The oviduct produces erythropoietin in an estrogen- and
oxygen-dependent manner
Seiji
Masuda,
Toshihiro
Kobayashi,
Mariko
Chikuma,
Masaya
Nagao, and
Ryuzo
Sasaki
Division of Integrated Life Science, Graduate School of
Biostudies, Kyoto University, Kyoto 606-8502, Japan
 |
ABSTRACT |
Previously, we showed that
erythropoietin (Epo) is produced in the mouse uterus, where Epo is
indispensable for estrogen (E2)-dependent angiogenesis.
Expression of uterine Epo mRNA is stimulated by E2 and
hypoxia. The hypoxic induction requires the presence of E2.
In the present study, we examined other female reproductive organs in
the mouse with respect to Epo mRNA expression and its stimuli
(E2 and hypoxia)-induced changes. Although Epo mRNA
expression was seen in the ovary and oviduct, the
E2-induced stimulation of Epo mRNA was found only in the
oviduct. The E2-induced stimulation in the oviduct was
transient and rapidly downregulated. Epo mRNA expression in the oviduct
was hypoxia inducible, in both the presence and the absence of
E2. E2-dependent production of Epo and its mRNA
expression were also found by use of cultured oviducts. The E2 action is probably mediated through the E2
receptor, and de novo protein synthesis is not required for
E2 induction of Epo mRNA. In the oviduct, the ampulla and
isthmus regions produce Epo.
hypoxia; ovary; isthmus; tamoxifen; ICI-182780
 |
INTRODUCTION |
A WELL-RECOGNIZED FUNCTION of erythropoietin (Epo) is
to increase the production of red blood cells by preventing apoptotic death of Epo-responsive erythroid precursor cells and by stimulating their proliferation and differentiation (reviewed in Refs. 18, 23, and
50). The fetal liver is the site of Epo production, which is essential
for fetal erythropoiesis (24, 46). The kidney is the major site of Epo
production in adults, and the kidney-derived Epo is responsible for the
stimulation of erythropoiesis in adults (reviewed in Refs. 18 and 23).
Epo production in these sites is stimulated under hypoxia, mainly
through the activation of Epo gene expression and partly through
stabilization of mRNA (reviewed in Refs. 5, 14, 36, 37, 40, 45).
The hypoxic activation of Epo gene expression is caused by binding of
the hypoxia-inducible factor-1 (HIF-1) to the hypoxia-responsive enhancer that lies in a region 120 bp 3' to the polyadenylation site.
HIF-1 is a heterodimer consisting of the subunit HIF-1
and the aryl
hydrocarbon receptor nuclear translocator (ARNT), both of which are
basic-helix-loop-helix proteins in the PAS (Per-AHR-ARNT-SIM) family of
transcription factors. Although ARNT, which is present in high amounts,
is not affected by oxygen levels, HIF-1
levels are very low under
normoxia, because HIF-1
is rapidly degraded via the
ubiquitin-proteasome pathway. Interaction of HIF-1
and the von
hippel-Lindau protein appears to be necessary for the degradation of
HIF-1
(31). Under hypoxia, HIF-1
is stabilized by an unknown
mechanism to form the active heterodimer with ARNT. Thus HIF-1
activation correlates well with the hypoxia-induced accumulation of the
HIF-1
subunit.
In addition to liver and kidney, recently two sites (brain and uterus)
have been shown to produce Epo with new physiological functions.
Neurons express the Epo receptor (EpoR) (8, 29, 32), and astrocytes
produce Epo (26, 27, 30). Thus the central nervous system has a
paracrine Epo/EpoR system, which is independent of the erythropoietic
system (3, 8, 25-30, 32). Epo infusion into the brain prevents
ischemia-induced death of cerebrocortical and hippocampal
neurons (38, 39). Evidence that endogenous brain Epo plays a critical
role in neuron survival under brain ischemia has been presented
(39). Consistent with the view that glutamate toxicity is a major cause
of ischemia-induced neuron death, Epo has been shown to protect
primary cultured cerebrocortical and hippocampal neurons from glutamate
toxicity (32). Low oxygen tension elevates Epo mRNA in the brain (43)
and enhances Epo production by the cultured astrocytes (26, 30), which
may be appropriate for the neuroprotective function of Epo
in the ischemic brain.
EpoR mRNA is expressed in endothelial cells from human umbilical vein,
bovine adrenal capillary, and rat brain capillary (2, 47). The
angiogenic activity of Epo was shown by the use of in vitro cultured
endothelial cells (1, 6, 15), but the in vivo significance of these
findings was not demonstrated. In healthy adults, blood vessel
formation is repressed, but an exception is the female reproductive
organ, where the active angiogenesis cyclically takes place for
remodeling of destroyed tissues. We have shown that there is another
paracrine Epo/EpoR system in the uterus and that Epo plays an important
role in the uterine angiogenesis via EpoR expressed in vascular
endothelial cells of the uterine endometrium (28, 49). Furthermore, Epo
production in the uterine tissue is stimulated by 17
-estradiol (or
estrogen, E2), an ovarian hormone. Because oxygen
concentration was thought to be a major regulator of Epo production,
E2 stimulation of uterine Epo production was surprising,
but it provided relevance of Epo function in an
E2-dependent cyclical angiogenesis in the uterus (49).
