The Erythrocyte Physiology, Metabolism, and Biochemical Disorders

John W. Harvey

Department of Physiological Sciences
College of Veterinary Medicine
University of Florida
Gainesville, Florida

Mammalian erythrocytes or red blood cells (RBCs) are anucleate cells that normally circulate for several months in blood despite limited synthetic capacities and repeated exposures to mechanical and metabolic insults. Their primary purpose is to carry hemoglobin (Hb), a heme-containing protein that accounts for more than 90% of the protein within RBCs (Quigley et al., 2004). The benefits of having Hb contained within cells, as opposed to free in plasma, include the much slower turnover in blood (free Hb has a half-life of only a few hours), the metabolic capability of RBCs to maintain iron in Hb in the functional ferrous state, and the ability to control Hb oxygen affinity by altering the concentrations of organic phosphates, especially 2,3-diphosphoglycerate (2,3DPG). In addition, the presence of free Hb in plasma in concentrations normally found in whole blood would exert an osmotic pressure several times greater than that normally exerted by plasma proteins, profoundly affecting the movement of fluid between the vascular system and tissues.

Most RBCs in normal dogs, cats, horses, cattle, and sheep occur in the shape of biconcave disks (discocytes). The degree of biconcavity is most pronounced in dogs and less so in cats and horses. RBCs from goats generally have a flat surface with little surface depression; a variety of irregularly shaped RBCs (poikilocytes) may be present in clinically normal goats (Jain, 1986). The apparent benefit of the biconcave shape is that it gives RBCs high surface area-to-volume ratios and allows for deformations that must take place as they circulate.

The RBC functions of oxygen transport, carbon dioxide transport, and hydrogen ion buffering are interrelated. Each Hb tetramer can bind four molecules of oxygen when fully saturated, forming oxyhemoglobin (OxyHb). Assuming a normal arterial pO₂ of 100 mmHg and an Hb concentration of 150 g/l (15 g/dl) in blood, the presence of Hb-containing RBCs increases the oxygen carrying capacity of blood approximately 70 times that which could be transported dissolved in plasma (West, 1985).

Approximately 10% of CO₂ is transported dissolved in blood, 5% to 10% is transported bound to amine groups of blood proteins, and 80% to 85% is transported in the form of bicarbonate in normal individuals (Hsia, 1998; Jensen, 2004). Carbonic acid is formed when dissolved CO₂ combines with water. This reaction occurs nonenzymatically but is accelerated by the presence of the carbonic anhydrase (CA), also called carbonate dehydratase, enzyme in RBCs. Bicarbonate is formed by the rapid spontaneous dissociation of carbonic acid as shown:

Hb potentiates the formation of bicarbonate by buffering hydrogen ions and shifting the equilibrium of the reaction to the right. Carbamino groups are formed by the combination of CO₂ with the terminal groups of proteins. The globin of Hb is the most important blood protein in this regard. The transportation of CO₂ from the tissues to the lungs as carbamino groups is potentiated because deoxyhemoglobin (DeoxyHb) binds twice as much CO₂ as OxyHb. The formation of carbamino groups can be represented as follows:

Hb is the most important protein buffer in blood because it occurs in high concentration, has a relatively low molecular weight, and has a large number of histidine residues with pK_a values close to 7.4, enabling them to function as effective buffers. It has about six times the buffering capacity of the plasma proteins. An additional factor of importance in contributing to the effectiveness of Hb as a blood buffer is the fact that DeoxyHb is a weaker acid than OxyHb. As a result, most of the H⁺ produced in the tissues under normal conditions is buffered as a direct result of the H⁺ uptake by DeoxyHb owing to an increase in effective pK_a of Hb following release of oxygen to the tissues (West, 1985).

Primitive hematopoietic stem cells (HSCs) appear to develop in the embryo from a common precursor cell for both endothelial and hematopoietic lineages. The first HSCs appear to develop as clusters of cells in the wall of the dorsal aorta, with subsequent development in the yolk sac, placenta, and fetal liver (Baron and Fraser, 2005). Beginning in midgestation and continuing throughout postnatal life, mammalian blood cells are produced continuously from HSCs within extravascular spaces of the bone marrow. HSCs are capable of proliferation, life-long self-renewal, and differentiation. HSCs replicate only once every 8 to 10 weeks (Abkowitz et al., 2002). The term hematopoietic progenitor cell (HPC) refers to cells that form colonies in bone marrow culture like HSCs but do not have long-term self-renewal capacities. HSCs and HPCs are mononuclear cells that cannot be distinguished morphologically from lymphocytes. The presence of a transmembrane glycoprotein termed cluster of differentiation antigen 34 (CD34) has been used to identify HSCs and early HPCs, but some HSCs (possibly inactive ones) lack CD34 (Gangenahalli et al., 2006). In addition, CD34 is also present on the surface of nonhematopoietic stem cells and vascular endothelial cells (Kucia et al., 2005; Wu et al., 2005). CD34 is believed to play a role in cell adhesion (Gangenahalli et al., 2006).

