, CD71 transferrin receptor and integrin α₄β₁.^141,142 The enrichment of these surface molecules in exosomes suggests that the exosomal pathway plays a major role in the clearance of these surface molecules. Other cellular components, such as lysosomes, endoplasmic reticulum, Golgi apparatus, ribosomes, and RNA, are also cleared during erythroid maturation. The precise mechanisms for the removal of these components are unclear. In Nix-deficient mice, the loss of CD71 and ribosomes are normal during erythroid maturation, suggesting that exosomal pathways play a major role in the clearance of CD71 or ribosomes.¹³⁸

MicroRNAs (miRNAs) are small 18- to 22-nucleotide noncoding RNAs that regulate gene expression by inhibiting protein translation or by destabilizing target mRNAs; they are important regulators of hematopoiesis. The role of miRNAs in regulation of erythropoiesis is being actively defined. Some miRNAs are mainly expressed in early stages of erythropoiesis, others in late stages, and some have biphasic expression during erythroid differentiation. Some appear to have erythroid specific expression.¹⁴³ The critical role of miRNAs and their relationship to the essential transcription factors regulating erythropoiesis is outlined by the report that the pivotal transcription factors for erythropoiesis, GATA-1 and NFE-2, directly regulate and control differentiation via miRNA-199b-5p. This miRNA then targets c-Kit, an important receptor on early erythroid cells.¹⁴⁴ Some miRNAs likely play a role in the commitment to erythroid versus megakaryocytic differentiation. Thus, miRNA-18a was reported to be upregulated during erythropoiesis and downregulated during megakaryopoiesis, while miRNA-145 was upregulated in megakaryopoiesis and downregulated in erythropoiesis. Their mRNA targets and their functional significance are being defined.¹⁴⁵ LIN28B and its targeted let-7 have regulated expression during the fetal-to-adult erythroid cell transition.¹⁴⁶ Another important erythroid development and function is regulated by miRNA-351. A complex regulatory transcriptional repressor system controlling heterochromatin formation exists composed of KRAB-ZFP–mediated repression that include cofactor KAP1 (KRAB-associated protein-1). Deletion of KAP1 in mice upregulates miRNA-351 and several other microRNAs, which then downregulate Nix and mitochondrial autophagy (see “Neocytolysis” in Chap. 33), resulting in expansion of mitochondria in erythroid progenitors and severe hypoproliferative anemia phenotype.¹⁴⁷

The red cell mass is maintained and regulated by the kidney and marrow, which, under steady-state conditions, precisely replaces cells lost by senescence. Red cell mass defines pathogenesis of anemia and polycythemia. The kinetics of red cell production and destruction helps establish their pathogenesis. A number of tests have been developed to measure the three main components of red cell kinetics: red cell mass, rate of red cell production, and rate of red cell destruction. Some of these tests are simple but indirect and only semiquantitative, such as hematocrit, reticulocyte count, haptoglobin, lactic dehydrogenase, and unconjugated bilirubin concentration. Examination of the marrow allows assessment of total cellularity and relative erythroid contribution but is limited in that the kinetics of cell production cannot be inferred from a single static image, obtained from a very small fraction of the whole marrow. These tests are very useful in the aggregate but can be supplemented by more complex but direct quantitation; however, most require use of radioisotopes.

HEMATOCRIT

Packed red cell volume is commonly referred as the hematocrit (Hct). It is measured as the percentage of the volume of whole blood that is made up by red blood cells. Historically, it was measured by centrifugation but modern counters measure Hct indirectly based on red cell count and the mean red cell volume (MCV). Total-body Hct is the volume of red cells in the body divided by the total blood volume. Blood Hct is the simplest and most widely used test for estimating the size of red cell mass. In most anemic patients, blood Hct gives an excellent approximation of total red cell mass and a functional estimation of the oxygen-carrying capacity and whole-blood viscosity. Its main drawback is that it is an indirect measure that is influenced by changes in plasma volume and may not reflect the size of the red cell mass in dehydrated patients. Dehydration usually is clinically apparent and in most cases can be taken into account when evaluating the significance of a specific Hct determination. Only direct measurement of red cell mass can differentiate between relative and absolute polycythemia. However, when the Hct is greater than 60 percent, almost all patients have an increase in total red cell mass.¹⁴⁸ The extent of the increase cannot be estimated accurately from a Hct measurement alone (Fig. 32–8).

