invited review
Developmental regulation of erythropoietin
and erythropoiesis
K. M. MORITZ,1 GAIK BEE LIM,2 AND E. M. WINTOUR1
Florey Institute of Experimental Physiology and Medicine,
University of Melbourne, Parkville, Victoria 3052, Australia;
and 2Department of Medicine, National University Hospital, Singapore 5000
1Howard
development; red blood cells; placenta; brain; mesonephros; metanephros
for all processes of life at the
cellular level (18, 68). Because red blood cells act as the
oxygen transporter in blood via the high-affinity hemoglobin molecule, erythropoiesis has to be tightly regulated so as to both maintain homeostasis and to meet
changes in oxygen supply and demand. Only one type of
fish (of the family Channichthyidae) does not have
hemoglobin, because of the loss of the b-globin gene
some 25 million years ago (see DEVELOPMENT OF ERYTHROPOIESIS IN FISH). The principal factor in the regulation
of erythropoiesis is a glycoprotein hormone named
erythropoietin (Epo). Although other factors may synergize with Epo (as discussed later), mice that are null
mutants for either the Epo gene or its receptor die
about day 13 of development (180), demonstrating the
critical importance of this one hormone. It is effective at
10211 M. The hormone is very sensitive to changes in
oxygen availability, and its levels are finely tuned by
changes in levels of oxygenation, by a classical negative
feedback loop. The story of erythropoiesis is therefore
very much the story of Epo and its regulation.
Erythropoiesis is the process whereby a fraction of
primitive multipotent hemopoietic stem cells becomes
committed to the red cell lineage, forming first burstforming units-erythroid (BFUe), then colony-forming
units-erythroid (CFUe), normoblasts, erythroblasts, reticulocytes, and ultimately the mature erythrocyte. The
oxygen-carrying pigment of mammalian erythrocytes,
OXYGEN IS REQUIRED
or red blood cells, is hemoglobin, four heme molecules
attached to four globin chains, which is synthesized in
the BFUes. Stem cells require one major growth factor
(Steele factor, Stem cell factor, c kit ligand) for their
replication and express the c-kit receptor. In the adult,
erythropoiesis requires the actions of interleukin-3
(IL-3) and Epo acting on their appropriate receptors.
ERYTHROPOIETIN: HISTORY
The concept that red blood cell production was regulated hormonally was first proposed in 1906 by Carnot
and Deflandre (78). Subsequent to that, however, it took
nearly half a century before the existence of such a factor, named erythropoietin, was proven conclusively. It
was another 20 years before the hormone was purified in amounts sufficient to deduce its amino acid
structure and 10 years after that the world was presented with the gene for Epo. In the 12 years since, many
advances have also been made in the understanding of
the mechanism of action of Epo and its regulation.
There have been many reviews about these topics in
the adult (36, 77, 78, 83, 86, 123, 138, 161, 184). This
review will concentrate on the development of erythropoiesis in the embryo/fetus/neonate and the sites at
which it occurs, as well as the ontogeny and regulation
of Epo during development.
0363-6119/97 $5.00 Copyright r 1997 the American Physiological Society
R1829
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Moritz, K. M., Gaik Bee Lim, and E. M. Wintour. Developmental
regulation of erythropoietin and erythropoiesis. Am. J. Physiol. 273
(Regulatory Integrative Comp. Physiol. 42): R1829–R1844, 1997.—It is
well established that erythropoiesis occurs first in the yolk sac, then in
the liver, subsequently moving to the bone marrow and, in rodents, the
spleen during development. The origin of the erythropoietic precursors
and some factors suggested to be important for the changing location of
erythropoiesis are discussed in this review. Until recently, the major site
of erythropoietin (Epo) production in the fetus was thought to be the liver,
but studies have shown now that the Epo gene is expressed strongly in
the fetal kidney, even in the temporary mesonephros. The metanephric
Epo mRNA is upregulated by anemia, downregulated by glucocorticoids,
and contributes substantially to circulating hormone levels in hemorrhaged ovine fetuses. Other sites of Epo and Epo receptor production,
likely to have important actions during development, are the placenta
and the brain.
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INVITED REVIEW
ERYTHROPOIESIS: MAMMALIAN
ROLE OF EXTRACELLULAR MATRIX IN ESTABLISHING
LIVER ERYTHROPOIESIS
The family of integrins is composed of cell surface
receptors that mediate interactions between cells and
the extracellular matrix (71). They are composed of two
units (a and b). Hemapoietic progenitor cells adhere
selectively to fibronectin, not to collagen, laminin, or
proteoglycans (71), and one member of the b1-integrin
receptors, very late activation antigen 4, has been
shown to be critical for erythropoiesis specifically (63).
In mice in which the b1-integrin gene has been deleted
by homologous recombination, the hematopoietic stem
cell can differentiate but not colonize the liver, although
yolk sac production is normal (67).
WHY DOES ERYTHROPOIESIS CEASE IN THE LIVER
TOWARD TERM?
There is some evidence that the suppression of
erythropoiesis in the liver is due to increasing glucocorticoid concentrations in fetal plasma as term approaches. Decapitation of rat fetuses, which thereby
removed the pituitary, at day 17.5 of gestation, resulted
in significantly increased liver erythropoiesis at day 21
than in control fetuses (10). These studies were ex-
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When and where do erythrocyte precursor cells originate? For some time now, it has been known that, in
humans, erythropoiesis occurs first in the yolk sac (3- to
4-wk-old human embryo), where relatively large, nucleated red blood cells (megaloblasts) are formed, containing predominantly, but not exclusively, embryonic hemoglobin (130). Embryonic hemoglobin (Hb Gower 1)
contains 2z- and 2e-globin chains. There are also small
amounts of Hb Gower 2 (a2e2 ) and Hb Portland (z2t2 ).
By 8 wk, the type of hemoglobin synthesized is predominantly fetal (a2t2 ) with small quantities of adult (a2b2 );
the liver is the predominant site of definitive erythropoiesis, and the cells are nonnucleated macrocytes. Erythropoiesis begins in the bone marrow by 11–12 wk and
ceases in the liver about the time of birth (178).
The sheep has been used widely as a developmental
animal model. It is particularly suitable for the study of
erythropoiesis, hemoglobin switching, and erythropoietin production, as the sites of erythropoiesis, and the
types of hemoglobin produced at different stages, most
closely resemble the situation in humans (175). Fetal
hemoglobin (a2t2 ) can first be detected at 26 days
(where term is 145–150 days) and completely replaces
embryonic Hb by 37 days, and synthesis of a2t2 terminates around birth. Adult hemoglobin synthesis (a2b2 )
begins ,1 mo before birth, but some cells contain fetal
hemoglobin until 3 wk after birth. The liver is the major
site of fetal erythropoiesis from 26 to 130 days, when
the bone marrow becomes the most important site. The
fetal spleen does not contain a store of red blood cells
until just after birth (17, 134).
In mice and rats the same sites of erythropoiesis
occur as in humans, but the embryonic hemoglobin
(z2e2 ) is replaced by adult hemoglobin (a2b2 ) without
the formation of fetal hemoglobin (a2t2 ). The yolk sac is
seeded by hematopoietic precursors by 8 days, and the
liver by 10 days, postcoitum.
More recently, the questions have concerned whether
there are independent cell precursors for embryonic
and definitive red blood cells and the origin of the cells
that colonize the yolk sac.
By establishing that both primitive and definitive
erythroid cells could synthesize, simultaneously, embryonic, fetal, and adult hemoglobins, investigators came
to the conclusion that there was no change in hemopoietic cell lineage, but rather a time-dependent change in
progenitor cell genetic programming (152). Similar
conclusions were drawn from studies on the differentiation, in vitro, of mouse embryonic stem cells (ES; 142).
Mouse ES are totipotent cells derived from the inner
cell mass of mouse blastocysts, which, if allowed to
differentiate, in vitro, can give rise to complex cystic
embryoid bodies that may also include visible blood
islands by day 8 of culture (34). Even before differentiation begins, these ES cells, in preculture medium, can
be shown by reverse transcription-polymerase chain
reaction to be expressing Epo and Epo receptor (EpoR)
genes (142). By day 8 of differentiation, erythroid BFUe
can be detected. When the EpoR was ‘‘knocked out,’’
embryonic primitive erythropoiesis occurred, but definitive cells did not colonize the liver. Homozygous mutants died by embryonic day 13.5 because of failure of
definitive erythropoiesis in the fetal liver. However,
primitive yolk sac erythropoiesis was not affected until
embryonic day 9.5, after which proliferation was impaired and the erythrocytes were smaller than normal
(98, 180).
However, the yolk sac is not the only preliver site of
development of multipotent hematopoietic progenitors
(CFU-S). It is now known that, in the mouse at least, an
intraembryonic preliver site of production of CFU-S is a
region designated the AGM [dorsal aorta, genital ridge/
gonads, and pro/mesonephros (114)]. Using an in vitro
organ culture system, these workers were able to show
that hematopoietic stem cells initiated autonomously
in the mouse AGM region, and they suggested that the
in vitro expansion potential of CFU-S from the AGM
was greater than that of yolk sac-derived cells. Hence
they concluded that the definitive adult cells that
colonized the fetal liver most likely derived from the
AGM region. The most recent evidence does suggest
that the precursors of primitive and definitive erythroid cells may be distinct types of cells (123).
What is the precursor of the stem cell? Some evidence
says it is cells that also have the potential to form
endothelial cells. Both blood island formation and
vasculogenesis do not occur when the vascular endothelial growth factor receptor Flk-1 is knocked out (147).
Both endothelial cells and erythroid precursors have
EpoRs, although of different affinities and, maybe,
structure (4). It has been suggested that endothelial
cells express more of the truncated receptor (EpoR-T),
which is more common in early erythroid precursors (5).
INVITED REVIEW
ROLE OF EPO/EPOR IN DEVELOPMENTAL
ERYTHROPOIESIS
In definitive or adult erythropoiesis, there is an
absolute requirement for Epo. The EpoRs first manifest
at the BFU-e stage and reach their highest numbers in
the CFU-e, after which receptor numbers decline.
Fetal erythroid colonies display different growth
characteristics from the adult. There is an accelerated
rate of maturation, resulting in erythrocytes that are
enlarged compared with adult erythrocytes and which
have a shorter life span of 70 days compared with 120
days in the adult. In studies of human umbilical cord
blood samples, both BFU-e and CFU-e reached maximal concentrations sooner than adult progenitors (69).
Moreover, human fetal BFU-e could be stimulated by
Epo alone, whereas the adult BFU-e required additional factors, such as IL-3 or GM-colony-stimulating
factor (CSF) (39). CFU-e also have different sensitivities to Epo at different stages of development. CFU-e
derived from 8.5-day mouse embryos were more sensitive to Epo than 13.5-day fetal liver and adult bone
marrow (187), although this effect was not observed in
human progenitors (148). The mechanisms by which
these effects occur are not well understood, but could
result from different forms of the EpoR being expressed
during fetal life or different proportions of more efficient receptors being expressed.
Role of Epo in ‘‘fetal’’ vs. ‘‘adult’’-type erythropoiesis.
In adult humans, an accelerated fetal-like erythropoiesis occurs when high concentrations of Epo are administered (64), and the mature erythrocytes have a relatively shorter lifespan. Studies in adult rats have
shown that recombinant human erythropoietin
(rHuEpo) stimulated newborn-like hemoglobin synthesis (1).
DEVELOPMENT OF ERYTHROPOIESIS IN FISH
With the exception of one group (Antarctic icefish),
all fish have some nucleated red blood cells containing
an oxygen-carrying pigment (48, 57, 139). In adult fish,
a spleen (sometimes located in the intestinal submucosa), the head-kidney (pronephros), and mesonephros
(42, 164) have been found to be sites of erythropoiesis.
