Emmanuel N. Dessypris
Stephen T. Sawyer
Concept of the Erythron
The entire process by which red cells are produced in the bone marrow is termed erythropoiesis. For descriptive purposes, this process can be divided into various stages, including the commitment of pluripotent stem cell progeny into erythroid differentiation, the erythropoietin-independent or early phase of erythropoiesis, and the erythropoietin-dependent or late phase of erythropoiesis. Under normal conditions, the whole process of erythropoiesis results in a red cell production rate such that the red cell mass in the body stays constant. This indicates that there are control mechanisms by which the size of the red cell mass is tightly regulated. These control mechanisms are better understood for the later rather than the earlier phases of erythropoiesis. The glycoprotein hormone erythropoietin (EPO) has been established as the major humoral regulator of red cell production (1,2). Erythropoiesis involves a great variety and number of cells at different stages of maturation, starting with the first stem cell progeny committed to erythroid differentiation and ending with the mature circulating red cell (Fig. 6.1). The whole mass of these erythroid cells has been termed the erythron (3), a concept that emphasizes the functional unity of the red cells, their morphologically recognizable marrow precursors, and the functionally defined progenitors of erythroid precursors. The concept of erythron as a tissue has thus far contributed significantly to the understanding of the physiology and pathology of erythropoiesis.
Erythroid Cells
Commitment of Stem Cell Progeny to Erythroid Differentiation
The work of Till and McCulloch (4) has provided experimental evidence for the presence in the bone marrow of cells capable of both self-renewal and production of progenies with potential for differentiation into red cells, granulocytes, and megakaryocytes. Injection of murine bone marrow cells into a sublethally irradiated syngeneic mouse leads to the formation of colonies of hematopoietic cells in the spleen of the recipient mouse after 8 to 12 days. The day 12 colonies contain cells of all three hematopoietic lineages and, in addition, cells that maintain their self-renewal capacity in the sense that they are capable of restoring hematopoiesis in a second group of sublethally irradiated mice.
The cell that gives rise to these colonies was termed colony-forming unit–spleen (CFU-S) and represents a cell identical or closely related to the multipotent hematopoietic stem cell (5). Recently, the pluripotent murine hematopoietic stem cell has been purified and characterized (6). Morphologically, the murine stem cells resemble small immature lymphocytes, and injection of as few as 30 of these cells can rescue 50% of sublethally irradiated mice. These stem cells can restore not only the hematopoiesis, but also the lymphopoiesis of the recipient mice (6).
Factors that affect commitment of stem cell progeny into a specific differentiation pathway are poorly understood and generally undefined (7,8). Although expression of lineage-specific transcription factors is the earliest molecular event associated with commitment toward a specific line, the stimuli responsible for their expression remain undefined. It is generally accepted that commitment and differentiation are irreversible events. A differentiated cell cannot regress to an undifferentiated stage or change into another differentiation pathway. Under normal conditions, once commitment occurs, differentiation proceeds fully to the stage of mature cell, which, in the case of blood cells, has a limited lifespan. Thus, differentiation is a process that leads to cell death. These concepts are generally well proven for the mature, morphologically recognizable hematopoietic cells and their progenitors, such as the marrow erythroblasts and their erythroid progenitors, but whether they apply equally well to more immature cells is less well known.
There are three major theories that address the process of commitment of stem cell progeny into a specific differentiation pathway (9). According to the stochastic theory, commitment is a random event that progressively restricts the potential for differentiation (4). This theory allows for regulatory factors to act only at later stages of hematopoiesis. This model derives its experimental support from the nonnormal (non-Poisson) distribution of lineage-specific cells in CFU-S–derived colonies (4) and in colonies derived from multilineage progenitor cells (10), as well as from the identification in vitro of progenitor cells with bipotent differentiation potential such as erythroid/megakaryocytic, erythroid/eosinophil, or neutrophil/erythroid (11,12,13). The second theory, that of the hemopoietic-inductive microenvironment, proposes that commitment of stem cell progeny to a specific pathway depends on the environment that surrounds each hematopoietic stem cell (14). This model is based on the sequential analysis of colony type in spleen colonies (CFU-S) and received further support by recent experiments with purified stem cells (6). The third theory proposes that commitment depends on humoral factors that compete among themselves at the stem cell progeny level in promoting differentiation toward one specific pathway (15,16).
Erythroid Progenitors
Erythroblasts in the bone marrow are generated from proliferating and differentiating earlier, more immature erythroid cells termed erythroid progenitors. These progenitor cells cannot be identified morphologically, but they are detectable functionally by their ability to form in vitro colonies of erythroblasts (17). The development of tissue culture techniques for cloning hematopoietic progenitor cells in semisolid culture media in vitro has led to the recognition and assay in the human and murine bone marrow of at least two erythroid progenitors, the colony-forming unit–erythroid (CFU-E) and the burst-forming unit–erythroid (BFU-E). Under the influence of EPO, these progenitors can grow in semisolid culture media and give rise to colonies of well-hemoglobinized erythroblasts.
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Figure 6.1. Schematic representation of the various stages of erythroid differentiation. BFU-E, burst-forming unit–erythroid; CFU-E, colony-forming unit–erythroid; CFU-GM, colony-forming unit–granulocyte-monocyte; CFU-MK, colony-forming unit–megakaryocyte; CFU-S, colony-forming unit–spleen. |
Colony-forming Unit–Erythroid
The CFU-E is an erythroid cell closely related to the proerythroblast (18). Under the influence of low concentrations of EPO, it gives rise (in 2 days in murine and in 5 to 8 days in human marrow) to colonies of 8 to 32 well-hemoglobinized erythroblasts (19,20,21) (Fig. 6.2). The clonal origin of these colonies has been demonstrated by glucose-6-phosphate dehydrogenase–isoenzyme analysis (22). Morphologically, CFU-E purified from progenitor cell cultures appear as immature cells with fine nuclear chromatin; a well-defined, large nucleolus; a high nuclear–cytoplasmic ratio; a perinuclear clear zone; and basophilic cytoplasm with pseudopods (23). On electron microscopy, this cell appears as a primitive blast with dispersed nuclear chromatin, a prominent nucleolus, and an agranular cytoplasm containing clumps of mitochondria and frequent pinocytotic vesicles (23). The number of CFU-E in the human marrow ranges from 50 to 400/1 × 105 light-density, nonadherent, nucleated cells and varies significantly with the methods used for cell separation and the culture conditions. The majority of the CFU-E are in a phase of active DNA synthesis (S phase) as demonstrated by a 70 to 90% killing of cells after short exposure to 3H-thymidine in vitro (3H-thymidine suicide) or after administration of cycle-specific chemotherapeutic agents in vivo (24,25,26). The size of the CFU-E compartment in intact animals depends on the levels of circulating EPO. Anemia associated with high EPO levels or administration of EPO leads to expansion of the CFU-E compartment, whereas transfusion-induced polycythemia leads to low EPO levels and to significant reduction of the CFU-E compartment (26). From a number of in vitro studies, it has been well established that the CFU-E is the most EPO-sensitive cell, carrying the highest density of EPO receptors (EPORs) on its surface, and it is also absolutely dependent on EPO for its survival. In the absence of EPO, CFU-E undergo programmed cell death (apoptosis) (23,27,28,29,30).
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Figure 6.2. A day 7 colony-forming unit–erythroid–derived colony of erythroblasts containing 16 cells. |
Burst-forming Unit–Erythroid
The BFU-E is an erythroid progenitor that is much more immature than the CFU-E, and it is more closely related to the multipotent hematopoietic stem cell, as indicated by its cell size and buoyant density and the relatively low percentage of these cells in active DNA synthesis (0 to 25%) (25,26,31,32). BFU-E can be separated from CFU-E by its slower velocity sedimentation at unit gravity (33). Morphologically, the BFU-E appears as a very immature blast cell with slightly oval, moderately basophilic cytoplasm with occasional pseudopods, very fine nuclear chromatin, and large nucleoli (29). On electron microscopy, the cytoplasm is abundant and contains polyribosomes, and the nucleus contains small amounts of clumped heterochromatin and prominent nucleoli (29). In the presence of EPO and under the influence of other factors acting on early hematopoietic cells, such as interleukin-3 (IL-3), granulocyte-macrophage colony-stimulating factor (GM-CSF), thrombopoietin, and stem cell factor, it gives rise in 5 to 7 days in mice and in 14 to 16 days in humans to clusters of many erythroid colonies containing a total of 500 to more than 5,000 well-hemoglobinized erythroblasts (Fig. 6.3). The clonal origin of the BFU-E–derived erythroblasts has been demonstrated by characterization of the type of hemoglobin produced by cells in single colonies in coculture experiments of bone marrow cells from a patient with homozygous hemoglobin C and marrow cells from another patient with homozygous hemoglobin S disease (34). The BFU-E can be considered as a progenitor of the CFU-E. Indeed, after 6 to 7 days in culture, cells generated from human BFU-E have all the functional characteristics of CFU-E (23). The concentration of BFU-E in the human bone marrow varies from 10 to 50/1 × 105 nucleated cells; however, this number fluctuates widely depending on the cell separation methods and the culture conditions. In contrast to CFU-E, BFU-E are also detectable in the peripheral blood at a concentration of 0.02 to 0.05% of light-density (<1.077), mononuclear blood cells (35,36,37). From both in vitro and in vivo experiments, it has been well established that the early stages of BFU-E proliferation and differentiation are EPO independent (26,29,32). BFUs-E can survive in vitro (for 48 to 72 hours) in the absence of EPO, but they are absolutely dependent on IL-3 for their survival (29). Only 20% of blood BFU-E express a very low density of EPORs detectable by autoradiography (29). The size of BFU-E compartment in the marrow of animals remains unaffected by the acute changes in the levels of circulating EPO induced by anemia or transfusional polycythemia (26). Anemia can induce an increase in the cycling of BFU-E without affecting their numbers (38,39), and in vitro EPO can induce BFU-E into DNA synthesis (40). In humans, chronic administration of EPO is associated with an increase in the concentration and cycling status of marrow BFU-E; however, these changes are also seen in granulocytic-monocytic, and megakaryocytic colony-forming units (CFU-GM, CFU-MK), and multilineage progenitors, indicating that, at the early progenitor cell level, the marrow responds to EPO as an organ in a non–lineage-specific manner (41). All this evidence indicates that the early stages of erythropoiesis at the BFU-E level are EPO independent, and dependence on EPO develops at a stage between BFU-E and CFU-E (29). The distinction between early (BFU-E) and late (CFU-E) erythroid progenitors, although valid, is by itself artificial. There are a variety of cells between BFU-E and CFU-E that form a continuum of erythroid progenitors at different stages of differentiation with properties between those of BFU-E and those of CFU-E. As an example, a subclass of erythroid progenitors termed mature BFUs-E has been described in human and murine marrow (24,25). These cells share properties from both CFU-E and BFU-E. They have a proliferative potential lower than BFU-E but higher than CFU-E, their cycling status is also intermediate between CFU-E and BFU-E, and they do not exhibit IL-3 dependence, but they show relative EPO dependence (24,25,28). Thus, it becomes clear that, during erythroid development, early progenitors of high proliferative potential in a relatively low cycling status with absolute dependence on IL-3 and responsiveness to but not dependence on EPO differentiate progressively through various stages into later progenitors of low proliferative potential with a high cycling status that are IL-3 independent and totally EPO dependent.
