Leave a comment

CHAPTER 29 PRODUCTION AND DESTRUCTION OF ERYTHROCYTES

CHAPTER 29 PRODUCTION AND DESTRUCTION OF ERYTHROCYTES
Williams Hematology

CHAPTER 29 PRODUCTION AND DESTRUCTION OF ERYTHROCYTES

ERNEST BEUTLER

The Red Cell Mass

Hematocrit

red cell labels

Plasma Labels
Red Cell Production

Effective Red Cell Production

Ineffective Red Cell Production

Total Erythropoiesis

Ferrokinetics

Transferrin Receptors

Phylogeny of Red Cell Production

Ontogeny of Red Cell Production

Kinetics of Red Cell Production

Regulation of Red Cell Production
Red Cell Destruction

Measurement of Red Cell Destruction

Mechanisms of Destruction

Senescence of Normal Erythrocytes

Fate of Destroyed Red Cells
Chapter References

The volume of red cells in the body (red cell mass) can be measured by labeling a sample of erythrocytes and estimating their dilution in the circulation. The red cell mass of normal women ranges from 23 to 29 ml/kg body weight; that of normal men from 26 to 32 ml/kg. The red cell mass is maintained by the production of erythrocytes in the marrow and, after they age, their destruction by the macrophages in the spleen, liver, and marrow. Red cell production can be estimated by performing ferrokinetic studies using radioactive 59Fe. The formation of red cells in the marrow is regulated largely by the hormone erythropoietin, which binds and cross-links its receptor which appears on developing erythroid cells. In the absence of erythropoietin these cells undergo apoptosis. Normal human red cells have a finite life span averaging 120 days with very little random destruction. The senescent changes in the red cell that mark it for destruction are not fully understood, but the exposure of phosphatidylserine on the membrane appears to be of major importance. The survival of red cells in the circulation can be measured by labeling with isotopes, particularly 51Cr, and assessing the disappearance of radioactivity from the circulation over time, by determining the disappearance of transfused allogeneic erythrocytes using immunologic markers, and by measuring the excretion of CO, a product of heme catabolism.

Acronyms and abbreviations that appear in this chapter include: ADP, adenosine diphosphate; AMP-deaminase, adenosine monophosphate-deaminase; BFU-E, burst forming units–erythroid; CFU-E, colony forming units–erythroid; CPM, counts per minute; DFP, diisopropylfluorophosphate; ELISA, enzyme-linked immunosorbent assay; EPO, erythropoietin; EPOR, EPO receptor; HIF-1, hypoxia-inducible factor-1; ICSH, International Committee on Standardization in Hematology; RCM, red cell mass.

THE RED CELL MASS
The red cell mass is maintained and regulated by the marrow, which under steady-state conditions precisely replaces cells lost by senescence, bleeding, or destruction. The red cell mass defines anemias and polycythemias, and the kinetics of red cell production and destruction helps to establish their pathogenesis. A number of tests have been developed to measure the three main components of red cell kinetics: the red cell mass, the rate of red cell production, and the rate of red cell destruction. Some of these are simple but indirect, such as the hematocrit, reticulocyte count, haptoglobin, lactic dehydrogenase and unconjugated bilirubin concentration. Examination of the marrow allows one to assess total cellularity and the relative erythroid contribution, but it is limited in that one cannot infer the kinetics of cell production from a single static image. These tests are very useful in the aggregate, but can be supplemented by more complex but direct quantitation made possible by the use of radioisotopes.
HEMATOCRIT
The hematocrit is the fractional volume of the blood that the erythrocytes occupy. It can be measured on a sample of blood, expressed either as a percentage or as a fraction, viz., the volume occupied by the erythrocytes in a ml of blood. The total body hematocrit is the volume of red cells in the body divided by the total blood volume. The blood hematocrit is the simplest and most widely used test by which to estimate the size of the red cell mass. In most anemic patients it gives an excellent approximation of the total red cell mass and a functional estimation of the oxygen-carrying capacity and whole blood viscosity. Its main drawback is that it is an indirect measure that is influenced by changes in the plasma volume and may not reflect the size of the red cell mass in dehydrated or polycythemic patients. Dehydration is usually clinically apparent and can in most cases be taken into account when evaluating the significance of a specific hematocrit determination. When the hematocrit is moderately elevated it may also not reflect the total red cell mass, and only the direct measurement of the red cell mass can differentiate between relative and absolute polycythemia. However, when the hematocrit is above 60 percent, virtually all patients have an increase in total red cell mass.1 The extent of the increase cannot be estimated accurately from a hematocrit measurement alone (see Fig. 30-2).
RED CELL LABELS
A more direct and accurate estimate of the size of the red cell mass is obtained from labeling a known volume of red cells and determining the dilution of this label in blood. Radioactive iron is an excellent label of red cells since it is biosynthetically incorporated into hemoglobin in vivo. In experimental animals it can be given to a donor animal and the donor’s cells can be transfused into the animal whose red cell volume is being assessed. However, the radiation exposure to the donor and the hazards of transfusing allogeneic cells preclude its use in humans. Thus, currently almost all methods used clinically employ labeling of autologous red cells in vitro by any one of a number of isotopes. If studies need to be carried out in radiosensitive individuals such as pregnant women, red cell labeling can be carried out by nonradioactive chromium2,3,4 and 5 or by biotin, which is then detected with streptavidin coupled to a fluorochrome.6 Among the isotopes available, 51Cr is the most widely used label, although 99mTc also is both convenient and accurate.7 Chromium in the form of the chromate ion (CrO2–) readily enters the red cell and binds to globin chains. Excess isotope in the incubation mixture can be removed by washing or by using ascorbic acid to reduce the chromate ion to a nonpermeant chromic ion. About 15 min after injection of a known amount of labeled cells a sample of blood is obtained; its volume, hematocrit, and radioactivity are determined; and the total red cell volume is calculated from the equation:

Sampling time is generally 15 min. Since 51Cr may also label white cells, if the white cell count is elevated (more than 25 × 109/liter) it is desirable to centrifuge and remove the buffy coat before labeling.
There is no theoretical objection to measuring the red cell mass by using labeled cells. It is independent of the hematocrit of the blood utilized to measure radioactivity, and replicate determination can be made with a coefficient of variation of approximately 1.5 percent.8 The principal problem lies in reporting the measured red cell mass. The total red cell mass can be expressed as a volume related to body surface (ml/m2) or as a volume related to body weight (ml/kg). A committee of the International Committee on Standardization in Hematology (ICSH)9 has extensively examined existing data and concluded that the most reproducible expressions of the red cell mass (RCM) are related to body surface area estimated from height and weight:

where RCM is the red cell mass, S the body surface area in square meters, and age the age in years.
These calculated values ± 25 percent included 98 percent of the measured male and 99 percent of the measured female values.9
Despite the ICSH recommendation, the most common method is to report red cell mass values in terms of milliliters per kilogram. This method of expression will, however, give erroneously low values in obese individuals because fat is hypovascular. A better method might be to express the red cell mass in terms of lean weight. In general lean weight is 20 percent less than actual weight in normal males and 25 percent less in normal females.7 However, estimation of lean weight in obese individuals is most inaccurate, and from a practical point of view it is probably best to report the red cell mass in terms of actual weight and make mental adjustments based on body configuration. In general, the red cell mass of normal females ranges from 23 to 29 ml/kg body weight. In normal males it is 26 to 32 ml/kg.9
PLASMA LABELS
The red cell mass can also be estimated from the plasma volume. Radioactive iodine (125I) is utilized to label albumin and measure its distribution volume.10 Other radioactive isotopes of iodine as well as of 99mTc have been used, but 125I has virtually supplanted all other plasma labels. Albumin labeled with radioactive iodine is commercially available, and a known amount is injected intravenously. Several blood samples are obtained within the first 15 min, centrifuged and the CPM per ml of plasma measured, plotted on semilogarithmic paper, and extrapolated to zero time. This is necessary, since in contradistinction to labeled red cells, labeled albumin is removed gradually, beginning immediately after injection. The plasma volume is calculated according to the equation:

The continuous exchange of intravascular with extravascular albumin is the major problem encountered when the plasma volume is measured with labeled albumin. Even with extrapolation to zero hour the plasma volume is somewhat larger than that measured with a strictly intravascular protein such as fibrinogen.11 Consequently, if the measurement of the plasma volume is used to calculate the size of the total red cell mass, it is a less reliable measure than determining the red cell mass directly with tagged red cells. This inaccuracy is further aggravated by the fact that the venous hematocrit used to calculate the red cell mass from the measured plasma volume does not reflect accurately the distribution of plasma and red cells in the body (see below). However, from a practical point of view, the results of estimating red cell mass from plasma volume are surprisingly accurate and have been advocated on the basis of simplicity and low cost.10,12
TOTAL-BODY HEMATOCRIT
When the total red cell mass is measured with labeled red cells it has been found that it is about 10 percent lower than that calculated from the plasma volume and the hematocrit of the peripheral blood. In fact, it is apparent that the mean hematocrit of blood in all of the vessels (total-body hematocrit) is somewhat lower than the hematocrit determined from blood obtained from large vessels.
Generally the ratio of total-body hematocrit as estimated by direct measurements of red cell volume and plasma volume to the large-vessel hematocrit ranges from 0.89 to 0.92.13 Consequently, when using the determined plasma volume to calculate red cell mass and total blood volume it is necessary to use a correction factor, and one of 0.90 is generally employed:

Recommended procedures for the determination and evaluation of blood volume are outlined by the International Committee for Standardization in Hematology.14
RED CELL PRODUCTION
Under normal circumstances most human red cells produced in the marrow live, or have the potential to live, a normal life span. Under certain conditions, however, a fraction of red cell production is ineffective, with destruction of nonviable red cells either within the marrow or shortly after the cells reach the blood.15
EFFECTIVE RED CELL PRODUCTION
Effective erythropoiesis is most simply estimated by determining the reticulocyte count. This count is usually expressed as the percentage of red cells that are reticulocytes, but it can also be expressed as the total number of circulating reticulocytes per unit of blood (absolute reticulocytes = reticulocyte percentage × red blood count). In order to use the reticulocyte percentage as a measure of the rate of red cell production the percentage may be corrected for the hematocrit, deriving what is designated the reticulocyte index16:

Since the red cell count is readily available in any electronic blood count printout it seems more logical to use the absolute reticulocyte count as the measure. The usefulness of the absolute reticulocyte count can be enhanced by taking into account the estimated life span of the reticulocytes. The life span of the reticulocytes in blood is about one day, but when red cell production is increased under conditions of erythropoietic stress, as for example with anemia, reticulocytes are released prematurely and circulate as reticulocytes for two to four days. Accordingly, an elevated reticulocyte count may give an erroneous impression of the actual rate of red cell production. In order to take this into account when estimating the rate of red cell production in anemic patients with high reticulocyte counts, it has been suggested that dividing the absolute reticulocyte count by two gives a more accurate estimate of red cell production.16
INEFFECTIVE RED CELL PRODUCTION
Ineffective erythropoiesis is suspected when the reticulocyte count is normal or only slightly increased despite erythroid hyperplasia of the marrow. It was first recognized as an entity from the study of incorporation of isotopes into fecal urobilin following the administration of labeled glycine, a precursor of heme.17 Two peaks were observed: an early one at 3 to 5 days and a late one at 100 to 120 days. It was suggested that one of the sources of the early-labeled peak was the hemoglobin of red cells that had never completed their development, having been destroyed either in the marrow or shortly after reaching the blood. Subsequent studies have revealed that in certain disorders, such as pernicious anemia, thalassemia, and sideroblastic anemia, ineffective erythropoiesis is a major component of total erythropoiesis. This component can be quantitated by measuring 15N-labeled glycine incorporation into the early bilirubin peaks,18,19 bilirubin turnover,20 or ferrokinetics.15 Calculated from bilirubin peaks and turnover, ineffective erythropoiesis under normal conditions amounts to about 4 to 12 percent of total erythropoiesis. Using ferrokinetic methods, ineffective erythropoiesis is calculated as the difference between total plasma iron turnover and erythrocyte iron turnover plus storage iron turnover (see below). The values estimated from such studies in normal subjects are somewhat higher, ranging from 14 to 34 percent.15 However, these results, both the high and the low, probably are misleading, since none of the methods actually measure cell death but only the turnover of heme and iron. It is possible that there is little premature death of cells in normal subjects but that much of the early release of bilirubin and iron is derived from the rim of hemoglobin extruded during enucleation of erythroblasts (see Chap. 22).
TOTAL ERYTHROPOIESIS
Total erythropoiesis, the sum of effective and ineffective red cell production, can be estimated from a marrow examination. Films or sections from marrow aspirates are first examined for relative content of fat and hematopoietic tissue. This gives an estimate of overall hematopoietic activity within the marrow space. Then a differential count is performed with determination of the ratio between granulocytic and erythroid precursors (the M/E ratio). In a normal adult, the ratio is about 3:1 to 5:1, and it can be used to estimate whether erythropoiesis is normal, increased, or decreased (see Chap. 3). It is only an approximation of total erythroid activity, since the ratio can be altered by changing the myeloid as well as the erythroid components and an aspirate or biopsy of a small segment of the marrow may not always reflect total marrow activity. However, when used in conjunction with determination of red blood cell count and reticulocyte count, it will under most circumstances provide qualitative information about the rate and effectiveness of the production of red blood cells. A more accurate quantitation of total erythropoiesis can be made by measuring the rate of production of red cells (ferrokinetics) or, in steady-state conditions, by measuring the rate of destruction of red cells (red cell life span, bilirubin production, or carbon monoxide excretion).
FERROKINETICS
In 1950, Huff and his associates first described a method for the measurement of the rate of production of red cells utilizing a simple model of iron metabolism21 (Fig. 29-1). In this method radioactive iron is complexed to transferrin in vitro and injected intravenously. Alternatively, 59Fe can be injected directly intravenously as the gluconate without preincubation with the patient’s own plasma, if enough unbound transferrin is available, since binding will be almost instantaneous. The rate of clearance of the transferrin-bound iron from the plasma (59Fe plasma T1/2) and the subsequent uptake in the red cells are measured. From these two values and from determinations of the plasma iron concentration and the plasma volume, the rate of formation of red cells can be calculated.15

FIGURE 29-1 The single dynamic pool model of iron metabolism. Radioactive iron injected into the plasma iron pool is cleared from the plasma as a single exponential and approximately 80 percent is incorporated into circulating blood cells.

The initial clearance of iron is exponential, and sampling during this period can be used to calculate the T1/2. In normal individuals this averages about 90 min; it is shorter in patients with hyperplasia of the erythropoietic tissue, and longer in patients with marrow hypoplasia (Fig. 29-2). The clearance rate, however, is not a direct measurement of erythropoietic activity because it is dependent on the size of the pool of unlabeled, circulating iron. Consequently calculation of the plasma iron turnover rate must include the plasma iron concentration. The clearance is expressed in milligrams of iron and the point of reference can be hemoglobin mass, blood volume, or weight, but a commonly used expression is as µg iron per deciliters of whole blood per day.

FIGURE 29-2 Iron clearance and iron utilization in normal subjects, patients with decreased effective red cell production (erythroid hypoplasia), and patients with ineffective red cell production.

Under normal conditions radioactive iron begins to be incorporated into newly formed red cells after a few days and reaches a maximum at about 10 to 14 days after injection (see Fig. 29-2). The normal utilization is 70 to 90 percent on the tenth to the fourteenth day, a value that is so high that a further increase has little significance. Decreased utilization, however, is an important finding and suggests that immature red cells are destroyed in the marrow before they are released to the circulation (ineffective erythropoiesis) or that serum iron is diverted to nonerythropoietic tissues (marrow hypoplasia) because of slow marrow uptake. The shape of the red cell utilization curve is also important, since an early and steep rise (rapid marrow transit time) suggests the presence of a high erythropoietin (EPO) level. Finally, an early rise in utilization with a subsequent fall off suggests hemolysis.
In the calculation of the utilization it is necessary to know the blood volume:

Using the plasma iron clearance and utilization of iron, the red cell turnover in milligram per deciliter blood for 24 h is calculated as follows:

The normal value of red cell iron turnover is 0.30 to 0.70 mg/dl blood for 24 h.15 This range fits very well with crude estimation of the iron used for maintaining the red cell mass in 1 dl of blood or 45 ml of packed red cells. The daily red cell production must equal the daily red cell destruction (45 ml/120 = 0.38 ml), assuming a red cell life span of 120 days, and since 1 ml of packed red cells contains about 1 mg of iron, a daily plasma iron turnover of 0.38 mg is needed by 1 dl of blood to maintain homeostasis.
The calculation of red cell iron turnover has provided useful information about the total volume and effectiveness of erythroid tissue (Table 29-1). However, when the serum iron concentration is elevated, it has given erroneous impressions of the state of erythropoiesis. Moreover, more prolonged sampling of plasma following an intravenous injection of 59Fe has shown that the clearance is not a single exponential but must be represented by several exponential components.22 This has led to the introduction of more complex models of iron kinetics with a single pool of plasma iron exchanging with a number of extravascular erythroid and nonerythroid pools. Careful analysis of such models has generated computer-supported methods calculating the degree and effectiveness of erythroid activity.23 Although possibly more accurate than the conventional method of calculating iron turnover, they appear to be too cumbersome for clinical use. Moreover, even these sophisticated methods may not give an accurate account of the state of erythropoiesis. It was found that despite a constant rate of red cell production, the plasma iron turnover increases with increasing plasma iron and transferrin saturation. This was first thought to be due to increased nonerythroid iron uptake and led to the introduction of various correction factors in the calculation of red cell iron turnover.22 However, the iron in plasma is present in two pools, a diferric and a monoferric transferrin pool, and the erythroid and nonerythroid receptors have a four times greater avidity for diferric transferrin than for monoferric transferrin. Consequently, the total plasma iron turnover depends on the degree of saturation and does not necessarily reflect the number of transferrin receptors, presumably a critical measure of erythropoietic capacity.24 In order to measure the number of these receptors, it has been proposed to adjust the plasma iron turnover equations for both nonerythroid uptake and degree of transferrin saturation and express the plasma turnover in terms of transferrin rather than iron.25 The normal erythroid uptake of transferrin is 60 ± 12 micromoles per liter of blood per day, a value that has been found to be appropriately decreased and increased in patients with hypoplastic and hyperplastic marrow.