Encouraged by these findings, we explored the expression of Epo mRNA
and the production of Epo in other female reproductive organs (ovary
and oviduct). Here we report changes of Epo mRNA contents in the mouse
oviduct upon E2 administration and/or exposure to a hypoxic
condition. The production of Epo protein and expression of Epo mRNA by
in vitro cultured oviducts were also examined.
 |
MATERIALS AND METHODS |
Animals.
Animals were maintained and handled in accordance with the guidelines
for the care and use of laboratory animals at Kyoto University. Outbred
mice of the ICR strain (Clea) were used for experiments at 3 wk of age.
E2 administration and/or hypoxic exposure.
To examine the effect of E2 administration and/or hypoxic
exposure on Epo mRNA levels in the ovary, oviduct, and uterus, we divided the mice into three groups. The animals in the first group received intraperitoneal administration of E2 and were left
under normoxia, those in the second group were given olive oil and
exposed to normobaric hypoxia (7% O2-93% N2),
and those in the third group were exposed to hypoxia immediately after
receiving E2 administration. E2 (from Research
Biochemicals International) was dissolved in olive oil. Mice were given
100 µl of E2 solution per animal (0.5 µg
E2/g body wt) by intraperitoneal injection. Olive oil was
given to control mice. Hypoxic stimulation was achieved by use of an air-tight cabinet into which a premixed gas was introduced. The gas
flow rate was adjusted so that 7% O2 was achieved ~30
min after the animals were placed into the cabinet. At different time points after E2 administration and/or hypoxic exposure, the
animals were anesthetized with ether, and then the tissues were quickly removed and frozen in liquid nitrogen until used for RNA extraction.
Standard cDNA fragments of Epo and
-actin.
Sequence coordinates of mouse Epo cDNA are based on the definition of
the transcription start site as +1 (42). A 451-bp fragment encompassing
272-722 of the mouse Epo cDNA was ligated into a vector pCR3.1-Uni
by use of the Eukaryotic TA Cloning Kit (Invitrogen). The resulting
plasmid was used as a standard Epo cDNA for PCR. As a standard
-actin cDNA, pAL41 (accession no. X03765) was used. These standard
cDNA fragments cover the 112-bp (Epo) and 261-bp (
-actin) nucleotide
sequences, which are identical to those that are amplified from reverse
transcription (RT)-derived cDNAs by use of the primers described in the
next two sections.
RT.
Total RNA was prepared from the frozen tissues according to the
protocol of the RNA Isolation System kit (Promega). RT was carried out
at 45°C for 60 min in 20 µl of RT mixture containing 1 µg total
RNA, 200 U SuperScript II (GIBCO BRL), 20 U RNase inhibitor (Takara),
0.5 mM of each dNTP, and 2.5 µM random nonamer primer. One microliter of cDNA product was used for real-time PCR.
Real-time PCR.
The PCR product of Epo mRNA-derived cDNA was quantified on real time,
which is accomplished by using a double dye-labeled fluorogenic
oligonucleotide probe (16) and an automated fluorescence-based system
for detection of PCR products. The probe is labeled at its 5' end with
a fluorogenic reporter dye, 6-carboxy-fluorescein (FAM), and at its 3'
end with a quencher dye, 6-carboxy-tetramethylrhodamine (TAMRA). The
nucleotide sequence in the probe
5'-(FAM)-TGCAGAAGGTCCCAGACTGAGTGAAAATA-3'-(TAMRA) corresponds to
fragments 397-425 in the mouse Epo cDNA. The Epo-specific sequences used for PCR were the forward primer 371F,
5'-GAGGCAGAAAATGTCACGATG-3', and the reverse primer 482R,
5'-CTTCCACCTCCATTCTTTTCC-3'. The forward and reverse primers
correspond to the nucleotides 371-391 and 462-482 in Epo
cDNA, respectively. They span exon/intron boundaries (exons II and III
in the 371F, and exons III and IV in the 482R); thus amplification of
contaminating genomic DNA is prevented. For PCR, we used TaqMan
Universal PCR Master Mix containing dUTP instead of dTTP (PE Applied
Biosystems, cat. no. 4304447). This master mixture also contains
uracil-N-glycosylase (UNG), which destroys any carryover PCR
product that might remain in PCR chambers. Before PCR was started, the
complete PCR mixture containing reverse-transcribed cDNA was treated in
a PCR chamber at 50°C for 2 min for the action of UNG, and then
treated at 95°C for 10 min, resulting in the inactivation of UNG
and activation of DNA polymerase. Then PCR, consisting of 50 cycles at
95°C for 15 s and at 60°C for 1 min, was performed. When DNA
polymerase engaged in extension of the primer reaches the
quencher-labeled nucleotide of the probe hybridized to cDNA, the
exonuclease activity of DNA polymerase excises the labeled nucleotide,
resulting in the emission of fluorescence. All procedures including
data analysis were performed on the ABI PRISM 7700 Sequence Detection
System (PE Applied Biosystems) using the software provided with the
instrument. This assay format allows real-time kinetic analysis of PCR
product generation, providing a broad linear dynamic range and ensuring
that quantification is based on analysis during the exponential
amplification. Messenger RNA of
-actin was also measured, and its
level was used as an internal control for normalization of the Epo mRNA
level. The
-actin sequence was amplified and detected using the
primers 5'-CTAGGCACCAAGGTGTGAT-3' and
5'-CAAACATGATCTGGGTCATC-3' and the probe
5'-(FAM)-TGGCACCACACCTTCTACAATGAG-3'-(TAMRA). When samples lacking RNA or reverse transcriptase in the reverse transcriptase reactions were subjected to PCR, each of these control reactions yielded no detectable fluorescence, excluding the possibility that the
contaminating DNA, including the RNA preparation-derived genomic DNA,
was amplified.