Blood cell production occurs throughout life in the bone marrow of adult animals because of the unique microenvironment present there. The hematopoietic microenvironment is a complex meshwork composed of stromal (fibroblast-like) cells, endothelial cells, adipocytes, macrophages, subsets of lymphocytes, natural killer cells, and osteoblasts; extracellular matrix components; and glycoprotein growth factors that profoundly affect HSC and HPC engraftment, survival, proliferation, and differentiation (Abboud and Lichtman, 2006).

Stromal cells and endothelial cells produce components of the extracellular matrix (ECM), including collagen fibers, basement membranes of vessels and vascular sinuses, proteoglycans, and glycoproteins. In addition to providing structural support, the ECM is important in the binding of hematopoietic cells and soluble growth factors to stromal cells and other cells in the microenvironment so that optimal proliferation and differentiation can occur by virtue of these cell-cell interactions.

Collagen fibers produced by stromal cells may not have direct stimulatory effects on hematopoiesis but rather are permissive, promoting hematopoiesis by forming an inert scaffolding around which the other elements of the microenvironment are organized. Hematopoietic cells can adhere to collagen types I and VI.

Adhesion molecules (most importantly β₁-integrins) on the surface of hematopoietic cells bind to ECM glycoproteins such as vascular cell adhesion molecule-1 (VCAM-1), hemonectin, fibronectin, laminin, tenascin, vitronectin, and thrombospondin. The spectrum of expression of adhesion molecules on hematopoietic cells that will differentially bind to these ECM glycoproteins varies with the type, maturity, and activation state of the hematopoietic cells. In addition to anchoring cells to a given microenvironmental niche, binding of adhesion molecules on hematopoietic cells also plays a role in cell regulation by direct activation of signal pathways for cell growth, survival, and differentiation or by modulating responses to growth factors.

Proteoglycans consist of a protein core with repeating carbohydrate glycosaminoglycans (GAGs) attached. Major proteoglycans in the marrow include heparan sulfate, chondroitin sulfate, hyaluronic acid, and dermatan sulfate. Proteoglycans enhance hematopoiesis by trapping soluble growth factors in the vicinity of hematopoietic cells and by strengthening the binding of hematopoietic cells to the stroma.

Hematopoietic cells develop in microenvironmental niches within the marrow. HSCs are concentrated near trabecular bone where osteoblasts help regulate their numbers (Yin and Li, 2006). Erythroid cells develop around macrophages, megakaryocytes form adjacent to sinusoidal endothelial cells, and granulocyte development is associated with stromal cells located away from the vascular sinuses (Abboud and Lichtman, 2006; Kaushansky, 2006a).

Proliferation of HSCs and HPCs cannot occur spontaneously but requires the presence of specific hematopoietic growth factors (HGFs) that may be produced locally in the bone marrow (paracrine or autocrine) or produced by peripheral tissues and transported to the marrow through the blood (endocrine). All cells in the hematopoietic microenvironment, including the hematopoietic cells themselves, produce HGFs or inhibitors of hematopoiesis. Some HGFs have been called poietins (erythropoietin and thrombopoietin). Other growth factors have been classified as colony-stimulating factors (CSFs) based on in vitro culture studies. Finally, some HGFs have been described as interleukins (ILs) (Kaushansky, 2006b).

Hematopoietic cells coexpress receptors for more than one HGF on their surface. The number of each receptor type present depends on the stage of cell activation and differentiation. Binding of an HGF to its receptor results in a series of enzymatic reactions that generate signals that promote the synthesis of molecules that inhibit apoptosis, the formation of cell-cycle regulators (cyclins), and the synthesis of additional HGFs and their receptors (Kaushansky, 2006b).

Early-acting HGFs are involved with triggering dormant (G_O) primitive HSCs to begin cycling. Stem cell factor (SCF), flt3 ligand (FL), and thrombopoietin are important early factors that act in combination with one or more other cytokines such as IL-3, IL-6, IL-11, and granulocyte-CSF (G-CSF).

Intermediate-acting HGFs have broad specificity. IL-3 (multi-CSF), granulocyte-macrophage-CSF (GM-CSF), and IL-4 support proliferation of multipotent HPCs. These factors also interact with late-acting factors to stimulate proliferation of a wide variety of committed progenitor cells. Late-acting HGFs have restricted specificity. Macrophage-CSF (M-CSF), G-CSF, erythropoietin, thrombopoietin, and IL-5 are more restrictive in their actions. They have their most potent effects on committed progenitor cells and later stages of development when cell lines can be recognized morphologically (Kaushansky, 2006b).