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Figure 6.3. A day 15 human bone marrow burst-forming unit–erythroid–derived burst (group of colonies) of erythroblasts containing over 1,000 cells. |
Erythroid Precursors
Erythroblast is a term first used by Ehrlich to refer to all forms of nucleated red cells, pathologic as well as normal. He classified erythroblasts into two main categories: A normal series, the normoblasts, and a pathologic series, the megaloblasts. He had observed the latter in pernicious anemia during relapse, as well as in early embryonic blood. The term megaloblast is used in this book in the pathologic sense given by Ehrlich. These abnormal cells are described and discussed in Chapter 40.
The least mature recognizable erythrocyte precursor cell is known as the pronormoblast or proerythroblast. Cells characteristic of subsequent stages of maturation are called normoblasts or erythroblasts. The various stages of maturation, in order of increasing maturity, are known as pronormoblasts, basophilic normoblasts, polychromatophilic normoblasts, and orthochromatic normoblasts. Morphologic characteristics of each stage, as seen with ordinary light microscopy after staining with Romanowsky dyes, are widely agreed on. Cytoplasmic maturation is assessed by the change in staining characteristics, as the deep blue color from the high RNA content of immature cells gives way to the red color characteristic of hemoglobin. Nuclear maturation is evaluated by the disappearance of nucleoli and the condensation of chromatin as nuclear activity ceases. The use of these features in defining stages of maturation is described in more detail in ensuing paragraphs. Electron microscopy has added a number of details to the understanding of normoblast structure (Fig. 6.4). At the pronormoblast stage, ferritin can be found in the cytoplasm. On electron microscopy, this large, iron-containing protein has the characteristic appearance of a tetrad, making positive identification possible. Ferritin may appear as isolated molecules within the cytoplasm, or it may be found in pinocytotic vesicles, or in larger structures (often surrounded by a membrane) that have been called siderosomes. The sources and metabolic fate of ferritin are discussed in Chapter 27. Its morphologic importance depends on the fact that the presence of ferritin helps to distinguish erythrocyte precursors from other immature cells.
The cytoplasm of erythrocyte precursors contains ribosomes, but for the most part, these remain free within the cytoplasm rather than being part of a well-defined endoplasmic reticulum. In the stages that are actively synthesizing hemoglobin, the ribosomes are found in units known as polyribosomes; these consist of two to eight individual ribosomes joined together by a strand of messenger RNA (mRNA). Ribosomal RNA accounts in large part for the deep blue staining that is characteristic of younger cells (pronormoblast to polychromatophilic erythroblast). Mitochondria in erythroid cells are round or oval, and the cristae are less distinct than in other cell lines. They are most numerous in the earlier stages of maturation and are located in an almost circular fashion around the nucleus. Many small vesicles that are approximately 50 nm in diameter are seen throughout the cytoplasm. They have a single membrane with an indistinct inner layer, and they sometimes contain ferritin particles (40). These vesicles are believed to arise by a process termed pinocytosis or rhopheocytosis (Fig. 6.4), whereby macromolecular substances are brought into the cell. The vesicle is formed from an invagination of the cell membrane, followed by closure to form a vacuole, which later separates from the membrane. Other cytoplasmic structures found in the young normoblast include a Golgi apparatus and occasional, randomly oriented microtubules. The latter are of unknown function but may represent remnants of the marginal band, a cytoskeletal structure characteristic of erythrocytes of lower species (43). Alternatively, they may be remnants of mitotic spindles (42).
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Figure 6.4. Schematic diagram of the ultrastructure of the normoblast, as visualized by electron microscopy. (Courtesy of Dr. Marcel Bessis.) |
Stages of Normoblastic Differentiation
The pronormoblast or proerythroblast is a round or oval cell of moderate to large size (14 to 19 μm diameter) (Fig. 6.5A and Fig. 6.6A). It possesses a relatively large nucleus, occupying perhaps 80% of the cell, and a rim of basophilic cytoplasm. The nucleus of the youngest cells in this group may differ little from that of the myeloblast. Nucleoli are present and may be prominent. There is a very thin or delicate membrane penetrated by pores, which connect the nucleoplasm to the cytoplasm (42). At this stage, only very small amounts of hemoglobin are present that cannot be detected by Giemsa stain. With the electron microscope at very high magnification (×100,000), ferritin molecules may be detected. As compared with that of myeloblasts and lymphoblasts, the cytoplasm has a tendency to be more homogeneous and condensed and may appear granular. A small, pale area may be found in the cytoplasm, probably corresponding to the Golgi apparatus (42). The nuclear chromatin is somewhat more coarse than that in myeloblasts or lymphoblasts. A number of mitochondria are seen in the cytoplasm with supravital stains or electron microscopy.
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Figure 6.5. Maturation of normoblasts as seen with transmission electron microscopy. Pronormoblast (A), basophilic normoblast (B), polychromatophilic normoblast (C),and orthochromatic normoblast (D). (Courtesy of Dr. Carl Kjeldsberg.) |
The basophilic normoblast or basophilic erythroblast is similar to the pronormoblast except that the nucleoli are no longer visible and the cell is somewhat smaller (12 to 17 μm in diameter) (Fig. 6.5B and Fig. 6.6B). Condensation of chromatin (formation of heterochromatin) begins in this stage. On light microscopy, the chromatin may appear coarse and granular; thus, there is little resemblance to the myeloblast. The nuclear structure may assume a wheel-spoke arrangement. The ribosomes reach their maximum number during this stage, and, as a consequence, the cytoplasm is deeply basophilic, even more so than in the pronormoblast. The color changes during subsequent stages reflect the increasing cytoplasmic concentration of acidophilic hemoglobin and the decreasing number with eventual disappearance of ribosomal RNA.
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Figure 6.6. Normoblasts. Pronormoblast (A); basophilic normoblast (B); early (C) and late (D) polychromatophilic normoblasts; orthochromatic normoblast with stippling (E).Magnification ×1,000; Wright stain. |
The first faint blush of hemoglobin, as indicated by one or more pink areas near the nucleus in dry fixed preparations, introduces the next stage, which is called the polychromatophilic normoblast or erythroblast (Fig. 6.5C and Fig. 6.6C and D). Increasing condensation of nuclear chromatin is observed during this stage. Irregular masses of chromatin are formed, which may stain very deeply. Nucleoli are not visible. The nucleus is smaller (7 to 9 μm) as is the cell as a whole (12 to 15 μm). The maximum number of mitochondria is found in the early phases of this stage, but as hemoglobin becomes more plentiful, mitochondria decrease in number.
When the cytoplasm possesses almost its full complement of hemoglobin, the cell is termed an orthochromatic normoblast or erythroblast (Fig. 6.5D and Fig. 6.6E). Strictly speaking, normoblasts are rarely orthochromatic in the sense that their color is the same as that of mature red cells, but this term is convenient in distinguishing the more acidophilic from the distinctly polychromatophilic stage. The orthochromatic normoblast is the smallest of the nucleated erythrocyte precursors (8 to 12 μm in diameter).
In this stage, the nucleus undergoes pyknotic degeneration, the chromatin becomes greatly condensed, and the nucleus shrinks. The nucleus may appear to be an almost homogenous mass. It may assume various bizarre forms such as buds, rosettes, clover leaves, or double spheres, or only a faint ring may remain. The changing pattern of the nuclear chromatin is not an artifact produced by fixation as it has been shown by studies with the electron microscope (44). Distortions of this process have been described. Finally, the nucleus is extruded (45) (Fig. 6.7).
After the nucleus has been extruded, the cell is known as a reticulocyte. These cells are somewhat larger than mature erythrocytes, perhaps 20% greater in volume (46). They contain certain cytoplasmic organelles, such as ribosomes, mitochondria, and the Golgi complex (Fig. 6.7C and D), and have special staining characteristics. Methyl alcohol or similar fixative agents used in staining cause a uniform precipitation of the ribosomal RNA. Such cells may appear uniformly blue or gray (diffuse basophilia), or various basophilic shades may be intermingled with pink-staining portions (polychromatophilia or polychromasia). Certain supravital staining techniques (see Chapter 1) cause the ribosomal RNA to precipitate or aggregate into a network of strands or clumps that have been termed reticulum. The presence of “reticulum” led to the term reticulocyte. Both terms are misleading because true endoplasmic reticulum does not exist in reticulocytes.
The “reticulum” may appear as a narrow band traversing the cell, it may be evenly distributed throughout the cell, or it may be so densely packed as to give the appearance of a nucleus. Generally speaking, the amount of reticulum in reticulocytes decreases as the cells mature, and, in “old” reticulocytes, only a few granules or scattered threads may be found (47). The shape and density of the network also depend, however, on a number of physical factors. Thus, the stronger the concentration of the dye, the larger and less broken up is the reticulum. Drying of the film tends to produce a fine reticulum (48). Heating tends to destroy the reticulum, with only rods and granules being demonstrable.
A change in the pH of the staining mixture toward the acid side results in a finely granular reticulum, whereas treatment with dilute alkali produces a stippled form (49).
With transmission electron microscopy, the mitochondria in reticulocytes are found to be grouped together, whereas the ribosomes are more evenly distributed (50) (Fig. 6.7C and D). There may or may not be visible ferritin molecules and pinocytotic vesicles in the cytoplasm. After supravital staining with cresyl blue, the ribosomes agglutinate into a network, and the mitochondria become swollen and distorted and their cristae disappear. As the reticulocyte matures, the various organelles decrease in number. Usually the mitochondria disappear first and the ribosomes last. At times, “autophagic vacuoles” (autophagosomes or secondary lysosomes) are found. These structures contain the degenerated organelles and may represent a mechanism whereby the unneeded cellular components are discarded.
The shape of the reticulocyte, as revealed by the scanning electron microscope, differs considerably from that of the mature erythrocyte. The reticulocyte is irregular and polylobulated, and regions of apparent cytoplasmic retraction may be found (Fig. 6.8). Only in the late stages does the bilaterally indented disc shape of the mature red corpuscle appear. Reticulocytes are more adhesive than adult corpuscles and move about in currents at a much slower rate than do mature cells (51). They appear to have a coating of globulin, at least part of which is transferrin (52). Their specific gravity is lower than that of adult corpuscles (53), and they tend to collect in the upper portions of suspensions of red cells. They vary in their resistance to hypotonic solutions (54). They have metabolic pathways that are lacking in mature red cells, including an intact tricarboxylic acid cycle (55).
Proliferation and Maturation of the Erythron
Within the erythron, cellular maturation and proliferation proceed simultaneously. All identifiable erythroid progenitors and the morphologically identifiable erythrocyte precursors are functionally destined to mature; thus, they are incapable of self-maintenance. Maintenance of the erythron at a given size and its expansion on demand are functions of the stem cell compartment (see Chapter 5). A scheme of the proliferation of the erythron and its various stages of development is presented in Figure 6.1. It takes approximately 12 to 15 days for a cell at the BFU-E stage to mature into erythroblasts. Within 6 to 8 days, a BFU-E proliferates and differentiates into a CFU-E, which needs another 5 to 7 days to proliferate and develop into basophilic erythroblasts, a period during which the CFU-E undergoes three to five successive divisions. Probably, three to five cell divisions also occur during the maturation of erythroid precursors (56). Thus, 8 to 32 mature red cells are derived from each pronormoblast. Cell division ceases at the stage of polychromatophilic erythroblasts. Orthochromatic normoblasts cannot synthesize DNA and, therefore, cannot divide.