TABLE 29-1 PLASMA RADIOACTIVE IRON CLEARANCE AND RED BLOOD CELL UPTAKE

TRANSFERRIN RECEPTORS
Under normal conditions diferric, and to a lesser extent monoferric, transferrin is bound to transferrin receptors and internalized. After having released its iron the transferrin-transferrin receptor complex is returned to the cell surface where the receptor is reanchored. However, some of the extracellular domain of the receptors with or without bound transferrin is released into the circulating blood, and its concentration in plasma is roughly proportional to cell-bound transferrin receptors and in turn to the total amount of erythropoietic tissue.26 Using monoclonal or polyclonal antibodies to the transferrin receptor in an ELISA technique, the normal concentration of circulating receptors is about 6-8 mg/L. It is appropriately increased in patients with hemolytic anemia and decreased in patients with hypoplastic anemia.26,27 The advantage in using measurements of transferrin receptors in determining erythroid activity is primarily the ease with which circulating receptor concentration can be measured. Serial measurements can be carried out without the need for steady-state conditions and without the need for laborious ferrokinetic studies. The disadvantages are that it only indirectly measures or estimates the total number of cell-bound receptors, that many nonerythroid cells contribute to the number of circulating receptors,28 and that a reduction in the intracellular concentration of iron up-regulates the synthesis of transferrin receptors.29 This latter feature is actually used to distinguish iron deficiency anemia with high receptor concentrations from the anemia of chronic disease with increased tissue iron and low receptor concentrations29,30 (see Chap. 38 and Chap. 41). As is true for all erythrokinetic studies, the measurement of transferrin receptors is of value in determining if the erythroid activity of a bone marrow aspirate or biopsy reflects total bone marrow activity or merely the activity of a small “hot pocket.”31 This question, however, can also be answered by a noninvasive magnetic resonance imaging of the whole skeleton.32
PHYLOGENY OF RED CELL PRODUCTION
Hemoglobin has been demonstrated in the most primitive animal forms, such as Paramecium and Tetrahymena, but the development of a cell especially designed to synthesize, carry, and protect respiratory pigments had to await the development of a circulatory system.33 Until then, some crustaceans, such as the Daphnia, were capable of developing a fairly sophisticated oxygen transport system without circulating red cells.34 The specific advantage derived from packaging hemoglobin in red cells is not related to viscosity, since the viscosity of blood is the same whether hemoglobin molecules are dissolved in plasma or concentrated in red cells.35 It appears more likely that the emergence of red cells is related to the protective and regulatory effect of intracellular compounds on hemoglobin and its oxygen affinity.
Circulating nucleated erythrocytes first appear in the worms of the phylum Nemertina and in the sessile marine creatures of the phylum Phoronida. Erythropoiesis in these primitive invertebrates takes place near or on the peritoneal surface, with endothelial cells acting as stem cells.33 In the phylum Annelida, which is considerably further up on the evolutionary scale, nonnucleated red cells are observed for the first time. However, the evolutionary advantage derived from denucleation appears to be slight, and nucleated red cells are observed in much further advanced animals, such as reptiles and birds.36 All mammalian erythrocytes are nonnucleated, even those in the most primitive forms such as the Australian duckbilled platypus.37
In the premammalian species, the spleen is the fundamental erythropoietic organ. In some fish, the kidneys are also involved in red cell production,38,39 but it is questionable if this is related to the development of a renal erythropoietic hormone. In the vertebrates, there is an evolutionary shift from the spleen to the liver and from the liver to the hollow bones. It appears that any organ with a relatively stagnant sinusoidal vascular system may serve as a site for red cell production and that the sinusoidal structure of the bone cavities in mammals renders these areas particularly well suited.40 The homeostatic regulation of blood or hemoglobin production has been studied in Daphnia.34 In these crustaceans, there is a balance between oxygen need and hemoglobin production. In the higher animals, this relationship is maintained by adjusting red cell production. Studies of birds,41 fish,42 and mammals43 indicate that red cell production is controlled by a humoral substance, erythropoietin, which is capable of adjusting red cell production to the demands for oxygen in the tissues. Studies of EPO isolated from a number of mammals indicate some biochemical variability, but still a considerable biologic similarity and genetic homology.44
ONTOGENY OF RED CELL PRODUCTION
A number of tissues in the mammalian species can support red cell production, but the environment inside the bone apparently is optimal for cellular proliferation and maturation. However, bone cavities do not develop until the fifth fetal month, and other, presumably less favorable, sites are responsible for red cell production during early embryonic life. In the human, blood cells are first formed outside the embryo in the numerous blood islands of the yolk sac.45 The cells formed here are very large and remain nucleated throughout their functional life span. During the second gestational month, they are slowly replaced by smaller, but still macrocytic, nonnucleated cells derived from hepatic erythropoiesis.46 The ontogeny of the hemoglobins is discussed in Chap. 7, Chap. 28, and Chap. 46.
During the next fetal months, the liver is the main red cell–producing organ. Splenic red cell production is presumed to be of importance between the third and the seventh fetal months. However, the spleen may sequester nucleated red cells formed elsewhere, and the presence of early erythroid cells in this organ does not conclusively indicate splenic erythropoiesis.47
At about the fifth fetal month, granulopoietic cells can be recognized in the central cartilaginous region of the bones, but the bone cavities only slowly become capable of supporting erythropoiesis. At the time of birth, the hepatic phase of blood cell production is finished, and all bone cavities are actively engaged in erythropoiesis. During the neonatal period, the volume of available marrow space is almost the same as the total volume of hematopoietic cells.48 This precarious balance continues for a few years until the growth of bones and bone cavities outstrips the growth of the hematopoietic mass. During the early years, the lack of reserve space forces reactivation of extramedullary foci in the liver and spleen whenever the hematopoietic system is challenged by blood loss, hypoxia, or hemolysis.49
During adult life, the expansion of marrow space continues, possibly by bone resorption, and there is a gradual increase in the amount of fatty tissue in all bone cavities. Because of the abundant marrow space, compensatory reactivation of extramedullary sites rarely takes place in later life, even during periods of prolonged and intense demand for additional blood cell formation. Extramedullary hematopoiesis during these years usually indicates inappropriate rather than compensatory blood formation.50
During late fetal life, the physiologic control of red cell production is probably tied to tissue hypoxia and release of EPO.51 The fetus is under continuous hypoxic stimulation (“Everest in utero”), and EPO has been demonstrated in the amniotic fluid of mothers with erythroblastotic babies. EPO can apparently not pass the placental barrier in sheep and higher animals, and infants born of mothers with various hematologic disorders varying from severe anemia to secondary polycythemia are usually born with the same degree of normal postpartum erythrocytosis. Since bilateral nephrectomy of fetal sheep fails to alter the rate of EPO production, while hepatectomy does, it appears that during fetal life this production is primarily extrarenal and hepatic.52 At the time of birth, there is a gradual,53 presumably intrinsically determined54 switch to renal production of EPO, and in the adult the kidney is responsible for 90 to 95 percent of total production.
KINETICS OF RED CELL PRODUCTION
STEM CELL AND PROGENITOR CELL POOLS
Mammalian nucleated red cells are characterized biochemically and morphologically by their continuous synthesis and accumulation of hemoglobin molecules. Because of relentless maturation, nucleated red cells must be derived from a stable compartment of cells capable of both differentiation and self-renewal. The existence of such cells, designated as stem cells, is discussed in Chap. 14.
Due to their capacity to grow luxuriously in culture, producing large bursts of hemoglobinized erythroblasts, the early progenitor cells committed to the erythroid cell line are designated as burst forming units–erythroid (BFU-E).55 In common with other early progenitors the BFU-Es express certain surface antigens such as CD-34 and receptors for a number of growth factors and cytokines.56 They also express a few receptors for EPO, but the mechanism of the activation of the EPO receptor (EPOR) gene that determines their commitment to the erythroid lineage is unknown.
The subsequent proliferation and maturation of the BFU-E terminate in the creation of CFU-E. This final erythroid progenitor cell receives its designation from its capacity to form a small erythroid colony consisting of 16 to 64 hemoglobinized erythroblasts.57 It now expresses a great number of EPORs, and it appears that its survival depends on the activation of these receptors by EPO.58
The in vivo composition of the progenitor cell pool is quite different from that observed in an in vitro culture. When cultured in a semi-solid medium in the presence of an optimal concentration of growth factors and EPO, each BFU-E will cause the formation of about 1000 CFU-Es before each of these latter cells differentiate into small erythroid colonies. In vivo, on the other hand, the number of CFU-Es is only three to five times greater than the number of BFU-Es.59 This suggests that stromal and accessory cells in the marrow microenvironment have a profound effect on the fertility and survival of progenitor cells. These effects of the microenvironment modulate the action of EPO and determine the number of CFU-Es available for subsequent differentiation to proerythroblasts. At the level of the CFU-E, the density of EPORs is maximal and their activation becomes crucial for survival and subsequent transformation into proerythroblasts. Cross-linking of the receptor by EPO is responsible for the survival of the cell and permits it to proceed with its preordained program of proliferation and maturation into red blood cells.
PRECURSOR CELLS
The creation of a normal red cell is the end result of an orderly transformation of a proerythroblast with a large nucleus and a volume of about 900 fl to a hemoglobinized anucleated disc with a volume of about 90 fl. Although the cytoplasmic maturation is continuous, the interposed mitotic divisions cause a stepwise reduction in cytoplasmic and nuclear volume, making it quite easy morphologically to recognize proerythroblasts, basophilic erythroblasts, polychromatophilic erythroblasts, orthochromatic erythroblasts, and reticulocytes.
Direct measurements of the number of marrow erythroblasts and reticulocytes have shown that for each proerythroblast there are about 50 erythroblasts and 113 reticulocytes (Table 29-2).60,61 This distribution would conform to the number of cells in a theoretic erythroid pyramid (Table 29-2, Fig. 29-3) in which each proerythroblast undergoes five mitotic divisions over a period of five days before it loses its nucleus and enters a three-day period of reticulocyte maturation. Undoubtedly there are some variations in the size and shape of these erythroid pyramids, but the question is if such varieties are random or play a role in the physiologic control of red cell production. When the production is suppressed, as in anemia of chronic renal disease, the erythroblastic distribution appears normal, with no morphologic or ferrokinetic evidence for the presence of ineffective erythroblasts or premature erythroblastic destruction.15 When production is increased, as in severe anemias, the erythroblastic pyramids also appear normal, with no evidence for additional mitotic divisions. Consequently it seems likely that the rate of red cell production depends on the number of erythroid pyramids formed, not on their shape.

TABLE 29-2 ERYTHROID POOLS

FIGURE 29-3 A theoretical model of the proliferation of the erythroid committed marrow cells, including their most important receptors.