1. Primitive Erythropoiesis

Primitive RBC production begins and predominates in the yolk-sac but also occurs later in the liver and bone marrow. Primitive RBCs are large (>400 fl in humans) generally nucleated cells with high nuclear-to-cytoplasmic ratios. Their nuclei have open (noncondensed) chromatin, and their cytoplasm contains predominantly embryonal Hb with high oxygen affinity (Segel and Palis, 2006; Tiedemann, 1977; Tsuji-Takayama et al., 2006). Like nonmammalian species, primitive RBCs enter blood as nucleated cells, but in contrast to nonmammalian species, enucleation can eventually occur in the circulation (Kingsley et al., 2004). Primitive RBCs appear to be generated in an erythropoietin (Epo)-independent manner, but their expansion and survival require Epo (Tsuji-Takayama et al., 2006). A switch to definitive erythropoiesis occurs during fetal development. Definitive erythropoiesis results in the production of smaller cells that generally extrude their nuclei before entering blood, produce fetal Hb (in some species) and adult Hb, and are highly dependent on Epo (Tsuji-Takayama et al., 2006).

2. Definitive Erythropoiesis

Oligopotent progenitor cells (including CFU-GEMM cells) are stimulated to proliferate and differentiate into BFU-E by SCF, IL-3, and GM-CSF in the presence of Epo. BFU-E proliferation and differentiation into CFU-E results from the presence of these same factors and may be further potentiated by additional growth factors. Epo is the primary growth factor involved in the proliferation and differentiation of CFU-E into rubriblasts, the first morphologically recognizable erythroid cells. CFU-E cells are more responsive to Epo than are BFU-E cells because CFU-E cells exhibit greater numbers of surface receptors for Epo (Sawada et al., 1990).

Marrow macrophages are important components of the hematopoietic microenvironment involved with erythropoiesis. Both early and late stages of erythroid development occur with intimate membrane apposition to central macrophages in so-called blood islands. Several adhesion molecules are important in forming these blood islands (Chasis, 2006). These central macrophages may regulate basal RBC production by producing both positive growth factors, including Epo, and negative factors such as IL-1, TNF-α, transforming growth factor-β, and interferon-α, –β, and –γ (Chasis, 2006; Weiss and Goodnough, 2005; Zermati et al., 2000). The finding that Epo can also be produced by erythroid progenitors suggests that these cells may support erythropoiesis by autocrine stimulation (Stopka et al., 1998). Although some degree of basal regulation of erythropoiesis occurs within the marrow microenvironment, humoral regulation is important, with Epo production occurring primarily within peritubular interstitial cells of the kidney and various inhibitory cytokines being produced at sites of inflammation throughout the body.

3. Erythropoietin

Epo is a 34 kDa glycoprotein hormone that exhibits a high degree of sequence homology among mammals (Wen et al., 1993). It is the principal HGF that promotes the viability, proliferation, and differentiation of erythroid progenitor cells expressing specific cell surface Epo receptors. The main mechanism used to achieve these effects is inhibition of apoptosis. The binding of Epo to its receptor results in autophosphorylation of the receptor and the activation of several kinases that initiate multiple signaling pathways (Eckardt and Kurtz, 2005). Early BFU-E cells do not express Epo receptors, but more mature BFU-E cells express Epo receptors and are responsive to Epo. Epo receptor copies on cell surfaces increase to maximum values in CFU-E cells and then decline in rubriblasts and continue to decrease in later stages of erythroid development (Porter and Goldberg, 1993; Prchal, 2006). Because of their Epo receptor density, CFU-E cells readily respond to Epo, promoting their proliferation, differentiation, and transformation into rubriblasts, the first morphologically recognizable erythroid cell type. High concentrations of Epo may accelerate rubriblast entry into the first mitotic division, shortening the marrow transit time and resulting in the early release of stress reticulocytes (Prchal, 2006).

Epo production in adult mammals occurs primarily within peritubular interstitial cells that are located within the inner cortex and outer medulla of the kidney. The liver is an extrarenal source of Epo in adults and the major site of Epo production in the mammalian fetus (Jelkmann, 2007). Bone marrow macrophages and erythroid progenitor cells themselves have also been shown to produce Epo, suggesting the possibility of short-range regulation of erythropoiesis (Stopka et al., 1998; Vogt et al., 1989).

The ability to deliver oxygen to the tissues depends on cardiovascular integrity, oxygen content in arterial blood, and Hb oxygen affinity. Low oxygen content in the blood can result from low partial pressure of oxygen (pO₂) in arterial blood, as occurs with high altitudes or with congenital heart defects in which some of the blood flow bypasses the pulmonary circulation. Low oxygen content in blood can also be present when arterial pO₂ is normal, as occurs with anemia and methemoglobinemia. An increased oxygen affinity of Hb within RBCs results in a decreased tendency to release oxygen to the tissues (McCully et al., 1999).