RED CELL MASS AND PLASMA VOLUME

A more direct and accurate estimate of the size of the red cell mass is obtained from labeling a known volume of red cells and determining the dilution of this label in blood. Radioactive iron is an excellent label of red cells because it is biosynthetically incorporated into hemoglobin in vivo. In experimental animals, radioactive iron can be given to a donor animal and the donor’s cells transfused into the animal whose red cell volume is being assessed. However, the radiation exposure to the donor and the hazards of transfusing allogeneic cells preclude its use in humans. Thus, almost all current clinical methods use labeling of autologous red cells in vitro by any one of a number of isotopes or biotin (Chap. 33). If studies must be performed in radiation-sensitive individuals, such as pregnant women, red cell labeling can be performed by nonradioactive chromium-123 or by biotin, which is detected with streptavidin coupled to a fluorochrome.¹⁴⁹ Among the isotopes available, chromium-51 (⁵¹Cr) is the most widely used label, although technetium-99m (^99mTc) is convenient and accurate.¹⁵⁰ Chromium in the form of the chromate ion (CrO₂⁻) readily enters the red cell and binds to globin chains. Excess isotope in the incubation mixture can be removed by washing or by using ascorbic acid to reduce the chromate ion to a nonpermeant chromic ion. Approximately 15 minutes after injection of a known amount of labeled cells, a sample of blood is obtained; its volume, Hct, and radioactivity are determined; and the total red cell volume is calculated from the equation:

where CPM = counts per minute. Sampling time is generally 15 minutes. Chromium also labels white cells; thus, one should centrifuge and remove the buffy coat before labeling if the white cell count is elevated (>25 × 10⁹/L).

No theoretical objection exists to measuring the red cell mass using labeled cells. It is independent of the Hct of the blood used to measure radioactivity, and replicate determination can be made with a coefficient of variation of approximately 1.5 percent.¹²⁶ The principal problem lies in reporting the measured red cell mass. The total red cell mass can be expressed as a volume related to body surface (mL/m²) or as a volume related to body weight (mL/kg). A committee of the International Committee on Standardization in Hematology (ICSH) has extensively examined existing data and concluded that the most reproducible expressions of red cell mass are related to body surface area estimated from height and weight¹⁵¹:

where RCM = red cell mass, S = body surface area in square meters, and Age = age in years. The calculated values, ±25 percent, included 98 percent of the measured male values and 99 percent of the measured female values.¹³

Despite the ICSH recommendation, the most common method is to report red cell mass values in terms of milliliters per kilogram. However, this method of expression gives erroneously low values in obese individuals because fat is hypovascular. A better method might be to express the red cell mass in terms of lean weight. In general, lean weight is 20 percent less than actual weight in normal males and 25 percent less in normal females.¹⁵⁰ However, estimation of lean weight in obese individuals is inaccurate. From a practical point of view, RCM probably is best reported in terms of actual weight, with mental adjustments made based on body configuration. In general, the RCM of normal females ranges from 23 to 29 mL/kg of body weight and of normal males ranges from 26 to 32 mL/kg of body weight.¹⁵¹

PLASMA LABELS

RCM also can be estimated from plasma volume. Radioactive iodine (¹²⁵I) is used to label albumin and measure its distribution volume.¹⁵² Other radioactive isotopes of iodine other than ^99mTc have been used, but ¹²⁵I has virtually supplanted all other plasma labels. Albumin labeled with radioactive iodine is commercially available, and a known amount is injected intravenously. Several blood samples are obtained within the first 15 minutes and centrifuged. CPM per milliliter of plasma is measured, plotted on semilogarithmic paper, and extrapolated to zero time. This procedure is necessary because, in contradistinction to labeled red cells, labeled albumin is removed gradually, beginning immediately after injection. Plasma volume is calculated according to the equation:

The continuous exchange of intravascular with extravascular albumin is the major problem encountered when plasma volume is measured with labeled albumin. Even with extrapolation to 0 hour, plasma volume is somewhat larger than that measured with a strictly intravascular protein such as fibrinogen.¹⁵³ Consequently, if measurement of the plasma volume is used to calculate the size of the total red cell mass, it is a less-reliable measure than determining red cell mass directly with tagged red cells. This inaccuracy is further augmented by the fact that the venous Hct used to calculate red cell mass from measured plasma volume does not reflect accurately the distribution of plasma and red cells in the body. However, from a practical point of view, the results of estimating RCM from plasma volume are surprisingly accurate and have been advocated based on simplicity and low cost.¹⁵²

When total RCM is measured with labeled red cells, the value is approximately 10 percent lower than that calculated from plasma volume and the Hct of blood. In fact, the mean Hct of blood in all of the vessels (total-body Hct) clearly is somewhat lower than the Hct measured from blood obtained from large vessels; these differences are a result of varying proportions of plasma in different size vessels.

Generally, the ratio of total-body Hct as estimated by direct measurements of red cell volume and plasma volume to the large-vessel Hct ranges from 0.89 to 0.92.¹⁵⁴ Consequently, when using the determined plasma volume to calculate RCM and total blood volume, a correction factor is necessary, and a value of 0.90 is generally used:

Recommended procedures for determination and evaluation of blood volume are outlined by the ICSH.¹⁵⁵

Under normal circumstances, most human red cells produced in the marrow live, or have the potential to live, a normal life span. Under certain conditions, however, a fraction of red cell production is ineffective; with destruction of nonviable red cells either within the marrow or shortly after the cells reach the blood.³⁸

EFFECTIVE RED CELL PRODUCTION

Effective erythropoiesis is most simply estimated by determining the reticulocyte count. Most modern automated counters measure reticulocytes by nucleic acid binding dyes such as thiazole orange using flow cytometry. These are fast reliable assays. This count usually is expressed as the percentage of red cells that are reticulocytes, but it also can be expressed as the total number of circulating reticulocytes per unit of blood (absolute reticulocyte count and corrected reticulocyte counts; equation 1 below).

Equation 1

A simple clinical method to estimate effective erythropoiesis uses the reticulocyte count to calculate the reticulocyte index (see equation 2 below).¹⁵⁶ This measurement depends on several assumptions: (1) the human red cell life span is approximately 100 days (actually approximately 115); (2) the life span is finite and, thus, the oldest 1 of 100 or 1 percent of red cells is removed (and replaced) each day; (3) the reticulocyte is identifiable as such in the blood for 1 day using supravital stain; and (4) the reticulocyte count of 1 percent in a person with a normal Hct represents normal red cell production and thus “1” is the basal reticulocyte index.

In anemic patients, two calculations are needed to measure the reticulocyte index and compare it to the normal of 1 in the basal state. To correct the reticulocyte percentage for the lower red cell count in anemic subjects, the reticulocyte percent is multiplied by the ratio of the patient’s Hct over the normal mean Hct, providing a corrected reticulocyte percent.

Conversion of the reticulocyte count (see equation 1 above) to the reticulocyte index (equation 2 below) is achieved by taking into account the estimated life span of reticulocytes. The life span of reticulocytes in blood in a normal individual is approximately 1 day. However, when red cell production is increased under conditions of erythropoietic stress, for example, in severe anemia, reticulocytes are released prematurely and circulate as reticulocytes for 2 to 4 days, except in situations with low EPO levels, as in renal insufficiency. These appear as polychromatophilic erythrocytes on Wright-stained blood film and are called stress or shift reticulocytes.

Equation 2

Accordingly, the corrected reticulocyte percent may give an erroneous impression of the actual rate of daily red cell production. To take this situation into account when estimating the rate of red cell production in anemic patients with high reticulocyte counts, dividing the corrected reticulocyte percent by a factor may provide a more accurate estimate of red cell production.¹⁵⁶ For simplicity, an average factor of 2 often is used; however, the factor depends on the degree of anemia: 1.5 in mild cases, 2.5 in moderate cases, and 3.0 in severe cases.