The hemopoietic cells contain receptors for growth
hormone (19) and contain a factor that competes with
human Epo in a radioimmunoassay (164).
During development, blood islands do form in a yolk
sac structure, before migrating to the ‘‘spleen’’’ and/or
kidney; there may also be some temporary hepatic
erythropoiesis.
The ice fish (family: Channichthyidae, suborder:
Notothenioidei) form an interesting group. The DNA
contains a-globin-like sequences, but because the
b-globin gene is not present in the genome, the fish do
not express any hemoglobin (23). These fish survive
because of physiological adaptations (scalelessness,
high skin respiration, and increased cardiac output and
blood volume) coupled with the fact that the waters of
the Antarctic are very well oxygenated because of the
extreme cold and turbulence. Heart ventricles, but not
atria or oxidative skeletal muscle, do contain myoglobin (150). It would be most interesting to know if the
Epo gene remains expressed in these fish.
EPO GENE AND PROTEIN: ADULT
The coding sequences for the Epo gene have now been
elucidated for at least 10 mammalian species, including
humans, two monkey species, rat, mouse, cat, dog,
sheep, and pig (52, 74, 99, 162). The Epo gene is highly
conserved between species. There is 80–82% amino
acid identity between the human sequence and that of
pig, sheep, mouse, and rat Epo genes (162). The singlecopy gene comprises five exons and four introns. Intron
I, the 58 flanking region and the 38 untranslated region
are also highly conserved between species. In 1993,
Galson et al. (55) compared several kilobases of flanking region in the mouse and human genes and found
several homologous regions over 3.3 kb of mouse and
5.5 kb of human 58 flanking sequence. These regions of
high homology are likely to play a role in the regulation
of Epo gene expression. The 3.9 kb of the 58 flanking
sequence was found to contain consensus recognition
sites for various cytokines and lymphokines, metalresponse elements, glucocorticoid-response elements,
and adenosine 38,58-cyclic monophosphate-response elements, as well as binding sites for AP1, nuclear factor
(NF)kb, and Sp1 transcription factors (70). The 38
flanking region included nitrogen-regulatory/oxygensensing consensus sequences, tissue-specific regulatory
elements, and binding sites for AP and Sp1 (90).
Epo is synthesized as a 193-amino acid polypeptide,
which includes a 27-amino acid residue leader sequence and a carboxy-terminal arginine residue removed during processing. The mature protein is 165amino acids residues long and extensively glycosylated,
with 40% carbohydrate content, resulting in a total
mass of 30.4 kDa. There are three N-linked carbohydrate chains and one O-linked carbohydrate chain
required for the correct processing and export of the
hormone (31, 35) and for its in vivo bioactivity (66).
Four conserved cysteine residues give rise to two
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tended (76), and similar results were obtained in
fetuses of bilaterally adrenalectomized mothers and
were reversed by cortisol treatment. In vitro, the effect
of dexamethasone on the number of CFUe was dependent on the concurrent Epo concentration (14). When
Epo values were .25 mU/ml, dexamethasone treatment inhibited CFUe formation. Erythroid precursors,
in rat liver, contain dexamethasone -binding sites with
characteristics of glucocorticoid receptors at days 15–16
of gestation (113). In chronically cannulated bilaterally
adrenalectomized fetal sheep, the amount of a-globin
mRNA in the liver at term was significantly greater
than in intact controls (Gunnersen and Wintour, unpublished results). Thus there is some support for the idea
that increasing levels of glucocorticoids may decrease
hepatic erythropoiesis, at least in rat and sheep fetuses. It is likely that there may be other mechanisms
that have a role in decreasing liver erythropoiesis and
these may differ between species.
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INVITED REVIEW
disulfide bridges. Although Epo has no significant
amino acid homology with any other protein apart from
the recently cloned megakaryocytopoietic hormone,
thrombopoietin (102), structurally it belongs to a family
of cytokines (9, 107). Other members of this family
include growth hormone, prolactin, the CSFs, and
interleukins. Recently a peptide of 20 amino acids, with
no sequence homology to Epo, has been found to
activate the Epo receptor (101, 179). This is discussed
in more detail later.
SITES OF SYNTHESIS: ADULT
REGULATION IN THE ADULT: OXYGEN SENSOR
Regulation of Epo gene expression occurs mainly at
the transcriptional level, although it is tissue specific,
developmentally controlled, and inducible. There are
no intracellular stores of the hormone. Because tissue
hypoxia is the major stimulus to Epo production, much
interest has centered around how this hypoxia is
actually sensed by the body (for review, see Refs. 18 and
137). In 1988, Goldberg et al. (58) proposed that the
oxygen sensor may be a heme protein that changed its
conformational state when insufficient oxygen was
present and thus transmitted the hypoxic signal. Other
studies have suggested a cytochrome P-450 protein
THE EPO RECEPTOR
The human and mouse EpoR have been cloned (for
reviews, see Refs. 104 and 184). In both, there are eight
exons and seven introns (see Fig. 1), encoding a 507/508
amino acid peptide (66 kDa). In the human gene, exons
1–5 encode the 251-amino acid extracellular domain,
exon 6 encodes a 20-amino acid membrane-spanning
region, and the 236-amino acid intracellular or cytoplasmic domain is encoded by exons 7 and 8. The exact
identity of the ligand-binding domain is not known, but
is thought to be encoded by the 58 portion of the fifth
exon and to stretch, probably, from asparagine 209 on
the extracellular domain to the WSXWS (Trp-Ser-X-TrpSer) motif (142). The intracellular signaling domain
appears to be between Gly 352 and Met 396 (141). In
the last 40–90 amino acids of the COOH terminal
cytoplasmic domain, there is a negative regulatory
domain, removal of which increases the efficiency of the
EpoR, such that the dissociation constant (Kd ) is now 1
pM instead of 100 pM (183). In fact, in ,30 members of
a Finnish family who have a benign elevation in
hematocrit, the EpoR gene contains a mutation that
truncates the receptor by 70 amino acids from the
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Kidney. The studies of Jacobson et al. (75) in 1957
established the kidney as the main source of circulating
Epo in the adult. Hybridization histochemistry studies
in the mouse to localize Epo mRNA (84, 88) confirmed
that the kidney is a primary site of Epo synthesis. The
cells that synthesize Epo in the kidney have been shown
to be interstitial cells with fibroblast-like characteristics (7) in the vicinity of the proximal tubules in humans
(38, 94), monkeys (50), mice (88), and sheep (28).
The other major site of Epo production in the adult is
the liver, but hepatic production cannot compensate for
loss of the kidneys in cases of chronic renal failure. The
contribution of hepatic Epo to circulating plasma levels
is 10–15% in anemic nephrectomized rats (41). The
hepatocytes are the major cell type producing Epo
mRNA in the liver (143), although with the use of a
transgenic mouse model (145) it was shown that cells
labeled as being Epo producing were hepatic epithelial
cells in the area around the central veins (85). Maxwell
et al. (112) have since found (1994) expression of Epo in
the fibroblast-like Ito cells that share some characteristics with the cells making Epo in the kidney.
Other sites of expression. Expression has also been
found in the brain (32, 108, 111, 122, 154, 182), the
testis (154), and the placenta (24). Hypoxia can increase mRNA in brain, but to a lesser extent than in
kidney. In these sites, particularly the brain and testes,
there is a barrier to the free diffusion of blood-borne
hormone or locally produced hormone. It is, therefore,
quite likely that the local production of Epo is required
for a purpose other than the classical one. For both the
brain and the testes, there is some evidence of local
effects (see discussion in EPO AND EPOR IN THE FETAL
BRAIN).
may be involved (46) or possibly the hydrogen peroxide
system (2, 47). A region in the 38 flanking region of the
Epo gene, an area that serves as a hypoxia-inducible
enhancer (15, 135), has also been identified. In 1992,
Semenza and Wang (146) identified a DNA-binding
protein, hypoxia-inducible factor 1 (HIF-1), that bound
to part of this region. HIF-1 has now been purified from
Hep3B and HeLa cells and characterized (157, 158).
HIF is a heterodimer transcription factor containing a
120-kDa HIF-a and a 90-kDa HIF-b subunit. The
HIF-b is identical to the aryl hydrocarbon nuclear
translocator sequence. The heterodimer HIF-1 binds to
a consensus sequence on the 38 enhancer of the Epo
gene, close to where the orphan receptor HNF-4 binds
to two tandem consensus steroid hormone response
elements, separated by 2 bp. Both HIF-a and HIF-b
bind to the transcriptional activator p300 and thus
trigger the Epo promoter to transcribe Epo mRNA (70,
87, 144).
There is also evidence for at least one, if not more,
negative regulatory element in the 58 promoter region
of the Epo gene. In 1994, with the use of antisense
oligonucleotides, Imagawa et al. (72) showed that binding to a 30-bp GATA element stimulated Epo gene
transcription even in the absence of hypoxia. This
implies that this element is normally a negative regulatory element. More recently, it has been shown that
GATA transcription factors can negatively regulate Epo
gene expression (73). In addition, in the 38 enhancer of
the human Epo gene, there is a response element for
the TR2 orphan receptor, which also suppresses Epo
gene expression (89). The TR2 orphan receptor is a
protein of 603 amino acids that shares homology with
members of the steroid/thyroid hormone receptor family. TR2 and Epo transcripts colocalized in kidney, liver,
and brain. Thus Epo gene expression is probably under
both positive and negative regulation.
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INVITED REVIEW
Fig. 1. Diagramatic representation of
various forms of the erythropoietin receptor (EpoR). EpoR-T, truncated receptor of Epo; EpoR-S, soluble EpoR; EC,
extracellular; IC, intracellular.
the Epo mimetic are required to produce the biological
effects of Epo. However, significant progress has been
made in understanding how the natural ligand Epo
might bind to the EpoR. It is quite probable that in the
future more active Epo-mimetics will be produced,
which may be of great clinical importance.
SIGNAL TRANSDUCTION
In recent years, much has been learned about the
subsequent signaling events following receptor dimerization. The signal transduction pathway includes the
Janus tyrosine protein kinase 2 (JAK2) and the phosphorylation and nuclear translocation of STAT1 and
STAT5 (signal transducing and activators of transcription proteins) (8, 79, 129, 177).
ONTOGENY OF EPO PRODUCTION IN THE FETUS
AND NEONATE
It had long been thought that, unlike in the adult in
which the kidney is the main Epo-synthesizing organ,
the liver is the predominant source of Epo in the fetus.
This was based on organ ablation studies in fetal sheep
(186) and newborn rats (60). More recent studies on
Epo mRNA expression in intact fetal sheep have shown
that the kidney is a major site of Epo production from
very early in gestation (41 days) (95, 174). In the ovine
fetus, the Epo gene is expressed in interstitial cells in
the vicinity of the proximal tubules in both the permanent metanephric kidney (28) and in the transient
mesonephric kidney (174) (see Fig. 2). This is also the
site of synthesis in human fetal kidney (94). In all
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COOH terminal (29, 103). One of this family became an
Olympic gold medalist in cross-country skiing (103).
At least two other forms of EpoR have been found.
One is soluble, as an alternative splicing event introduces a stop codon before the TM sequence. In the
mouse, the EpoR may bind Epo, but in the human it is
thought not to do so, as it lacks exon 5 (65, 141). There
have also been reports of EpoR-T that lacks most of the
cytoplasmic region except for 56 amino acids encoded
by exon 7 and intron 7 (122, 123). This truncated
receptor is predominantly expressed in immature erythroid cells and can transmit a mitogenic signal, but
fails to prevent apoptosis, and thus cells expressing
EpoR-T may not mature to functional erythrocytes
(122). The full EpoR is found on late-stage progenitors
(see Fig. 1).