Epo production is stimulated by tissue hypoxia, which is mediated by hypoxia-inducible factors (HIFs) that are heterodimers consisting of α and β subunits. An α subunit denoted 2α is most important in Epo production, at least for definitive erythropoiesis. Both α and β subunits are continuously translated, but α subunits are labile and regulated by tissue oxygen levels. At normal tissue oxygen levels in tissue (pO₂ > 36 mmHg), α subunits are hydroxylated by prolyl hydroxylases, polyubiquitinated, and removed by proteasomal degradation. When tissue oxygen levels are low (pO₂ < 36 mmHg), α subunits are no longer hydroxylated and degraded, allowing them to translocate into the nucleus and combine with β subunits to form heterodimeric transcription factors. These HIF heterodimers activate the transcription of the Epo gene, and many other target genes, by binding to the hypoxia responsive elements (HREs) in their promoter/enhancer regions. Binding to the Epo gene results in increased Epo synthesis when tissue hypoxia is present (Gruber et al., 2007; Jelkmann, 2007).

Other tissues also exhibit Epo receptors, and Epo also stimulates nonhematopoietic actions including promoting proliferation and migration of endothelial cells, enhancing neovascularization, stimulating the production of modulators of vascular tone, and exerting cardioprotective and neuroprotective effects (Jelkmann, 2007).

Rubriblasts are large cells (approximately 900 fl in humans) that are continuously generated from progenitor cells in the extravascular space of the bone marrow. The division of a rubriblast initiates a series of approximately 5 divisions over a period of 3 to 5 days to produce about 32 metarubricytes that are no longer capable of division (Prchal, 2006). These divisions are called maturational divisions because there is a progressive maturation of the nucleus and cytoplasm concomitant with the divisions. Each division yields a smaller cell with greater nuclear condensation and increased Hb synthesis. An immature RBC, termed a reticulocyte, is formed following extrusion of the nucleus (Harvey, 2001).

Erythroid precursors have iron requirements that far exceed the iron requirements of any other cell type because of the need for Hb synthesis. Developing erythroid cells generally extract about 75% of the iron circulating in plasma (Smith, 1997).

1. Transferrin and Transferrin Receptors

FIGURE 7-1 Summary of metabolic activities of the erythroid series. Maturation progresses from left to right. The time intervals indicated are for cattle. From Kaneko, 1980, with permission.

2. Intracellular Iron Transport

A direct interorganelle transfer of iron occurs between endosomes and mitochondria (Sheftel et al., 2007). Most iron within mitochondria is incorporated into protoporphyrin IX to form heme, but some mitochondrial iron is used to synthesize iron-sulfur complexes that are important prosthetic groups for numerous proteins involved in electron transfer, metabolic, and regulatory processes (Lill et al., 2006). Some iron is presumably released from endosomes into a cytoplasmic labile iron pool (LIP). Various chaperone molecules have been proposed, but the nature of the iron in the LIP remains enigmatic (Beutler, 2006).

3. Ferritin

Iron not required for iron-sulfur complex or heme synthesis is stored as ferritin within the cytoplasm. Each ferritin molecule is composed of a protein shell of 24 apoferritin subunits surrounding a central core containing as many as 4000 iron atoms as ferric hydroxide (Arosio and Levi, 2002). Individual ferritin molecules can be visualized by electron microscopy, and large aggregates of ferritin molecules can be visualized by light microscopy when stained for iron using the Prussian blue reaction. When membrane bound, ferritin aggregates have been called siderosomes (Bessis, 1973). Iron stored as ferritin is not available for Hb synthesis in erythroid cells (Ponka et al., 1998).

4. Iron Regulation of Transferrin Receptor and Ferritin Expression

5. Siderotic Inclusions in Erythroid Cells

Anucleated RBCs containing siderotic (iron-positive) inclusions are called siderocytes. Nucleated siderocytes have been called sideroblasts in human hematology, in which terminology used for RBC precursors is generally different from that conventionally used in veterinary hematology (Bottomley, 2004). Siderotic inclusions in erythroid cells may consist of cytoplasmic ferritin aggregates or iron-loaded mitochondria. Ferritin aggregates can occur normally in nucleated erythroid cells of humans (Cartwright and Deiss, 1975), dogs (Feldman et al., 1981), and pigs (Deiss et al., 1966), but the presence of iron-loaded mitochondria is a pathological finding (Cartwright and Deiss, 1975). Electron microscopy is used to definitively identify the nature of siderotic inclusions (Fresco, 1981; Hammond et al., 1969); however, the location of iron-positive inclusions in a ring around the nucleus of a nucleated siderocyte (termed ringed sideroblast in human hematology) strongly suggests the presence of iron-loaded mitochondria (Bottomley, 2004). Conditions resulting in the pathological iron accumulation in mitochondria may induce the synthesis of a novel mitochondrial ferritin (Torti and Torti, 2002).