Copy numbers of Epo or
-actin mRNA-derived cDNA were calculated from
the standard curves of PCR drawn by the use of the standard Epo or
-actin cDNA fragments. The lower limit of quantitative detection by
the present method was 30 copies of cDNA.
Culture of the oviduct.
The oviducts were removed from 3-wk-old mice. The unilateral oviduct
was cultured in a phenol red-free DMEM supplemented with 10%
charcoal-treated fetal calf serum containing the test substance, and
the contralateral oviduct was cultured in the same medium but without
the test substance, as a control. These media were incubated in a humid
5% CO2 atmosphere at 37°C. Epo protein secreted in the
culture media was measured by a sandwich-type enzyme-linked immunoassay
(EIA) by use of two monoclonal antibodies that bind Epo at different
epitopes (33), and the tissue was used for total RNA preparation.
Recombinant human Epo, produced and isolated as described previously
(12, 13), was used as a standard. This assay measures Epo at a
concentration as low as 1 pg/ml.
To identify the oviductal Epo production site, the infundibulum,
ampulla, and isthmus were removed from the oviduct under a microscope
and cultured in the presence or absence of E2 for 8 h. Epo
protein secreted in the culture media was measured by EIA.
 |
RESULTS |
In vivo induction of Epo mRNA in female reproductive organs by
E2 and hypoxia.
Previously (unpublished observations), we showed with
ovariectomized mice that Epo mRNA in the uterus was E2
inducible but not hypoxia inducible in the absence of E2.
In the present experiments, to avoid the effects of the internal
E2, we used 3-wk-old mice before commencement of cyclical
synthesis of E2 in the ovaries. E2 was given
intraperitoneally, and then the ovary and oviduct were removed to
measure Epo mRNA at 1 h after E2 injection. Figure 1 shows the results. Epo mRNA was detected
in both the ovary and oviduct from E2-uninjected animals,
and the Epo mRNA level in the oviduct was threefold greater than that
in the ovary. Furthermore, Epo mRNA in the oviduct was highly
E2 inducible (4- to 5-fold induction), whereas that in the
ovary slightly increased upon E2 injection. Therefore, we
examined the Epo expression in the oviduct.

View larger version (12K):
[in this window]
[in a new window]
|
Fig. 1.
In vivo effect of estradiol (E2) on expression of
erythropoietin (Epo) mRNA in ovary and oviduct. Animals were
administered 0.5 µg E2/g body wt or olive oil. At 1 h
after injection, ovary and oviduct were removed for quantitative
measurement of Epo and -actin mRNAs by real-time RT-PCR (see
MATERIALS AND METHODS). The ordinate indicates multiple
increases in induction of Epo mRNA over control levels (olive oil),
which were 29,600 copies/2 oviducts and 11,700 copies/2 ovaries, and
defined as 1. Bars are means ± SE (n = 3). Statistical
significance of differences was determined with Student's
t-test. * P < 0.05, significantly different from
values of controls (values in the absence of E2).
|
|
To examine the effects of E2 and hypoxia on Epo mRNA levels
in the oviduct, mice were given E2 or the vehicle and
immediately exposed to hypoxia (7% O2) or kept under
normoxia. At different time points the oviducts were removed to measure
Epo mRNA. Figure 2A shows the
results. When animals received E2 and were kept under normoxia, Epo mRNA was increased eightfold at 2 h and decreased at 4 h.
When animals were exposed to hypoxia without E2 injection, a significant induction was seen at 1 and 2 h. Administration of
E2 potentiated the induction of Epo mRNA by hypoxia. The
oviduct of animals that received both stimuli showed the highest
induction (30-fold) of Epo mRNA at 4 h, but the level was markedly
decreased at 8 h.

View larger version (14K):
[in this window]
[in a new window]
|
Fig. 2.