As the erythroblast matures, its synthetic activities increase rapidly and become targeted to the production of all of the proteins characteristic of mature red blood cells, particularly globin. Eventually 95 percent of all protein in the red cell is hemoglobin, almost all hemoglobin A (a2b2), with only small amounts of hemoglobin F (a2g2) and hemoglobin A2 (a2d2). Hemoglobin F is unequally distributed, with certain cells, designated as F cells, containing up to 25 percent of their total hemoglobin as F hemoglobin. There is a sharp decline in the density of EPORs on early erythroblasts, and they are absent on the more mature forms. On the other hand, the number of receptors for transferrin increases sharply, reflecting the increased demands for iron in heme synthesis.
The microenvironment may be of importance for the proliferation and maturation of erythroblasts. However, in situ secreted growth factors and cytokines appear to be of less importance for precursor cells than for progenitor cells. Intercellular adhesion molecules are of course needed to secure the structural integrity of the marrow, and fibronectin is of special importance for erythroblasts.62 The loss of fibronectin receptors heralds the migration of reticulocytes into blood, but some reticulocytes remain sticky even after release and are temporarily sequestered by the spleen. Since erythroid colonies developed in vitro consist almost entirely of nucleated red cells, enucleation may primarily be induced by stromal or endothelial cells.
REGULATION OF RED CELL PRODUCTION
FEEDBACK CONTROLS
Under physiologic conditions, the circulating red cell mass is maintained at an optimal size by appropriate adjustments in the rate of red cell production. Red cell destruction and red cell loss may influence the size of the red cell mass, but these are not physiologic variables, and the spleen of humans,63 unlike that of dogs and race horses,64 does not serve as a reservoir of red cells. The feedback signals that adjust the rate of production of red cells to the need for red cells could be generated from tissues serviced by red cells (functional feedback) or from the red cells themselves (end-product feedback).
FUNCTIONAL FEEDBACK—ERYTHROPOIETIN
History The red cell mass is a large organ designed largely for the purpose of transporting oxygen to the tissues. Thus the size of the red cell mass and the rate of red cell production must be closely related to supply and demand for oxygen in the tissues. Toward the end of the 19th century French mountaineers and physiologists had established that a low tissue tension of oxygen would stimulate the rate of red cell production.65 However, the mode of stimulation was hotly debated. In 1906 the French Sorbonne professor, Dr. Paul Carnot, and his associate, Mademoiselle DeFlandre, suggested that hypoxia generates a humoral factor capable of stimulating red cell production.66 On the other hand the famous biochemist, Friederich Miescher, proposed that marrow hypoxia directly stimulates red cell production.67 Both hypotheses, unfortunately, were based on very questionable experimental data and subsequent attempts to clarify the picture brought more heat than light. Finally, in 1950, Kurt Reissmann in an ingenious study on parabiotic rats provided strong support for the existence of an indirect humoral mechanism.68 A few years later Erslev demonstrated convincingly that the plasma from anemic rabbits and primates contains a red cell–stimulating factor.69,70 It was appropriately named EPO, and it became generally accepted that it was involved in the regulation of red cell production. In 1957 Jacobson and coworkers71 found that it was produced by the kidney, a finding that raised the tempting possibility that if it only could be isolated in adequate amounts it might be of therapeutic benefit to patients with renal anemia.
Structure Purification of EPO provided some partial sequences that led to cloning of the gene and permitted mass production of the recombinant protein.72 The EPO gene contains five exons, four introns, and functionally important 5′ and 3′ untranslated sequences.73,74 There is 80 to 90 percent homology between the human gene and genes for mouse and monkey EPO. The cDNA codes for a chain of 193 amino acids including a 27–amino acid leader peptide. One amino acid apparently is lost during processing, leaving the mature circulating EPO with 165 amino acids. EPO and its recombinant form are heavily glycosylated a-globulins with a molecular mass of 34,000 daltons and a specific activity of about 200,000 IU/mg.73,74 Sixty percent of the molecular weight is contributed by amino acids, while 40 percent is made up of carbohydrate. Although the recombinant form is synthesized by hamster cells, the carbohydrate structure is probably almost the same as in the natural human hormone, and so far no instances of antibody production have been observed in patients receiving recombinant EPO.
The amino acids form a single chain with two internal disulfide bonds that have been shown to be necessary for biologic activity. A biologically active 20–amino acid peptide has been found to be required for the cross-linking (dimerization) of the EPOR, which activates it.75 Dimerization of the receptor with biologic activity can also be accomplished by a totally unrelated synthetic 20–amino acid cyclic peptide. Thus, the biologic effect of EPO can be mimicked by totally unrelated synthetic compounds that dimerize the receptor.76,77 There is extensive homology among EPO from various species but also enough differences so that it is possible to raise antibodies against human EPO in rodents, antibodies vital for the performance of radioimmune assays. EPO has four linkages to carbohydrates, three to asparagine and one to serine.73,74 The importance of the carbohydrate moiety appears to lie in its action on cellular processing and secretion. Furthermore, its terminal sialic acids prevent EPO from being incorporated and catabolized by hepatocytes in vivo. In vitro, however, sialated and desialated EPO have the same biologic activity.
Regulation The classic study by Jacobson and coworkers in 1957 suggested strongly that the kidney was the organ of production.71 Using molecular probes for EPO mRNA it became possible to pinpoint the synthesis to cortical interstitial cells78,79 of endothelial or fibroblastic lineage.80 The cells appear to function in an all-or-none fashion, with the overall production of mRNA dependent on the number of cells activated.81
Hypoxia is the obvious initiating cause of EPO gene activation in these cells and such activation depends on the presence of an enhancer sequence, “the hypoxic-responsive element,” positioned in the 3′ tail of the gene,82 which responds to the hypoxia-inducible factor-1 (HIF-1). HIF-1 is a heterodimer composed of a and b subunits. Its activity is primarily determined by hypoxia-induced stabilization of HIF-1a, which is otherwise rapidly degraded in oxygenated cells.83
Certain 5′ sequences located 6000 to 12,000 bp upstream also affect gene transcription. These sequences are not hypoxia-sensitive but appear necessary for tissue and cellular specificity.84 Hepatic production is contributed primarily by hepatocytes but is much less than the renal production.85 In rodents it may contribute 10 to 15 percent of total EPO circulating in plasma, but probably even less in humans. During fetal life, however, hepatic EPO production is of major importance for red cell production, and anephric fetal sheep and anephric neonatal rats produce normal amounts of EPO and red cells.52 At the time of birth there is a gradual and irreversible switch from hepatic to renal production.53 Interestingly, however, regenerating hepatic tissue such as that found in rats after partial hepatectomy86 or in humans after injury caused by hepatitis apparently synthesizes more EPO than normal adult hepatic tissue.87 Hematopoietic progenitors also produce erythropoietin.88
Metabolism The production of EPO is regulated almost exclusively at the level of transcription, and it is not stored but secreted immediately.89 Circulating recombinant EPO and presumably native EPO as well have a T1/2 of 4 to 12 h with a volume of distribution slightly larger than that of the plasma volume.90 The linking of EPO molecules into dimers and trimers greatly increases their T1/2 and augments their biologic activity by >26-fold.91 A small amount of EPO is excreted in the urine but this only can account for 10 percent of total body EPO turnover.91 EPO may be consumed by erythropoietic tissue,92 but the half-life of EPO is about the same in animals with marrow hypoplasia as in those with marrow hyperplasia.93 It seems likely that removal of the terminal sialic acid, which will expose galactose residues, subjects the molecule to degradation via hepatic galactose receptors. This results in rapid clearance of EPO from the circulation via hepatic galactose receptors. However, no definite proof for this hypothesis exists.
Method of Action The EPOR belongs to the cytokine receptor superfamily. It is composed of a single chain which is dimerized by EPO, an alteration that activates the receptor and initiates a cascade of signaling that includes activation of the JAK2 protein.75,94 An alternatively spliced form of the receptor appears to have a dominant negative effect, i.e., it inhibits the response of the receptor.95 Erythropoietin acts both as a mitogen and as a survival factor preventing apoptosis or premature cell death and permitting cells to proceed with programmed proliferation and maturation.58,96 The fact that some primitive erythroid cells are produced in the EPOR knockout mouse97 suggests that the role of EPO is that of an expansion factor for the erythroid lineage, not that of a factor that determines commitment of precursors.
The BFU-Es contain only a few EPORs and the number of BFU-Es is not influenced materially by the presence or absence of EPO. However, receptor density and EPO dependency increase gradually as the progenitor cells mature, culminating at the level of the CFU-E.98 At that level EPO is necessary for the survival of the CFU-Es and their transformation to proerythroblasts, and the number of transformed cells determines to a great extent the number of red cells produced. The proerythroblasts also contain EPORs, which in the presence of high levels of EPO, may accelerate their entry into the first mitotic division. This may lead to a shortened marrow transit time of erythroblasts15 and result in the release of still unfinished reticulocytes, so-called stress reticulocytes.99 However, after having reached the stage of hemoglobin synthesis it seems unlikely that EPO will change the overall composition of each erythroblastic pyramid since, as stated before, the ratio between early and late erythroblasts is approximately the same in patients with renal disease (low EPO production) or nonrenal anemia (high EPO production).
Although EPO undoubtedly is the main regulator of red cell production, the size of the progenitor cell pool and the number and responsiveness of the EPORs must also play a role. However, it is not clear whether or how these factors are physiologically regulated.
END-PRODUCT FEEDBACK
Products released through the destruction of red cells have been thought to influence or even control the rate of red cell production. Supporting this hypothesis is the impression that anemia due to hemolysis is associated with a more pronounced erythroid hyperplasia and reticulocytosis than blood loss anemia of the same severity. Part of this difference may be related to the more chronic nature of many types of hemolytic anemia with the accompanying expansion of marrow and progenitor cell pools, to a selective destruction of nonreticulated red cells, to a shift of the reticulocyte pool from the marrow to the circulation, and to readily available iron from destroyed red cells. However, in vitro studies have shown a stimulatory effect of hemin on erythropoiesis,100 and such a mechanism might constitute a feedback loop, enhancing erythropoiesis when red cell destruction is increased.
The observations and considerations discussed in this chapter, in addition to information about the adaptability of oxygen affinity (see Chap. 28) have led to the construction of an operational feedback circuit that appears capable of adjusting and maintaining the red cell mass at an optimal size for oxygen transport (see Fig. 40-2). This circuit is based on a feedback between the marrow and the kidney mediated in one direction by oxygen and in the opposite by EPO.
RED CELL DESTRUCTION
MEASUREMENT OF RED CELL DESTRUCTION
RED CELL LIFE SPAN
The original method for the measurement of the red cell life span consisted in the transfusion of cells that were compatible but identifiable immunologically—the Ashby technique.101 During World War II and shortly after, this method was used extensively, but in recent years it has been completely replaced by techniques based on labeling of autologous blood.
In 1946 Shemin and Rittenberg demonstrated that the incorporation of 15N-labeled glycine into heme could be utilized to measure the life span of the red cells.102 Since then a number of other isotopic methods have been developed. These can be divided into three groups: (1) those that label a cohort of cells, (2) those that label cells randomly, and (3) those that use indirect measurements such as the rate of production of red cells or the rate of heme breakdown. The first two classes yield information about the nature of the shortening of the red cell life span, age-dependent or random. The last group yields only mean life span.
COHORT METHODS
Cohort methods depend on the biosynthetic incorporation of the label into the developing red cells. In these methods a group of cells of approximately the same age is labeled. The labels used are glycine-containing labeled nitrogen (15N),102 radioactive carbon (14C),103 or radioactive iron, either 55Fe or 59Fe.104,105 and 106 The main disadvantage of cohort labeling is the need for prolonged periods of sampling, especially if the life span is only moderately reduced (Fig. 29-4). In addition, isotopes from destroyed red cells may be reutilized, making it difficult to interpret results.

FIGURE 29-4 Red cell life span measured by cohort labeling or random labeling. When red cells are labeled randomly with 51Cr there is a daily 1 percent elution which needs to be corrected for in the calculation of total red cell life span.