Except for iron deficiency, disorders in heme synthesis have the potential to cause excess iron accumulation in mitochondria (Beutler, 1995b; Fairbanks and Beutler, 1995). Experimental pyridoxine deficiency and experimental chronic copper deficiency have both resulted in mitochondrial iron overload in nucleated erythroid cells in bone marrow of deficient pigs (Hammond et al., 1969; Lee et al., 1968a). Drugs or chemicals reported to cause siderocytes or nucleated siderocytes in dogs include chloramphenicol (Harvey et al., 1985), lead, hydroxyzine, zinc (Harvey, 2001), and an oxazolidinone antibiotic (Lund and Brown, 1997).

Hb is a tetrameric protein consisting of four polypeptide globin chains each of which contains a heme prosthetic group within a hydrophobic pocket. The molecule consists of two identical alpha and two nonalpha chains that are generally classified as beta chains in adults.

1. Heme Synthesis and Metabolism

3. Control of Hb Synthesis

Even though three different pathways are required for Hb synthesis in RBC precursors and reticulocytes, minimal intermediates (iron, globin chains, or heme) accumulate in the cytoplasm of these cells. Several positive and negative feedback mechanisms account for the balanced production of these Hb components. As already discussed, an increase in the LIP limits the uptake of additional iron by decreasing TfR1 synthesis. The availability of iron also limits and, thereby controls, heme synthesis. Free “uncommitted” heme inhibits iron uptake by erythroid cells and consequently heme synthesis. In addition, free heme is essential for the synthesis of globin chains at both the transcriptional and translational levels (Koury and Ponka, 2004). Consequently, globin synthesis does not occur in the absence of heme.

4. Hb Types in Animals

Hb types are different in animal and human embryos than in fetuses or in adults. Embryonal Hbs are composed of either one or two pairs of peptide chains not found in adult Hbs (Kitchen and Brett, 1974). In higher primates, embryonal Hbs are replaced by fetal Hb composed of 2 α chains and 2 γ chains (HbF) in the fetus, followed by Hbs with 2 α chains and 2 β chains (HbA) or 2 α chains and 2 δ chains (HbA₂) after birth. The γ globin gene was deleted during evolution in artiodactyls (even-toed ungulates). The γ globin gene functions as an embryonic gene in other nonartiodactyl mammals. Among nonprimate mammals, a specific fetal globin gene is expressed only in artiodactyls. In this lineage, a duplicated β globin gene (β^F) is expressed in the fetus, rather than a γ globin gene. The entire β globin gene locus was duplicated in cattle and HbB sheep and triplicated in goats and HbA sheep during evolution (Gumucio et al., 1996). Most fetal Hb in cattle () is replaced by adult Hb types during the first 3 weeks after birth (Lee et al., 1971). Embryonal Hbs are replaced by adult Hb types during the fetal period in cats, dogs, horses, and pigs. Consequently, Hb types present in fetuses are identical to those found in adults in these species (Bunn and Kitchen, 1973; Kitchen and Brett, 1974).

FIGURE 7-2 Changes in proportion of β globin types in Hb from five newborn goats during the first 120 days of life. Fetal Hb contains β^F. HbC contains β^C. HbD differs from HbA by a single amino acid substitution in position 21 of their β chains, β^D and β^A, respectively. Modification of a figure by Huisman et al., 1969.

The switch from production of fetal Hb to the production of adult Hb appears to result from an inherent programming of hematopoietic stem cells (Wood et al., 1985). In contrast, HbC synthesis appears to be mediated by Epo, which is low at birth and increases after birth as the hematocrit decreases (see Section V.C) (Barker et al., 1980; Huisman et al., 1969). Adult sheep RBCs contain little or no HbC, but adult goat RBCs may have as much as 10% HbC (Huisman et al., 1969). Sheep and goats that synthesize HbC during the neonatal period will also increase HbC synthesis in anemic adults and in nonanemic adults treated with Epo (Vestri et al., 1983; Yu et al., 2005). Most goats and sheep with HbA have 12 genes in the β-globin locus, including the β^C gene. Cattle and sheep with HbB have 8 genes in the β-globin locus and lack the β^C gene (Garner and Lingrel, 1988). HbC was not reported to occur in three breeds of Omani goats after birth even though they were reported to be either homozygous for HbA or heterozygous for HbA and HbB (Johnson et al., 2002). The composition of the β-globin locus was not investigated in these animals.