Temporal patterns of induction of Epo mRNA in oviduct (A) and
uterus (B) by E2 and/or hypoxia. , Animals were
exposed to hypoxia (7% O2) immediately after
administration of 0.5 µg E2/g body weight
(E2/Hx). , Animals were exposed to hypoxia after
administration of olive oil ( E2/Hx). , Animals
were left under normoxia after administration of 0.5 µg
E2/g body wt (E2/Nx). At indicated time points,
tissues were removed for measurement of Epo mRNA. The ordinate shows
multiples of increase in (fold) induction over control level (at
time 0). Bars are means ± SE (n = 4). Statistical
significance of differences was determined with ANOVA followed by
Dunnett's test. * P < 0.05, ** P < 0.01, significantly different from values of controls (time 0),
respectively.
|
|
For comparison, Epo mRNA in the uterus was also assayed (Fig.
2B). There was a marked increase under normoxia at 2 h after E2 injection, but at 4 h the level markedly decreased.
Without E2 injection, the uterine Epo mRNA level was
unchanged despite the continuous hypoxia. However, uterine Epo mRNA
became hypoxia inducible in the presence of E2. These
results, obtained by the use of prepuberal young mice, were similar to
those obtained by the use of ovariectomized adult animals (unpublished
observations). A remarkable difference between uterus and oviduct was
that the oviductal Epo mRNA of E2-uninjected animals is
hypoxia inducible, whereas the uterus does not respond to hypoxia
unless E2 is given.
Induction of Epo mRNA and production of Epo by cultured oviducts.
To examine Epo production in the in vitro cultured oviducts and its
induction by E2, the unilateral oviduct was cultured with E2 for 8 h. The contralateral oviduct was cultured without
E2 as a control. Epo protein was detectable in the medium
after culture without E2, and E2 stimulated Epo
production in a dose-dependent manner (Fig.
3A). Figure 3B shows the
production of Epo with culture over time.

View larger version (10K):
[in this window]
[in a new window]
|
Fig. 3.
E2-dependent production of Epo in cultured oviduct.
A: dose dependency; B: time dependency. In (A),
oviducts were cultured for 8 h with E2 at concentrations
indicated. In (B), oviducts were cultured with 100 nM
E2 for times indicated. Epo secreted in culture media was
measured. Bars represent means ± SE (n = 4). Statistical
significance of differences was determined with ANOVA followed by
Dunnett's test. * P < 0.05 and ** P < 0.01, significantly different from values of controls, respectively.
|
|
Figure 4 shows the effects of the
E2 receptor (ER) antagonist and of inhibitors of protein
and RNA synthesis on the E2-induced production of Epo.
Addition of ICI-182780, a specific antagonist of the ER, decreased Epo
production. Cycloheximide and actinomycin D completely inhibited Epo
production, demonstrating that Epo secreted into the culture media was
newly synthesized during culture. Tamoxifen, which is an E2
antagonist in breast tissues but acts as an agonist in female
reproductive organs (10a), stimulated Epo production with a potency
similar to E2 (data not shown). ICI-182780 repressed
tamoxifen-induced Epo production.

View larger version (12K):
[in this window]
[in a new window]
|
Fig. 4.
Effect of an E2 antagonist and inhibitors of protein and
RNA synthesis on Epo production by cultured oviducts. Oviducts were
cultured for 8 h with 100 nM E2, 100 µM ICI-182780 (ICI,
an E2 antagonist), 10 µM actinomycin D (Act D), or 200 µM cycloheximide (CHX). Epo secreted in culture media was measured.
Bars represent means ± SE (n = 4). Statistical significance
of differences was determined with ANOVA followed by Tukey-Kramer's
test. ** P < 0.01, significantly different from each
other.
|
|
In addition to oxygen and E2, Epo production has been shown
to be influenced by some other substances. Thyroid hormones enhance hypoxia-induced Epo production by HepG2 and the perfused rat kidney (10); retinoic acid promotes Epo production in HepG2 and vitamin A-depleted rats (33); and insulin, insulin-like growth factor (IGF) I,
and IGF-II stimulate Epo production by cultured astrocytes (27). We
examined the stimulation of Epo production in the cultured oviducts by
various ligands for nuclear receptors, including progesterone, testosterone, 3,3',5-triiodo-L-thyronine,
L-thyroxine, all-trans retinoic acid, and vitamin
D3. The oviducts were cultured for 8 h in the presence of
individual compounds at 100 nM. The addition of these compounds had no
significant effect on Epo production (data not shown), indicating that
low oxygen and E2 are specific stimulators for Epo
production in the oviduct.
Next, we investigated the expression of Epo mRNA in cultured oviducts.
As Fig. 5 shows, Epo mRNA was clearly
induced by E2 at 2 h after culture, reached the maximum at
4 h, and markedly decreased at 8 h. The E2-induced
expression of Epo mRNA in cultured oviducts was repressed by ICI-182780
and actinomycin D (Fig. 6). In contrast,
Epo mRNA was superinduced by cycloheximide in both the presence and
absence of E2, indicating that de novo protein synthesis is
not needed for induction of Epo mRNA.

View larger version (20K):
[in this window]
[in a new window]
|
Fig. 5.