EpoR (R129C) has a point mutation in the extracellular domain of the EpoR (183) in which the arginine at
position 129 is changed to a cystine. This leads to a
receptor that is constitutively active in the absence of
the ligand.
One of the most important breakthroughs in determining the mode of action of Epo in binding to its
receptor has come from the recent finding of an Epomimetic peptide (101, 160, 179). This Epo mimetic is a
cyclic (disulfide bonded; cys p6 to cys p15) 20-amino
acid peptide, which bears no sequence homology to Epo.
It functions as a dimer (2 3 2.1 kDa), binds to and
causes dimerization of two EpoR molecules, and functions both in vitro (on human bone marrow) and in vivo
(in mice) as a biologically active Epo. The Kd for the
EpoR is 1,000 times lower (200 nM) than Epo itself
(Kd 5 200 pM), so that much larger concentrations of
R1834
INVITED REVIEW
mammals, there is a period during which these transient mesonephric kidneys and the early metanephric
kidneys are present simultaneously and, during this
period (18–54 days in the sheep), both kidneys contain
Epo mRNA, as shown in Figs. 2 and 3.
By 60 days, the ovine fetal kidney expresses the Epo
gene at a level 10-fold higher than in the liver at the
same age (95). Taking into account that the liver weighs
five times as much as the combined kidney weight, it
would seem as if both organs contribute significantly to
the circulating plasma levels. At ,80 days of gestation,
expression in the fetal liver begins to decline, but
expression remains high in the kidney until after 100
days. By 140 days, Epo mRNA levels in the liver were
only 15% of 60-day levels. In the kidney, expression of
Epo mRNA at 140 days was 10% of that at 60 days (see
Fig. 4).
This change in expression of the Epo gene was
reflected in plasma Epo levels (see Fig. 5), with concentrations being significantly lower at 130–145 days than
at 100 days (118, 120, 176). In the rat, fetal plasma Epo
levels declined from day 17 to day 21 of pregnancy; this
decline was accompanied by a large decline in amniotic
fluid Epo (22). It should be noted that Epo does not
cross the ovine placenta either from fetus to ewe (167)
or from ewe to fetus (168), and thus fetal plasma Epo
concentrations in the sheep reflect fetal production and
clearance. This appears to differ from the mouse where
placental transfer was demonstrated by Sawyer et al.
(140) in 1989. However, in mice that were null mutants
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Fig. 2. Epo gene expression in mesonephros (meso) and metanephros
(meta) and the liver of fetuses at 41 days (d) of gestation. Expression
in 60-day fetal liver and metanephros are shown for comparison. Top:
a Bioimaging analyzer-generated image; Bottom: ratio of Epo mRNA
to standard (Std; 5 µg total RNA; 5 fg standard) expressed as
means 6 SE. [From Wintour et al. (174) with kind permission from
W.B. Saunders Company.]
for Epo, because of homologous recombination, the
fetus dies at 13.5 days of development, indicating that
maternal Epo is not sufficient to maintain adequate
erythropoiesis and viability of the developing mouse.
Widness et al. (165) have reported that Epo clearance
rates are significantly greater in the late-gestation
ovine fetus than in the adult, which could account for
lower levels in the fetus than in the adult. The sites at
which Epo are metabolized/cleared are not known even
in the adult. It has been claimed that metabolic clearance rate and/or half-life (t 5 1⁄2) of rhEpo in adult rats
or sheep is not affected by the numbers of erythropoietic precursors in the bone marrow (133) or by removal
of the kidneys or liver (172). In adult rats, the metabolic
clearance rate and t‰ of rhEpo are not affected by
pregnancy (59). One major unanswered question, therefore, is how and where Epo is cleared in the fetus.
In human fetuses, Epo mRNA is also detected in the
kidney at midgestation (125), and plasma Epo levels
have been reported to increase significantly throughout
development (155), although there was no correlation
with maternal Epo values (92). However, the samples
from early to midgestation were taken by fetoscopy,
whereas those at term were from cord blood after delivery.
Widness et al. (165) reported that cord blood Epo values
were higher following labor when compared with cesarean section. Therefore there may not be a real increase
in plasma Epo with gestational age if samples were
collected by the same method, e.g., fetoscopy. In fact,
other studies (51) showed quite low levels of fetal Epo
throughout human development from 20 to 40 wk,
levels being generally less than 5 mU/ml.
It is interesting that Epo levels should be relatively
low during gestation when erythropoiesis is occurring
at a rapid rate. In the ovine fetus, blood volume is
increasing at the rate of 7 ml/day over the last third of
gestation to keep up with the rapid growth of the fetus
(12). Hematocrit is maintained over this period so the
fetus must be producing some 50 3 109 extra red blood
cells daily. It thus would appear that Epo is either more
efficient at stimulating erythropoiesis during fetal development or that some other factor is synergizing with
Epo to increase its effectiveness. Also, it may be that
Epo acts as a paracrine hormone in the liver as this is
both a site of Epo production and erythropoiesis during
development. Hepatocyte growth factor (HGF) in the
presence of Epo has been demonstrated to stimulate
hemopoietic progenitors to form colonies in vitro, an
effect that is synergized by stem cell factor (54). The
HGF receptor mRNA is highly expressed in embryonic
erythroid cells, and HGF mRNA was also produced by
the embryonic liver of the mouse (54). A recent report
presents evidence that thrombopoietin may have a
proliferative effect on erythroid cells in vitro (128).
Activin, which is known to be formed by the placenta,
can also potentiate the effects of Epo on BFUe and
CFUe formation in vitro (115, 185). The IGF family
have also been reported to stimulate erythropoiesis in
vitro (6, 16), but IGF-I was not effective in preventing
the normal postnatal decline in hematocrit in the
neonatal lamb in vivo (119).
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Epo concentrations are significantly higher in smallfor-gestational age fetuses (92, 151), and this is associated with fetal acidemia and with fetal erythroblastosis. Umbilical cord plasma Epo was also higher in
Fig. 4. Relative levels of Epo gene expression in liver (open bars) and
kidney (hatched bars) across gestation in the ovine fetus as determined by competitive reverse transcription-polymerase chain reaction.
Fig. 5. Plasma Epo levels (open bars) and hematocrit (hatched bars)
in sheep at various stages of development.
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Fig. 3. Light microscope image of mesonephros at 41 days of gestation in the ovine fetus, hybridized with sense (A)
and antisense (B) 35S-labeled riboprobes for ovine Epo. Arrows indicate a group of interstitial cells in the vicinity of
the proximal tubules (P) expressing the Epo gene. Light (C)- and dark (D)-field micrographs of a section of ovine
metanephros from a 60-day gestation fetus using the same probe. The sites of Epo gene expression are groups of
interstitial cells labeled with black dots (arrows in C) or white dots in the dark field. All sections were
counterstained with hematoxylin-eosin. Original magnification 3400. [From Wintour et al. (174).]
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cautious, however, with respect to widespread use of
rhEpo in premature babies.
EPO AND EPOR IN THE FETAL BRAIN
It is of great interest that Epo may have an important
role in the fetus/neonate during brain development. In
adult rats it was shown that whole brain contained
small amounts of Epo mRNA (,5% of that in the normoxic kidney) that could be increased by hypoxia, but
was insignificant with respect to the 200-fold increase
induced by hypoxia in renal mRNA content (154). The
rat pheochromocytoma cell line, PC-12 cells, expresses
an EpoR, the mRNA of which is indistinguishable from
that expressed by rat splenic erythroid precursors, but
which has a very low affinity (Kd 5 16 nM) compared
with either of those on rat cells (Kd 5 95 pM for
high-affinity sites; Kd 5 1.9 nM for low-affinity sites),
suggesting that some accessory protein may alter the
ligand-binding affinity (111). These PC-12 cells responded to Epo by increasing intracellular calcium ion
and monoamine (dopamine, dihydroxyphenylacetic acid,
and homovanilic acid but not 3-methyl-dopamine) concentrations.
Primary cultured cells from the septal region of
15-day embryonic mice responded to high levels (5–10
U/ml) of rhEpo with a 50% increase in choline acetyltransferase activity 2 days later (82). Comparable
effects occurred in the cholinergic hybridoma cell line
SN6.10.2.2. What was of even greater significance,
potentially, was that in vivo both low dose (4 daily
injections of 12.5 U/injection) or high dose (4 daily
injections of 125 U/injection) of rhEpo injected into the
brains of rats with unilateral fimbria/fornix resections
increased the survival rate of septal cholinergic neurons (82). In the brain, the Epo produced was slightly
smaller than renal Epo (30.3 vs. 30.5 kDa), which is
thought to be due to less sialylation of brain Epo, and
this Epo, in in vitro tests, was more potent than serum
Epo at stimulating erythropoiesis, but less potent as a
neuronal growth factor. As a neuronal growth factor,
Epo worked in the nanomolar range, whereas onetenth or less stimulated erythropoiesis. Epo production
in the brain and placenta probably has more relevance
to fetal development than to physiological actions in
the adult (111). Specific Epo-binding sites were found in
mouse brains, with the highest intensity of labeling
occurring in the internal capsule, corpus callosum,
zona incerta, fimbria hippocampus, and mamillothalamic tract (32). Moderate labeling also occurred in the
hippocampus, pyramidal cells of the cortex, brain stem,
mesencephalon, lateral posterior thalamic nuclei, and
granular layer of the cerebellum. Both Epo and EpoR
mRNAs were detected in these mouse brains. Very
recently, human and monkey brains have also been
shown to express both Epo and the EpoR (108), and it
was also demonstrated that mouse astrocytes are a
source of Epo that can be increased .100-fold by
hypoxia. The sites at which human brain tissue seemed
to contain Epo and EpoR mRNA were the temporal
cortex, amygdala, and hippocampus. However, brain
capillary endothelial cells also contain abundant (10,300
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infants of diabetic mothers (169) and in infants of
mothers consuming .300 g of ethanol per week (61).
Interestingly, infants with renal agenesis have elevated
serum Epo concentrations compared with reference
values (166). Neonates with congenital kidney diseases
such as polycystic kidney diseases or severe hydronephrosis have normal values of hematocrit, hemoglobin,
and serum Epo. It has also been demonstrated that
very low birth weight infants have significantly greater
plasma clearance and distribution volume compared
with adults (171).
In healthy children, serum Epo decreases after birth
(35.6 mU/ml, range 17–56 mU/ml) and reaches a nadir
during the first 2 mo (11.5 mU/ml). Epo levels subsequently increase slightly and remain constant between
2 mo and adolescence (18.8 mU/ml, range 7–47 mU/ml)
(37). The period of changing Epo levels after birth
occurs in other species as well, such as the sheep (176).
Neonatal lambs may exhibit a wide range of plasma
Epo concentrations in the month after birth although
levels are always higher than in the unstimulated fetus
or adult (see Fig. 5). After 3–4 wk of age, the plasma
Epo concentrations start to decline and by 8–10 wk
have reached the basal adult level (176). This occurs
despite the well-documented fall in hematocrit that
occurs after birth in all species studied. In the neonatal
lamb, this decline in hematocrit occurs because of a
constant increase in plasma volume with only a slight
increase in red cell volume over weeks 3–7 (119). The
red blood cells are thus diluted in an increasingly larger
plasma volume.