An example follows: A patient with autoimmune hemolytic anemia has a Hct of 10 and reticulocyte count of 70 percent. The marrow cannot increase production by 70-fold. To measure the approximate true increase, we calculate the reticulocyte index as follows: corrected reticulocyte percent = 70 × 10/45 = 15, and the reticulocyte index = 15/3= 5 × basal. Thus, marrow erythroid production in response to this severe anemia has increased fivefold, a plausible response to this severity of hemolytic anemia.

INEFFECTIVE RED CELL PRODUCTION

Ineffective erythropoiesis is suspected when the reticulocyte count is normal or only slightly increased despite erythroid hyperplasia of the marrow. Ineffective erythropoiesis was first recognized as an entity from the study of isotope incorporation into fecal urobilin following administration of labeled glycine, a precursor of heme.¹⁵⁷ Two peaks were observed: an early peak at 3 to 5 days and a late peak at 100 to 120 days. One of the sources of the early labeled peak was suggested to be the hemoglobin of red cells that had never completed their development, having been destroyed either in the marrow or shortly after reaching the blood. Subsequent studies revealed that in certain disorders, such as pernicious anemia, thalassemia, and sideroblastic anemia, ineffective erythropoiesis is a major component of total erythropoiesis. This component can be quantitated by measuring ¹⁵N-labeled glycine incorporation into the early bilirubin peaks¹⁵⁷ or ferrokinetics.³⁸ Calculated from bilirubin peaks and turnover, ineffective erythropoiesis under normal conditions amounts to approximately 4 to 12 percent of total erythropoiesis. Using ferrokinetic methods, ineffective erythropoiesis is calculated as the difference between total plasma iron turnover and erythrocyte iron turnover plus storage iron turnover (see “Ferrokinetics”). The values estimated from such studies in normal subjects are higher, ranging from 14 to 34 percent.³⁸ However, the results, both high and low, probably are misleading because none of the methods actually measures cell death, only the turnover of heme and iron. It is possible that little premature death of cells occurs in normal subjects, but much of the early release of bilirubin and iron is derived from the rim of hemoglobin extruded during enucleation of erythroblasts (Chap. 31).

TOTAL ERYTHROPOIESIS

Total erythropoiesis, which is the sum of effective and ineffective red cell production, can be estimated from a marrow examination. Films or sections from marrow aspirates and biopsies are first examined for relative content of fat and hematopoietic tissue. This examination gives an estimate of overall hematopoietic activity within the marrow space. A differential count then is performed, determining the ratio between granulocytic and erythroid precursors (M:E ratio). In a normal adult, the ratio is approximately 3:1 to 5:1. The ratio can be used to estimate whether erythropoiesis is normal, increased, or decreased (Chap. 3). The ratio is only an approximation of total erythroid activity because the ratio can be altered by changing the myeloid and erythroid components, and an aspirate or biopsy of a small segment of the marrow may not always reflect total marrow activity. These assumptions are valid as long as the marrow reflects the steady-state, if the marrow is recovering from aplasia, or is developing aplasia, it will not accurately reflect output of mature red cells. However, when used in conjunction with determination of red blood cell count and reticulocyte count, under most circumstances the ratio provides qualitative information about the rate and effectiveness of red blood cell production. A more accurate quantitation of total erythropoiesis can be made by measuring the rate of production of red cells (ferrokinetics) or, in steady-state conditions, the rate of destruction of red cells (red cell life span, bilirubin production, carbon monoxide excretion).