RANDOM LABEL METHODS
The random label methods are the Ashby differential agglutination technique,101 which uses an immunologic marker, and isotopic techniques employing chromium (50Cr, 51Cr, or 53Cr)4,107; diisopropylfluorophosphate (DFP) labeled with 32P,108 3H,114 or 14C109; 14C cyanate110; or biotin.111,112 By far the most commonly used isotope for red cell life spans is chromium, which penetrates the red cell membrane as the chromate ion and binds to the b and g chains of globin. Unfortunately, these bonds are not covalent and there is a continuous elution of the isotope, varying from 0.5 to 2.9 percent per day.113 DFP, on the other hand, is irreversibly bound to red cell cholinesterase. There is some elution of unbound DFP during the first two to three days of study, but after that DFP disappearance closely matches red cell destruction.114,115 Nevertheless, because sample preparation is somewhat complicated, this label is not commonly used.
To accurately calculate red cell life span using a random label method requires steady-state conditions or that corrections can be made for concurrent blood loss or blood transfusion. Fortunately, it is usually possible to gain an accurate estimate of red cell half-life by sampling three times a week for one to two weeks.
In the normal human the red cell life span is finite and about 120 days, with very little random destruction, i.e., loss irrespective of age (0.06 to 0.4 percent per day). In some mammalian species the amount of random destruction is much greater.116 The survival curve of randomly labeled human red cells should consequently be nearly linear from day 0 to day 120 with a half-life of 60 days. When 51Cr is used as the label, about 1 percent of label elutes per day and the survival curve becomes exponential with a half-life of about 30 days (see Fig. 29-4). For clinical use, the red cell life span is usually expressed as chromium T1/2 and compared to the normal of 30 days.
Since merely expressing the red cell life span measured by chromium as chromium T1/2 will not give information as to the character of destruction, senescence versus random, it has been recommended that in addition a correction factor for chromium elution be used and the data recorded using linear coordinates.117 If the data lie on a straight line the destruction is by senescence and the life span can be calculated as twice the half-life. If the data indicate exponential disappearance and it is necessary to use a semilogarithmic paper in order to depict the data on a straight line, the destruction is random and the life span is 1.44 times half-life. One objection to this method is that the degree of chromium elution is not a constant but varies from day to day and from disease to disease.113 Furthermore, the best fit of data is rarely linear or exponential but somewhere in between. Computer-assisted methods can resolve ambiguities, but the inherent biologic and technical variations in measuring red cell life span are such that it is better to rely on chromium T1/2 with intuitive adjustments based on clinical findings.
INDIRECT METHODS
There are two approaches to the calculation of the red cell life span by indirect methods: from a measurement of the rate of production of red cells utilizing radioactive iron, or from a measurement of the rate of breakdown of heme to bilirubin118 and carbon monoxide.119 Both of these compounds are derived almost exclusively from catabolized hemoglobin, and measurements of their rate of production have provided useful information about the red cell life span. There are probably too many variables that affect the bilirubin level to make it a reliable, quantitative measurement of red cell destruction. The measurement of CO production was formerly very tedious, but with the development of newer technology, it has become more practical. An advantage of the measurement of blood CO as an indication of the rate of red cell destruction is that it gives the rate of destruction at a single point in time.
IN SITU LOCALIZATION OF RED CELL PRODUCTION AND DESTRUCTION
As part of routine erythrokinetic studies both radioactive iron and radioactive chromium have been used for studying the localization of red cell production and red cell destruction. This is accomplished by positioning probes for external counting over the sacrum, liver, spleen, and heart and measuring the distribution of radioactivity in the body.120
In a normal subject 59Fe injected IV is cleared rapidly from the plasma, and within 24 h about 85 percent of the radioactivity can be accounted for in the marrow. The liver and the spleen divide the remaining 15 percent. Over the next 10 days the marrow radioactivity decreases gradually due to the release into circulating blood of red cells labeled with radioactive hemoglobin. Patterns showing different uptake and distribution of the radioactive iron have been found for various hematologic disorders.16 In hypersplenism the trapping and destruction of iron-labeled cells in the spleen will increase splenic radioactivity rapidly, and in patients with erythroid hypoplasia the distribution of radioactive iron between liver and marrow is reversed (Fig. 29-5).

FIGURE 29-5 Tissue distribution of 59Fe in normal subjects, hypersplenic patients, and anemic patients with ineffective and effective erythropoiesis. The radioactivity is expressed on the ordinate as a ratio relative to the radioactivity measured in the same organ 15 min after the intravenous administration of the isotope. (Redrawn from Hillman and Finch.16)

More effective methods demonstrating in situ erythropoiesis involve imaging marrow, liver, and spleen with a 99mTc sulfur colloid or Indium-111.121 Although these isotopes label primarily the monocyte-macrophage system, their uptake is similar to that of 59Fe and they can be used as surrogate markers to estimate the distribution of erythroid tissue.
Probing for 51Cr-labeled red cells will give very characteristic tissue distribution of radioactivity and is especially used to demonstrate the degree of red cell sequestration and destruction in an enlarged spleen122 (Fig. 29-6). This method has been used to predict the results of elective splenectomy, but the utility of this approach has been challenged.123 The in situ localization of red cell sequestration or destruction can also be determined by following the tissue distribution of 59Fe-labeled red cells, especially if the red cell life span is very short.

FIGURE 29-6 Tissue distribution of red cells labeled with 51Cr in normal subjects and in patients with hypersplenism. The radioactivity of blood is given on the ordinate in percent of the radioactivity found 15 min after injection. The radioactivity of the spleen and liver is expressed as a ratio relative to the radioactivity determined simultaneously over the precordium. (Redrawn from Jandl et al.122)

MECHANISMS OF DESTRUCTION
INTRAVASCULAR DESTRUCTION
If the red cell membrane is breached in the circulation, the red cell is destroyed. This mode of demise of the erythrocyte occurs at a low frequency normally but may be the predominant mode of destruction in some hemolytic disorders, e.g., paroxysmal nocturnal hemoglobinuria (Chap. 36), where the complement complex creates holes in the red cell membrane, and in cardiac valve hemolysis (Chap. 50) and microangiopathic hemolytic anemia (Chap. 51), where the shear stress may be so strong as to break open the membrane.
EXTRAVASCULAR DESTRUCTION
Most commonly the life of the red cell comes to an end when it is ingested by a macrophage. Clearly, signals that allow the macrophage to distinguish the younger normal red cell from a damaged or senescent cell must exist. Such signals consist of decreased deformability and/or altered surface properties.
DECREASED DEFORMABILITY
The red cell does not circulate as the biconcave disc that we are accustomed to observing under the microscope. Instead, it is normally greatly distorted by the shear stresses in the circulation, and such distortion is an absolute requirement for the red cell to be able to negotiate the narrow slits that separate the splenic pulp from the sinuses (Chap. 5). The deformability of the erythrocyte can be measured clinically using the ektocytometer, an instrument which displays the diffraction pattern of a red cell suspension under shear stress.124 The red cell membrane, a lipid bilayer, bends readily but has very little capacity to stretch. Thus, deformability is largely a function of the excess red cell membrane intrinsic to the biconcave disc shape of the cell and to some extent of the viscosity of the hemoglobin solution within the cell. As the red cell loses membrane it assumes a spherical shape and loses its ability to deform. Hereditary spherocytosis and hereditary elliptocytosis are prototypic of hemolytic anemias in which decreased deformability as a result of a decreased surface/volume ratio plays a key role in red cell destruction (Chap. 43). However, loss of membrane plays a role in many types of pathologic hemolysis, including autoimmune hemolytic anemia (Chap. 55). In sickle cell disease and hemoglobin C disease (Chap. 47) the internal viscosity of the cell is increased. Loss of water from the red cell, as may occur when the membrane is damaged and leaks potassium, as in hereditary xerocytosis (Chap. 44), also markedly impairs the deformability of the cell.
ALTERED SURFACE PROPERTIES
The surface of the red cell membrane can be altered by binding of antibodies to surface antigens, by binding of complement components, and by chemical alterations, particularly oxidation of membrane components. IgG-coated red cells125 and red cells coated by C3126,127 are bound and sphered by mononuclear cells in vitro, and it is likely that they are similarly damaged by macrophages in the body of patients with autoimmune hemolytic anemia (Chap. 55).
In vitro oxidation of red cells with phenylhydrazine or ADP plus iron causes clustering of band 3 protein in the membrane. Although the physiologic significance of this is far from clear, it has been suggested that the clustered proteins serve as a recognition site for the binding of IgG.128,129 Oxidative damage to the membrane may play a role in the removal of sickle cells (Chap. 47) and thalassemic cells from the circulation (Chap. 46).
SENESCENCE OF NORMAL ERYTHROCYTES
The classical 1949 studies of London et al.130 in which the heme in erythrocytes was labeled with 15N glycine established that most red cells have a finite life span between 100 and 140 days.131
METHODOLOGIC CONSIDERATIONS
Labeling a cohort of human erythrocytes with 59Fe and centrifuging the cells in a density gradient demonstrates that reticulocytes and young red cells are less dense than mature red cells.132,133 However, at the end of the life span of the labeled cohort, radioactivity is fairly evenly distributed throughout red cells of all densities with only a slight tendency of the radioactivity to be concentrated in the more dense cells. Unfortunately, most studies of the properties of senescent cells in the past have been based upon the characteristics of the most dense fraction of erythrocytes, using various fractionating techniques. Although some investigators still regard density separation to be a valid way of isolating aged red cells,134,135 and 136 there is now a large body of evidence that indicates that the most dense fraction of red cells is only slightly enriched with old erythrocytes.137,138 and 139
There are two animal models and one human disease model that provide cells that are truly aged. In mice, in vivo aged cells have been produced by serial transfusion, maintaining polycythemia to suppress virtually all erythropoiesis.140 In other species, particularly the rabbit, red cells have been labeled with traces of biotin, which allows them to be recovered from the circulation.141 The human model is transient erythroblastopenia of childhood (Chap. 32), a disorder in which there is cessation of all erythropoiesis for several months. The use of the latter model has been criticized because of the fact that this disorder is not fully understood and that the red cells in the circulation may not be entirely normal.142 However, the results that have been obtained are consistent with those obtained in animal models and are probably reliable.
PROPERTIES OF AGED CELLS
While the activities of a large number of enzymes, including hexokinase, glucose-6-phosphate dehydrogenase, and pyruvate kinase, are higher in reticulocytes than in mature erythrocytes, they do not continue to decline during the aging of the erythrocyte.141,143 Pyrimidine-5′-nucleotidase144,145 and AMP-deaminase146,147 and 148 appear to be exceptions to this rule in that there is continuing decline of enzyme activity throughout the life span of the red cell. The density and deformability of the aged cells in erythroblastopenia of childhood are normal.137 Fluorescent sorting of NN erythrocytes transfused into humans shows that the most dense fractions are only minimally enriched with old cells,149 and biotinylated aged cells of rabbits have been found to have only a modestly decreased surface area, volume, cell water, and density and therefore slightly decreased deformability.138,150
The amount of immunoglobulin on red cell membranes has been reported to increase with aging of the cells,151,152 and it has been proposed that such accumulation of immunoglobulin mediates removal of senescent erythrocytes. However, immunoglobulin levels on aged, biotinylated rabbit cells are not increased153 and the fact that red cell life span has never been demonstrated to be prolonged in a-gamma-globulinemic patients casts serious doubt upon the concept that immunoglobulins mediate removal of senescent red-cells.
The exposure of phosphatidylserine is one of the signals that allows macrophages to recognize apoptotic cells. It is likely that this is, indeed, the signal by which macrophages recognize senescent erythrocytes.112,154 A model that has been developed suggests that the average time during which phosphatidylserine is exposed is only 0.3 to 0.5 days, so that few cells with increased exposure of the phospholipid are in the circulation at any time. It is not yet clear whether this is the only or even the primary signal that indicates that a cell has reached the end of its life span, but it is the only major difference between senescent and nonsenescent erythrocytes that has been documented clearly.
FATE OF DESTROYED RED CELLS
INTRAVASCULAR DESTRUCTION
Hemoglobin When red cells are destroyed in the vascular compartment the hemoglobin escaping into the plasma is bound to haptoglobin. Each molecule of haptoglobin, a dimeric glycoprotein, can bind two hemoglobin b dimers.155 The haptoglobin-hemoglobin complex is cleared from the plasma with a T1/2 of 10 to 30 min.156 After the complex is carried to the liver parenchyma157 the heme of the hemoglobin is converted to iron and biliverdin by heme oxygenase and the biliverdin is further catabolized to bilirubin. CO is released in the course of cleavage of heme by heme oxygenase.
Free haptoglobin, in contrast to the hemoglobin-haptoglobin complex has a T1/2 of 5 days, and when large amounts of the rapidly turned over haptoglobin-hemoglobin complex are formed the haptoglobin content of the plasma is depleted. The haptoglobin content of the plasma is diminished not only in the plasma of patients undergoing frank intravascular hemolysis but also from the plasma of patients who, like those with sickle cell disease, have accelerated red cell destruction occurring primarily within macrophages. Presumably there is either enough intravascular hemolysis in such hemolytic disorders to lower the plasma haptoglobin level or enough leakage from the phagocytic cells into the plasma to bind to haptoglobin. Thus the measurement of plasma haptoglobin levels has some usefulness in diagnosing the presence of hemolysis.
Heme Free heme that is released into the circulation is bound in a 1:1 ratio to the plasma glycoprotein hemopexin, which is cleared from the plasma with a T1/2 of 7 to 8 h.158 The heme is delivered to the liver where it is converted to bilirubin. When the capacity of hemopexin to bind heme has been saturated, excess heme may bind to albumin to form methemalbumin.159
EXTRAVASCULAR DESTRUCTION
Red cells that are engulfed by phagocytic cells are degraded within lysosomes into lipids, protein, and heme. The proteins and lipids are reprocessed in their respective catabolic pathways, and the heme is cleaved by a microsomal heme oxygenase160 into iron and biliverdin. The latter is catabolized to bilirubin.
BILIRUBIN EXCRETION
Regardless of the site of destruction of hemoglobin, one of the final products is bilirubin, and this is excreted through the bile into the gastrointestinal tract where it is converted to urobilinogens by bacterial reduction.161 A small fraction of urobilinogen is reabsorbed and excreted into the urine. Thus, the fecal and urinary urobilinogen excretion have been used as an indicator of the rate of hemolysis, but is only uncommonly used for this purpose in modern practice because the collections are cumbersome and because alternatively degradative pathways detract severely from the accuracy of the estimates of the rate of heme catabolism.
This chapter is based in part on chapters in previous editions of this text by Drs. Nathaniel Berlin and Allan J. Erslev.
CHAPTER REFERENCES