In normal babies, this period is known as the physiological anemia of infancy (27). In premature babies,
however, the anemia is more profound and persists
longer and can lead to a clinical condition termed
’’anemia of prematurity‘‘ (153), which is characterized
by low reticulocyte counts and an inadequate Epo
response. These infants frequently require blood transfusions and in recent years there has been a number of
clinical trials using rhEpo to prevent or alleviate the
anemia. Results of early studies were conflicting, with
some supporting the use of exogenous Epo to treat the
anemia (11, 40) and other studies finding that Epo
treatment had low efficacy and was required at high
doses to be effective (62, 148, 173). In a study on rhesus
monkeys, it was in fact shown that doses that were
effective at increasing Hb values in adults were not
effective in infant monkeys (56). A cost-benefit analysis
carried out in the United States in 1994 (149) showed
that at that time there were no cost savings with
routine use of rhEpo with supplemental red blood cell
transfusions compared with transfusions alone. This
was supported by another study (45) that pointed out
that rhEpo can only become a more cost-efficient treatment if the efficacy of rhEpo can be increased and/or the
cost of the treatment lowered. A pilot study, however,
has shown that the timing of the treatment is important. Infants with very low birth weights had significantly lower transfusion requirement if treatment was
begun at 3 days of age (105). Many clinicians remain
INVITED REVIEW
proliferation of one form of central nervous system
progenitor cells (20).
EPO AND EPOR IN THE PLACENTA
Epo mRNA was detected in the human placenta by
reverse transcription-polymerase chain reaction, but
there was insufficient mRNA to be detected by Northern blot even using 50 µg total RNA loaded (24). It was
quite surprising that relatively large amounts of immunoreactivity were detected by immunohistochemistry
on a wide variety of placental tissue-villous cytotrophoblasts, endovascular and intravascular cytotrophoblasts, cytotrophoblast cell columns, and synctiotrophoblasts. Epo is not stored in the adult kidney or liver (78,
83). The finding that EpoR mRNA is expressed by
placental tissues, in amounts detectable by Northern
analysis, and that the EpoR antibody labeled all the
same tissues as the Epo antibody, makes one wonder
whether previously detected Epo immunoreactivity was
actually due to Epo antibody binding to Epo protein
that was attached to the EpoR (25). Therefore, until
good hybridization histochemistry studies are carried
out with human placenta, the site of Epo synthesis in
this tissue must be regarded as unproven. An independent study showing that endothelial cells of human
umbilical cord do contain EpoR mRNA suggests that
one site of Epo action, whether derived from a local or
systemic source, could be placental vasculature (5). The
role of Epo in the placenta is still unknown. It could
contribute to the increase in maternal plasma Epo that
occurs during normal human and rat pregnancy (26,
30). The increase seems to be due to increased production rather than reduced metabolic clearance rate, at
least in rats (59).
REGULATION OF SYSTEMIC EPO IN THE FETUS
In the ovine fetus, it has been demonstrated that a
standard hemorrhage of 20% fetal body weight over a
10-min period elicits a greater plasma Epo response
earlier in gestation (100–110 days) than later in gestation at 140 days (118, 120). If the fetus is nephrectomized at ,100 days, basal levels of Epo over the
following week do not change. If the fetus is then
hemorrhaged, however, the liver is not able to compensate fully for loss of the kidneys in terms of Epo
production. The plasma level reached at 24 h posthemorrhage was only 19 mU/ml in nephrectomized fetuses
compared with 70 mU/ml in the intact fetuses (97).
Expression of hepatic Epo mRNA was increased to the
same extent in both groups (see Fig. 6). This indicates
that although the liver may produce sufficient Epo for
maintaining adequate erythropoiesis in, for example,
cases of renal agenesis, it can only increase production
to a limited extent during periods of erythropoietic
stress. It also demonstrates that the renal mRNA is
translated into protein.
Studies have also indicated that levels of Epo gene
expression during development may be dependent on
fetal glucocorticoid status. Maternal treatment for 2–3
days with a synthetic glucocorticoid, dexamethasone,
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sites per cell) low-affinity (Kd 5 860 pM) binding sites
for iodinated rhEpo (181). What is now required are
good hybridization histochemistry studies to localize
exact cellular expression sites of Epo and EpoR mRNA.
When the human EpoR gene was incorporated into the
genome of transgenic mice at relatively low levels
(4.7 3 1024 to 17.6 3 1024 ng/µg mRNA), human EpoR
transcripts were detected in the brains of the adult
mice. No endogenous (mouse) EpoR mRNA was detected in adult brains, but very high levels (4.1 3 1022
ng/µg mRNA) were found in the brains of mouse
embryos at embryonic day 10, which decreased with
development, disappearing by embryonic day 16 (100).
This work suggested strongly that the Epo/EpoR system might have a more important role in the developing rather than the adult brain. The human fetus at
13–17 weeks of gestation, which covers the period of
maximum neurogenesis (33), expresses EpoR mRNA in
the spinal cord, tentatively identified as ependymal
cells enclosing the central canal (93). Epo protein has
been measured by enzyme-linked immunosorbent assays in the CSF of preterm and term neonates at levels
higher than in infants or adults (80). These workers
have also reported that Epo and EpoR mRNAs are
present in brains of human fetuses from 5 to 24 wk of
gestation (81). This is a very exciting new area.
The finding that Epo mRNA has been detected in the
hippocampus and amygdala of humans, monkeys, and
mice and can be upregulated by hypoxia/anemia may
have significant potential importance for the fetus,
because these areas are all vulnerable to damage by
intrauterine hypoxic episodes (106). It has therefore
been suggested that Epo may play some role in brain
development. Very recently a decapeptide, LVV-hemorphin 7, was isolated from sheep brain; LVV-hemorphin
7 binds to the angiotensin IV (ANG IV) receptor (AT4 )
and has an identical sequence to part of the sheep
b-globins (116). It has been suggested that this peptide
is indeed the natural ligand for the ANG IV receptorAT4. The synthesis of globins, by mouse neurons, has
been reported (126). It was claimed that neuronal cells
of the adult hippocampus, basal ganglia, and parietal
and cerebellar cortex contained immunoreactive globin. Because mRNA for the EpoR has been found in the
primate cortex, cerebellum, hypothalamus, hippocampus, and nucleus caudatus (108), it is tempting to
speculate that Epo may control the production of the
globin precursor of LVV-hemorphin 7 in the brain.
Because the AT4 receptor appears to be involved in
regulation of neurite outgrowth in cultured embryonic
chicken sympathetic neurons (117), there may be some
role for EpoR in the developing brain via this peptide
LVV-hemorphin 7.
It is of note that the knockout of genes, such as GATA
2 and 3, which arrests erythropoiesis, also leads to
brain abnormalities (127). As noted earlier, the normal
intracellular signaling of the EpoR involves association
of a membrane proximal region with JAK2 that is
autophosphorylated. JAK2 is found in embryonic mouse
brain, and activation of a JAK/STAT (signal transducers and activators of transcription) pathway leads to
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at 60 days of gestation, leads to a decline in Epo
expression in both the kidney and liver, but does not
affect maternal Epo mRNA expression in either organ.
At 80 days of gestation, a similar infusion only decreases renal mRNA. Cortisol infusion directly to the
Fig. 8. Effect of intravenous cortisol infusion (230 µg/h for 48 h) into 5
ovine fetuses at 100–110 days of gestation compared with 7 control
fetuses. Top: Bioimaging analyzer-generated image; B: mean 6 SE of
ratio of Epo mRNA to standard. **P , 0.01. [From Lim et al. (96),
copyright The Endocrine Society.]
fetus for 48 h at 100–110 days also decreases renal Epo
mRNA (see Fig. 7). Adrenalectomy in late gestation is
associated with a fivefold increase in renal Epo mRNA,
an effect that can be reversed by replacement of cortisol
Fig. 7. Effect of cortisol infusion (F) into 3 bilaterally adrenalectomized (Adx) fetuses for 5 days preceding tissue
collection at 141 days of gestation on Epo gene expression, compared with Epo expression in 6 control and 7
bilaterally adrenalectomized fetuses. Top: Bioimaging analyzer-generated image; Bottom: mean 6 SE of ratio of
Epo mRNA to standard in each group of fetuses. *P , 0.05. [From Lim et al. (96), copyright The Endocrine Society.]
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Fig. 6. Epo mRNA/standard ratio in liver (A) and kidney (B) of ovine
fetuses at 3 stages of gestation: 75 (4C, 4H), 110 (4C, 4H, 6HN), and
140 days (4C, 4H), where C is control, H is hemorrhaged, and HN is
nephrectomized and hemorrhaged. Values are means 6 SE. Ratios
are corrected for fixed amount of total RNA (10 µg) and standard (10
fg). *P , 0.05; **P , 0.01. [From Lim et al. (97), with kind permission from Elsevier Science Ireland Ltd., Bay 15K, Shannon Industrial
Estate, Co. Clare, Ireland.]
INVITED REVIEW
to the fetus (96, see Fig. 8). It is possible that the effect
of steroids is via an interaction with a GATA trancription factor.
ROLE OF GATA TRANSCRIPTION FACTORS
IN REGULATION OF BOTH ERYTHROPOIESIS AND EPO
UNANSWERED QUESTIONS
The regulation of developmental erythropoiesis is
obviously of great interest to both clinicians and scientists alike. This review demonstrates that although our
knowledge has increased dramatically over the last
5–10 years, many more questions are constantly being
generated. The recent findings of Epo in the placenta
and brain lead us to wonder as to its role in these
tissues. Very little is still known about how hypoxia
induces Epo. What is the oxygen sensor? Why is Epo
mRNA present in such a large amount early in gesta-
tion in both liver and kidney and then why does it
decline? What is the role of the GATA transcription
factors in the regulation of Epo production? Are there
forms of the EpoR in the fetus that vary from those in
the adult? Is there a greater proportion of a more active
EpoR in the fetus and a change to a large proportion of
less active EpoR in the neonate?
All the works of the authors have been supported by the block
grant to the Institute from NHMRC, Australia.
Address for reprint requests: M. Wintour, Howard Florey Institute
of Experimental Physiology and Medicine, Univ. of Melbourne,
Parkville 3052, Australia.
REFERENCES
1. Abdelrahman, H. G., K. K Srivastava, and M. C. Datta.
Stimulation of newborn-like hemoglobin synthesis in adult rats
by recombinant human erythropoietin and suppression by
asprin. Acta Haematol. 96: 214–220, 1996.
2. Acker, H. Mechanisms and meaning of cellular oxygen sensing
in the organism. Respir. Physiol. 95: 1–10, 1994.
3. Aird, W. C., J. D. Parvin, P. A. Sharp, and R. D. Rosenberg.
The interaction of GATA-binding proteins and basal transcription factors with GATA-box-containing core promoters. J. Biol.
Chem. 269: 883–889, 1994.
4. Anagnostou, A., E. S. Lee, N. Kessimian, R. Levinson, and
M. Steiner. Erythropoietin has a mitogenic and positive chemotactic effect on endothelial cells. Proc. Natl. Acad. Sci. USA 87:
5978–5982, 1990.
5. Anagnostou, A., Z. Liu, S. Manfred, K. Chin, E. S. Lee, N.
Kessimian, and C. T. Noguchi. Erythropoietin receptor mRNA
expression in human endothelial cells. Proc. Natl. Acad. Sci.
USA 91: 3974–3978, 1994.
6. Aron, D. C. Insulin-like growth factor-1 and erythropoiesis.
Biofactors 3: 211–216, 1992.
7. Bachmann, S., M. Le Hir, and K. Eckardt. Co-localization of
erythropoietin mRNA and ecto-58-nucleotidase immunoreactivity in peritubular cells of rat renal cortex indicates that
fibroblasts produce erythropoietin. J. Histochem. Cytochem. 41:
335–341, 1993.