Considerable heterogeneity of Hb types occurs in adult animals. Two or more Hb types are reported to occur in domestic animal species (Braend, 1988; Kitchen, 1974; Kohn et al., 1998, 1999; Pirastru et al., 2003; Rando and Masina, 1985). Most polymorphism of animal Hbs is determined genetically and is usually caused by multiple amino acid interchanges (Kitchen, 1974). Hb types were initially classified based on electrophoretic mobility in gels. Electrophoretic differences can result from differences in α or β chain structure. For example, HbA and HbB in sheep are attributable to differences in β chains (Garner and Lingrel, 1988), but HbA and HbB in goats are attributed to differences in α chains (Huisman and Kitchens, 1968; Pieragostini et al., 2005). Electrophoretic mobility in gels is not sensitive enough to identify many α and β chain variants, and distinctly different Hb tetramers can migrate with similar mobilities, resulting in misleading information. For example, HbB () and HbD () in goats have almost the same mobility at alkaline pH (Pieragostini et al., 2005).

Nongenetic alterations in Hb structure can also contribute to apparent Hb heterogeneity. Examples include the N-acetylation of beta chains in cat HbB (Taketa et al., 1972) and glycosylation of Hb, a function of intracellular glucose concentration and RBC life span (Higgins et al., 1982; Rendell et al., 1985). Increased glycosylation of Hb has been reported in diabetic dogs and cats (Elliott et al., 1997, 1999). Similar to the glycosylation of Hb by glucose, cyanate combines spontaneously and irreversibly with Hb to form carbamylated Hb. Cyanate forms spontaneously from urea. Dogs with chronic renal failure and long-term increases in urea concentration have larger amounts of carbamylated Hb than dogs with acute renal failure (Heiene et al., 2001).

1. Formation

Most reticulocytes are formed from metarubricytes within the extravascular space of the bone marrow by a process of nuclear extrusion that requires functional microtubules (Chasis et al., 1989) and is likened to mitosis (Bessis, 1973; Simpson and Kling, 1967). The erythroblast macrophage protein (Emp) associates with F-actin and is important in denucleation of metarubricytes, as well as blood island formation (Soni et al., 2006). Extruded nuclei quickly expose phosphatidylserine on their surfaces, which promotes their binding to, and phagocytosis by, blood island macrophages (Yoshida et al., 2005).

Reticulocyte cytoplasm contains ribosomes, polyribosomes, and mitochondria necessary for the completion of Hb synthesis. Reticulocytes derive their name from a network or reticulum that appears when stained with basic dyes such as methylene blue and brilliant cresyl green. That network is not preexisting but is an artifact formed by the precipitation of ribosomal ribonucleic acids and proteins. As reticulocytes mature, the amount of ribosomal material decreases until only a few basophilic specks can be visualized with reticulocyte staining procedures. These mature reticulocytes have been referred to as type IV (Houwen, 1992) or punctate reticulocytes (Alsaker, 1977; Perkins et al., 1995). To reduce the chance that a staining artifact would result in misclassifying a mature RBC as a punctate reticulocyte using a reticulocyte stain, the cell being evaluated should have two or more discrete blue granules that are visible without requiring fine focus adjustment of the cell to be classified as a punctate reticulocyte.

2. Metabolism

Reticulocyte metabolism and maturation have been reviewed by Rapoport (1986) and are summarized here. Immature reticulocytes continue to synthesize protein (primarily globin chains) with residual mRNA, tRNA, and rRNA formed before denucleation. About 30% of total Hb is synthesized after nuclear extrusion (Geminard et al., 2002). Synthesis of fatty acids is minimal, but phospholipids are synthesized from preformed fatty acids. Substrates for protein and lipid synthesis and for energy metabolism are provided from endogenous sources (breakdown of mitochondria and ribosomes) as well as from plasma. The reticulocyte can synthesize adenine and guanine nucleotides de novo (Rapoport, 1986).

Most ATP is generated in reticulocytes by oxidative phosphorylation in mitochondria. Glucose is the major substrate, but amino acids and fatty acids can also be utilized for energy (Rapoport, 1986).

3. Maturation into RBCs

Although loss of membrane components accounts for much of the change in membrane protein composition during reticulocyte maturation, certain proteins such as protein 4.1 and glycophorin C increase because they are still being synthesized in reticulocytes (Chasis et al., 1989). These membrane alterations result in increased mechanical stability of blood reticulocyte and RBC membranes compared to marrow reticulocyte and nucleated erythroid cell membranes (Waugh et al., 2001).

Mitochondria undergo degenerative changes in a programmed death phenomenon (mitoptosis) owing to 15-lipoxygenase attack and subsequent ATP-dependent proteolysis (Geminard et al., 2002). Degenerating mitochondria are either digested or extruded following entrapment in structures resembling autophagic vacuoles (Simpson and Kling, 1968). The polysomes separate into monosomes and decrease in number and disappear as reticulocytes mature into RBCs. The degradation of ribosomes appears to be energy dependent; it presumably involves proteases and RNAases (Rapoport, 1986).

4. Species Differences in Marrow Release

Reticulocyte maturation begins in the bone marrow and is completed in the peripheral blood and spleen in dogs, cats, and pigs. As reticulocytes mature, they lose the surface receptors needed to adhere to fibronectin and thrombospondin components of the extracellular matrix, presumably facilitating their release from the bone marrow (Telen, 2000). Residual adhesion molecule receptors on newly released reticulocytes may explain their tendency to concentrate in the reticular meshwork of the spleen (Patel et al., 1985).

Reticulocytes become progressively more deformable as they mature, a characteristic that also facilitates their release from the marrow (Waugh et al., 2001). To exit the extravascular space of the marrow, reticulocytes press against the abluminal surfaces of endothelial cells that make up the sinus wall. Cytoplasm thins and small pores (0.5 to 2 μm) develop in endothelial cells, which allow reticulocytes to be pushed through by a small pressure gradient across the sinus wall (Lichtman and Santillo, 1986; Waugh, 1991). Pores apparently close after cell passage.

Relatively immature aggregate-type reticulocytes are released from canine bone marrow; consequently, most of these cells appear polychromatophilic when viewed following routine blood film staining procedures (Laber et al., 1974). Absolute reticulocyte counts oscillate with a periodicity of approximately 14 days in some dogs, suggesting that canine erythropoiesis may have a homeostatically controlled physiological rhythm (Morley and Stohlman, 1969). Reticulocytes are normally not released from feline bone marrow until they mature to punctate-type reticulocytes; consequently, few or no aggregate reticulocytes (<0.4%), but up to 10% punctate reticulocytes, are found in blood from normal adult cats (Cramer and Lewis, 1972). The high percentage of punctate reticulocytes results from a long maturation time with delayed degradation of ribosomes (Fan et al., 1978). Reticulocytes are generally absent in peripheral blood of healthy adult cattle and goats, but a small number of punctate types (0.5%) may occur in adult sheep (Jain, 1986). Based on microscopic examination of blood films stained with new methylene blue, equine reticulocytes are absent from blood normally and rarely released in response to anemia. However, low numbers of reticulocytes have been reported in the blood of normal and anemic horses using an Advia 120 (Siemens Medical Solutions Diagnostics, Tarrytown, New York) automated analyzer (Cooper et al., 2005). Either the instrument is more sensitive than microscopic evaluation, or values reported in normal horses represent “noise” in the instrument.

5. “Stress” Reticulocytes

Except for horses, increased numbers of reticulocytes are released in response to anemia, with better responses to hemolytic anemia than to hemorrhage. When the degree of anemia is severe, basophilic macroreticulocytes, or so-called stress reticulocytes, may be released into blood. It is proposed that a generation in the maturation sequence is skipped and immature reticulocytes, about twice the normal size, are released (Rapoport, 1986). Increased Epo results in a diminution in the adventitial cell and endothelial cell barrier separating marrow hematopoietic cells from the sinus, thereby potentiating the premature release of stress reticulocytes from the marrow (Chamberlain et al., 1975). Although a portion of these macroreticulocytes apparently is rapidly removed from the circulation (Noble et al., 1990), it is clear from studies in cats that some can mature into macrocytic RBCs with relatively normal life spans (Weiser and Kociba, 1982).

1. Ineffective Erythropoiesis

2. Vitamin and Mineral Deficiencies

Folate is required for normal DNA synthesis. Folate deficiency impairs the activity of the folate-requiring enzyme thymidylate synthase (Jandl, 1987). Not only is deoxythymidylate triphosphate (dTTP) synthesis decreased, but deoxyuridylate triphosphate (dUTP) accumulates secondarily in the cell such that some becomes incorporated into DNA in place of dTTP. Cycles of excision and attempts to repair these copy errors, with limited thymidine available, result in chromosomal breaks and malformations and slowing of the S phase in the cell cycle. Consequently, erythroid precursors are often large with deranged-appearing nuclear chromatin; such cells are classified as megaloblastic cells. Folate deficiency in people causes macrocytic anemia because fewer divisions occur as a result of retarded nucleic acid synthesis in the presence of normal protein synthesis (Jandl, 1987). Possible causes of folate deficiency include dietary deficiency, impaired absorption, and drugs that interfere with folate metabolism.

Macrocytic anemias resulting from folate deficiency are rarely reported in animals. A possible case was reported in a dog on anticonvulsant therapy (Lewis and Rebar, 1979), but serum folate was not measured. Megaloblastic precursors were present in bone marrow of cats with experimental dietary folate deficiency, but hematocrits and mean cell volumes (MCVs) remained normal (Thenen and Rasmussen, 1978). However, macrocytic anemia with dyserythropoiesis was reported in a cat that appeared to have both a folate-deficient diet and defective folate absorption (Myers et al., 1996). Macrocytic anemia has been reported in folate-deficient pigs (Bush et al., 1956), but not lambs (Stokstad, 1968).

Vitamin B₁₂ (cobalamin) deficiency in people causes hematological abnormalities similar to folate deficiency because vitamin B₁₂ is necessary for normal folate metabolism in humans (Chanarin et al., 1985). In contrast, vitamin B₁₂ deficiency does not cause macrocytic anemia in any animal species (Chanarin et al., 1985). Anemia has been reported in some experimental animal studies, but RBCs were of normal size (Stokstad, 1968; Underwood, 1977), although slight increases in MCV have been reported in B₁₂-deficient goats fed diets deficient in cobalt (Mgongo et al., 1981). Cobalamin deficiency has been reported secondary to an inherited malabsorption of cobalamin in giant schnauzer dogs (Fyfe, 2000). Affected animals have normocytic, nonregenerative anemia with increased anisocytosis and poikilocytosis, neutropenia with hypersegmented neutrophils, and giant platelets. Megaloblastic changes in the bone marrow were particularly evident in the myeloid cell line. The malabsorption of cobalamin in these dogs apparently results from the absence of an intrinsic factor-cobalamin receptor in the apical brush border of the ileum. No blood or bone marrow abnormalities were recognized in kittens fed a B₁₂-deficient diet for several months (Morris, 1977), but a normocytic nonregenerative anemia was present in a cobalamin-deficient cat that probably had an inherited disorder of cobalamin absorption (Vaden et al., 1992).

A number of disorders exhibit macrocytic anemias with megaloblastic abnormalities in the marrow that mimic findings in human folate or cobalamin deficiency but have had normal serum levels of these vitamins when measured. Examples include cats infected with the feline leukemia virus (Dunn et al., 1984; Weiser and Kociba, 1983a), cattle with congenital dyserythropoiesis (Steffen et al., 1992), and myelodysplastic syndromes (Harvey, 2001). In addition, some miniature and toy poodles exhibit macrocytosis without anemia and variable megaloblastic abnormalities in the bone marrow with normal serum folate and cobalamin values (Canfield and Watson, 1989).

Abnormalities in heme or globin synthesis can result in the formation of microcytic hypochromic RBCs. Cellular division is normal, but Hb synthesis is delayed; consequently, one or more extra divisions occur in RBC development, resulting in smaller cells than normal.

Pyridoxine, vitamin B₆, is required for the first step in heme synthesis. Although natural cases of pyridoxine deficiency have not been documented in domestic animals, microcytic anemias with high serum iron values have been produced experimentally in dogs (McKibbin et al., 1942), cats (Bai et al., 1989; Carvalho da Silva et al., 1959), and pigs (Deiss et al., 1966) with dietary pyridoxine deficiency. Erythroid cells with iron-loaded mitochondria, secondary to impaired heme synthesis, have been demonstrated in pigs fed a pyridoxine-deficient diet (Hammond et al., 1969).

With the exception of young growing animals, iron deficiency in domestic animals usually results from blood loss. Milk contains little iron; consequently, nursing animals can easily deplete their body iron store as they grow (Furugouri, 1972; Harvey et al., 1987; Holter et al., 1991; Siimes et al., 1980). Microcytic RBCs are produced in response to iron deficiency (Holman and Drew, 1966; Holter et al., 1991; Reece et al., 1984), but a low MCV may not develop postnatally in species where the MCV is above adult values at birth (Weiser and Kociba, 1983b). The potential for development of severe iron deficiency in young animals appears to be less in species that begin to eat food at an early age.

Selective erythroid aplasia occurs in cats infected with FeLV subgroup C (FeLV-C), but not in cats infected only with subgroups A or B (Riedel et al., 1986). CFU-E numbers are markedly decreased but BFU-E numbers are normal in infected cats, indicating that FeLV-C inhibits differentiation of BFU-E into CFU-E (Abkowitz, 1991). The cell surface receptor for FeLV-C is called FLVCR. This receptor has recently been demonstrated to be a heme exporter (Quigley et al., 2004). Free heme is toxic to cells, and it is hypothesized that the binding of FeLV-C to FLVCR on CFU-E progenitor cells inhibits heme export from these cells, resulting in their destruction (Quigley et al., 2004).

High doses of chloramphenicol cause reversible erythroid hypoplasia in some dogs (Watson, 1977) and erythroid aplasia in cats (Watson and Middleton, 1978). Marked erythroid hypoplasia has been reported in dogs, cats, and horses given recombinant human erythropoietin (Cowgill et al., 1998; Piercy et al., 1998; Woods et al., 1997). Antibodies made against this human recombinant glycoprotein apparently cross-react with the animals’ endogenous erythropoietin.

FIGURE 7-3 Schematic model of the organization of the RBC membrane skeleton.