1.
Pearson TC, Botterill CA, Glass UH, Wetherley-Mein G: Interpretation of measured red cell mass and plasma volume in males with elevated venous PCV values. Scand J Haematol 33:68, 1984.

2.
Drysdale HC, Emerson PM, Holmes A: An improved method for the measurement of red cell survival using non-radioactive chromium. J Clin Pathol 32:655, 1979.

3.
Heaton WAL, Hanbury CM, Keegan TE, Pleban P, Holmes S: Studies with nonradioisotopic sodium chromate: I. Development of a technique for measuring red cell volume. Transfusion 29:696, 1989.

4.
Silver HM, Seebeck MA, Cowett RM, Patterson KY, Veillon C: Red cell volume determination using a stable isotope of chromium. Journal of the Society For Gynecologic Investigation 4:254, 1997.

5.
Sioufi HA, Button LN, Jacobson MS, Kevy SV: Nonradioactive chromium technique for red cell labeling. Vox Sang 58:204, 1990.

6.
Cavill I, Trevett D, Fisher J, Hoy T: The measurement of the total volume of red cells in man: a non-radioactive approach using biotin. Br J Haematol 70:491, 1988.

7.
Jones J, Mollison PL: A simple and efficient method of labelling red cells with 99m Tc for determination of red cell volume. Br J Haematol 38:141, 1978.

8.
Chaplin H Jr: Precision of red cell volume measurement using P32 labeled cells. J Physiol 123:22, 1954.

9.
Pearson TC, Guthrie DL, Simpson J, et al: Interpretation of measured red cell mass and plasma volume in adults: Expert Panel on Radionuclides of the International Council for Standardization in Haematology. Br J Haematol 89:748, 1995.

10.
Fairbanks VF, Klee GG, Wiseman GA, et al: Measurement of blood volume and red cell mass: Re-examination of 51Cr and 125I methods. Blood Cells Mol Dis 22:169, 1996.

11.
Larson RA: Studies of the body hematocrit phenomenon: Dynamic hematocrit of large vessel and initial distribution space of albumin and fibrinogen in the whole body. Scand J Clin Lab Invest 22:189, 1998.

12.
Fairbanks VF: Measurement of blood volume and red cell mass: Re-examination of 51Cr and 125I methods—Commentary. Blood Cells Mol Dis 22:186C, 1996.

13.
Button LN, Gibson II JG, Walter CW: Simultaneous determination of the volume of red cells and plasma for survival studies of stored blood. Transfusion 5:143, 1965.

14.
International Committee for Standardization in Haematology: Recommended methods for measurement of red-cell and plasma volume. J Nucl Med 21:793, 1980.

15.
Finch CA, Deubelbeiss K, Cook JD, et al: Ferrokinetics in man. Medicine (Baltimore) 49:17, 1970.

16.
Hillman RS, Finch CA: Erythropoiesis: normal and abnormal. Semin Hematol 4:327, 1967.

17.
London IM, West R, Shemin D, Rittenberg D: On the origin of bile pigment in normal man. J Biol Chem 184:351, 1950.

18.
Samson D, Halliday D, Nicholson DC, Chanarin I: Quantitation of ineffective erythropoiesis from the incorporation of [15N] delta-aminolaevulinic acid and [15N] glycine into early labelled bilirubin: II. Anaemic patients. Br J Haematol 34:45, 1976.

19.
Samson D, Halliday D, Nicholson DC, Chanarin I: Quantitation of ineffective erythropoiesis from the incorporation of [15N] delta-aminolaevulinic acid and [15N] glycine into early labelled bilirubin: I. Normal subjects. Br J Haematol 34:33, 1976.

20.
Berk PD, Blaschke TF, Scharschmidt BF, Waggoner JG, Berlin NI: A new approach to quantitation of the various sources of bilrubin in man. J Lab Clin Med 87:767, 1976.

21.
Huff RI, Hennessey TG, Austin RE: Plasma and red cell iron turnover in normal subjects and in patients having various hematopoietic disorders. J Clin Invest 29:1041, 1950.

22.
Cook JD, Marsaglia G, Eschbach JW, Funk DD, Finch CA: Ferrokinetics: a biologic model for plasma iron exchange in man. J Clin Invest 49:197, 1970.

23.
Ricketts C, Cavill I, Napier JA, Jacobs A: Ferrokinetics and erythropoiesis in man: an evaluation of ferrokinetic measurements. Br J Haematol 35:41, 1977.

24.
Bauer W, Stray S, Huebers H, Finch C: The relationship between plasma iron and plasma iron turnover in the rat. Blood 57:239, 1981.

25.
Beguin Y: The soluble transferrin receptor: biological aspects and clinical usefulness as quantitative measure of erythropoiesis. Haematologica 77:1, 1992.

26.
Huebers HA, Beguin Y, Pootrakul P, Einspahr D, Finch CA: Intact transferrin receptors in human plasma and their relation to erythropoiesis. Blood 75:102, 1990.

27.
Flowers CH, Skikne BS, Covell AM, Cook JD: The clinical measurement of serum transferrin receptor. J Lab Clin Med 114:368, 1989.

28.
Morishita Y, Kataoka T, Towatari M, et al: Up-regulation of transferrin receptor gene expression by granulocyte colony-stimulating factor in human myeloid leukemia cells. Cancer Res 50:7955, 1990.

29.
Kuiper-Kramer PA, Huisman CMS, Van der Molen-Sinke J, Abbes A, Van Eijk HG: The expression of transferrin receptors on erythroblasts in anaemia of chronic disease, myelodysplastic syndromes and iron deficiency. Acta Haematol (Basel) 97:127, 1997.

30.
Ferguson BJ, Skikne BS, Simpson KM, Baynes RD, Cook JD: Serum transferrin receptor distinguishes the anemia of chronic disease from iron deficiency anemia. J Lab Clin Med 119:385, 1992.

31.
Kansu E, Erslev AJ: Aplastic anaemia with ‘hot pockets.’ Scand J Haematol 17:326, 1976.

32.
Steiner RM, Mitchell DG, Rao VM, Schweitzer ME: Magnetic resonance imaging of diffuse bone marrow disease. Radiol Clin North Am 31:383, 1993.

33.
Scott RB: Comparative hematology: the phylogeny of the erythrocyte. Blut 12:340, 1966.

34.
Fox HM: The hemoglobin of Daphnia. Proc R Soc Lond (Biol) 135:195, 1948.

35.
Schmidt-Nielsen K, Taylor CR: Red blood cells: why or why not? Science 162:274, 1968.

36.
Andrew W: Comparative Hematology. Grune & Stratton, New York, 1965.

37.
Bolliger A: Observations on the blood of a monotreme Tachyglossus aculeatus. Aust J Sci 22:257, 1959.

38.
Jordan HE: Comparative hematology, in Handbook of Hematology, p 703. Hoeber-Harper, New York, 1938.

39.
Iorio RJ: Some morphologic and kinetic studies of the developing erythroid cells of the common gold fish Carassius auratus. Cell Tissue Kinet 2:319, 1969.

40.
Robb-Smith AHT: The Growth of Knowledge of the Functions of the Blood, edited by RG Macfarlane, AHT Robb-Smith. Academic Press, New York, 1961.

41.
Rosse WF, Waldmann TA: Factors controlling erythropoiesis in birds. Blood 27:654, 1966.

42.
Zanjani ED: Humoral factors influencing erythropoiesis in the fish (Blue Gourami-Trichogaster trichopteras). Blood 33:573, 1969.