8. Barber, D. L., and A. D. D’Andrea. Erythropoietin and
interleukin-2 activate distinct JAK kinase family members.
Mol. Cell. Biol. 14: 6505–6514, 1994.
9. Bazan, J. F. Haemopoietic receptors and helical cytokines.
Immunol. Today 11: 350–354, 1990.
10. Bearn, J. G. The influence of the rat foetal pituitary on foetal
liver deoxyribonucleic acid and foetal liver haematopoietic
tissue. Acta Anat. (Basel) 68: 239–250, 1967.
11. Bechensteen, A. G., P. Hågå, S. Halvorsen, A. Whitelaw, K.
Liestol, R. Lindemann, J. Grogaard, M. Hellebostad, O. D.
Saugstad, M. Gronn, L. Daae, H. Refsum, and E. Sundal.
Erythropoietin, protein, and iron supplementation and the
prevention of anaemia of prematurity. Arch. Dis. Child. 69:
19–23, 1993.
12. Bell, R. J., and E. M. Wintour. The effect of maternal water
deprivation on ovine fetal blood volume. Q. J. Exp. Physiol. 70:
95–99, 1985.
13. Berry, M., F. Grosveld, and N. Dillon. A single point mutation is the cause of the Greek form of hereditary persistence of
fetal haemoglobin. Nature 358: 499–502, 1992.
14. Billat, C. L., J.-M. Felix, and R. L. Jacquot. In vitro and in
vivo regulation of hepatic erythropoiesis by erythropoietin and
glucocorticoids in the rat fetus. Exp. Hematol. 10: 133–137,
1982.
15. Blanchard, K. L., A. M. Acquaviva, D. L. Galson, and H. F.
Bunn. Hypoxic induction of the human erythropoietin gene:
cooperation between the promoter and enhancer, each of which
contains steroid receptor response elements. Mol. Cell. Biol. 12:
5373–5385, 1992.
16. Boyer, S. H., T. R. Bishop, O. C. Rogers, A. N. Noyes, L. P.
Frelin, and S. Hobbs. Roles of erythropoietin, insulin-like
growth factor I, and unidentified serum factors in promoting
maturation of purified murine erythroid colony-forming units.
Blood 80: 2503–2512, 1992.
Downloaded from http://ajpregu.physiology.org/ by 10.220.33.6 on June 18, 2017
The family of GATA transcription factors contains
proteins that bind to the motif T/A (GATA) A/G in the
DNA of promoter or enhancer of many genes. The
family now includes six members (GATAs 1–6), all
related by a highly conserved DNA-binding region,
comprising two zinc fingers. The first one to be described (GATA1) was first described in chicken erythroid cells as an ‘‘erythroid-specific factor’’ (Eryf 1; 44)
and in human erythroid cell lines as NF-E1 or GF-1 (44,
110, 157). It is a basic 38-kDa protein comprising 304
amino acids with zinc fingers between amino acids
110–134 and 164–188 (43). Mammalian GATA1 contains 413 amino acids. The carboxyl finger is thought to
be sufficient for DNA binding, with the amino finger
possibly specifying location. GATA-1 is found in the
hematopoietic system and Sertoli cells of the testes.
GATA1 and SP1 (which activates the CCACCC box)
appear to cooperate with other transcriptional factors
to activate or inhibit transcription (49). GATA1 is found
in the yolk sac where primitive erythropoiesis is occurring, but the level of GATA1 mRNA vs. globin mRNA is
much higher in the fetal liver where definitive erythropoiesis occurs (163). Evidence suggests that GATA1
represses embryonic h-globin and may also repress
fetal g-globin production (13, 136). In some genes (e.g.,
mouse Epo), the core promoter contains a GATA motif
instead of the consensus TATAAA site (3) and binding of
GATA1 can inhibit transcription initiation. Although it
was not clearly apparent in early studies of chimeric
mice containing GATA1-deleted male embryonic stem
cells (131, 132, 159), it is now established that GATA
2/2 embryos die of anemia at 9.5–10.5 days of embryonic development, and both primitive and definitive
cells die at the proerythroblast stage (53). Similarly,
GATAs 2 and 3 have been shown to be essential for
erythropoiesis (126).
In the 58 promoter region of the Epo gene, there is a
negative regulatory element at 230 bp that contains a
GATA site (72). Further studies by this group established the role of GATA factors in the regulation of Epo
(73). Glucocorticoid inhibition of mouse GATA function
has been demonstrated by the interaction of the steroid
receptor ligand complex and GATA protein (21).
R1839
R1840
INVITED REVIEW
38. Eckardt, K., M. Möllman, R. Neumann, R. Brunkhorst,
H.-U. Burger, G. Lonnemann, H. Scholz, G. Keusch, B.
Buchholz, U. Frei, C. Bauer, and A. Kurtz. Erythropoietin in
polycystic kidneys. J. Clin. Invest. 84: 1160–1166, 1989.
39. Emerson, S. G., S. Thomas, J. L. Ferrara, and J. L.
Greenstein. Developmental regulation of erythropoiesis by
hematopoietic growth factors: analysis on populations of BFU-E
from bone marrow, peripheral blood and fetal liver. Blood 74:
49–55, 1989.
40. Emmerson, A. J. B., H. J. Coles, C. M. M. Stern, and
Pearson. Double blind trial of recombinant human erythropoietin in preterm infants. Arch. Dis. Child. 68: 291–296, 1993.
41. Erslev, A. J., J. Caro, E. Kansu, and R. Silver. Renal and
extrarenal erythropoietin production in anaemic rats. Br. J.
Haematol. 45: 65–72, 1980.
42. Esteban, M. A., J. Meseguer, A. G. Ayala, and B. Agulleiro.
Erythropoiesis and thromobopoiesis in the head-kidney of the
Sea Bass (Dicentrarchus labrax L.): an ultrastructural study.
Arch. Histol. Cytol. 52: 407–419, 1989.
43. Evans, T., and G. Felsenfeld. The erythroid-specific transcription factor eryfl: a new finger protein. Cell 58: 877–885, 1989.
44. Evans, T., M. Reitman, and G. Felsenfeld. An erythrocytespecific DNA-binding factor recognizes a regulatory sequence
common to all chicken globin genes. Proc. Natl. Acad. Sci. USA
85: 5976–5980, 1988.
45. Fain, J., P. Hilsenrath, J. A. Widness, R. G. Strauss, and
A. H. Mutnick. A cost analysis comparing erythropoietin and
red cell transfusions in the treatment of anaemia of prematurity. Transfusion 35: 936–943, 1995.
46. Fandrey, J. Role of cytochrome P450 in the control of the
production of erythropoietin. Life Sci. 47: 127–134, 1990.
47. Fandrey, J., S. Frede, and W. Jelkmann. Role of hydrogen
peroxide in hypoxia-induced erythropietin production. Biochem. J. 303: 507–510, 1994.
48. Fänge, R. Fish blood cells. In: Fish Physiology. New York:
Academic, 1992, p. 1–54.
49. Fischer, K.-D., A. Haese, and J. Nowock. Cooperation of
GATA-1 and Sp1 can result in synergistic transcriptional
activation or interference. J. Biol. Chem. 268: 23915–23923,
1996.
50. Fisher, J. W., S. Koury, T. Ducey, and S. Mendel. Erythropoietin production by interstitial cells of hypoxic monkey kidneys.
Br. J. Haematol. 95: 27–32, 1996.
51. Forestier, F., F. Daffos, N. Catherine, M. Renard, and J.
Andreux. Developmental hematopoiesis in normal human
fetal blood. Blood 77: 2360–2363, 1991.
52. Fu, P., B. Evans, G. B. Lim, K. M. Moritz, and E. M.
Wintour. The sheep erythropoietin gene: molecular cloning and
effect of hemorrhage on plasma erythropoietin and renal/liver
messenger RNA in adult sheep. Mol. Cell. Endocrinol. 93:
107–116, 1993.
53. Fujiwara, Y., C. P. Browne, K. Cunniff, S. C. Goff, and S. H.
Orkin. Arrested development of embryonic red cell precursors
in mouse embryos lacking transcription factor GATA-1. Proc.
Natl. Acad. Sci. USA 93: 12355–12358, 1996.
54. Galimi, F., G. P. Bagnara, L. Bonsi, E. Cottone, A. Follenzi,
and A. Simeone. Hepatocyte growth factor induces proliferation and differentiation of multipotent and erythroid hemopoietic progenitors. J. Cell Biol. 127: 1743–1754, 1994.
55. Galson, D. L., C. C. Tan, P. J. Ratcliffe, and H. F. Bunn.
Comparison of the human and mouse erythropoietin genes
shows extensive homology in the flanking regions. Blood 82:
3321–3326, 1993.
56. George, J. W., C. A. Bracco, K. M. Shannon, G. J. Davis,
I. L. Smith, R. H. Phibbs, and A. G. Hendrickx. Age-related
differences in erythropoietic response to recombinant human
erythropoietin: comparison in adult and infant rhesus monkeys.
Pediatr. Res. 28: 567–571, 1990.
57. Glomski, C. A., J. Tamburlin, and M. Chainani. The phylogenetic odyssey of the erythrocyte. III. Fish, the lower vertebrate experience. Histol. Histopathol. 7: 501–528, 1992.
58. Goldberg, M. A., S. P. Dunning, and H. F. Bunn. Regulation
of the erythropoietin gene: evidence that the oxygen sensor is a
heme protein. Science 242: 1412–1415, 1988.
Downloaded from http://ajpregu.physiology.org/ by 10.220.33.6 on June 18, 2017
17. Brace, R. A. Blood volume and its measurements in chronically
catheterized sheep fetus. Am. J. Physiol. 244 (Heart Circ.
Physiol. 13): H487–H494, 1983.
18. Bunn, H. F., and R. O. Poyton. Oxygen sensing and molecular
adaptation to hypoxia. Am. Physiol. Rev. 76: 839–885, 1996.
19. Calduch-Giner, J. A., A. Sitjà-Bobadilla, P. AlvarezPellitero, and J. Pérez-Sánchez. Evidence for a direct action
of GH on haemopoietic cells of a marine fish, the gilthead sea
bream (Sparus aurata). J. Endocrinol. 146: 459–467, 1995.
20. Cattaneo, E., C. De Fraja, L. Conti, B. Reinach, L. Bolis, S.
Govoni, and E. Liboi. Activation of the JAK/STAT pathway
leads to proliferation of ST14A central nervous system progenitor cells. J. Biol. Chem. 271: 23374–23379, 1996.
21. Chang, T.-J., B. M Scher, S. Waxman, and W. Scher.
Inhibition of mouse GATA-1 function by the glucocorticoid
receptor: possible mechanism of steroid inhibition of erythroleukemia cell differentiation. Mol. Endocrinol. 7: 528–542, 1993.
22. Clemons, G. K., S. L. Fitzsimmons, and D. DeManincor.
Immunoreactive erythropoietin concentrations in fetal and
neonatal rats and the effects of hypoxia. Blood 68: 892–899,
1986.
23. Cocca, E., M. Ratnayake-Lecamwasam, S. K. Parker, L.
Camardella, M Ciaramella, G. di Prisco, and H. W. Detrich III. Genomic remnants of a-globin genes in the hemoglobinless antarctic icefishes. Proc. Natl. Acad. Sci. USA 92: 1817–
1821, 1995.
24. Conrad, K. P., D. F. Benyo, A. Westerhausen-Larsen, and
T. M. Miles. Expression of erythropoietin by the human placenta. FASEB J. 10: 760–766, 1996.
25. Conrad, K. P., T. M. Miles, and D. Fairchild Benyo.
Evidence for expression of the erythropoietin receptor by trophoblast cells of the human placenta (Abstract). J. Soc. Gynecol.