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Figure 6.7. Formation of reticulocytes. A: Normoblast expelling nucleus. B: Normoblast nucleus after expulsion, with rim of cytoplasm. C: Reticulocyte immediately after expulsion of nucleus. D: Reticulocyte. (Courtesy of Dr. Carl Kjeldsberg.) |
Two events may decrease the theoretic yield of cells. One of these is the death of the cell before or shortly after its release from the marrow (ineffective erythropoiesis) (Chapter 26). The second is a skipped cell division, a phenomenon that results in a large hemoglobin-poor cell (Chapter 26). These events occur only to a limited extent in normal subjects but may occur much more frequently under pathologic circumstances.
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Figure 6.8. Reticulocyte as seen by scanning electron microscope. |
The biochemical events that occur at the stem cell progeny during its commitment to erythroid differentiation are unknown. The same holds true for the earlier identifiable erythroid progenitor BFU-E. This cell is totally IL-3 dependent and shows a small number of EPORs (29). Within 72 hours in culture, these cells become fully dependent on EPO (mature BFUs-E) and, in its presence, proliferate and differentiate into CFUs-E (23,29). At this stage, a number of differentiation events can be detected. From studies in murine erythroid cells, it has been established that EPO induces an increase in the synthesis of RNA and that this is closely followed by the induction of murine β-globin gene transcription (57). Other biochemical events associated with terminal erythroid differentiation include increased uptake of calcium and glucose, synthesis of transferrin receptors, increased iron uptake, hemoglobin synthesis, and appearance of erythrocyte membrane proteins (band 3 and 4.1) (58,59,60,61). Hemoglobin synthesis continues as the cell matures further into the stage of basophilic erythroblast, and, at the polychromatophilic erythroblast stage, enough hemoglobin has accumulated in the cytoplasm to give the cell the mild acidophilic reaction detected by Romanowsky stains. Hemoglobin synthesis continues through the orthochromatic stage and persists at a very low rate in the reticulocyte after denucleation. Mature red cells, being devoid of ribosomes, are unable to synthesize hemoglobin.
As previously noted, morphologic evidence of nuclear degeneration (heterochromatin formation) can be seen as early as the basophilic normoblast stage. By the orthochromatic stage, the nucleus is completely inactive, unable to synthesize either DNA or RNA. The factors leading to cessation of nuclear activity are not fully understood, but there is evidence that they may be related to intracellular hemoglobin concentration (62). Hemoglobin is found within the nucleus, possibly gaining entrance through pores in the nuclear membrane (62,63,64). After reaching a critical concentration (possibly 20 g/dl) (62), nuclear hemoglobin may react with nucleohistones, thereby bringing about chromosomal inactivation and nuclear condensation. According to this hypothesis, the number of cell divisions and the ultimate erythrocyte size are related to the rate of hemoglobin synthesis. For example, microcytic cells are produced in iron deficiency because it takes longer to reach the critical hemoglobin concentration and the generation time is unaffected; hence, more cell divisions occur before nuclear inactivation, and the resulting cell is small. In contrast, the macrocytes observed when erythropoiesis is stimulated may be the end results of an EPO-induced acceleration of hemoglobin synthesis, which in turn leads to an earlier onset of nuclear degeneration and a reduced number of cell divisions. Also consistent with this hypothesis is the observation that the mean corpuscular hemoglobin concentration remains relatively constant in a variety of mammalian species, even though erythrocyte size varies greatly (65).
After the nucleus degenerates, it is extruded from the cell (66). This process, which has been observed in living normoblasts by phase contrast microscopy (67), is completed in 5 to 60 minutes. During the extrusion process, mitochondria and cytoplasmic vesicles accumulate near the nuclear border (42,68). The role of these structures in nuclear extrusion is not entirely clear, but supravital staining with Janus green B, a mitochondrial toxin, inhibits denucleation (66). The extruded nucleus carries with it a rim of cytoplasm, including ribosomes, hemoglobin, and occasional mitochondria.
Enucleation is a process similar to cytokinesis and does not seem to depend on either the presence of extracellular matrix proteins or accessory cells (69). Among the various cytoskeletal proteins, filamentous actin plays an important role in the process of enucleation, accumulating between the extruding nucleus and incipient reticulocyte. Supporting the major role of filamentous actin in the process of enucleation is the fact that low concentrations of cytochalasin D cause complete inhibition of enucleation (69).
Within the marrow, denucleation may sometimes occur as the erythroblast traverses the endothelial cell that forms the sinus wall (70). The normoblast cytoplasm and small organelles (ribosomes and mitochondria) squeeze through endothelial, cytoplasmic pores 1 to 4 μm in diameter, but the more rigid nucleus cannot conform to this pore size. The nucleus thus becomes caught and “pitted” from the cell. Passage through the endothelial pores is not essential to enucleation, however, because the whole process can be observed in vitro (67,69). Soon after denucleation, the nucleus is engulfed by a macrophage. The cell may remain within the marrow as a reticulocyte for several days. Factors controlling release into the circulation are discussed in Chapter 5. After release, the reticulocyte may be sequestered for 1 to 2 days in the spleen (71). Here, additional maturation may occur, and the composition of the membrane lipids may be altered. As the reticulocyte matures to an adult erythrocyte, it loses its ability to synthesize hemoglobin (72). Both particulate and soluble RNA fractions appear to be catabolized by a ribonuclease. The resulting oligonucleotides are probably further degraded by phosphodiesterases and phosphatases to pyrimidine nucleotides. A specific pyrimidine 5′-nucleotidase found in reticulocytes dephosphorylates these nucleotides, and the free pyrimidine bases then can leak out of the cell (73). If the pyrimidine 5′-nucleotidase is lacking because of hereditary deficiency (73) or lead poisoning (74), RNA degradation is greatly retarded, and basophilic stippling due to retained RNA aggregates becomes very prominent.
Biosynthesis of Hemoglobin
Because hemoglobin accounts for approximately 90% of the dry weight of the erythrocyte, the biosynthesis of hemoglobin is intimately related to erythropoiesis. As detailed in the previous section, many of the morphologic criteria used in staging the maturation of erythrocyte precursors are related to hemoglobin production and content. Furthermore, the initial events associated with the differentiation of CFUs-E into erythrocyte precursors include the activation of genes relating to hemoglobin synthesis (57).
Three complex metabolic pathways are required for synthesis of hemoglobin, corresponding to the three structural components of hemoglobin: Protein (globin), protoporphyrin, and iron. The first two of these are discussed in the pages to follow. Iron metabolism is described in Chapter 27.
Globin Synthesis
Globin Genes and the Structure of Chromatin
Distinct structural genetic loci exist for each of the known normal polypeptide chains in hemoglobin. Thus, there are α, β, γ, δ, and ∊ genes. In most human populations, the α genetic locus is duplicated, and there are four (two pairs of) identical α genes in normal subjects (75,77). There are also at least two different pairs of γ genes, one (Gγ) coding for a γ-chain with glycine at position 136 (H14) and another (Aγ) coding for a γ-chain with alanine at the same position (78). In contrast, there appears to be only one pair of genes coding for β- and δ-chains, respectively. (See also Chapter 26.)
Genetic evidence has indicated that the α and β genes are not linked, and it is now clear that they are located on different chromosomes (79). The α-gene cluster (approximately 30 kb) is located on the short arm of chromosome 16 and contains also the locus encoding for the ζ-chain (80). The β-gene cluster (approximately 50 kb) is located on chromosome 11 and includes the genes for the Gγ-, Aγ-, δ-, and ∊-globins (80,81). A schematic representation of the α- and β-gene clusters is shown in Figure 6.9. Only 5 to 10% of the genetic material in erythroblasts is transcriptionally active, and the globin structural genes are included in this fraction (82). These active genetic regions of DNA make up the open portion, or euchromatin, of nuclear material, whereas unexpressed genes are included in the condensed, or heterochromatin, fraction. Differentiation of erythroid progenitors to erythroblasts is accompanied by the activation of the genes involved in erythroid differentiation, including the globin genes (57,83). The modulation of expression of genes is imposed by chromatin structure, which includes not only strands of DNA, but also histone and nonhistone proteins.
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Figure 6.9. Organization of the human globin gene clusters on chromosomes 16 and 11. Solid areas within genes represent coding sequences; open areas represent intervening sequences. Each cluster includes pseudogenes (Ψζ, Ψα, Ψβ), which have sequence homology to functional genes but include mutations that prevent their expression. |
The structure of chromatin by electron microscopy has been compared to beads on a string (82,83). The beads represent nucleosomes, each of which consists of eight histone molecules, an octamer, associated with approximately 200 base pairs of DNA. These nucleosomes are distributed randomly along the DNA double helix and are joined by “spacer regions” of DNA, 40 to 100 base pairs long, associated with a specific histone (H1). In general, the association with histones is believed to limit accessibility to the genetic material by RNA polymerase, thereby preventing or retarding transcription. However, most current evidence suggests that active globin genes in erythroid cells are incorporated into structures at least resembling nucleosomes (82). Genes that are not transcriptionally active are closely associated with chromatin and may have an increased number of methylated nucleotides (84,85,86). Under these conditions, the genes are insensitive to nuclease digestion. Activation of these genes is associated with modification of the chromatin structure induced by transacting factors. Thus, genes associated with erythroid development are resistant to nuclease digestion in nonerythroid or early noncommitted hematopoietic cells. As the cell matures within the erythroid line of differentiation, these genes are progressively activated as indicated by the appearance of nuclease-sensitive sites in areas encoding for globin genes that include also a number of regulatory sequences (82,83,87,88).
Transcription and Messenger RNA Processing
Synthesis of RNA takes place under the influence of large complex enzymes (or groups of enzymes) called RNA polymerases (89). Three such enzymes (or groups of enzymes) have been described: Type I(A), which transcribes the genes for most ribosomal RNA and is found largely in the nucleolus; type II(B), which transcribes genes for “unique sequence” RNAs, including the globin messenger; and type III(C), which transcribes genes for transfer RNA (tRNA) plus a low-molecular-weight (MW) (S) fraction of ribosomal RNA. Types II(B) and III(C) RNA polymerase are found in the nucleoplasm outside of the nucleolus. Of the two DNA strands in the double helix, only one, the “sense strand,” appears to be translated in vivo.
Globin mRNA, like most eukaryotic mRNAs, is synthesized in a precursor form that is two to three times as long as the molecule that ultimately serves as the template for protein synthesis (75,82,83). These precursor molecules are called heterogeneous nuclear RNAs. They have relatively short half-lives, on the order of 15 to 30 minutes.
The heterogeneous nuclear RNA molecule undergoes “processing” to be converted into the final mRNA (90). Processing includes at least three posttranscriptional events: “Capping” at the 5′ end of the molecule, polyadenylation at the 3′ end, and “splicing,” which results in removal of so-called intervening sequences or introns. The last are untranslated sequences of unknown function that interrupt a translated sequence (91). In mouse β-globin heterogeneous nuclear RNA, for example, a long intervening sequence occurs between the codons for amino acids 104 and 105. This “unnecessary” segment must be “edited out.” The final mRNA molecule contains 675 to 750 nucleotides.
The primary structure of mRNA can be divided into four regions: The 5′ untranslated region (which includes the cap), the translated or coding region, the 3′ untranslated region, and the polyadenosine region. The “cap” is characterized by an atypical 5′-triphosphate-5′ linkage with guanosine-5-triphosphate (GTP) and methylation of adjacent nucleotides. This structure appears to be essential for maximal translational activity, presumably because it affects the initiation of translation; however, its exact role has not been determined. The cap is followed by an untranslated region of 36 nucleotide bases in α-globin mRNA and 53 bases in β-globin mRNA. The difference may explain the observation that β-mRNA is translated more efficiently than α-mRNA (82,83). Normally, this relatively inefficient translation of the α-chain is compensated for by an increased amount of α-mRNA.
The translated sequence begins with an initiator sequence of three bases (AUG) followed by a sequence of triplet codons, each of which corresponds to an amino acid in globin, according to the genetic code. The translated sequence ends with a terminator codon (UAA), which is followed by a noncoding area of undefined function.
Finally, the molecule ends with a polyadenosine region, of variable length, that probably affects the stability or half-life of the molecule. The number of adenosine residues appears to decrease as mRNA ages (92); in older reticulocytes, mRNA may contain little or no polyadenosine.
Translation
Translation is a ribosomal process whereby a polypeptide chain is synthesized according to the pattern provided by the sequence of codons in mRNA. The process has been divided into four major stages: Activation, initiation, elongation, and termination. The activation step involves formation of an ester linkage between amino acids and specific tRNAs in the cytoplasm. This process requires adenosine triphosphate, magnesium, and specific enzymes known as aminoacyl-tRNA synthetases. Initiation of polypeptide chain formation occurs when a special, methionine-bearing, initiator tRNA (tRNAmetF) becomes aligned with the initiator codon (AUG) in mRNA on the ribosome (82,83). This complex step requires at least eight protein initiation factors (93). One of these (erythrocyte initiation factor [eIF]-2) forms a complex with GTP and tRNAmetF, which attaches to the small (40S) ribosomal subunit. Next, mRNA and the larger (60S) ribosomal subunit are added under the influence of other initiation factors. The methionine molecule that begins the polypeptide chain at the N-terminal is eventually cleaved from the protein.
As the polypeptide chain elongates, specific amino acid–bearing tRNAs become attached to the ribosomes (82,83). Each tRNA has an anticodon sequence corresponding to the specific codon in mRNA. As each tRNA is positioned, its amino acid becomes attached by a peptide bond to another amino acid previously bound to the ribosome. Formation of the peptide bond is a property of peptidyl transferase activity associated with the large ribosomal subunit. At least two factors are required for ribosomal binding of tRNA. The other (“translocase”) is necessary for the movement of codons along the ribosome, a process using energy from GTP. A single mRNA molecule may have several ribosomes attached to form polyribosomes. When the terminator codon (UAA) is reached, the polypeptide chain is completed and released from the ribosome. This termination step requires GTP and one or more “releasing factors” (93).
Tetramer Formation
Once the polypeptides are released from the ribosomes, they quickly and spontaneously form αβ dimers and α2β2 tetramers. Apparently, no enzymes or cofactors are involved in this process.
Regulation of Globin Synthesis
Heme is of particular importance in controlling the rate of globin synthesis (94,95). It stimulates globin synthesis in intact reticulocytes and cell-free systems, and, in its absence, polyribosomes disaggregate (96,97,98). The major effect of heme is exerted on the chain-initiation step in translation. In the absence of heme, an inhibitor of globin synthesis accumulates (99,100). This inhibitor acts by phosphorylating the initiation factor (eIF-1) that promotes binding of tRNAmetF to ribosomes (101,102). Heme not only inactivates the inhibitor (102), but also stimulates an enzyme that dephosphorylates eIF-2 (82).
Heme may also exert effects on transcription, on mRNA processing, or on both (103). In mouse erythroleukemia cells, hemin behaves as an inducer. The role of heme in mammalian protein synthesis appears to extend beyond the erythrocyte; heme-dependent factors for the initiation of protein synthesis have been demonstrated in liver and brain (104,105).
Heme Biosynthetic Pathway
Strictly defined, heme is the ferrous iron complex of protoporphyrin IX; however, the term heme is also used in the generic sense to indicate iron protoporphyrin IX without regard to the oxidation state of the iron. Thus, hemoglobin, peroxidase, and cytochrome c are all “heme” proteins even though the iron is ferrous in hemoglobin, ferric in peroxidase, and either ferric or ferrous in cytochrome c (106,107,108,109,110,111).
Protoporphyrin IX is a tetrapyrrole, a complex structure (“macrocycle”) made up of four pyrrole rings joined together (Fig. 6.10). The four pyrrole rings are designated A, B, C, and D. At the periphery of the tetrapyrrole, there are eight sites where side chains may be located. Positions 1 and 2 are on the A ring, 3 and 4 on the B ring, and so forth. The four bridge carbons (methene groups) are designated α, β, γ, and δ. Thus, protoporphyrin IX may be described as 1,3,5,8 methyl, 2,4-vinyl-6,7, propionic acid–porphyrin, or porphine. Porphine, the so-called parent compound, is a hypothetical tetrapyrrole with only hydrogen atoms at the eight peripheral positions.
The designation protoporphyrin IX arises from the fact that, of the 15 possible isomers of protoporphyrin, the ninth one that Hans Fischer (who developed the foundations of porphyrin chemistry) and his coworkers synthesized was the same as the protoporphyrin that they prepared from heme. The IX isomer is the only one found in nature. Porphyrins, by definition, are cyclicly conjugated tetrapyrroles. As such, they have a number of common properties. They are very stable, essentially flat molecules. The macrocyclic ring itself has little or no affinity for water. Porphyrins might be compared to phonograph records—both are flat, have a hole in the middle, and tend to stack and stick together. In the case of phonograph records, the sticking together is largely electrostatic; in stacked porphyrins, the attractive forces are the consequence of πbonding (interchange of π electrons) as well as van der Waals forces.
All porphyrins are intensely colored and have many common light absorption characteristics. All have an extremely intense absorption band at approximately 400 nm, the so-called Soret band. In addition, they all have four absorption bands in the visible region of the spectrum. All porphyrins fluoresce, but fluorescence is characteristically lost when metals are bound to form metalloporphyrins. Exceptions include Mg-porphyrins and Zn-porphyrins, which fluoresce despite their metal content (Chapter 30). Of all the known porphyrins, only five are of importance in human physiology and pathophysiology: Uroporphyrin (two isomers), coproporphyrin (two isomers), and protoporphyrin (one isomer).
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Figure 6.10. The chemical structure of protoporphyrin IX. |
If, under appropriate conditions, a porphyrin is reduced to the fullest extent (e.g., with sodium amalgam or sodium borohydride), a total of six hydrogen atoms are added to the molecule, four to the bridge carbon positions and two to the pyrroline rings. (Compare the structures of protoporphyrinogen III and protoporphyrin IX in Figure 6.11.) The fully reduced compounds are called porphyrinogens. They are colorless, do not fluoresce, cannot bind metal ions, and are extremely unstable with regard to oxidation. Only the fully reduced compounds, uroporphyrinogen and coproporphyrinogen, are true intermediates in heme biosynthesis. Once these porphyrinogens are oxidized to their corresponding porphyrins, they can no longer function as substrates for the heme biosynthetic enzymes and must eventually be excreted in the urine and stool.
Uroporphyrinogen and coproporphyrinogen can occur in four isomeric forms. Of these, only two are known to occur naturally in mammalian tissues, namely, the I and III isomer forms. Without exception, all biologically functional tetrapyrroles are derived from uroporphyrinogen III. Uroporphyrinogen I and coproporphyrinogen I are useless by-products of heme synthesis. They are biologic anomalies and can be thought of as biochemical mistakes (see Chapter 30). Once formed, most uroporphyrinogen I is enzymically decarboxylated to coproporphyrinogen I and excreted as the oxidized compound, coproporphyrin I.
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Figure 6.11. Heme biosynthetic pathway. Ac, acetate; ALA, δ-aminolevulinic acid; CoA, coenzyme A; CoAS, succinyl-CoA; CoASH, uncombined coenzyme A; COPRO’GEN, coproporphyrinogen; PBG, porphobilinogen; PLP, pyridoxal 5′-phosphate; Pr, propionate; PROTO’GEN, protoporphyrinogen; URO’GEN, uroporphyrinogen; Vi, vinyl. (Modified from Bottomley SS, Eberhard Muller U. Pathophysiology of heme synthesis. Semin Hematol 1988;25:282.) |
The difference between the type I and type III isomers is apparent on examination of the D ring. In the type I isomer, the 7 and 8 positions are occupied by acetate and propionate, respectively (as are positions 1 and 2, 3 and 4, and 5 and 6). In the type III isomer, the order in the D ring is reversed, and propionate and acetate are at positions 7 and 8, respectively. In the case of protoporphyrin IX, propionate is at position 7, and position 8 is occupied by a methyl group derived from the decarboxylation of an acetate group.
The first committed step in the biosynthesis of heme (Fig. 6.11) is the condensation of glycine and succinyl coenzyme A to yield δ-aminolevulinic acid (ALA). This reaction is highly exergonic, because it involves the cleavage of the thio-ester bond of succinyl coenzyme A and is essentially irreversible. These features suggest that the enzyme catalyzing this reaction (ALA synthase) plays a key regulatory role in the biosynthesis of heme. Indeed, considerable experimental evidence supports this contention.
In the next reaction, two molecules of ALA undergo condensation, catalyzed by the enzyme ALA dehydrase or dehydratase, to yield the monopyrrole porphobilinogen with the concomitant loss of two molecules of water. This is the only known biologic reaction in which two identical molecules combine in such a way that the product is not simply a dimer but a distinctly different molecule. The product, porphobilinogen, is the primary building block for all natural tetrapyrroles, including the hemes, the chlorophylls, and the vitamin B12 derivatives (cobalamins).
The third reaction occurs in the presence of a pair of enzymes acting in concert (uroporphyrinogen I synthase or porphobilinogen deaminase and uroporphyrinogen III cosynthase or uroporphyrin III synthase). Four molecules of porphobilinogen condense to yield first an intermediate compound called hydroxymethylbilane (HMB), which, under the action of uroporphyrin III synthase, is rapidly converted to the first macrocyclic precursor of all tetrapyrroles, uroporphyrinogen III. While remaining fully reduced, uroporphyrinogen III then undergoes a series of four decarboxylation steps catalyzed by the enzyme uroporphyrinogen decarboxylase. All four acetate side chains are sequentially decarboxylated at positions 1, 3, 5, and 8, yielding in turn CO2 and methyl groups. All of the intermediate decarboxylation products have been isolated and identified. The final product of this series of decarboxylation reactions is coproporphyrinogen III, again, a fully reduced tetrapyrrole.
Coproporphyrinogen III is transported by an unknown mechanism from the cytosol into the mitochondrion for the three subsequent reactions that yield heme. First, the propionic acid side chains at positions 2 and 4 are oxidatively decarboxylated by the enzyme coproporphyrinogen oxidase, resulting in the formation of vinyl groups. The product of this reaction is protoporphyrinogen IX. Protoporphyrinogen IX undergoes spontaneous oxidation to protoporphyrin in vitro, but, in vivo, an enzyme, protoporphyrinogen IX oxidase, is required to catalyze this reaction.
Protoporphyrin IX next combines with ferrous iron to yield the final product, the metalloporphyrin ferrous-protoporphyrin IX (heme). This reaction is catalyzed by the enzyme ferrochelatase or heme synthase.
Biosynthesis of δ-Aminolevulinic Acid
The first committed precursor of protoporphyrin IX is an aminoketone, ALA (112). Isotopic labeling studies demonstrated that the precursors of ALA are succinyl-coenzyme A (CoA) and glycine (113). Succinyl-CoA is generated mainly by the oxidative decarboxylation of α-ketoglutarate in the citric acid cycle. The direct formation of ALA from succinyl-CoA and glycine was first demonstrated in avian erythrocytes and in photosynthetic bacteria (114,115).
The enzyme responsible for catalyzing the condensation of succinyl-CoA and glycine is ALA synthase. The products of the condensation reaction are ALA, CO2, and uncombined CoA. That pyridoxal phosphate is a cofactor in the reaction was first suggested by nutritional studies in pigs (116). Subsequently, the specific defect in heme biosynthesis caused by pyridoxine deficiency was demonstrated (117,118). The deficient synthesis of heme from glycine in erythrocytes from pyridoxine-deficient animals was corrected by the addition of pyridoxal phosphate (118), whereas the synthesis of heme from exogenously supplied ALA was normal in the pyridoxine-deficient cell and was unaffected by the addition of pyridoxal phosphate (118).
In mammalian cells, ALA synthase first appears in the cytosol, where it is synthesized on polyribosomes (119). The enzyme is then transported by an unknown mechanism to its functional site inside the mitochondria. In the process, the enzyme is modified so that the apparent MW of the mitochondrial enzyme is much less than that of the cytosol enzyme. The cytosol enzyme is probably a precursor of the mitochondrial enzyme and either is multimeric or exists in association with other proteins (120).
The genes encoding for human ALA synthase have been identified and cloned (121). There are two forms of this enzyme, one specific for erythroid cells and the other present in all other tissues. The erythroid-specific enzyme is encoded by a gene present in chromosome X, whereas the nonerythroid form of the enzyme is encoded by a gene on chromosome 3 (122,123). An alternative way of generating ALA similar to one found in plants may exist in mammals. Plants form ALA directly from a five-carbon precursor by a transamination reaction (104,124) starting with α-ketoglutaric acid. This reaction is not confined to plants because an enzyme catalyzing such a transamination has been found in mammalian liver, as well as in plants, algae, and bacteria. The mammalian enzyme, alanine-dioxovalerate aminotransferase, has been highly purified from mitochondria (104). The capacity of this enzyme to form ALA is far greater than that of ALA synthase from the same mitochondria, but its physiologic significance in mammals remains unknown.
Biosynthesis of Porphobilinogen
The monopyrrole porphobilinogen is well established as a precursor of the hemes, the chlorophylls, and the cobalamins (125). Because all known forms of life require at least one of these classes of tetrapyrroles, it follows that porphobilinogen is biologically ubiquitous. Porphobilinogen is formed by the condensation of two molecules of ALA and the loss of two water molecules. The enzyme that catalyzes this reaction is ALA dehydrase.
Throughout the animal kingdom, ALA dehydrase is a soluble enzyme found in the cytosol (107). It is abundant in tissues such as bone marrow and liver where heme biosynthesis is active. It is also active in mature, circulating erythrocytes, even though these cells are not actively synthesizing heme. Its persistence results from the enzyme’s inherent stability. It is inhibited by heavy metals, particularly lead, and under properly controlled conditions can serve as an index of environmental pollution by lead (126,127).
The mammalian enzyme is an octamer of 31-kd subunits containing eight atoms of zinc required for stability and activity (122). Free sulfhydryl groups (–SH) are also essential for activity of ALA dehydrase from all sources (120,128), and these –SH groups seem to be protected by the zinc. Three isoforms of this enzyme have been reported, and the gene encoding for the enzyme has been identified and located on chromosome 9 (129,130).
The quantity of enzyme present in most organisms greatly exceeds the amount needed by the organism to synthesize ALA (110). Thus, it would appear not to play a regulatory role in heme synthesis. However, several of the characteristics of ALA dehydrase are similar to those “expected” of regulatory enzymes. It is, for instance, inhibited by heme (107,131), and its kinetic properties suggest that the binding of the first molecule of ALA to the enzyme surface influences the affinity of the enzyme for the second molecule of ALA (132).
Biosynthesis of Uroporphyrinogens I and III
Porphobilinogen is a rather unstable, chemically reactive molecule. Within a few hours, a solution of porphobilinogen exposed to air and light develops a deep orange-red color. The color results from the formation of porphobilin, a poorly defined mixture of mono-, di-, and tripyrrolic oxidation products (107). This phenomenon can be observed in the urine of patients with acute intermittent porphyria, who excrete large quantities of porphobilinogen (Chapter 30).
If porphobilinogen is incubated in solution at an acid pH, nonenzymic condensation occurs and uroporphyrinogen is formed (107). All four possible isomers of uroporphyrinogen are formed under these conditions. The reaction is often referred to as a “head-to-tail” condensation because of the apparent orientation of the precursor molecules. The penultimate reaction product apparently is an open-chain tetrapyrrole called HMB. On loss of the amino group attached to ring A, the –CH2+ can attack the free α-position on the D-ring pyrrole, thus forming the tetrapyrrole macrocycle. In vivo at neutral pH, this series of reactions is catalyzed by the cytosolic enzyme HMB synthase, formerly known as uroporphyrinogen I synthase or porphobilinogen deaminase. Two tissue-specific isoenzymes have been reported for HMB synthase, an erythroid tissue–specific and a nonerythroid form, both of which are products of a single gene located on chromosome 11q23 (107,133,134,135). In vivo, however, this enzyme works in concert with a second enzyme, uroporphyrinogen III cosynthase (136), to form uroporphyrinogen III, the precursor of all known functional tetrapyrroles.
The molecular mechanism by which uroporphyrinogen III cosynthase effects the “turning around” of the D ring has been studied intensively (107,137,138). Uroporphyrinogen III cosynthase does not actually isomerize uroporphyrinogen I: The cosynthase does not use uroporphyrinogen I as a substrate nor does it use porphobilinogen. The molecular mechanism involved in this reaction has been clarified by nuclear magnetic resonance spectroscopy (125,137,138,139,140). Studies involving this technique have made it apparent that HMB synthase first catalyzes the head-to-tail condensation of four porphobilinogen molecules, yielding the aminomethyl tetrapyrrole. The tetrapyrrole is then deaminated, yielding a macrocycle that has been termed preuroporphyrinogen (139,140). Two structures for preuroporphyrinogen have been proposed (139,141). Filtration experiments have shown that preuroporphyrinogen is released from the surface of uroporphyrinogen I synthase and serves as the substrate for uroporphyrinogen III cosynthase (140). The cosynthase opens the bond linking the methylene bridge carbon to the D ring, allowing the D ring to rotate 180 degrees about the carbon–nitrogen bond linking rings A and D. A new carbon–carbon bond is formed linking rings C and D (1,3-sigmatropic shift). The carbon–nitrogen bond then opens, and a new carbon–carbon bond is formed linking rings A and D (1,5-sigmatropic shift) and yielding uroporphyrinogen III (140).
Biosynthesis of Coproporphyrinogen III
The formation of coproporphyrinogen III is accomplished by the enzymic decarboxylation of the four acetic acid side chains of uroporphyrinogen III (108), a reaction catalyzed by what has been assumed to be a single cytosolic enzyme, uroporphyrinogen decarboxylase. The decarboxylation proceeds in a clockwise fashion, starting with the acetic acid on the D ring of uroporphyrinogen III and continuing with the successive decarboxylations of the acetic acid residues on rings A, B, and C (142).
The precise definition of the substrate for the enzyme, as well as for the reaction product, remains unsettled. Partially decarboxylated porphyrins containing seven, six, and five carboxyl groups are demonstrable in small amounts in both the urine and feces of humans. All of these partially decarboxylated porphyrinogens also can serve as substrates for uroporphyrinogen decarboxylase (142). Although uroporphyrinogen I and uroporphyrinogen III appear to be used equally well as substrates (143,144,145,146), there does appear to be isomer preference for the 7-carboxyl substrate (the I isomer is the preferred) (145,147) and the 5-carboxyl substrate (the III isomer is the preferred) (114). Multiple “substrates” and multiple “reaction products” suggest that multiple enzymes may be involved in the four decarboxylation steps. However, studies with purified preparations of uroporphyrinogen decarboxylase have shown that this enzyme catalyzes all four decarboxylations (146,148,149,150,151).
Biosynthesis of Protoporphyrinogen IX
The formation of protoporphyrinogen IX from coproporphyrinogen III is catalyzed by the enzyme coproporphyrinogen oxidase, a mitochondrial enzyme distributed in two fractions, a major fraction in the intermembrane space and a minor one in the intermembrane–matrix complex (152). This enzyme has an absolute requirement for molecular oxygen (153,154). It sequentially and oxidatively decarboxylates the propionic acid side chains in rings A and B of coproporphyrinogen III to form vinyl groups (155). At the same time, the propionic acid groups in pyrrole rings C and D are not decarboxylated. It has been assumed that they are essential for binding of the substrate to coproporphyrinogen oxidase (120), but more recent studies make this assumption unlikely (156). For coproporphyrinogen III to be metabolized, it must cross the outer mitochondrial membrane. It is not known if crossing the membrane is the result of a passive transfer or an energy-requiring step.
Biosynthesis of Protoporphyrin IX
The product of the coproporphyrinogen III oxidase reaction is protoporphyrinogen IX. To serve as a substrate for the final enzyme in the pathway, heme synthase (ferrochelatase), protoporphyrinogen IX must first be oxidized to protoporphyrin IX, the only time in the heme biosynthetic pathway that the oxidized porphyrin serves as substrate. Although protoporphyrinogen IX is easily oxidized nonenzymatically to protoporphyrin in vitro, an enzyme is required to catalyze this reaction in vivo. A membrane-associated, mitochondrial, oxidizing enzyme—protoporphyrinogen IX oxidase—was first found in yeast (156) and later was demonstrated in mammalian cells, including rat liver, human fibroblasts, reticulocytes, and leukocytes (158). The enzyme is heat labile and is inactivated by proteolytic digestion.
Biosynthesis of Heme
The insertion of ferrous iron into protoporphyrin IX to form heme is catalyzed by the enzyme heme synthase (ferrochelatase, protoheme ferrolyase, EC 4.99.1.1). Heme synthase is either tightly bound to or is an integral part of the inner mitochondrial membrane (159). Heme synthase activity has been partially purified from avian erythrocytes (159,160), human and rat liver (159,161), rabbit reticulocytes (162), human bone marrow (163), microorganisms (164), and plant tissues (165,166).
Although no cofactors have clearly been demonstrated to be required by heme synthase, a number of studies have suggested a role for pyridoxal-5-phosphate (167). It seems clear that the in vivo substrates for heme synthase are ferrous iron and protoporphyrin. However, in vitro, the enzyme catalyzes the incorporation of several metals (iron, cobalt, and zinc) into several dicarboxylic porphyrins (protoporphyrin, mesoporphyrin, and deuteroporphyrin) (107). Under strictly anaerobic conditions, the ferrous iron–protoporphyrin chelate can be formed nonenzymatically (157,168). This finding has led to the speculation that nonenzymic formation of heme may occur in vivo, but most evidence supports a catalytic role for the enzyme heme synthase. In addition, there is evidence that heme synthase may play a regulatory role in the biosynthesis of heme.
Regulation of the Heme Biosynthetic Pathway
δ-Aminolevulinic Acid Synthase
The regulation of a biosynthetic pathway is generally effected at the first enzymatic reaction synthesizing a precursor compound committed to ultimate incorporation into the final product (169). Frequently, such reactions are strongly exergonic and essentially irreversible. These generalizations hold true for the heme biosynthetic pathway. Control of the pathway is exerted through ALA synthase. The amount of ALA synthase present is regulated by induction and repression of enzyme synthesis (120). In higher organisms, this enzyme has a short half-life, allowing a rapid response to changes in the demand for heme and, thus, ALA (170).
Most studies involving the induction of mammalian ALA synthase have used hepatic tissue as the source of the enzyme, but ALA synthase has also been shown to be inducible in Friend leukemic cells (171,172), adrenal gland (173), kidney (174,175), and bone marrow (176). Hepatic ALA synthase may be induced by a number of chemicals, drugs, and nonglucocorticoid steroids (120). The amount of enzyme may increase by a factor as great as 300 (177). Conclusive immunochemical data have demonstrated that inducing agents do not merely increase the activity of preformed ALA synthase; rather, they bring about an increase in the absolute amount of enzyme (178). Compounds that inhibit protein synthesis block the expected effects of inducing agents (162,179). In erythroid cells, however, the erythroid-specific δ-ALA synthase is constitutively produced at such high levels that this step in heme synthesis does not seem to be rate limiting, and enzymes involved in later steps of the heme biosynthetic pathway may play a more important regulatory role (180).
Heme plays a critical central role in the regulation of ALA synthase activity. Heme directly inhibits the catalytic activity of the preformed enzyme in a classic example of negative feedback inhibition (108,181). In addition, the concentration of heme within the cell appears to regulate the synthesis of this enzyme (182). When the amount of heme is high, new synthesis of the enzyme is repressed. When the amount of heme is low, new synthesis of the enzyme is induced. Thus, agents that interfere with the synthesis of heme can lead to the induction of ALA synthase, and agents that induce the synthesis of hemoproteins can produce a similar effect (183,184). Agents that exert these effects on the induction of ALA synthase are clinically important, for they can precipitate acute attacks in patients with acute intermittent porphyria and other closely related disorders of porphyrin metabolism (Chapter 30).
Secondary Control Mechanisms
The regulation of heme biosynthesis appears to be sufficiently important in nature that additional control points have been incorporated during the process of evolution. Although the actual in vivo importance of the several secondary control mechanisms has not been fully established, available evidence suggests that they may play some role.
δ-Aminolevulinic Acid Dehydrase
ALA dehydrase from many species is inhibited by heme (185). When ALA synthase is present in excess, ALA dehydrase becomes the regulatory enzyme in the heme biosynthetic pathway. The demonstration of an alternative route for the synthesis of ALA (see above) indicates consistency in nature, because control points come after rather than before “committed” compounds are formed.
Uroporphyrinogen I Synthase
Uroporphyrinogen I synthase, partially purified from human erythrocytes, consumes porphobilinogen and forms uroporphyrinogen I in accordance with classic Michaelis-Menten kinetics. The addition of uroporphyrinogen III cosynthase increases the affinity for the substrate of the dual enzymatic system, but the maximum velocity of the reaction leading to the synthesis of uroporphyrinogen III is decreased (138). The changes in affinity for the substrate of the dual enzyme system suggest that a conformational modification of uroporphyrinogen I synthase is produced on its association with the cosynthase (186). These findings, coupled with studies using crude liver homogenates, suggest that enhanced porphyrin synthesis from porphobilinogen occurs when cosynthase is inactivated (187). Thus, the activity of uroporphyrinogen I synthase increases when the activity of the cosynthase decreases. The mechanism of this regulatory effect of cosynthase on uroporphyrinogen I synthase is unknown.
Heme Synthase
This enzyme also exhibits characteristics suggesting that it may play a regulatory role in heme biosynthesis. Heme synthase is subject to both substrate inhibition by protoporphyrin IX (188) and feedback inhibition by heme (189). Heme synthase can also be induced by several agents (190,191). Finally, studies using Friend leukemia cells (172) and cultured human skin fibroblasts (191) have suggested a rate-limiting step in the formation of heme at the level of heme synthase.
Control of Erythropoiesis
It is evident that a well-balanced mechanism exists that maintains the erythron within “normal” limits and mediates the response to a variety of normal and abnormal situations. In broad outlines, this control system operates in the following manner. Alterations in the concentration of hemoglobin in the blood lead to changes in tissue oxygen tension within the kidney. In response to hypoxia, the kidney secretes a hormone called EPO. This hormone induces erythroid progenitor cells to differentiate into pronormoblasts, thereby bringing about expansion of the erythroid marrow and an increase in red cell production. This, in turn, leads to an increase in the size of the erythron and an increase in tissue oxygen levels. Each of the major steps in this process is discussed in greater detail in the sections that follow.
Tissue Oxygen
Tissue oxygen tension depends on the relative rates of oxygen supply and demand. Oxygen supply is a complex function of interacting but semi-independent variables, including (a) blood flow, (b) blood hemoglobin concentration, (c) hemoglobin oxygen saturation, and (d) hemoglobin oxygen affinity. Each of these functions may be altered to compensate for a deficiency in one of the others. For example, in severe anemia, cardiac output and respiratory rate may increase, and hemoglobin oxygen affinity may be reduced through the 2,3-diphosphoglycerate effect. Conversely, in respiratory insufficiency, secondary polycythemia occurs.
Despite the influence of cardiovascular and respiratory adjustments, tissue oxygen tension decreases roughly in proportion to the degree of anemia. Conversely, induced polycythemia of moderate degree leads to normal or increased tissue oxygen tension and to increased tolerance to hypoxia. These changes occur despite the increase in blood viscosity that accompanies polycythemia, suggesting that peripheral vascular resistance decreases to compensate for increased viscosity. However, with advanced degrees of polycythemia, the increase in viscosity may be great enough to negate the advantages of increased oxygen-carrying capacity.
Tissue hypoxia is the fundamental stimulus to erythropoiesis, as was first suggested by Miescher in 1893. This concept has been amply confirmed (1). However, it now seems clear that hypoxia does not exert its effects by a direct action on the marrow, as Miescher believed, but instead acts by inducing the elaboration of a hormone, EPO. The nature of the tissue oxygen receptors (or oxygen sensor) has only recently been understood. These sensors are located within the kidney because production of EPO can be induced by renal artery constriction or by hypoxic perfusion of the isolated kidney.
Erythropoietin
Structure of Erythropoietin
EPO is a glycoprotein hormone produced by the kidney, and it is the major humoral regulator of red cell production. EPO was originally purified from the urine of patients with aplastic anemia (192). It has an MW of 34,000 daltons as determined electrophoretically and contains 30% carbohydrate, of which 11% consists of sialic acid, 11% total hexose, and 8% N-acetylglucosamine (193). The potency of EPO is expressed in units, with one unit being equal to the amount of EPO present in one tenth of the International Reference Preparation (194). This unit had been originally defined as the amount of EPO that produced in the starved rat the same erythropoietic response (increase in serum EPO level) as 5 μmol of cobalt (1). The potency of the purified human urinary EPO has been determined to be 70,400 U/mg of protein or 50,000 U/mg of total weight (192).
Purification of human urinary EPO has allowed the isolation and cloning of the human genomic DNA encoding for human EPO (195). The gene encoding EPO has been localized on human chromosome 7 (7 pter-q22) (196). The EPO gene exists as a single copy in a 5.4-kb DNA fragment and contains four introns and five exons for the 193–amino acid polypeptide (195). The gene also includes a 27–amino acid signal peptide (leader sequence), which is cleared during EPO secretion, and a 166–amino acid peptide with an MW of 18,398 (195). The C-terminal arginine is absent from both recombinant and natural hormone, presumably because of posttranslational modification by a carboxypeptidase (197). The human EPO contains four cysteine residues linked by disulfide bonds, which, when reduced or alkylated, lead to significant loss of activity (199,200).
Recombinant EPO synthesized by mammalian cells is highly glycosylated, and the carbohydrate structure of the recombinant hormone is similar to but distinct from that of the natural hormone (197). Recombinant EPO has an MW of 30,000 by velocity sedimentation (200) and contains approximately 39% carbohydrate (201); however, recombinant EPO migrates slightly slower on sodium dodecyl sulfate–electrophoresis than urinary EPO, which suggests a slightly higher apparent MW of 40,000 compared to an MW of 34,000 of the native urinary EPO (202). The discrepancy between the electrophoretically obtained apparent MWs of natural and recombinant hormone is almost certainly due to distinctly different glycosylation of recombinant EPO in the Chinese hamster ovary cell line compared to the glycosylation of native EPO in kidney cells. Recombinant EPO must have additional carbohydrate, less negatively charged carbohydrates, or both compared to urinary EPO to result in reduced electrophoretic mobility.
Glycosylation of the hormonal peptide is absolutely necessary for its in vivo activity. The bulk of glycosylation of EPO is at a single site of N-linked carbohydrate. Asialated EPO and nonglyco-sylated recombinant EPO produced in bacteria have no activity in vivo, which can be at least partially attributed to rapid clearance of the hormone by the liver via the galactose receptors of hepatic cells (200,203).
The importance of glycosylation of EPO to EPO’s in vivo activity and half-life led to modification of the EPO gene/protein to make a more effective pharmaceutical. The gene coding for EPO was modified by adding a second site of N-linked glycosylation, such that when the gene is expressed in Chinese hamster ovary cells, the amount of carbohydrate attached to the modified EPO peptide is almost doubled. The new product was called darbepoetin or novel erythroid-stimulating protein. Darbepoetin is highly related to EPO, such that it binds to the same receptor, but it has a longer in vivo half-life than recombinant EPO so that fewer injections per week than recombinant EPO are required for therapeutic use (204).
Studies on the amino acid sequences of human and murine EPO have shown a very high degree of conservation of the molecule structure in these two species (205,206). Which part of the EPO molecule is primarily responsible for its function is not known, although results of studies with monoclonal antibodies have indicated that the amino acids in positions 99 to 129 may play an important role in the binding of EPO to cell membrane (207).
Site and Regulation of Erythropoietin Production
Almost 50 years ago, Jacobson et al. established that the kidney is the major organ of EPO production in adult rats (208). Also, humans with end-stage renal failure were found to have low serum EPO concentrations, which were restored to normal after successful renal transplantation (210). The cloning of the murine EPO gene has allowed studies on the production of EPO-specific mRNA in anemic mice. Induction of anemia leads, within 1 hour, to the appearance of EPO-encoding mRNA in the kidney and liver of anemic mice and rats (210,212). After bleeding, the EPO-mRNA in the kidney increases 500 to 1,000 times compared to normal kidney, whereas the liver produces only 7% of the total EPO-mRNA (211). In no tissue other than the kidneys and the liver is EPO-mRNA detectable, even in the presence of severe anemia (213). These changes in EPO-mRNA were followed by parallel changes in serum EPO concentration determined by radioimmunoassay, indicating that EPO production in response to anemia represents de novo synthesis rather than release of preformed hormone (212). More recent evidence indicates that the anemia-induced increase in EPO-mRNA is to a great extent due to an increased transcription (213). Murine EPO-mRNA was detected by ribonuclease protection assay at 14 days of gestation in the liver and almost a week later in the kidneys, which assume a major role in EPO production after birth (214). In cases of paraneoplastic erythrocytosis, EPO-mRNA has been detected in the neoplastic cells (215,216).
Specialized cells producing EPO have been identified in renal and hepatic parenchyma by the technique of in situ hybridization using radioactive probes specific for EPO-mRNA (Fig. 6.12) (217,218,219). These rare EPO-producing cells are found in the interstitium of the renal parenchyma, outside the tubular basement membrane, mostly in the inner cortex and outer medulla. The nature of these interstitial peritubular EPO-producing cells remains controversial; however, the bulk of experimental evidence indicates that these cells are fibroblastlike type I interstitial cells (220,221). In liver, mRNA is detected in hepatocytes. The number of the interstitial renal EPO-producing cells increases in response to anemia, indicating that increased demands for EPO are met by an increase in the number of EPO-producing cells rather than by an increased synthesis of EPO by a preset number of cells (212). Thus, it appears that EPO is synthesized de novo in response to hypoxia and that there is no detectable storage of the hormone.
Increased levels of circulating EPO do not seem to exert any negative effect on EPO production.
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Figure 6.12. In situ hybridization of murine kidney cells using a probe specific for murine erythropoietin (EPO)-mRNA. EPO-producing cells are found in the interstitium, covered heavily with grains. Arrows indicate peritubular interstitial cells not producing EPO. (Courtesy of Dr. Stephen Koury.) |
The mechanism by which hypoxia leads to EPO synthesis has only recently been understood. Results of several studies have shown that the EPO gene contains sequences that are oxygen sensitive and are involved in the regulation of EPO gene expression (222). By the use of plasmid constructs, transgenic mice, and cell transfection, it has been shown that these oxygen-sensitive sequences, located at the region flanking the 3′ end of the EPO gene, can confer to cells the ability to respond to hypoxia by an increase of the protein encoded by the reporter gene (223). In the same area of the EPO gene, an enhancer has been identified with sequences suggesting that it belongs to the family of the nuclear hormone receptor superfamily (224). The ligand for this oxygen-sensitive enhancer was identified as a protein of 120 kd termed hypoxia-inducible factor 1 (HIF-1) (225,226,227). This DNA-binding protein is tightly regulated by the oxygen tension and is considered to be the physiologic regulator of EPO transcription (228).
HIFs are heterodimeric, helix-loop-helix, transcription factors consisting of two subunits, HIF-1α and HIF-1β. The concentration and transcriptional activity of HIF-1α increase in a geometric fashion on exposure to hypoxia. In addition to EPO, a large number of genes (glucose transporters, glycolytic enzymes, vascular endothelial growth factors, and many others) are activated during hypoxia to aid the cell in adapting to hypoxic conditions (229). HIF-1α is constitutively expressed under normoxic conditions, but it is rapidly degraded via the ubiquitin proteosome complex after it is tagged with the protein of von Hippel-Lindau. Binding of the protein of von Hippel-Lindau to HIF-1α requires hydroxylation of the latter by a proline hydroxylase (230,231,232,233). Prolylhydroxylase is an oxygen and iron-dependent enzyme. Under hypoxic condition, little or no proline hydroxylation takes place; thus, the protein of von Hippel-Lindau does not bind to HIF-1α, which accumulates, heterodimerizes with HIF-1β, and recruits the p300/CREB-binding protein transcriptional coactivator, and the whole complex binds to EPO enhancer to promote EPO gene transcription. Recruitment of p300 is inhibited by asparagine hydroxylation that is catalyzed by an oxygen-sensitive asparaginyl-hydroxylase (234). It seems that these two amino acid hydroxylases, by their dependence on normal intracellular oxygen for their function, act as the oxygen sensor in the EPO-producing interstitial cells in the kidney, and by regulating the function of HIF-1 at least at two distinct points (233), they ultimately control EPO synthesis and production.
Action of Erythropoietin
Erythropoietin Receptors
EPO binds to specific molecules on the cell surface, the EPORs. Expression of both EPO and EPORs is necessary for adult life. Deletion of either the genes that code for EPO or the EPOR in mice results in identical phenotypes of fetal death at day 12 to 13 as a result of severe anemia (235). Arguably, the most important control point of erythropoiesis is the interaction of EPO with the receptor for EPO. The activation of the EPOR generates an intracellular signal in immature erythroid cells that promotes the survival of these cells that would otherwise undergo apoptosis. EPO may also promote proliferation. A direct instructive effect of EPO on either primitive hematopoietic cells or mature erythroid cells to direct differentiation is not supported by most data.
Specific EPORs are expressed on hematopoietic cells that respond to EPO and have been identified on human (23) and murine erythroid cells (236), on erythroleukemic cell lines (236), in fetal liver tissue rich in erythroid elements, in mouse and rat placenta (215,238), and in megakaryocytes (238,239). The density of EPORs on erythroid cells is relatively small (approximately 1,000 molecules per cell) and varies correlating with the cell’s responsiveness to and dependence on EPO (239). EPORs are detectable by autoradiography on human BFUs-E; their density increases as the BFU-E matures to CFU-E (29). Erythroid cells at a stage between CFU-E and proerythroblast seem to have the highest density of EPORs, which decrease as the proerythroblast matures and eventually disappear at the stage of orthochromatic erythroblast (29,238). EPORs are not expressed on reticulocytes or red cells.
Whereas functional EPORs were detected in mouse and rat placenta (215,238), these placental EPORs apparently cannot function to transport significant amounts of EPO from the maternal circulation to the circulation of the fetus. The deletion of the EPO gene in a mouse fetus results in severe anemia and death at days 12 to 13 postinception (235). If sufficient EPO from the mother were transported across the placenta to the fetus, one would expect normal birth of the mice missing the EPO gene (EPO-/-) but death by anemia shortly afterward. Thus, the fetal death of EPO-/- mice proves that fetal EPO production is necessary for survival. The detection of receptors for EPO in megakaryocytes (238,239) is understandable because EPO concentration can affect platelet levels. In contrast, the physiologic significance of the presence of receptors for EPO in neurons, the kidneys, endothelial cells, embryonic muscles, and breast cancer cells is not yet established. It is doubtful that expression of receptors for EPO in nonerythroid tissues is required for normal embryonic development because tissue-specific expression of the EPO receptor under control of an erythroid promoter in EPOR-/- embryonic stem cells results in apparently normal mice. This likely proves that EPOR expression is only required in erythroid cells for normal development (240).
Cloning of the murine erythroleukemia EPOR gene revealed a DNA sequence encoding a 507–amino acid peptide of 62 kd with a single membrane–spanning domain in a direction such that the amino-terminal site is located extracellularly and the carboxy-terminal site is located intracytoplasmically (241). The EPOR undergoes glycosylation and phosphorylation before being incorporated into the cell membrane as a 70- to 78-kd protein (241). Cells transfected with this gene express both high- and low-affinity EPORs (241). The human EPOR gene has been localized in the long arm of chromosome 19 (242). After the cloning of the EPOR gene, other receptor genes were cloned that were similar in sequence, such that it is now recognized that the EPOR is related to a large family of receptors, the hematopoietic receptor superfamily, that includes receptors for interleukins-2 through -9, granulocyte colony-stimulating factor, and granulocyte-macrophage colony-stimulating factor, and receptors for other factors such as growth hormone and prolactin that do not modulate hematopoietic cells (243).
After binding of EPO to its receptor, the hormone is rapidly endocytosed and degraded (239,244). Whether the receptor is also degraded or recycles to the membrane is not known, although experimental evidence favors degradation (236). Crystallographic studies confirm that, as is the case for other members of the hematopoietic superfamily of receptors, one molecule of EPO may simultaneously bind to two receptors for EPO. It is clear that activation of the receptors by the hormone leads to formation of receptor homodimers (245); however, evidence suggests that EPOR dimers likely exist even before the binding of EPO. EPO binding shifts and stabilizes an active receptor conformation that brings the two EPORs in closer contact (246,247).
Tyrosine phosphorylation of the EPOR (248) is the first observable event after EPO binding. Because the EPOR lacks a kinase domain, a tyrosine protein kinase must associate with the receptor. The tyrosine protein kinase JAK2, a member of the Janus family of kinases, is the primary EPOR-associated kinase being able to bind to conserved sequence of amino acids found in cytoplasmic domains of the EPOR (249,250) and other receptors related in a sequence to the receptors for EPO. JAK2 is also implicated as the only significant EPOR-interacting kinase because deletion of the JAK2 kinase gene in mice results in fetal death on day 12 to 13 associated with severe anemia that mirrors the phenotype after either deletion of the EPO gene or the gene coding the receptor for EPO (251). A single amino acid mutation (Val617Phe) located in the regulatory pseudokinase domain of this kinase results in constitutive activation of JAK2. Such a somatic mutation has been detected in the majority of patients with polycythemia vera and in a lower frequency in patients with other myeloproliferative disorders (252,253).
To summarize the current view of this research, two EPOR molecules bind to one EPO molecule. JAK2 kinase molecules associated with each receptor are activated by the physical process of bringing the inactive (or low-activity) kinases in close proximity by a shift in receptor conformation, such that these low-activity kinases cross-phosphorylate each other to gain full activity. The activated JAK2 kinase then phosphorylates all eight tyrosine residues of the EPOR cytoplasmic domain. These phosphorylated tyrosine residues of the receptor become docking sites for signaling molecules that may be phosphorylated by JAK2 to become active, such as the signal-transducing activators of transcription (STAT) proteins. STAT1 and STAT5 become phosphorylated, dimerize, and are transported to the nucleus to mediate gene transcription. Alternatively, the translocation of active signaling molecules from the cytoplasm to the plasma membrane via docking to the phosphorylated EPOR activates a signaling cascade. Examples include the translocation of phosphatidylinositol-3-kinase (PI3-kinase) from the cytoplasm to phosphorylate plasma membrane lipids at the cell surface (254).
Domains of the receptor for EPO, the specific tyrosine residues of the receptor, or both are apparently important in activation of distinctive signaling pathways. For example, tyrosine residue 343 is required for EPO-dependent activation of STAT5 signaling (256), whereas the distal end and tyrosine residue 479 of the EPOR are required for activation of both PI3-kinase and MAP kinase cascade. The importance of EPOR tyrosine phosphorylation is difficult to reconcile with a study that showed that transgenic mice expressing receptors for EPO without any cytoplasmic tyrosine residues are essentially normal regarding erythropoiesis and hematocrit (257). Thus, the tyrosine residues of the receptor for EPO are likely important in more subtle ways than previously believed. This is suggested also by the fact that all eight tyrosines are conserved in both number and alignment between the murine and human EPOR genes. If these residues served no important purpose, one would expect differences in either tyrosine numbers or alignment between the mouse and human genes.
The distal end of the EPOR acts as a negative regulatory domain to which tyrosine protein phosphatases dock to phosphorylated tyrosine residues to attenuate EPO signaling. This is suggested by the findings in a transgenic mouse created to express a truncated human receptor EPO, replacing the murine EPOR. This mouse was found to have a severe erythrocytosis, mimicking the disease state noted in humans with polycythemia vera who have elevated red cell mass due to mutations in the negative regulatory domain of the EPOR gene (258).
The role of the transcription factor, STAT5, in signaling after phosphorylation and activation by binding to the receptor for EPO is controversial. STAT5 is expressed from two very similar genes, STAT5a and STAT5b. In transgenic mice in which both genes for STAT5a and STAT5b were deleted or “knocked out” (STAT5a/b-/- mice), STAT5a/b was found to be neither essential nor significant for erythropoiesis (259). However, other investigators obtained and studied the same STAT5a/b-/- mice and reported a marked fetal anemia in the STAT5-/- mice and lesser but significant anemia in newborn and adult STAT5-/- mice (260). Erythropoiesis in STAT5a/b-/- mice was then re-examined by the initial investigators, who reported that the hematocrit was slightly reduced at fetal and newborn stages, but comparison of photographs of the STAT5-/- fetus did not appear to be significantly different from the wild-type fetus (258). Thus, STAT5 seems to have some role in erythropoiesis, different from the initial conclusion that STAT5 has no effect, but it also plays a less significant role in fetal erythropoiesis than was initially claimed (260). EPO is known to weakly activate STAT1 and to strongly activate STAT5 in primary erythroid cells, such that STAT1 activation may partially compensate for STAT5 and explain why deletion of STAT5 in mice results in a relatively mild inhibition of erythropoiesis (261). A potential target of STAT5 is the Bcl-Xl gene. Bcl-Xl is an antiapoptotic protein believed to be essential for EPO-dependent survival. Two recent studies with Bcl-Xl–deficient mice confirmed that expression of Bcl-Xl is required for normal erythropoiesis, but they showed that Bcl-Xl promotes the survival of mature erythroid cells that no loner depend on EPO for survival (262,263). Thus, Bcl-Xl must be regulated by factors other than EPO-activated STAT5.
A large body of evidence now shows that both the MAP kinase cascade and PI3-kinase cascade are activated after EPO activation of the EPOR. Although EPO may act to (a) promote the survival of immature progenitors by preventing apoptosis or programmed cell death, (b) drive the proliferation of progenitor cells, and (c) direct the differentiation of progenitor cells along erythroid maturation, it is not clear if some intracellular signaling pathways distinctly activate only one of these events. Some data suggest that the EPO-dependent activation of the MAP kinase pathway is more important in directing proliferation, whereas the PI3-kinase pathway is equally important in survival and proliferation (254). The central role of PI3 kinase in signaling downstream of the receptor for EPO is suggested from the surprising ability of a constitutively active Akt (a kinase directly downstream of PI3 kinase) to partially rescue erythroid development (CFU-E) when expressed in JAK2-/- fetal liver cells (255).
Mechanism of Action
EPO is a hormone that promotes erythroid differentiation (1). The role of EPO during the very early stages of erythropoiesis is still undefined. Cell lines with features of multipotent hematopoietic progenitor cells (264,265) and purified human blood BFUs-E (29) express a small number of EPORs, suggesting a possible role of EPO for their survival and differentiation. Although, in vivo, the BFU-E pool is unaffected by acute changes of the level of serum EPO and BFUs-E are EPO independent for their survival, they can respond to EPO by increasing their cycling, which is part of the process of erythroid differentiation (40). Chronic administration of recombinant EPO (rEPO) to humans with end-stage renal disease results in global stimulation of the bone marrow with an increase in the concentration and cycling of all types of hematopoietic progenitors, but this effect is most likely indirect (41).
The erythroid cell that is the most sensitive to EPO is a cell between the CFU-E and the proerythroblast (29,238), and this erythroid cell can be considered the primary target of EPO action. These cells express the highest density of EPORs on their membrane and are absolutely dependent on EPO for their survival (29,266,267). Studies on murine splenic erythroid cells infected with the anemia strain of Friend virus (268) have shown that binding of EPO is followed by a series of biochemical events, including increase in Ca2+ uptake (60), internalization of the hormone (244), increase in total RNA synthesis (269), glucose and iron uptake (239), rate of transcription of the α- and β-globin gene (211,270), expression of transferrin receptors (59), and eventually increase of hemoglobin synthesis as well as of membrane bands 3 and 4.1 (267,271). All of these changes result in an increased rate of erythroid differentiation, ending with an increase in the reticulocyte production and an eventual increase in the red cell mass.
One of the most impressive effects of EPO on the erythroid cells is the ability of the hormone to maintain the viability of these cells irrespective of any effect on cycling and differentiation (266,267). In the absence of EPO, erythroid cells die. It has been shown that EPO retards the cleavage of DNA that occurs normally in CFU-E (272). In the absence of EPO, DNA cleavage is rapid and proceeds to cell death. The pattern of rapid DNA cleavage occurring in erythroid cells deprived of EPO is characteristic of cells undergoing apoptosis (programmed cell death) (272). In the presence of EPO, cell death is avoided and the erythroid cells are allowed to differentiate and form red cells. These findings suggest that the hormone promotes erythroid differentiation simply by allowing cell survival, which is a prerequisite for both cell proliferation and maturation. This model also suggests that, under normal conditions, a large number of generated CFU-E are not surviving, but, under conditions of high EPO levels, expansion of the erythroid marrow is seen simply by allowing survival of more CFU-E, resulting in an increased rate of red cell production. Within the same context, once the red cell mass is restored to normal, the ensuing decrease of EPO levels leads to a rapid turn-off of erythropoiesis by allowing programmed cell death to occur (272).
The observation that relatively immature erythroid progenitor cells continue to develop in fetal mice in which either the gene for EPO or the gene coding for the EPOR has been deleted (235) supports the model that EPO acts primarily by promoting survival of more mature erythroid cells and that EPO has no or a less significant role in proliferation of erythroid precursors or in directing erythroid differentiation of immature hematopoietic cells. The abundant erythroid cells (proerythroblasts near the CFU-E stage of erythroid development) from the spleens of either EPO-/- or EPOR-/- mice undergo apoptosis or programmed death unless these spleen cells are either cultured in vitro in EPO (EPO-/- mice) or forced by transfection with complementary DNA to express the EPOR (EPOR-/-mice) and then cultured in the presence of EPO. Thus, neither EPO nor receptors for EPO are necessary for the proliferation and differentiation of stem cells and early progenitor cells into relatively mature erythroid cells. Both EPO and the receptors for EPO, however, are absolutely required for erythroid cells to survive the transition from CFU-E/proerythroblasts to mature erythro-blasts, suggesting a clear role of EPO in directing the survival of these cells.
In addition to erythroid cells, EPO affects megakaryocytes and their progenitors CFUs-MK. EPO acts as a colony-stimulating factor for murine CFU-MK (273), whereas in humans, it potentates the effect of megakaryocyte colony-stimulating factors present in lymphocyte-conditioned medium (274). It also promotes the differentiation of murine megakaryocytes (275), which express EPORs (276), and, when injected at high doses into mice, it increases platelet production (277). In patients with end-stage renal disease treated with EPO, a minor increase in the platelet count, averaging approximately 30,000/μl, has been noted (278). EPO does not seem to contribute in a significant way to the regulation of platelet counts because anephric patients can maintain a normal platelet count (278). Its various effects on megakaryopoiesis and thrombopoiesis may be explained by the extensive homology between EPO and thrombopoietin, the major humoral regulator of platelet mass (279).
Assays for Erythropoietin and Levels in Health and Disease
The presence of EPO in serum, urine, or other body fluids can be detected by bioassays or immunoassays. Historically, EPO was detected by the polycythemic mouse assay in which the serum sample was injected with 59Fe into polycythemic mice and the amount of 59Fe incorporated into newly released red cells was measured (1,280,281). This method has low sensitivity and is affected by factors influencing erythropoiesis in vivo that are present in the injected sample. In vitro bioassays have been described (280,281), but none has received wide clinical application.
Radioimmunoassays for EPO were developed initially by using purified human urinary EPO as antigen and standard (282,283,284,285,286,287); these have been replaced by purified human recombinant EPO (284,285,286). Using the same principles of immunologic detection of the protein, an enzyme-linked immunosorbent assay was developed, which is commercially available and currently used as the method for determining serum EPO concentrations.
The immunoassays have the advantages of being quick, accurate, relatively inexpensive, and capable of quantifying very low EPO levels ordinarily not detectable by bioassays. Normal serum EPO levels, although variable with the type of assay, usually range between 5 and 30 mU/ml. An inverse correlation has been established between the logarithm of serum EPO concentration and the concentration of hemoglobin in the blood (282,285); however, the magnitude of the increase in the serum EPO concentration in response to anemia is variable among individuals. A disadvantage of the immunoassays is that they detect immunoreactive but not necessarily bioactive hormone. Thus, in renal failure, when serum EPO levels are low or undetectable by bioassays, the immunoassays detect higher levels (284,286,287).
The clinical utility of a single serum EPO measurement has not yet been well established. A low value in the presence of anemia, not associated with renal failure, indicates an abnormal response to anemia, but a normal or elevated level of EPO does not allow any firm conclusion as to whether the increase is appropriate for the severity of anemia. A better approach in interpreting serum EPO levels is a comparison of EPO response to various degrees of anemia between groups of patients with anemia of established etiology. Using such an approach, a blunted EPO response has been described in patients with rheumatoid arthritis (288), solid and hematopoietic malignancies (289,290), sickle cell disease (291), and acquired immunodeficiency syndrome (292) and in premature neonates (293). Conflicting results have also been published regarding the use of serum EPO levels in the differential diagnosis of polycythemia (76,294,295,296). In the presence of an increased red cell mass, an elevated serum EPO value indicates that the polycythemia is secondary to chronic stimulation of the marrow by the hormone, which excludes the diagnosis of polycythemia vera. A normal value in an individual case of polycythemia, however, cannot rule out secondary polycythemia. In this group of patients with polycythemia, there is such an overlap in serum EPO levels that a single EPO value cannot always distinguish with confidence between polycythemia rubra vera and secondary polycythemia (295).
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