FERROKINETICS

In 1950, Huff and associates¹⁵⁸ described a method for measuring the rate of red cell production utilizing a simple model of iron metabolism (Fig. 32–9; Chap. 42). In this method, radioactive iron is complexed to transferrin in vitro and injected intravenously. Alternatively, ⁵⁹Fe can be injected directly intravenously as the gluconate without preincubation with the patient’s own plasma, providing enough unbound transferrin is available, because binding is almost instantaneous. The rate of clearance of the transferrin-bound iron from the plasma (⁵⁹Fe plasma T_1/2) and the subsequent uptake in the red cells are measured. From these two values and from determinations of plasma iron concentration and plasma volume, the rate of formation of red cells can be calculated.³⁸

The initial clearance of iron is exponential, and sampling during this period can be used to calculate T_1/2. In normal individuals, initial clearance averages approximately 90 minutes. Initial clearance is shorter in patients with hyperplasia of the erythropoietic tissue and longer in patients with marrow hypoplasia (see Fig. 32–9). However, the clearance rate is not a direct measurement of erythropoietic activity because it depends on the size of the pool of unlabeled, circulating iron. Consequently, calculation of the plasma iron turnover rate must include the plasma iron concentration. Clearance is expressed in milligrams of iron. The point of reference can be hemoglobin mass, blood volume, or weight, but a commonly used expression is micrograms of iron per deciliters of whole blood per day:

Under normal conditions, radioactive iron is incorporated into newly formed red cells after a few days and reaches a maximum approximately 10 to 14 days after injection (see Fig. 32–9). Normal utilization is 70 to 90 percent on day 10 to 14, a value that is so high that further increases have little significance. However, decreased utilization is an important finding and suggests immature red cells are destroyed in the marrow before they are released to the circulation (ineffective erythropoiesis) or that serum iron is diverted to nonerythropoietic tissues (marrow hypoplasia). The shape of the red cell utilization curve also is important. An early and steep rise (rapid marrow transit time) suggests a high EPO level. Finally, an early rise in utilization with a subsequent fall off suggests hemolysis.

When calculating utilization, the blood volume must be known:

Using the plasma iron clearance and utilization of iron, the red cell turnover in milligrams per dL blood for 24-hours is calculated as follows:

The normal value of red cell iron turnover is 0.30 to 0.70 mg/dL blood per 24 hours.³⁸ This range fits very well with a crude estimation of the iron used for maintaining the red cell mass in 1 dL of blood or 45 mL of packed red cells. The daily red cell production must equal the daily red cell destruction (45 mL/120 = 0.38 mL), assuming a red cell life span of 120 days. Because 1 mL of packed red cells contains approximately 1 mg of iron, a daily plasma iron turnover of 0.38 mg is needed by 1 dL of blood to maintain homeostasis.

Calculating red cell iron turnover has provided useful information about the total volume and effectiveness of erythroid tissue (Table 32–2). However, an elevated serum iron concentration gives erroneous impressions of the state of erythropoiesis. Moreover, more prolonged sampling of plasma following an intravenous injection of ⁵⁹Fe has shown that clearance is not a single exponential, but must be represented by several exponential components.¹⁵⁹ This finding has led to the introduction of more complex models of iron kinetics with a single pool of plasma iron exchanging with a number of extravascular erythroid and nonerythroid pools. Careful analysis of such models has generated computer-supported methods calculating the degree and effectiveness of erythroid activity.¹⁶⁰ Although possibly more accurate than the conventional method of calculating iron turnover, the models appear to be too cumbersome for clinical use. Moreover, even these sophisticated methods may not give an accurate account of the state of erythropoiesis. Despite a constant rate of red cell production, the plasma iron turnover was found to increase with increasing plasma iron and transferrin saturation. This finding was first thought to result from increased nonerythroid iron uptake and led to the introduction of various correction factors in the calculation of red cell iron turnover.¹⁶⁰ However, the iron in plasma is present in two pools, a diferric and a monoferric transferrin pool (Chap. 42), and the erythroid and nonerythroid receptors have a four times greater avidity for diferric transferrin than for monoferric transferrin. Consequently, total plasma iron turnover depends on the degree of saturation and does not necessarily reflect the number of transferrin receptors, presumably a critical measure of erythropoietic capacity.¹⁶¹ To measure the number of transferrin receptors, adjusting the plasma iron turnover equations for both nonerythroid uptake and degree of transferrin saturation and expressing the plasma turnover in terms of transferrin rather than iron have been proposed.¹⁶² Normal erythroid uptake of transferrin is 60 ± 12 μmol/L of blood per day, a value that has appropriately decreased and increased in patients with hypoplastic and hyperplastic marrow.