Invest. 4: 228, 1997.
26. Cotes, P. M., and C. E. Canning. Changes in serum immunoreactive erythropoietin during the menstrual cycle and normal
pregnancy. Br. J. Obstet. Gynaecol. 90: 304–311, 1990.
27. Dallman, P. R. Anemia of prematurity. Annu. Rev. Med. 32:
143–160, 1981.
28. Darby, I. A., B. A. Evans, P. Fu, G. B Lim, K. M. Moritz, and
E. M. Wintour. Erythropoietin gene expression in fetal and
adult sheep kidney. Br. J. Haematol. 89: 266–270, 1995.
29. De la Chapelle, A., A. L. Träskelin, and E. Juvonen.
Truncated erythropoietin receptor causes dominantly inherited
benign human erythrocytosis. Proc. Natl. Acad. Sci. USA 90:
4495–4499, 1993.
30. Del Valle, G. O., M. D. Mosher, and K. P Conrad. Serum
immunoreactive erythropoietin and red blood cells mass during
pregnancy in conscious rats. Am. J. Physiol. 265 (Regulatory
Integrative Comp. Physiol. 34): R399–R403, 1993.
31. Delorme, E., T. Lorenzini, J. Griffin, F. Martin, F. Jacobsen, T. Boone, and S. Elliott. Role of glycosylation on the
secretion and biological activity of erythropoietin. Biochemistry
31: 9871–9872, 1992.
32. Digicaylioglu, M., S. Bichet, H. H. Marti, R. H. Wenger, L.
Rivas, C. Bauer, and M. Gassman. Localization of specific
erythropoietin binding sites in defined areas of the mouse brain.
Proc. Natl. Acad. Sci. USA 92: 3717–3720, 1995.
33. Dobbing, J., and J. Smart. Vulnerability of developing brain
and behaviour. Br. Med. Bull. 30: 164–168, 1974.
34. Doetschman, T. C., H. Eistetter, M. Katz, W. Schmidt, and
R. Kemler. The in vitro development of blastocyst-derived
embryonic stem cell lines: formation of visceral yolk sac, blood
islands, and myocardium. J. Embryol. Exp. Morphol. 87: 27–32,
1985.
35. Dube, S., J. W. Fisher, and J. S. Powell. Glycosylation at
specific sites of erythropoietin is essential for biosynthesis,
secretion and biological function. J. Biol. Chem. 263: 17516–
17521, 1988.
36. Eckardt, K. Erythropoietin: oxygen-dependent control of erythropoiesis and its failure in renal disease. Nephron 67: 7–23,
1994.
37. Eckardt, K., W. Hartmann, U. Vetter, F. Pohlandt, R.
Burghardt, and A. Kurtz. Serum immunoreactive erythropoietin of children in health and disease. Eur. J. Pediatr. 149:
459–463, 1990.
INVITED REVIEW
80. Juul, S. E., J. Harcum, Y. Li, and R. D. Christensen.
Erythropoietin is present in the cerebrospinal fluid of neonates
(Abstract). Pediatr. Res. 39: 1715, 1996.
81. Juul, S. E., Y. Li, D. A. Calhoun, and R. D. Christensen.
Erythropoietin and its receptor are expressed in the central
nervous system of first and second trimester human fetuses
(Abstract). Pediatr. Res. 39: 1301, 1996.
82. Konishi, Y., D.-H. Chui, H. Hirose, T. Kunishita, and T.
Tabira. Trophic effect of erythropoietin and other hematopoietic factors on central cholinergic neurons in vitro and in vivo.
Brain Res. 609: 29–35, 1993.
83. Koury, M. J., and M. C. Bondurant. The molecular mechanism of erythropoietin action. Eur. J. Biochem. 210: 649–663,
1992.
84. Koury, S., M. Bondurant, and M. Koury. Localization of
erythropoietin synthesizing cells in murine kidneys by in situ
hybridization. Blood 71: 524–527, 1988.
85. Koury, S. T., M. C. Bondurant, M. J. Koury, and G. L.
Semenza. Localization of cells producing erythropoietin in
murine liver by in situ hybridization. Blood 77: 2497–2503,
1991.
86. Krantz, S. B. Erythropoietin. Blood 77: 419–434, 1991.
87. Kvietikova, I., R. H. Wenger, H. H. Marti, and M. Gassman.
The hypoxia-inducible factor-1 DNA recognition site is cAMPresponsive. Kidney Int. 51: 564–566, 1997.
88. Lacombe, C., J. Da Silva, P. Bruneval, N. Foutnier, N.
Wendling, J. Casadevall, J. Camilleri, B. Bariety, B. Varet,
and P. Tambourin. Peritubular cells are the site of erythropoietin synthesis in the murine hypoxic kidney. J. Clin. Invest. 81:
620–623, 1988.
89. Lee, H.-J., W.-J. Young, C. C. Shih, and C. Chang. Suppression of the human erythropoietin gene expression by the TR2
orphan receptor, a member of the steroid receptor superfamily.
J. Biol. Chem. 271: 10405–10412, 1996.
90. Lee-Huang, S., J.-J. Lin, H. F. Kung, L. Lee, and P.
Lee-Huang. The 38 flanking region of the human erythropoietinencoding gene contains nitrogen-regulatory/oxygen-sensing consensus sequences and tissue-specific transcriptional regulatory
elements. Gene 137: 203–210, 1993.
91. Lee-Huang, S., J. J. Lin, H. F. Kung, L. Lee, and P.
Lee-Huang. The human erythropoietin-encoding gene contains a CAAT box, TATA boxes and other transcriptional
regulatory elements in its 58 flanking region. Gene 128: 227–
236, 1993.
92. Lemery, D. J., J. Santolaya, A. F. Serre, S. Denoix, G. H.
Besse, P. C. Vanlieferinghen, M. J. Bezou, G. Gaillard, and
B. Jacquetin. Serum erythropoietin in small for gestational
age fetuses. Biol. Neonate 65: 89–93, 1994.
93. Li, Y., S. E. Juul, J. A. Morris-Wiman, D. A. Calhoun, and
R. D. Christensen. Erythropoietin receptors are expressed in
the central nervous system in mid-trimester human fetuses.
Pediatr. Res. 40: 376–380, 1996.
94. Liapis, H., J. Roby, T. P. Birkland, R. M. Davila, D. Ritter,
and W. C. Parks. In situ hybridization of human erythropoietin in pre- and postnatal kidneys. Pediatr. Pathol. Lab. Med.
15: 875–883, 1995.
95. Lim, G. B., K. Jeyaseelan, and E. M. Wintour. Ontogeny of
erythropoietin gene expression in the sheep fetus: effect of
dexamethasone at 60 days of gestation. Blood 84: 460–466,
1994.
96. Lim, G. B., M. Dodic, L. Earnest, K. Jeyaseelan, and E. M.
Wintour. Regulation of erythropoietin gene expression in fetal
sheep by glucocorticoids. Endocrinology 137: 1658–1663, 1996.
97. Lim, G. B., K. Moritz, K. Jeyaseelan, and E. M. Wintour.
Effect of hemorrhage and nephrectomy on erythropoietin gene
expression in the ovine fetus. Mol. Cell. Endocrinol. 117:
101–109, 1996.
98. Lin, C.-S., S.-K. Lim, V. D’Agati, and F. Costantini.
Differential effects of an erythropoietin receptor gene disruption on primitive and definitive erythropoiesis. Genes Dev. 10:
154–164, 1996.
99. Lin, F.-K., C.-H. Lin, P.-H. Lai, J. K. Browne, J. C. Egrie, R.
Smalling, G. M. Fox, K. K. Chen, M. Castro, and S. Suggs.
Monkey erythropoietin gene: cloning expression and compari-
Downloaded from http://ajpregu.physiology.org/ by 10.220.33.6 on June 18, 2017
59. Gough, S. R., M. D. Mosher, and K. P. Conrad. Metabolism
of erythropoietin in conscious pregnant rats. Am. J. Physiol. 268
(Regulatory Integrative Comp. Physiol. 37): R1117–R1120, 1995.
60. Gruber, D. F., J. R. Zucali, J. Wleklinski, V. Larussa, and
E. A. Mirand. Temporal transition in the site of rat erythropoietin production. Exp. Hematol. 5: 399–407, 1977.
61. Halmesmaki, E., K. A. Teramo, J. A. Widness, G. K. Clemons, and O. Ylikorkala. Maternal alcohol abuse is associated
with elevated fetal erythropoietin levels. Obstet. Gynecol. 76:
219–222, 1990.
62. Halperin, D. S., P. Wacker, G. Lacount, M. Felix, J.-F.
Babel, M. Aapro, and M. Wyss. Effects of recombinant human
erythropoietin in infants with the anemia of prematurity. J.
Pediatr. 116: 779–786, 1990.
63. Hamamura, K., H. Matsuda, Y. Takeuchi, S. Habu, H.
Yagita, and K. Okumura. A critical role of VLA-4 in erythropoiesis in vivo. Blood 87: 2513–2517, 1996.
64. Hanicki, Z., W. Sulowicz, M. Stepniewski, M. Kuzniewski,
A. Krasniak, J. Kopec, U. Cieszkowska, and K. Stolarska.
Recombinant human erythropoietin stimulates synthesis of
fetal haemoglobin in haemodialyzed patients with anaemia due
to end-stage kidney. Contrib. Nephrol. 11: 11–14, 1990.
65. Harris, K. W., and J. C. Winkelmann. Enzyme-linked immunosorbent assay detects a potential soluble form of the erythropoietin receptor in human plasma. Am. J. Hematol. 52: 8–13,
1996.
66. Higuchi, M., M. Oh-eda, H. Kuboniwa, K. Tomonoh, Y.
Shimonoka, and N. Ochi. Role of sugar chains in the expression of the biological activity of human erythropoietin. J. Biol.
Chem. 267: 7703–7709, 1992.
67. Hirsch, E., A. Iglesias, A. J. Potocnik, U. Hartmann, and
R. Fässler. Impaired migration but not differentiation of
haematopoietic stem cells in the absence of b1 integrins. Nature
380: 171–175, 1996.
68. Hochachka, P. W., L. T. Buck, C. J. Doll, and S. C. Land.
Unifying theory of hypoxia tolerance: molecular/metabolic defense and rescue mechanisms for surviving oxygen lack. Proc.
Natl. Acad. Sci. USA 93: 9493–9498, 1996.
69. Holbrook, S. T., R. D. Christensen, and G. Rothstein.
Erythroid colonies derived from fetal blood display different
growth patterns from those derived from adult marrow. Pediatr.
Res. 24: 605–608, 1988.
70. Huang, L. E., V. Ho, Z. Arany, D. Krainc, D. Galson, D.
Tendler, D. M. Livingston, and H. F. Bunn. Erythropoietin
gene regulation depends on heme-dependent oxygen sensing
and assembly of interacting transcription factors. Kidney Int.
51: 548–552, 1997.
71. Hynes, R. O. Integrins: versatility, modulation, and signaling
in cell adhesion. Cell 69: 11–25, 1992.
72. Imagawa, S., T. Izumi, and Y. Miura. Positive and negative
regulation of the erythropoietin gene. J. Biol. Chem. 269:
9038–9042, 1994.
73. Imagawa, S., M. Yamamoto, and Y. Miura. GATA transcription factors negatively regulate erythropoietin gene expression.
Acta Haematol. 95: 248–256, 1996.