43.
Erslev AJ: Control of red cell production. Ann Rev Med 11:315, 1959.

44.
Shoemaker C, Mitsock LD: Murine erythropoietin gene: Cloning expression and human gene homology. Mol Cell Biol 6:849, 1986.

45.
Le Douarin NM: Cell migrations in embryos. Cell 38:353, 1984.

46.
Hoyes AD, Riches DJ, Martin BGH: The fine structure of haematopoiesis in the human fetal liver. J Anat 115:99, 1973.

47.
Rosenberg M: Fetal hematopoiesis. Blood 33:66, 1969.

48.
Hudson G: Bone marrow volume in the human foetus and newborn. Br J Haematol 11:446, 1965.

49.
Brannon D: Extramedullary hematopoiesis in anemia. Bull Johns Hopkins Hosp 41:104, 1927.

50.
Erslev A: Medullary and extramedullary blood formation. Clin Orthop 52:25, 1967.

51.
Finne PH: Erythropoietin production in fetal hypoxia and in anemic uremic patients. Ann NY Acad Sci 149:497, 1968.

52.
Zanjani ED, Poster J, Burlington H, et al: Liver as the primary site of erythropoietin formation in the fetus. J Lab Clin Med 89:640, 1977.

53.
Zanjani ED, Ascensao JL, McGlare PG, et al: Studies on the liver to kidney switch of erythropoietin production. J Clin Invest 67:1183, 1981.

54.
Flake AW, Harrison MR, Adzick NS, Zanjani ED: Erythropoietin production by the fetal liver in an adult environment. Blood 70:542, 1987.

55.
Stephenson IK, Axelrod AA, McLeod DL, Shreeve MM: Induction of hemoglobin-synthesizing cells by erythropoietin in vitro. Proc Natl Acad Sci USA 65:1542, 1971.

56.
Civin CI, Loken MR: Cell surface antigens on human marrow cells: dissection of hematopoietic development using monoclonal antibodies and multiparameter flow cytometry. Int J Cell Cloning 5:267, 1987.

57.
Testa NG: Structure and regulation of the erythroid system at the level of progenitor cells. Crit Rev Oncol Hematol 9:17, 1989.

58.
Fisher JW: Erythropoietin: physiologic and pharmacologic aspects. Proc Soc Exp Biol Med 216:358, 1997.

59.
Adamson JW, Torok-Storb B, Lin N: Analysis of erythropoiesis by erythroid colony formation in culture. Blood Cells 4:89, 1978.

60.
Donohue DM, Reiff RH, Hanson ML, Betson Y, Finch CA: Quantitation measurement of the erythrocytic and granulocytic cells of marrow and blood. J Clin Invest 37:1571, 1958.

61.
Finch CA, Harker LA, Cook JD: Kinetics of the formed elements of human blood. Blood 50:699, 1977.

62.
Goltry KL, Patel VP: Specific domains of fibronectin mediate adhesion and migration of early murine erythroid progenitors. Blood 90:138, 1997.

63.
Prankerd TAJ: The spleen and anemia. BMJ 2:517, 1963.

64.
Baker CH, Remington JW: Role of the spleen in determining total body hematocrit. Am J Physiol 198:906, 1960.

65.
Erslev AJ: Blood and Mountains, in Blood, Pure and Eloquent, edited by MM Wintrobe, p 257. McGraw-Hill, New York, 1980.

66.
Carnot P, Deflandre C: Sur l’activité hématopoiétique des serum au cours de la régénération du sang. Acad Sci Med 3:384, 1906.

67.
Miescher F: Über die Beziehungen Zwischen Meereshohe und Beschaffenheit des Blutes. Koresp Bltt Schweitz Aerzte 24:809, 1893.

68.
Reissmann KR: Studies on the mechanism of erythropoietic stimulation in parabiotic rats during hypoxia. Blood 5:372, 1950.

69.
Erslev AJ: Humoral regulation of red cell production. Blood 8:349, 1953.

70.
Erslev AJ, Lavietes PH, van Wagenen G: Erythropoietic stimulation induced by “anemia” serum. Proc Soc Exp Biol Med 83:548, 1953.

71.
Jacobson LO, Goldwasser E, Fried W, Plzak L: Role of the kidney in erythropoiesis. Nature 179:633, 1957.

72.
Lappin TR, Rich IN: Erythropoietin—the first 90 years. Clin Lab Haematol 18:137, 1996.

73.
Jelkmann W: Erythropoietin: structure, control of production, and function. Physiol Rev 72:449, 1992.

74.
Jelkmann W, Metzen E: Erythropoietin in the control of red cell production. Anat Anz 178:391, 1996.

75.
Yoshimura A, Misawa H: Physiology and function of the erythropoietin receptor. Curr Opin Hematol 5:171, 1998.

76.
Livnah O, Stura EA, Johnson DL, et al: Functional mimicry of a protein hormone by a peptide agonist: the EPO receptor complex at 2.8 A. Science 273:464, 1996.

77.
Wrighton NC, Farrell FX, Chang R, et al: Small peptides as potent mimetics of the protein hormone erythropoietin. Science 273:458, 1996.

78.
Koury ST, Boudurant MC, Koury MJ: Localization of erythropoietic synthesizing cells in murine kidneys by in situ hybridization. Blood 71:524, 1988.

79.
Lacombe C, Da Silva J-L, Bruneval P, Fournier J-G: Peritubular cells are the site of erythropoietin synthesis in the murine hypoxic kidney. J Clin Invest 81:620, 1988.

80.
Koury ST, Koury MJ, Bondurant MC, Caro J, Graber SE: Quantitation of erythropoietic-producing cells in kidneys of mice by in situ hybridization: Correlation with hematocrit, renal erythropoietin mRNA, and serum erythropoietin concentration. Blood 71:645, 1989.

81.
Bachmann S, Le Hir M, Eckardt K-U: Co-localization of erythropoietin mRNA and Eco-5′-nucleotidase immunoreactivity in peritubular cells of rat renal cortex indicates that fibroblasts produce erythropoietin. J Histochem Cytochem 41:335, 1993.

82.
Wenger RH, Kvietikova I, Rolfs A, Camenisch G, Gassmann M: Oxygen-regulated erythropoietin gene expression is dependent on a CpG methylation-free hypoxia-inducible factor-1 DNA-binding site. Eur J Biochem 253:771, 1998.

83.
Huang LE, Gu J, Schau M, Bunn HF: Regulation of hypoxia-inducible factor 1alpha is mediated by an O2-dependent degradation domain via the ubiquitin-proteasome pathway. Proc Natl Acad Sci USA 95:7987, 1998.

84.
Semenza GL, Dureza RC, Traystman MD, Gearhart JD, Antonarakis SE: Human erythropoietin gene expression in transgenic mice: multiple transcription initiation sites and cis-acting regulatory elements. Mol Cell Biol 10:930, 1990.

85.
Schuster SJ, Koury ST, Bohrer M, Salceda S, Caro J: Cellular sites of extrarenal and renal erythropoietin production in anaemic rats. Br J Haematol 81:153, 1992.

86.
Naughton BA, Kaplan SM, Roy M, et al: Hepatic regeneration and erythropoietin production in the rat. Science 196:301, 1977.

87.
Brown S, Caro J, Erslev AJ, Murray T: Spontaneous increase in erythropoietin and hematocrit value associated with transient liver enzyme abnormalities in an anephric patient undergoing hemodialysis. Am J Med 68:280, 1980.

88.
Stopka T, Zivny JH, Stopkova P, Prchal JF, Prchal JT: Human hematopoietic progenitors express erythropoietin. Blood 91:3766, 1998.

89.
Fandry J, Bunn HF: In vivo and in vitro regulation of erythropoietin mRNA: measurement by competitive polymerase reaction. Blood 81:617, 1993.

90.
Flaharty KK, Caro J, Erslev A, et al: Pharmacokinetics and erythropoietic response to human recombinant erythropoietin in healthy men. Clin Pharmacol Ther 47:557, 1990.

91.
Sytkowski AJ, Lunn ED, Davis KL, Feldman L, Siekman S: Human erythropoietin dimers with markedly enhanced in vivo activity. Proc Natl Acad Sci USA 95:1184, 1998.

92.
Sawyer ST, Krantz SB, Goldwasser E: Binding and receptor-mediated endocytosis of erythropoietin in Friend virus-infected erythroid cells. J Biol Chem 262:5554, 1987.

93.
Piroso E, Erslev AJ, Flaharty KK, Caro J: Erythropoietin life span in rats with hypoplastic and hyperplastic bone marrows. Am J Hematol 36:105, 1991.

94.
Joneja B, Wojchowski DM: Mitogenic signaling and inhibition of apoptosis via the erythropoietin receptor Box-1 domain. J Biol Chem 272:11176, 1997.

95.
Nakamura Y, Nakauchi H: A truncated erythropoietin receptor and cell death: a reanalysis. Science 264:588, 1994.

96.
Spivak JL, Pham T, Isaacs M, Hankins WD: Erythropoietin is both a mitogen and a survival factor. Blood 77:1228, 1991.

97.
Lin CS, Lim SK, D’Agati V, Costantini F: Differential effects of an erythropoietin receptor gene disruption on primitive and definitive erythropoiesis. Genes Dev 10:154, 1996.

98.
Sawyer ST, Penta K: Erythropoietin cell biology. Hematol Oncol Clin North Am 8:895, 1994.

99.
Noble NA, Xu Q-P, Hoge LL: Reticulocytes: II. Reexamination of the in vivo survival of stress reticulocytes. Blood 75:1877, 1990.

100.
Mayeux P, Felix JM, Billat C, Jacquot R: Induction by hemin of proliferation and of differentiation of progenitor erythroid cells responsible for erythropoietin. Exp Hematol 14:801, 1986.

101.
Ashby W: The determination of the length of life of transfused blood corpuscles in man. J Exp Med 29:267, 1919.

102.
Shemin D, Rittenberg D: Life span of human red blood cell. J Biol Chem 166:627, 1946.

103.
Berlin NI, Meyer LM, Lazarus M: Life span of the rat red blood cell as determined by glycine-2-C14. Am J Physiol 165:565, 1951.

104.
Beutler E, Dern RJ, Alving AS: The hemolytic effect of primaquine: IV. The relationship of cell age to hemolysis. J Lab Clin Med 44:439, 1954.

105.
Birgens HS, Hansen OP, Henriksen JH, Wantzin P: Quantitation of erythropoiesis in myelomatosis. Scand J Haematol 22:357, 1979.