Expression of Epo mRNA in cultured oviducts is induced by
E2. Oviducts were cultured with or without 100 nM
E2 for indicated times, and Epo mRNA was measured. Open and
filled columns, Epo mRNA in oviducts cultured without ( ) and
with E2 (+), respectively. Bars are means ± SE (n
= 3). Statistical significance of differences was determined with
ANOVA followed by Tukey-Kramer's test. * P < 0.05 and
** P < 0.01, respectively.
|
|

View larger version (18K):
[in this window]
[in a new window]
|
Fig. 6.
Effect of an E2 antagonist and inhibitors of protein and
RNA synthesis on E2-induced expression of Epo mRNA in
cultured oviducts. Oviducts were cultured for 8 h with 100 nM
E2, 100 µM ICI, 10 µM Act D, or 200 µM CHX. Epo mRNA
in culture oviducts was measured. Bars represent means ± SE (n
= 3). Statistical significance of differences was determined with
ANOVA followed by Tukey-Kramer's test. ** P < 0.01, significantly different from each other.
|
|
The oviduct is composed of three morphologically distinguishable
segments. The funnel-shaped abdominal end of the oviduct is called the
infundibulum. The expanded intermediate segment below the infundibulum
is the ampulla. The isthmus is the slender medial third near the
uterine wall. To clarify which segment is responsible for the oviductal
Epo production, oviducts were separated into the three segments, and
they were cultured with or without E2. Epo protein in the
culture media was assayed (Fig. 7). The ampulla and isthmus produced Epo in the absence of E2, and
the production was stimulated by E2. Epo in the culture
media of the infundibulum was almost undetectable in the presence and
absence of E2.

View larger version (13K):
[in this window]
[in a new window]
|
Fig. 7.
Ampulla and isthmus produce Epo in an E2-dependent manner.
Each part of the oviduct was dissected and cultured in the presence or
absence of E2 for 8 h. Secreted Epo was measured by enzyme
immunoassay. Filled columns, Epo protein secreted into media cultured
with 100 nM E2. Open columns, Epo protein when cultured
without E2. Bars represent means ± SE (n = 4).
Statistical significance of differences was determined with Student's
t -test. * P < 0.05 and *** P
< 0.001, significantly different from values of controls (values
in absence of E2), respectively.
|
|
 |
DISCUSSION |
There are at least four sites (liver, kidney, brain, and uterus) of Epo
production (28). Epo produced by the liver and kidney stimulates
erythropoiesis, brain Epo prevents ischemic neuron death, and uterine
Epo plays a critical role in the E2-dependent cyclical
angiogenesis in the uterus. Whereas Epo production in the kidney,
liver, and brain is hypoxia inducible, that in the uterus is
E2 inducible. These findings of Epo production and a new
function in the uterus prompted us to explore the possibility that
other female reproductive organs (ovary and oviduct) may produce Epo.
Although both the ovary and oviduct expressed Epo mRNA, expression of
Epo mRNA in the oviduct is highly E2 inducible; therefore,
we examined the properties of Epo production by the oviduct. The in
vitro cultured oviducts were also found to express Epo mRNA and secrete
Epo into the culture media, both of which were E2 inducible.
Tamoxifen (an ER agonist in female reproductive organs) elevates the
expression of oviductal Epo, and ICI-182780 (a specific ER antagonist)
counteracts the E2-induced expression. These results have
made it very likely that E2 acts through ER.
E2-induced accumulation of the oviductal Epo mRNA does not
require de novo protein synthesis, and the accumulation is inhibited by
actinomycin D, suggesting that this accumulation is largely due to the
transcriptional activation of the Epo gene by E2. In the
5' flanking region of both mouse and human Epo genes, the palindromic
consensus sequence of the E2 response element is not
present, but some consensus half-sites, and the sequences homologous to
the consensus half-site, exist. Four half-sites separated from each
other by >100 bp have been shown to confer E2
responsiveness to target genes synergistically (20). Expression of the
reporter gene flanked by a 5' flanking region of the Epo gene was
activated by E2, and this activation required ER-
(unpublished observations). Thus E2-induced accumulation of
oviductal Epo mRNA is at least partly attributable to the
transcriptional activation of the Epo gene.
Downregulation of the oviductal Epo mRNA that occurs shortly after
injection of E2 is due to the loss of the cellular response to E2 but not to the metabolic depletion of E2.
One possibility to account for this loss is that ER was rapidly
degraded by the addition of E2. In fact, stimulation of
proteolytic degradation of ER by E2 was seen in mouse
uterus and breast cancer cell lines (11, 17, 35). It is also possible
that the downregulation of E2-induced transcriptional
activation of the Epo gene is caused by E2-induced
modulation of transcriptional factors associated with ER. Co-activators
such as SRC1/NCoA1, TIF2/GRIP1, AIBI/pCIP/ACTR, p300/CBP, NcoA62,
PBP/TRAP220, and/or co-repressors such as TIF-1 and RIP140 (4, 7, 19,
22, 34, 41, 44) have been shown to be involved in
E2-induced regulation of gene transcription through ER. In
addition, BRCA1 inhibits E2 signaling by blocking the
transcriptional activation function of AF-2 in ER-
(9). Thus a
variety of modes are possible for the downregulation of E2-induced transcriptional activation. Although the
mechanism of the downregulation remains to be studied, the short-lived
stimulatory effect of E2 on expression of the oviductal Epo
gene may be critical for ensuring a yet unknown function of oviductal
Epo without significant perturbation of erythropoiesis.
The oviduct provides the appropriate environment for fertilization of
the ovum released from the ovary and embryonic development, and it
transports the embryo to the uterus. The oviduct can be divided into
three segments, infundibulum, ampulla, and isthmus. The infundibulum
secures oocytes extruded from the ovary. The ampulla is the site of
fertilization. Ciliated cells in the isthmus propel embryos toward the
uterus. The isthmus is thought to be an important region for the
capacitation of spermatozoa. The ampulla and isthmus are the Epo
production sites in the oviduct. Papers have appeared (43, 48)
suggesting that Epo might be involved in sperm formation. Epo mRNA has
been detected in the rat testis under normoxia, and its level is
increased by hypoxia (43). Epo stimulates epididymal sperm maturation
and sperm-fertilizing activity in rats (48). We speculate that Epo in
the oviduct plays a role in fertilization, including sperm
capacitation. Further studies are clearly needed to elucidate the
physiological function of Epo in the oviduct.
 |
ACKNOWLEDGEMENTS |
This work was supported by Grants-in-Aid from the Ministry of
Education, Science and Culture of Japan and from the "Research for
the Future" program in The Japan Society for the Promotion of Science.
 |
FOOTNOTES |
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: R. Sasaki,
Division of Integrated Life Science, Graduate School of Biostudies,
Kyoto Univ., Kyoto 606-8502, Japan (E-mail:
rsasaki{at}kais.kyoto-u.ac.jp).
Received 7 September 1999; accepted in final form 14 January 2000.
 |
REFERENCES |
1.
Anagnostou, A,
Lee ES,
Kessimian N,
Levinson R,
and
Steiner M.
Erythropoietin has a mitogenic and positive chemotactic effect on endothelial cells.
Proc Natl Acad Sci USA
87:
5978-5982,
1990[Abstract].
2.
Anagnostou, A,
Liu Z,
Steiner M,
Chin K,
Lee ES,
Kessimian N,
and
Noguchi CT.
Erythropoietin receptor mRNA expression in human endothelial cells.
Proc Natl Acad Sci USA
91:
3974-3978,
1994[Abstract].
3.
Assandri, R,
Egger M,
Gassmann M,
Niggli E,
Bauer C,
Forster I,
and
Gorlach A.
Erythropoietin modulates intracellular calcium in a human neuroblastoma cell line.
J Physiol (Lond)
516:
343-352,
1999[Abstract/Free Full Text].
4.
Barrett, TJ,
and
Spelsberg TC.
Steroid receptors at the nexus of transcriptional regulation.
J Cell Biochem Suppl
30-31:
185-193,
1998.
5.
Bunn, HF,
and
Poyton RO.
Oxygen sensing and molecular adaptation to hypoxia.
Physiol Rev
76:
839-885,
1996[Abstract/Free Full Text].
6.
Carlini, RG,
Reyes AA,
and
Rothstein M.
Recombinant human erythropoietin stimulates angiogenesis in vitro.
Kidney Int
47:
740-745,
1995[ISI][Medline].
7.
Chen, JD,
and
Li H.
Coactivation and corepression in transcriptional regulation by steroid/nuclear hormone receptors.
Crit Rev Eukaryot Gene Expr
8:
169-190,
1998[ISI][Medline].
8.
Digicaylioglu, M,
Bichet S,
Marti HH,
Wenger RH,
Rivas LA,
Bauer C,
and
Gassmann M.
Localization of specific erythropoietin binding sites in defined areas of the mouse brain.
Proc Natl Acad Sci USA
92:
3717-3720,
1995[Abstract/Free Full Text].
9.
Fan, S,
Wang J-A,
Yuan R,
Ma R,
Meng Q,
Edros MR,
Pestell RG,
Yuan F,
Auborn KJ,
Goldberg ID,
and
Rosen EM.
BRCA1 inhibition of estrogen receptor signaling in transfected cells.
Science
284:
1354-1356,
1999[Abstract/Free Full Text].
10.
Fandrey, J,
Pagel H,
Frede S,
Wolff M,
and
Jelkmann W.
Thyroid hormones enhance hypoxia-induced erythropoietin in vitro.
Exp Hematol
22:
272-277,
1994[ISI][Medline].
10a.
Gallo, MA,
and
Kaufman D.
Antagonistic and agonistic effects of tamoxifen: significance in human cancer.
Semin Oncol
24, Suppl1:
71-81,
1997.
11.
Gibson, MK,
Nemmers LA,
Beckman WC, Jr,
Davis VL,
Curtis SW,
and
Korach KS.
The mechanism of ICI 164,384 antiestrogenicity involves rapid loss of estrogen receptor in uterine tissue.
Endocrinology
129:
2000-2010,
1991[Abstract].
12.
Goto, M,
Akai K,
Murakami A,
Hashimoto C,
Tsuda E,
Ueda M,
Kawanishi G,
Takahashi N,
Ishimoto A,
Chiba H,
and
Sasaki R.
Production of recombinant human erythropoietin in mammalian cells: host-cell dependency of the biological activity of the cloned glycoprotein.
Bio/Technology
6:
67-71,
1988[ISI].
13.
Goto, M,
Murakami A,
Akai K,
Kawanishi G,
Ueda M,
Chiba H,
and
Sasaki R.
Characterization and use of monoclonal antibodies directed against human erythropoietin that recognize different antigenic determinants.
Blood
74:
1415-1423,
1989[Abstract].
14.
Guillemin, K,
and
Krasnow MA.
The hypoxic response: huffing and hifing.
Cell
89:
9-12,
1997[ISI][Medline].
15.
Haller, H,
Christel C,
Dannenberg L,
Thiele P,
Lindschau C,
and
Luft FC.
Signal transduction of erythropoietin in endothelial cells.
Kidney Int
50:
481-488,
1996[ISI][Medline].
16.
Heid, CA,
Stevens J,
Livak KJ,
and
Williams PM.
Real time quantitative PCR.
Genome Res
6:
986-994,
1996[Abstract].
17.
Horigome, T,
Ogata F,
Golding TS,
and
Korach KS.
Estradiol-stimulated proteolytic cleavage of the estrogen receptor in mouse uterus.
Endocrinology
123:
2540-2548,
1988[Abstract].
18.
Jelkmann, W.
Erythropoietin: structure, control of production, and function.
Physiol Rev
72:
449-489,
1992[Free Full Text].
19.
Jenster, G.
Coactivators and corepressors as mediators of nuclear receptor function: an update.
Mol Cell Endocrinol
143:
1-7,
1998[ISI][Medline].
20.
Kato, S,
Tora L,
Yamaguchi J,
Masushige S,
Bellard M,
and
Chambon P.
A far upstream estrogen response element of the ovalbumin gene contains several half-palindromic 5'-TGACC-3' motifs acting synergistically.
Cell
68:
731-742,
1992[ISI][Medline].
22.
Klein-Hitpass, L,
Schwerk C,
Kahmann S,
and
Vassen L.
Targets of activated steroid hormone receptors: basal transcription factors and receptor interacting proteins.
J Mol Med
76:
490-496,
1998[ISI][Medline].
23.
Krantz, SB.
Erythropoietin.
Blood
77:
419-434,
1991[ISI][Medline].
24.
Lin, C-S,
Lim S-K,
D'Agati V,
and
Costantini F.
Differential effects of an erythropoietin receptor gene disruption on primitive and definitive erythropoiesis.
Genes Dev
10:
154-164,
1996[Abstract].
25.
Liu, C,
Shen K,
Liu Z,
and
Noguchi CT.
Regulated human erythropoietin receptor expression in mouse brain.
J Biol Chem
272:
32395-32400,
1997[Abstract/Free Full Text].
26.
Marti, HH,
Wenger RH,
Rivas LA,
Straumann U,
Digicaylioglu M,
Henn V,
Yonekawa Y,
Bauer C,
and
Gassmann M.
Erythropoietin gene expression in human, monkey and murine brain.
Eur J Neurosci
8:
666-676,
1996[ISI][Medline].
27.
Masuda, S,
Chikuma M,
and
Sasaki R.
Insulin-like growth factors and insulin stimulate erythropoietin production in primary cultured astrocytes.
Brain Res
746:
63-70,
1997[ISI][Medline].
28.
Masuda, S,
Nagao M,
and
Sasaki R.
Erythropoietic, neurotrophic, and angiogenic functions of erythropoietin and regulation of erythropoietin production.
Int J Hematol
70:
1-6,
1999[ISI][Medline].
29.
Masuda, S,
Nagao M,
Takahata K,
Konishi Y,
Gallyas F, Jr,
Tabira T,
and
Sasaki R.
Functional erythropoietin receptor of the cells with neural characteristics: comparison with receptor properties of erythroid cells.
J Biol Chem
268:
11208-11216,
1993[Abstract/Free Full Text].
30.
Masuda, S,
Okano M,
Yamagishi K,
Nagao M,
Ueda M,
and
Sasaki R.
A novel site of erythropoietin production: oxygen-dependent production in cultured rat astrocytes.
J Biol Chem
269:
19488-19493,
1994[Abstract/Free Full Text].
31.
Maxwell, PH,
Wiesener MS,
Chang-W G,
Clifford SC,
Vaux EC,
Cockman ME,
Wykoff CC,
Pugh CW,
Maher ER,
and
Ratcliffe PJ.
The tumor suppressor protein VHL targets for oxygen-dependent proteolysis.
Nature
399:
271-275,
1999[ISI][Medline].
32.
Morishita, E,
Masuda S,
Nagao M,
Yasuda Y,
and
Sasaki R.
Erythropoietin receptor is expressed in rat hippocampal and cerebral cortical neurons, and erythropoietin prevents in vitro glutamate-induced neuronal death.
Neuroscience
76:
105-116,
1997[ISI][Medline].
33.
Okano, M,
Masuda S,
Narita H,
Masushige S,
Kato S,
Imagawa S,
and
Sasaki R.
Retinoic acid up-regulates erythropoietin production in hepatoma cells and in vitamin A-depleted rats.
FEBS Lett
349:
229-233,
1994[ISI][Medline].
34.
Parker, MG.
Transcriptional activation by oestrogen receptors.
Biochem Soc Symp
63:
45-50,
1998[Medline].
35.
Pink, JJ,
and
Jordan VC.
Models of estrogen receptor regulation by estrogens and antiestrogens in breast cancer cell lines.
Cancer Res
56:
2321-2330,
1996[Abstract].
36.
Porter, DL,
and
Goldberg MA.
Regulation of erythropoietin production.
Exp Hematol
21:
399-404,
1993[ISI][Medline].
37.
Ratcliffe, PJ,
O'Rourke JF,
Maxwell PH,
and
Pugh CW.
Oxygen sensing, hypoxia-inducible factor-1 and the regulation of mammalian gene expression.
J Exp Biol
201:
1153-1162,
1998[Abstract/Free Full Text].
38.
Sadamoto, Y,
Igase K,
Sakanaka M,
Sato K,
Otsuka H,
Sakaki S,
Masuda S,
and
Sasaki R.
Erythropoietin prevents place navigation disability and cortical infarction in rats with permanent occlusion of the middle cerebral artery.
Biochem Biophys Res Commun
253:
26-32,
1998[ISI][Medline].
39.
Sakanaka, M,
Wen TC,
Matsuda S,
Masuda S,
Morishita E,
Nagao M,
and
Sasaki R.
In vivo evidence that erythropoietin protects neurons from ischemic damage.
Proc Natl Acad Sci USA
95:
4635-4640,
1998[Abstract/Free Full Text].
40.
Semenza, GL.
Hypoxia-inducible factor 1 and the molecular physiology of oxygen homeostasis.
J Lab Clin Med
131:
207-214,
1998[ISI][Medline].
41.
Shibata, H,
Spencer TE,
Onate SA,
Jenster G,
Tsai SY,
Tsai MJ,
and
O'Malley BW.
Role of co-activators and co-repressors in the mechanism of steroid/thyroid receptor action.
Recent Prog Horm Res
52:
141-164,
1997[ISI][Medline].
42.
Shoemaker, CB,
and
Mitsock LD.
Murine erythropoietin gene: cloning, expression, and human gene homology.
Mol Cell Biol
6:
849-858,
1986[ISI][Medline].
43.
Tan, CC,
Eckardt K-U,
Firth JD,
and
Ratcliffe PJ.
Feedback modulation of renal and hepatic erythropoietin mRNA in response to graded anemia and hypoxia.
Am J Physiol Renal Fluid Electrolyte Physiol
263:
F474-F481,
1992[Abstract/Free Full Text].
44.
Torchia, J,
Glass C,
and
Rosenfeld MG.
Co-activators and co-repressors in the integration of transcriptional responses.
Curr Opin Cell Biol
10:
373-383,
1998[ISI][Medline].
45.
Wenger, RH,
and
Gassmann M.
Oxygen(es) and the hypoxia-inducible factor-1.
Biol Chem
378:
609-616,
1997.
46.
Wu, H,
Liu X,
Jaenisch R,
and
Lodish HF.
Generation of committed erythroid BFU-E and CFU-E progenitors does not require erythropoietin or the erythropoietin receptor.
Cell
83:
59-67,
1995[ISI][Medline].
47.
Yamaji, R,
Okada T,
Moriya M,
Naito M,
Tsuruo T,
Miyatake K,
and
Nakano Y.
Brain capillary endothelial cells express two forms of erythropoietin receptor mRNA.
Eur J Biochem
239:
494-500,
1996[Abstract].
48.
Yamamoto, Y,
Sofikitis N,
and
Miyagawa I.
Effects of erythropoietin, bromocryptine and hydrazine on testicular function in rats with chronic renal failure.
Andrologia
29:
141-144,
1997[ISI][Medline].
49.
Yasuda, Y,
Masuda S,
Chikuma M,
Inoue K,
Nagao M,
and
Sasaki R.
Estrogen-dependent production of erythropoietin in uterus and its implication in uterine angiogenesis.
J Biol Chem
273:
25381-25387,
1998[Abstract/Free Full Text].
50.
Youssoufian, H,
Longmore G,
Neumann D,
Yoshimura A,
and
Lodish HF.
Structure, function, and activation of the erythropoietin receptor.
Blood
81:
2223-2236,
1993[ISI][Medline].
Am J Physiol Endocrinol Metab 278(6):E1038-E1044
0193-1849/00 $5.00
Copyright © 2000 the American Physiological Society