74. Jacobs, K., C. Shoemaker, R. Rudersdorf, S. D. Neill, R. J.
Kauffman, A. Mufson, J. Seehra, S. S. Jones, R. Hewick,
E. F. Fritsch, M. Kawakite, T. Shimizu, and T. Miyake.
Isolation and characterization of genomic and cDNA clones of
human erythropoietin. Nature 313: 806–810, 1985.
75. Jacobson, L. O., E. Goldwasser, W. Fried, and L. Plzak.
Role of the kidney in erythropoiesis. Nature 179: 633–634,
1957.
76. Jacquot, R., and J. Nagel. Glucocorticoids and evolution of
haemopoietic tissue in the liver of the rat foetus. Ann. Immunol.
(Inst. Pasteur): 127 C: 895–903, 1976.
77. Jelkmann, W. Erythropoietin: structure, control of production,
and function. Physiol. Rev. 72: 449–489, 1992.
78. Jelkmann, W. Biology of erythropoietin. Clin. Invest. Med. 72:
S3–S10, 1994.
79. Jiang, N., T. C. He, A. Miyajima, and D. M. Wojchowski.
The box1 domain of the erythropoietin receptor specifies janus
kinase 2 activation and functions mitogenically within an
interleukin 2 b-receptor chimera. J. Biol. Chem. 271: 16472–
16476, 1996.
R1841
R1842
100.
101.
102.
103.
104.
106.
107.
108.
109.
110.
111.
112.
113.
114.
115.
116.
117.
118.
son with the human erythropoietin gene. Gene 44: 201–209,
1986.
Liu, Z. Y., K. Chin, and C. T. Noguchi. Tissue specific
expression of human erythropoietin receptor in transgenic
mice. Dev. Biol. 166: 158–169, 1994.
Livnah, O., E. A. Stura, D. L. Johnson, S. A. Middleton,
L. S. Mulcahy, N. C. Wrighton, W. J. Dower, L. K. Jolliffe,
and I. A. Wilson. Functional mimicry of a protein hormone by a
peptide agonist: the Epo receptor complex at 2.8 Å. Science 273:
464–471, 1996.
Lok, S., K. Kaushansky, R. D. Holly, J. L. Kuijper, C. E.
Lofton-Day, P. J. Oort, F. J. Grant, M. D. Heipel, S. K.
Burkhead, and J. M. Kramer. Cloning and expression of
murine thrombopoietin cDNA and stimulation of platelet production in vivo. Nature 369: 565–568, 1994.
Longmore, G. D. Erythropoietin receptor mutations and olympic glory. Nat. Genet. 4: 108–110, 1993.
Longmore, G. D., S. S. Watowich, D. J. Hilton, and H. F.
Lodish. The erythropoietin receptor: its role in hematopoiesis
and myeloproliferative diseases. J. Cell Biol. 123: 1305–1308,
1993.
Maier, R. F., M. Obladen, P. Scigalla, O. Linderkamp, G.
Duc., and G. Hieronimi. The effect of epoetin beta (recombinant human erythropoietin) on the need for transfusion in
very-low-birth-weight infants. N. Engl. J. Med. 330: 1173–1178,
1994
Mallard, E. C., C. E. Williams, A. J. Gunn, M. I. Gunning,
and P. D. Gluckman. Frequent episodes of brief ischemia
sensitize the fetal sheep brain to neuronal loss and induce
striatal injury. Pediatr. Res. 33: 61–65, 1993.
Manavalan, P., D. L. Swope, and R. M. Withy. Sequence and
structural relationships in the cytokine family. J. Protein Chem.
11: 321–331, 1992.
Marti, H. H., R. H. Wenger, L. Rivas, U. Straumann, M.
Digicaylioglu, V. Henn, Y. Yonekawa, C. Bauer, and M.
Gassman. Erythropoietin gene expression in human, monkey
and murine brain. Eur. J. Neurosci. 8: 666–676, 1996.
Martin, D. I. K., and S. H. Orkin. The transcriptional
activation and DNA binding by the erythroid factor GF-1/NF-E1/
Eryf 1. Genes Dev. 4: 1886–1890, 1990.
Masuda, S., M. Nagao, K. Takahata, Y. Konishi, F. Gallyas,
Jr., T. Tabira, and R. Sasaki. Functional erythropoietin
receptor of the cells with neural characteristics. J. Biol. Chem.
268: 11208–11216, 1993.
Masuda, S., M. Okano, K. Yamagishi, M. Nagao, M. Ueda,
and R. Sasaki. A novel site of erythropoietin production. J.
Biol. Chem. 269: 19488–19493, 1994.
Maxwell, P. H., D. J. P. Ferguson, M. K. Osmond, C. W.
Pugh, A. Heryet, B. G. Doe, M. H. Johnson, and P. J.
Ratcliffe. Expression of a homologously recombined erythropoietin-SV-40 T antigen fusion gene in mouse liver: evidence for
erythropoietin production by Ito cells. Blood 84: 1823–1830,
1994.
Mayeux, P., C. Billat, J. M. Felix, and R. Jacquot. Evidence
for glucocorticosteroid receptors in the erythroid cell line of fetal
rat liver. J. Endocrinol. 96: 311–319, 1983.
Medvinsky, A., and E. Dzierzak. Definitive hematopoiesis is
autonomously initiated by the AGM region. Cell 86: 897–906,
1996.
Meunier, H., C. Rivier, R. M. Evans, and W. Vale. Gonadal
and extragonadal expression of inhibin a, bA, and bB subunits in
various tissues predicts diverse functions. Proc. Natl. Acad. Sci.
USA 85: 246–251, 1988.
Moeller, I., R. A. Lew, F. A. O. Mendelsohn, A. I. Smith,
M. E. Brennan, T. J. Tetaz, and S. Y. Chai. The globin
fragment, LVV-haemorphin-7, is an endogenous ligand for the
AT4 receptor in the brain. J. Neurochem. 68: 2530–2537, 1997.
Moeller, I., D. H. Small, G. Reed, J. W. Harding, and F. A. O.
Mendelsohn. Angiotensin IV inhibits neurite outgrowth in
cultured embryonic chicken sympathetic neurones. Brain Res.
725: 61–66, 1996.
Moritz, K., E. Cooper, and E. M. Wintour. The effect of
haemorrhage on erythropoietin concentration in the mature
ovine fetus. J. Dev. Physiol. (Eynsham) 17: 157–161, 1992.
119. Moritz, K. M., P. C. Owens, and E. M. Wintour. Changes in
blood and red cell volume in the neonatal lamb and the effect of
insulin-like growth factor I. Clin. Exp. Pharmacol. Physiol. 23:
134–139, 1996.
120. Moritz, K. M., and E. M. Wintour. The effect of graded
haemorrhage on erythropoietin production in the immature
ovine fetus. Clin. Exp. Pharmacol. Physiol. 19: 503–508, 1992.
121. Nakamura, Y., N. Komatsu, and H. Nakauchi. A truncated
erythropoietin receptor that fails to prevent programmed cell
death of erythroid cells. Science 257: 1138–1141, 1992.
122. Nakamura, Y., and H. Nakauchi. A truncated erythropoietin
receptor and cell death: a reanalysis. Science 264: 588–589,
1994.
123. Nakano, T., H. Kodama, and T. Honjo. In vitro development
of primitive and definitive erythrocytes from different precursors. Science 272: 722–724, 1996.
124. Ogawa, M. Differentiation and proliferation of hematopoietic
stem cells. Blood 81: 2844–2853, 1993.
125. Ohls, R. K. The erythropoietin gene is expressed in midtrimester human kidney (Abstract). Blood 88, Suppl. 1: 566,
1996.
126. Ohyagi, Y., T. Yamada, and I. Goto. Hemoglobin as a novel
protein developmentally regulated in neurons. Brain Res. 635:
323–327, 1994.
127. Pandolfi, P. P., M. E. Roth, A. Karis, M. W. Leonard, E.
Dzierzak, F. G. Grosveld, J. D. Engel, and M. H. Lindenbaum. Targeted disruption of the GATA3 gene causes severe
abnormalities in the nervous system and in fetal liver haematopoiesis. Nat. Genet. 11: 40–44, 1995.
128. Papayannopoulou, T., M. Brice, D. Farrer, and K. Kaushansky. Insights into the cellular mechanisms of erythropoietinthrombopoietin synergy. Exp. Hematol. 24: 660–669, 1996.
129. Penta, K., and S. T. Sawyer. Erythopoietin induces the
tyrosine phosphorylation, nuclear translocation, and DNA binding of STAT1 and STAT5 in erythroid cells. J. Biol. Chem. 270:
31282–31287, 1995.
130. Peschle, A. Human ontogenic development: studies on the
hemopoietic system and the expression of homeo box genes.
Ann. NY Acad. Sci. 511: 101–116, 1987.
131. Pevny, L., C. S. Lin, V. D’Agati, C. M. Simon, S. H. Orkin,
and F. Constantini. Development of hematopoietic cells lacking transcription factor GATA-1. Development 121: 163–172,
1996.
132. Pevny, L., M. C. Simon, E. Robertson, W. H. Klein, S. H.
Tsai, V. D’Agati, S. H. Orkin, and F. Constantini. Erythroid
differentiation in chimaeric mice blocked by a targeted mutation in the gene for transcription factor GATA-I. Nature 349:
257–260, 1991.
133. Piroso, E., A. J. Erslev, K. K. Flaharty, and J. Caro.
Erythropoietin life span in rats with hypoplastic and hyperplastic bone marrows. Am. J. Hematol. 36: 105–110, 1991.
134. Potocnik, S. J., and E. M. Wintour. Development of the
spleen as a red blood cell reservoir in lambs. Reprod. Fertil. Dev.
8: 311–315, 1996.
135. Pugh, C. W., B. L. Ebert, O. Ebrahim, and P. J. Ratcliffe.
Characterisation of functional domains within the mouse erythropoietin 3’ enhancer conveying oxygen-regulated responses in
different cell lines. Biochim. Biophys. Acta 1217: 297–306,
1994.
136. Raich, N., C. H. Clegg, J. Grofti, P.-H. Roméo, and G.
Stamatoyannopoulos. GATA1 and YY1 are developmental
repressors of the human I-globin gene. EMBO J. 14: 801–809,
1995.
137. Ratcliffe, J. G., B. L. Ebert, J. D. Firth, J. M. Gleadle, P. H.
Maxwell, M. Nagao, J. F. O’Rourke, C. W. Pugh, and S. M.
Wood. Oxygen regulated gene expression: erythropoietin as a
model system. Kidney Int. 51: 514–526, 1997.
138. Ratcliffe, P. J. Molecular biology of erythropoietin. Kidney Int.
44: 887–904, 1993.
139. Ruud, J. T. Vertebrates without erythrocytes and blood pigment. Nature 173: 848–850, 1954.
140. Sawyer, S. T., S. B. Krantz, and K. Sawada. Receptors for
erythropoietin in mouse and human erythroid cells and placenta. Blood 74: 103–109, 1989.
Downloaded from http://ajpregu.physiology.org/ by 10.220.33.6 on June 18, 2017
105.
INVITED REVIEW
INVITED REVIEW
159. Weiss, M. J., G. Keller, and S. H. Orkin. Novel insights into
erythroid development revealed through in vitro differentiation
of GATA-12 embryonic stem cells. Genes Dev. 8: 1184–1197,
1994.
160. Wells, J. A. Hormone mimicry. Science 273: 449–450, 1996.
161. Wells, J. A., and A. M. de Vos. Hematopoietic receptor
complexes. Annu. Rev. Biochem. 65: 609–634, 1996.
162. Wen, D., J.-P. R. Boissel, T. E. Tracy, R. H. Gruninger, L. S.
Mulcahy, J. Czelusniak, M. Goodman, and H. F. Bunn.
Erythropoietin structure-function relationships: high degree of
sequence homology among mammals. Blood 82: 1507–1516,
1993.
163. Whitelaw, E., S.-F. Tsai, P. Hogben, and S. Orkin. Regulated
expression of globin chains and the erythroid transcription
factor GATA-1 during erythropoiesis in the developing mouse.
Mol. Cell. Biol. 10: 6596–6601, 1990.
164. Wickramasinghe, S. N. Erythropoietin and the human kidney: evidence for an evolutionary link from studies of Salmo
Gairdneri. Comp. Biochem. Physiol. A Physiol. 104A: 63–65,
1993.
165. Widness, J. A., G. K. Clemons, J. F. Garcia, W. Oh, and R.
Schwartz. Increased immunoreactive erythropoietin in cord
serum after labor (Abstract). Am. J. Obstet. Gynecol. 148: 194,
1984.
166. Widness, J. A., C. K. Clemons, and A. F. Phillips. Erythropoieten levels and erythropoiesis at birth in infants with Potter’s
Syndrome. J. Pediatr. 117: 155–158, 1990.
167. Widness, J. A., T. A. Malone, and R. A. Mufson. Impermeability of the ovine placenta to 35S-recombinant erythropoietin.
Pediatr. Res. 25: 649–651, 1989.
168. Widness, J. A., S. T. Sawyer, R. L. Schmidt, and D. H.
Chestnut. Lack of maternal to fetal transfer of 125I-labeled
erythropoietin in sheep. J. Dev. Physiol. (Eynsham) 15: 139–
143, 1991.
169. Widness, J. A., J. B. Susa, J. F. Garcia, D. B. Singer, P.
Sehgal, W. Oh, R. Schwartz, and H. C. Schwartz. Increased
erythropoiesis and elevated erythropoietin in infants born to
diabetic mothers and in hyperinsulinemic rhesus fetuses. J.
Clin. Invest. 67: 637–642, 1981.
170. Widness, J. A., P. Veng-Pedersen, N. B. Modi, R. L. Schmidt,
and D. H. Chestnut. Developmental differences in erythropoietin pharmacokinetics: increased clearance and distribution in
fetal and neonatal sheep. J. Pharmacol. Exp. Ther. 261: 977–
984, 1992.
171. Widness, J. A., P. Veng-Pedersen, C. Peters, L. Periera,
R. L. Schmidt, and L. S. Lowe. Erythropoietin pharmacokinetics in premature infants: developmental, nonlinearity, and
treatment effects. J. Appl. Physiol. 80: 140–148, 1996.
172. Widness, J. A., P. Veng-Pedersen, R. L. Schmidt, and C.
Peters. In vivo 125I-erythropoietin pharmacokinetics are unchanged following anesthesia, nephrectomy and hepatectomy
in sheep. J. Pharmacol. Exp. Ther. In press.
173. Wilimas, J. A., and W. M. Gist. Erythropoietin not yet a
standard treatment for anemia of prematurity. Pediatrics 95:
9–10, 1995.
174. Wintour, E.M., A. Butkus, L. Earnest, and S. Pompolo. The
erythropoietin gene is expressed strongly in the mammalian
mesonephric kidney. Blood 88: 3349–3353, 1996.
175. Wintour, E. M., A. Butkus, G. Clemons, and K. Moritz.
Erythropoiesis and haemoglobin switching in the fetus and
neonate. Proc. Aust. Physiol. Pharmacol. Soc. 22: 44–52, 1991.
176. Wintour, E. M., G. K. Clemons, A. Butkus, A. Horvath, K.
Moritz, and M. K. Towstoless. Immunoreactive erythropoietin concentration and haemoglobin type in the perinatal period
in sheep of various haemoglobin genotypes. J. Dev. Physiol.
(Eynsham) (14: 259–265, 1990.
177. Witthuhn, B. A., F. W. Quelle, O. Silvennoinen, T. Yi, B.
Tang, O. Miura, and J. N. Ihle. JAK2 associates with the
erythropoietin receptor and is tyrosine phosphorylated and
activated following stimulation with erythropoietin. Cell 74:
227–236, 1993.
178. Wood, W. G. Haemoglobin synthesis during human fetal development. Br. Med. Bull. 32: 282–287, 1976.
Downloaded from http://ajpregu.physiology.org/ by 10.220.33.6 on June 18, 2017
141. Schimmenti, L. A., G. Blechert, K. W. Harris, and J. C.
Winkelmann. Localization of an essential ligand binding determinant of the human erythropoietin receptor to a domain
N-terminal to the WSXWS motif: implications for soluble
receptor function. Exp. Hematol. 23: 1341–1346, 1995.
142. Schmitt, R. M., E. Bruyns, and H. R. Snodgrass. Hematopoietic development of embryonic stem cells in vitro: cytokine and
receptor gene expression. Genes Dev. 5: 728–740, 1991.
143. Schuster, S., S. Koury, M. Bohrer, S. Salceda, and J. Caro.
Cellular sites of extrarenal and renal erythropoietin production
in anemic rats. Br. J. Haematol. 81: 153–159, 1992.
144. Semenza, G. L., F. Agani, G. Booth, J. Forsythe, N. Iyer,
B.-H. Jiang, S. Leung, R. Roe, C. Wiener, and A. Yu.
Structural and functional analysis of hypoxia-inducible factor 1.
Kidney Intern. Med. 51: 553–555, 1997.
145. Semenza, G. L., M. D. Traystman, J. D. Gearhart, and S.
Antonarakis. Polycythemia in transgenic mice expressing the
human erythropoietin gene. Proc. Natl. Acad. Sci. USA 86:
2301–2305, 1989.
146. Semenza, G. L., and G. L. Wang. A nuclear factor induced by
hypoxia via de novo protein synthesis binds to the human
erythropoietin gene enhancer at a site required for transcriptional activation. Mol. Cell. Biol. 12: 5447–5454, 1992.
147. Shalaby, F., J. Rossant, T. P. Yamaguchi, M. Gertsenstein,
X. F. Wu, M. L. Breitman, and A. C. Schuh. Failure of
blood-island formation and vasculogenesis in Flk-1 deficient
mice. Nature 376: 62–66, 1995.
148. Shannon, K. M., W. C. Mentzer, R. I. Abels, P. Freeman, N.
Newton, D. Thompson, S. Sniderman, R. Ballard, and
R. H. Phibbs. Recombinant human erythropoietin in the
anemia of prematurity: results of a placebo-controlled pilot
study. J. Pediatr. 118: 949–955, 1991.
149. Shireman, T. I., P. E. Hilsenrath, R. G. Strauss, J. A.
Widness, and A. H. Mutnick. Recombinant human erythropoietin vs. transfusions in the treatment of anemia of prematurity.
A cost of benefit analysis. Arch. Pediatr. Adolesc. Med. 148:
582–588, 1994.
150. Sidell, B. D., M. E. Vayda, D. J. Small, T. J. Moylan, R. L.
Londraville, M.-L. Yuan, K. J. Rodnick, Z. A. Eppley, and
L. Costello. Variable expression of myoglobin among the
hemoglobinless Antarctic icefishes. Proc. Natl. Acad. Sci. USA
94: 3420–3424, 1997.
151. Snijders, R. J. M., A. Abbas, O. Melby, R. M. Ireland, and
K. H. Nicolaides. Fetal plasma erythropoietin concentration in
severe growth retardation (Abstract). Am. J. Obstet. Gynecol.
168: 615, 1993.
152. Stamatoyannopoulos, G., P. Constantoulakis, M. Brice, S.
Kurachi, and T. Papayannopoulou. Coexpression of embryonic, fetal, and adult globins in erythroid cells of human
embryos: relevance to the cell-lineage models of globin switching. Dev. Biol. 123: 191–197, 1987.
153. Stockman, J. A., and F. A. Oski. Physiological anaemia of
infancy and the anemia of prematurity. Clin. Hemorheol. 7:
3–18, 1978.
154. Tan, C. C., K. Eckardt, J. D. Firth, and P. J. Ratcliffe.
Feedback modulation of renal and hepatic erythropoietin mRNA
in response to graded anemia and hypoxia. Am. J. Physiol. 263
(Renal Fluid Electrolyte Physiol. 32): F474–F481, 1992.
155. Thomas, R. M., C. E. Canning, P. M. Cotes, D. C. Linch,
C. H. Rodeck, C. E. Rossiter, and E. R. Huehns. Erythropoietin and cord blood haemoglobin in the regulation of human
fetal erythropoiesis. J. Obstet. Gynaecol. Res. 90: 795–800,
1983.
156. Tsai, S. F., D. I. Martin, L. I. Zon, A. D. D’Andrea, G. G.
Wong, and S. H. Orkin. Cloning of cDNA for the major
DNA-binding protein of the erythroid lineage through expression in mammalian cells. Nature 339: 446–451, 1989.
157. Wang, G. L., B.-H. Jiang, E. A. Rue, and G. L. Semenza.
Hypoxia-inducible factor 1 is a basic-helix-loop-helix-PAS heterodimer regulated by cellular O2 tension. Proc. Natl. Acad. Sci.
USA 92: 5510–5514, 1995.
158. Wang, G. L., and G. L. Semenza. Purification and characterization of hypoxia-inducible factor 1. J. Biol. Chem. 270: 1230–
1237, 1995.
R1843
R1844
INVITED REVIEW
179. Wrighton, N. C., F. X. Farrell, R. Chang, A. K. Kashyap,
F. P. Barbone, L. S. Mulcahy, D. L. Johnson, R. W. Barrett,
L. K. Jolliffe, and W. J. Dower. Small peptides as potent
mimetics of the protein hormone erythropoietin. Science 273:
458–463, 1996.
180. Wu, H., X. Liu, R. Jaenisch, and H. F. Lodish. Generation of
committed erythroid BFU-E and CFU-E progenitors does not
require erythropoietin or the erythropoietin receptor. Cell 83:
59–67, 1995.
181. Yamaji, R., T. Okada, M. Moriya, M. Naito, T. Tsuruo, K.
Miyatake, and Y. Nakano. Brain capillary endothelial cells
express two forms of erythropoietin receptor mRNA. Eur. J.
Biochem. 239: 494–500, 1996.
182. Yasuda, Y., M. Nagao, M. Okano, S. Masuda, R. Sasaki, H.
Konishi, and T. Tanimura. Localization of erythropoietin and
erythropoietin-receptor in postimplantation mouse embryos.
Dev. Growth Diff. 35: 711–722, 1993.
183. Yoshimura, A., G. Longmore, and H. F. Lodish. Point
mutation in the exoplasmic domain of the erythropoietin receptor resulting in hormone-independent activation and tumorigenicity. Nature 348: 647–649, 1990.
184. Youssoufian, H., G. Longmore, D. Neumann, A. Yoshimura,
and H. F. Lodish. Structure, function, and activation of the
erythropoietin receptor. Blood 81: 2223–2236, 1993.
185. Yu, J., L.-E. Shao, V. Lemas, A. L. Yu, J. Vaughan, J. Rivier,
and W. Vale. Importance of FSH-releasing protein and inhibin
in erythrodifferentiation. Nature 330: 765–767, 1981.
186. Zanjani, E. D., and J. L. Ascensao. Erythropoietin. Transfusion 29: 47–57, 1989.
187. Zimmermann, F., and I. N. Rich. The sensitivity of in vitro
erythropoietic progenitor cells to different erythropoietin reagents during development and the role of cell death in culture.
Exp. Hematol. 24: 330–339, 1996.
Downloaded from http://ajpregu.physiology.org/ by 10.220.33.6 on June 18, 2017