106.
Weinstein IM, Beutler E: The use of Cr-51 and Fe-59 in a combined procedure to study erythrocyte production and destruction in normal human subjects and in patients with hemolytic or aplastic anemia. J Lab Clin Med 45:616, 1955.

107.
Heaton WA: Evaluation of posttransfusion recovery and survival of transfused red cells. Transfusion Medicine Reviews 6:153, 1992.

108.
Cohen JA, Warringa MGPJ: The fate of P32 labeled diisopropyl fluorophosphonate in the human body and its use as a labeling agent in study of turnover of blood plasma and red cells. J Clin Invest 33:459, 1954.

109.
Milner PF, Charache S: Life span of carbamylated red cells in sickle cell anemia. J Clin Invest 52:3161, 1973.

110.
Eschbach JW, Korn D, Finch CA: 14C cyanate as a tag for red cell survival in normal and uremic man. J Lab Clin Med 89:823, 1977.

111.
Wardrop KJ, Tucker RL, Anderson EP: Use of an in vitro biotinylation technique for determination of posttransfusion viability of stored canine packed red blood cells. Amer J Vet Res 59:397, 1998.

112.
Boas E, Forman L, Beutler E: Phosphatidyl serine exposure and red cell viability in red cell ageing and in hemolytic anemia. Proc Natl Acad Sci USA 85:3077, 1998.

113.
Bentley SA, Glass HI, Lewis SM, Szur L: Elution correction in 51Cr red cell survival studies. Br J Haematol 26:179, 1974.

114.
Cline MJ, Berlin NI: Simultaneous measurement of the survival of two populations of erythrocytes with the use of labelled diisopropyl fluorophosphate. J Lab Clin Med 61:249, 1963.

115.
McCurdy PR, Sherman AS: Irreversibly sickled cells and red cell survival in sickle cell anemia: a study with both DF32P and 51CR. Am J Med 64:253, 1978.

116.
Eadie GS, Brown IW Jr: Red blood cell survival studies. Blood 8:1110, 1953.

117.
International Committee for Standardization in Haematology: Recommended method for radioisotope red-cell survival studies. Br J Haematol 45:659, 1980.

118.
Berlin NI, Berk PD: Quantitative aspects of bilirubin metabolism for hematologists. Blood 57:983, 1981.

119.
Doyle J, Vreman HJ, Stevenson DK, et al: Does vitamin C cause hemolysis in premature newborn infants? Results of a multicenter double-blind, randomized, controlled trial. J Pediatr 130:103, 1997.

120.
ICSH panel on diagnostic applications of radioisotopes in hematology: Recommended methods for surface counting to determine sites of red cell destruction. Br J Haematol 30:249, 1975.

121.
Datz FL, Taylor AJ: The clinical use of radionuclide bone marrow imaging. Semin Nucl Med 15:239, 1985.

122.
Jandl JH, Greenberg MS, Yonemoto RH, Castle WB: Clinical determination of the sites of red cell sequestration in hemolytic anemias. J Clin Invest 35:842, 1956.

123.
Ferrant A, Cauwe F, Michaux JL, et al: Assessment of the sites of red cell destruction using quantitative measurements of splenic and hepatic red cell destruction. Br J Haematol 50:591, 1982.

124.
Johnson RM, Ravindranath Y: Osmotic scan ektacytometry in clinical diagnosis. J Pediatr Hematol Oncol 18:122, 1996.

125.
Lo Buglio AA, Cotran RS, Jandl JH: Red cells coated with immunoglobulin g: Binding and sphering by mononuclear cells in man. Science 158:1582, 1967.

126.
Jandl JH, Tomlinson AS: The destruction of red cells by antibodies in man: II. Pyrogenic, leukocytic and dermal responses to immune hemolysis. J Clin Invest 37:1202, 1958.

127.
Lutz HU, Stammler P, Kock D, Taylor RP: Opsonic potential of C3b-anti-band 3 complexes when generated on senescent and oxidatively stressed red cells or in fluid phase, in Red Blood Cell Aging, p 367. Plenum Press, New York, 1991.

128.
Low PS, Waugh SM, Zinke K, Drenckhahn D: The role of hemoglobin denaturation and band 3 clustering in red blood cell aging. Science 227:531, 1985.

129.
Beppu M, Mizukami A, Nagoya M, Kikugawa K: Binding of anti-band 3 autoantibody to oxidatively damaged erythrocytes. Formation of senescent antigen on erythrocyte surface by an oxidative mechanism. J Biol Chem 265:3226, 1990.

130.
London IM, Shemin D, West R, Rittenberg D: Heme synthesis and red blood cell dynamics in normal humans and in subjects with polycythemia vera, sickle-cell anemias, and pernicious anemia. J Biol Chem 179:463, 1949.

131.
Bratosin D, Mazurier J, Tissier JP, et al: Cellular and molecular mechanisms of senescent erythrocyte phagocytosis by macrophages. A review. Biochimie 80:173, 1998.

132.
Borun ER, Figueroa WG, Perry SM: The distribution of Fe59 tagged human erythrocytes in centrifuged specimens as a function of cell age. J Clin Invest 36:676, 1957.

133.
Luthra MG, Friedman JM, Sears DA: Studies of density fractions of normal human erythrocytes labeled with iron-59 in vivo. J Lab Clin Med 94:879, 1979.

134.
Piomelli S, Seaman C: Mechanism of red blood cell aging: Relationship of cell density and cell age. Am J Hematol 42:46, 1993.

135.
Lutz HU, Stammler P, Fasler S, Ingold M, Fehr J: Density separation of human red blood cells on self-forming Percoll gradients: Correlation with cell age. Biochim Biophys Acta 1116:1, 1992.

136.
Piccinini G, Minetti G, Balduini C, Brovelli A: Oxidation state of glutathione and membrane proteins in human red cells of different age. Mech Ageing Dev 78:15, 1995.

137.
Linderkamp O, Friederichs E, Boehler T, Ludwig A: Age dependency of red blood cell deformability and density: Studies in transient erythroblastopenia of childhood. Br J Haematol 83:125, 1993.

138.
Dale GL, Norenberg SL: Density fractionation of erythrocytes by Percoll/hypaque results in only a slight enrichment for aged cells. Biochim Biophys Acta 1036:183, 1990.

139.
Beutler E: Isolation of the aged. Blood Cells 14:1, 1988.

140.
Ganzoni AM, Oakes R, Hillman RS: Red cell aging in vivo. J Clin Invest 50:1373, 1971.

141.
Suzuki T, Dale GL: Senescent erythrocytes: isolation of in vivo aged cells and their biochemical characteristics. Proc Natl Acad Sci USA 85:1647, 1988.

142.
Haram S, Carriero D, Seaman C, Piomelli S: The mechanism of decline of age-dependent enzymes in the red blood cell. Enzyme 45:47, 1991.

143.
Zimran A, Forman L, Suzuki T, Dale GL, Beutler E: In vivo aging of red cell enzymes: Study of biotinylated red blood cells in rabbits. Am J Hematol 33:249, 1990.

144.
Beutler E, Hartman G: Age-related red cell enzymes in children with transient erythroblastopenia of childhood and hemolytic anemia. Pediatr Res 19:44, 1985.

145.
Beutler E: The relationship of red cell enzymes to red cell life-span. Blood Cells 14:69, 1988.

146.
Dale GL, Norenberg SL: Time-dependent loss of adenosine 5′-monophosphate deaminase activity may explain elevated adenosine 5′-triphosphate levels in senescent erythrocytes. Blood 74:2157, 1989.

147.
Paglia DE, Valentine WN, Nakatani M, Brockway RA: AMP deaminase as a cell-age marker in transient erythroblastopenia of childhood and its role in the adenylate economy of erythrocytes. Blood 74:2161, 1989.

148.
Dale GL, Norenberg SL, Suzuki T, Forman L: Altered adenine nucleotide metabolism in senescent erythrocytes from the rabbit. Prog Clin Biol Res 319:259, 1989.

149.
Clark MR, Corash L, Jensen RH: Density distribution of aging, transfused human red cells. Blood 74(Suppl 1):217a, 1989.

150.
Waugh RE, Narla M, Jackson CW, et al: Rheologic properties of senescent erythrocytes: Loss of surface area and volume with red blood cell age. Blood 79:1351, 1992.

151.
Kay MM, Marchalonis JJ, Schluter SF, Bosman G: Human erythrocyte aging: cellular and molecular biology. Transfus Med Rev 5:173, 1991.

152.
Sheiban E, Gershon H: Recognition and sequestration of young and old erythrocytes from young and elderly human donors: in vitro studies. J Lab Clin Med 121:493, 1993.

153.
Dale GL: Does surface bound immunoglobulin mediate erythrocyte death? Commentary. Blood Cells 14:36, 1988.

154.
Connor J, Pak CC, Schroit AJ: Exposure of phosphatidylserine in the outer leaflet of human red blood cells. Relationship to cell density, cell age, and clearance by mononuclear cells. J Biol Chem 269:2399, 1994.

155.
Nagel RL, Gibson QH: The binding of hemoglobin to haptoglobin and its relation to subunit dissociation of hemoglobin. J Biol Chem 246:69, 1971.

156.
Garby L, Noyes WD: Studies on hemoglobin metabolism: I. The kinetic properties of the plasma hemoglobin pool in normal man. J Clin Invest 38:1479, 1959.

157.
Hershko C: The fate of circulating hemoglobins. Br J Haematol 29:199, 1975.

158.
Sears DA: Disposal of plasma heme in normal man and patients with intravascular hemolysis. J Clin Invest 49:5, 1970.

159.
Rosen H, Sears DA: Spectral properties of hemopexinheme: The Schumm test. J Lab Clin Med 74:941, 1969.

160.
Maines MD: The heme oxygenase system: a regulator of second messenger gases. Annu Rev Pharmacol Toxicol 37:517, 1997.

161.
Elder G, Gray CH, Nicholson DG: Bile pigment fate in gastrointestinal tract. Semin Hematol 9:71, 1972.
Books@Ovid
Copyright © 2001 McGraw-Hill
Ernest Beutler, Marshall A. Lichtman, Barry S. Coller, Thomas J. Kipps, and Uri Seligsohn
Williams Hematology

Advertisements

Leave a Reply

Fill in your details below or click an icon to log in:

WordPress.com Logo

You are commenting using your WordPress.com account. Log Out / Change )

Twitter picture

You are commenting using your Twitter account. Log Out / Change )

Facebook photo

You are commenting using your Facebook account. Log Out / Change )

Google+ photo

You are commenting using your Google+ account. Log Out / Change )

Connecting to %s

%d bloggers like this: