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HEMATOPOIESIS
HIF-2␣ regulates murine hematopoietic development in an
erythropoietin-dependent manner
Marzia Scortegagna, Kan Ding, Quiyang Zhang, Yavuz Oktay, Michael J. Bennett, Michael Bennett,
John M. Shelton, James A. Richardson, Orson Moe, and Joseph A. Garcia
Erythropoiesis in the adult mammal depends critically on erythropoietin, an inducible cytokine with pluripotent effects.
Erythropoietin gene expression increases
under conditions associated with lowered oxygen content such as anemia and
hypoxia. HIF-1␣, the founding member of
the hypoxia-inducible factor (HIF) alpha
class, was identified by its ability to bind
and activate the hypoxia-responsive enhancer in the erythropoietin regulatory
region in vitro. The existence of multiple
HIF alpha members raises the question of
which HIF alpha member or members
regulates erythropoietin expression in
vivo. We previously reported that mice
lacking wild-type HIF-2␣, encoded by the
EPAS1 gene, exhibit pancytopenia. In this
study, we have characterized the etiology
of this hematopoietic phenotype. Molecular studies of EPAS1-null kidneys reveal
dramatically decreased erythropoietin
gene expression. EPAS1-null as well as
heterozygous mice have impaired renal
erythropoietin induction in response to
hypoxia. Treatment of EPAS1-null mice
with exogenous erythropoietin reverses
the hematopoietic and other defects. We
propose that HIF-2␣ is an essential regulator of murine erythropoietin production.
Impairments in HIF signaling, involving
either HIF-1␣ or HIF-2␣, may play a prominent role in conditions involving altered
hematopoietic or erythropoietin homeostasis. (Blood. 2005;105:3133-3140)
© 2005 by The American Society of Hematology
Introduction
Hematopoiesis involves complex interactions of specialized cell
types and molecular signaling events that vary according to
development.1 The major sites of murine hematopoietic development depend on the particular developmental or postnatal stage.2,3
Erythropoiesis, that aspect of hematopoiesis concerned with generation of erythrocytes, occurs in 2 distinct developmental phases
characterized by primitive erythropoiesis in the yolk sac blood
islands and definitive erythropoiesis in the fetal liver or bone
marrow later in development.1,4,5 Besides location, primitive and
definitive erythropoiesis also differ in their growth factor or
cytokine requirements.
Erythropoietin was first described as an endocrine regulator of
erythropoiesis produced in the kidneys; later studies revealed a
paracrine role of this cytokine in global hematopoiesis and other
aspects of mammalian physiology.6-9 Primitive erythropoiesis in
the yolk sac is erythropoietin-independent, but requires other
cytokines such as vascular endothelial growth factor (VEGF) and
c-Kit.10-12 In contrast, definitive erythropoiesis in the fetal liver and
in the adult bone marrow is erythropoietin-dependent as evident by
gene disruption studies of the erythropoietin13 or erythropoietin
receptor14 gene. The temporal distinction of primitive and definitive erythropoiesis is in part due to developmental timing of
erythropoietin and erythropoietin receptor gene expression.15
The sites of definitive erythropoiesis are also sites of erythropoietin production.16 These sites include resident macrophages in the
adult bone marrow as well as in the fetal liver blood islands,17 the
predominant sites of definitive erythropoiesis in the adult and
neonate, respectively. Erythropoietin produced in these locations
may function as a paracrine growth factor for global hematopoiesis.
However, despite its production in and impact on the bone marrow
compartment, the kidney is the major production site of circulating
erythropoietin in the adult mammal.18
Although the kidney has long been recognized as the source of
endocrine erythropoietin,18 the exact site of renal erythropoietin
production has been a topic of debate.6 Molecular and transgenic
mouse studies provide evidence for erythropoietin production in at
least 2 different candidate cell types, the proximal tubular cells19,20
and the peritubular or interstitial cells.21-24 Transgenic mice generated using a human erythropoietin transgene demonstrated erythropoietin production in the renal peritubular interstitial cell population,25 whereas transgenic mice using a mouse erythropoietin
promoter-lacZ transgene revealed lacZ expression in the proximal
tubular cells.26 Another transgenic mouse strain resulting from
homologous recombination of a simian virus 40 (SV40) epitopetagged erythropoietin transgene localized erythropoietin production to the fibroblast interstitial population.27 Hence, several cell
populations are candidate sites for erythropoietin production in
the kidney.
Renal erythropoietin production is regulated primarily at the transcriptional level and is markedly induced by anemia or global hypoxia.28
From the Departments of Internal Medicine, Pathology, and Molecular Biology,
University of Texas Southwestern Medical Center; and the Department of
Pathology, Children’s Medical Center, Dallas, TX.
the American Cancer Society, and the Donald W. Reynolds Foundation.
Submitted May 3, 2004; accepted December 27, 2004. Prepublished online as
Blood First Edition Paper, December 30, 2004; DOI 10.1182/blood-2004-05-1695.
Supported by grants to J.G. from the National Institutes of Health, National Heart,
Lung, Blood Institute (NIH/NHLBI); NIH/National Institute of Mental Health
(NIMH) Conte Center, the American Heart Association (AHA)/Texas Affiliate,
BLOOD, 15 APRIL 2005 䡠 VOLUME 105, NUMBER 8
M.S. and K.D. contributed equally to this study.
Reprints: Joseph A. Garcia, UT Southwestern Medical Center, 5323 Harry Hines
Blvd, Dallas, TX 75390-8573; e-mail: [email protected].
The publication costs of this article were defrayed in part by page charge
payment. Therefore, and solely to indicate this fact, this article is hereby
marked ‘‘advertisement’’ in accordance with 18 U.S.C. section 1734.
© 2005 by The American Society of Hematology
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3134
BLOOD, 15 APRIL 2005 䡠 VOLUME 105, NUMBER 8
SCORTEGAGNA et al
Identifying the molecular mechanisms controlling erythropoietin gene
expression was enabled by the identification of a hepatic cell line that
expressed erythropoietin in a hypoxia-dependent manner.29,30 Investigators defined the enhancer element in the erythropoietin gene responsible
for hypoxia induction31-33 and used DNA affinity chromatography to
identify31 and purify34 a heterodimer protein complex, hypoxia inducible factor 1 (HIF-1). HIF-1 consists of 2 members of the basic
helix-loop-helix (bHLH): Per-Arnt-Sim (PAS) domain containing transcription factor family, HIF-1 alpha (HIF-1␣) and HIF-1 beta (HIF-1␤),
with the latter previously identified as the aryl hydrocarbon nuclear
translocator (ARNT).35
Biologic specificity of bHLH-PAS domain transcription factors
is dictated by the alpha component whereas biologic activity is
conferred by the beta component. With the use of bioinformatics,
additional members of the HIF alpha family were identified.
Endothelial PAS domain protein 1 (EPAS1),36 also known as
HIF-2␣, is closely related to HIF-1␣ in composition. Although
early studies of HIF-2␣ primarily evaluated whether biologic
actions of HIF-2␣ were similar to those of HIF-1␣, more recent
investigations have highlighted unique aspects for HIF-2␣ or
HIF-1␣.37-40 Evidence for dual requirements of HIF-1␣ and HIF-2␣
in the in vivo regulation of HIF target genes in the adult mouse has
not been presented to date.
Targeted disruption of HIF members provided insights into their
roles in hematopoiesis. Embryos lacking ARNT have defects in
yolk sac vascularization, suggesting a defect in primitive hematopoiesis.41 Differentiation assays of ARNT⫺/⫺ embryonic stem (ES)
cells suggest ARNT is essential for global hematopoiesis.42 Further
characterization with ARNT⫺/⫺ ES cell assays suggested that the
absence of VEGF might be responsible for the global hematopoietic defect. Since VEGF is required for primitive erythropoiesis
and since VEGF is a target gene for HIF-1, the ARNT⫺/⫺ ES cell
experiments suggest a role for HIF members in primitive erythropoiesis. Moreover, the ineffectiveness of exogenous erythropoietin
upon ARNT⫺/⫺ ES cell differentiation constitute further evidence
that the ARNT⫺/⫺ hematopoietic phenotype may involve abnormal
primitive erythropoiesis.42
Chimeric and conditional knockout mice have provided some
insights into the role of HIF-1 in B-cell development43 and myeloid
cell function,44 respectively. Examination of HIF-1␣⫹/⫺ mice
revealed a minor impairment in chronic continuous hypoxiastimulated erythrocytosis,45 whereas a brief period of intermittent
or continuous hypoxia treatment revealed a lack of induction of
renal erythropoietin gene expression.9 These results suggest a role
for HIF-1 in erythropoietin gene expression in the adult mouse and
hence in definitive erythropoiesis. However, a specific role for
HIF-1 in global hematopoiesis has been hampered by the unavailability of HIF-1␣⫺/⫺ mice.
The generation of adult mice globally lacking HIF-2␣ (EPAS1⫺/⫺
mice) allowed biologic roles for HIF-2␣ to be defined in the
context of the intact organism.46,47 One aspect of the EPAS1⫺/⫺
phenotype is hematopoietic deficiency.47 Transplantation experiments using irradiated EPAS1⫺/⫺ mice as recipients of syngeneic
EPAS1⫹/⫹ hematopoietic stem cells revealed that the hematopoietic
defect in EPAS1⫺/⫺ mice is due to altered bone marrow microenvironment function and/or to effects of the EPAS1⫺/⫺ state on
expression of systemic mediators.47 Gene expression of VEGF
members and VEGF receptors in the heterogeneous bone marrow
cell population is not significantly different between EPAS1⫺/⫺ and
EPAS1⫹/⫹ mice, implying that abnormalities in VEGF signaling
may not be responsible for the hematopoietic defect in EPAS1⫺/⫺
mice.47 We reasoned that a systemic mediator of hematopoietic
development might be impaired in EPAS1⫺/⫺ mice. Given the
potential role of erythropoietin in global hematopoiesis and the
likely role of HIF members in erythropoietin regulation, we were
particularly interested in determining whether abnormalities in
erythropoietin signaling might be responsible for the hematopoietic
defect observed in EPAS1⫺/⫺ mice.
Materials and methods
Animal studies
We generated F1 hybrid mice by crossing 129⫹/⫺ female mice with C57⫹/⫺
male mice carrying a single mutant EPAS1 allele as described.47 Mice
housed in a standard 12:12 light/dark cycle were fed ad lib with 4%
fat-containing chow (mating pairs) or 11% fat-containing chow (pregnant
females and newborn mice up to one month old). Polymerase chain reaction
(PCR) genotyping was performed as described.47
We performed hypoxia experiments using either short-term, mediumterm, or long-term treatment protocols. For short-term hypoxia treatments,
mice were housed in a 13-L plexiglass container and exposed to air
mixtures using flow-through air delivery by pressurized tanks at a rate of 2
L per minute. For room air (RA) treatments, age- and gender-matched mice
were exposed to 6 RA/RA cycles with each cycle consisting of 5 minutes of
RA (21% oxygen/79% nitrogen) followed by 5 minutes of RA (21%
oxygen/79% nitrogen). For short-term intermittent hypoxia (STIH), ageand gender-matched mice were exposed to 6 RA/hypoxia cycles with each
cycle consisting of 5 minutes of RA (21% oxygen/79% nitrogen) followed
by 5 minutes of hypoxia air mixture (6% oxygen/94% nitrogen). Mice were
harvested for tissues 1 hour after completion of the STZH treatment
protocol (2 hours after the initiation of the first hypoxia cycle) for RNA or
harvested immediately for protein. For short-term continuous hypoxia
(STCH) or medium-term continuous hypoxia (MTCH), age- and gendermatched mice were exposed to hypoxia air mixture (6% oxygen/94%
nitrogen) for 2 hours or 4 hours, respectively. Mice were harvested
for tissues immediately after completion of the STCH or MTCH
treatment protocol.
For long-term continuous hypoxia (LTCH), age- and gender-matched
mice were housed individually in cages contained within a large plexiglass
chamber. Mice were exposed for 12 days to hypoxia air mixture (10%
oxygen/94% nitrogen) using flow-through air delivery by pressurized tanks
at a rate of 2 L per minute. Carbon dioxide scrubbers were present to keep
levels below 0.5% as assessed by daily checks. Cage changes were made
halfway through the hypoxia treatment protocol and mice were fed ad lib
with standard chow.
We performed rescue experiments with intraperitoneal injections on
Mondays, Wednesdays, and Fridays of low-dose48 (500 U/kg body
weight) or high-dose49 (5000 U/kg body weight) recombinant human
erythropoietin (Epogen; Amgen, Thousand Oaks, CA) prepared in
phosphate-buffered saline (PBS). Erythropoietin injections with 500
U/kg began after postnatal day 30 and increased to 5000 U/kg after
postnatal day 50. Age- and gender-matched EPAS1⫺/⫺ and EPAS1⫹/⫹
mice were used in the rescue experiments. We obtained approval for all
experimental protocols from the UT Southwestern Medical Center
Institutional Animal Care and Use Committee.
Gross pathologic and histologic studies
For whole mouse and kidney weights, we collected information from ageand gender-matched EPAS1⫺/⫺ and EPAS1⫹/⫹ mice. Organ–to–bodyweight ratios were calculated from wet organ weights measured from
exsanguinated mice normalized to the respective body weight obtained
prior to organ harvest.
For ␤-galactosidase staining, we used EPAS1⫹/⫹, EPAS1⫹/⫺ or EPAS1⫺/⫺
mice perfused with 4% parafomaldehyde/PBS by cardiac puncture. We
removed and postfixed select organs for 1 additional hour prior to
subsequent manipulations. We embedded samples in paraffin for further
sectioning when indicated. For ␤-galactosidase staining, we stained kidney
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BLOOD, 15 APRIL 2005 䡠 VOLUME 105, NUMBER 8
vibratome sections, embedded the stained sections in paraffin for sectioning, and imaged the resultant sections (⫻40 fields). We performed
hematoxylin and eosin (H&E) staining on paraformaldehyde-fixed, paraffinembedded samples (⫻20 fields, kidney), oil red O staining on frozen
sections (⫻40, liver), and Giemsa staining on decalcified bone marrow left
in situ (⫻40).
For in situ analyses, kidneys were removed from paraformaldehydeperfused EPAS1⫹/⫹ mice maintained under STCH conditions as described.
Sagittal-oriented paraffin sections (5 ␮m) were placed on slides for
35S-labeled RNA radioactive in situ hybridization. The antisense or sense
complementary RNA (cRNA) probes used were generated from pGEM
T-easy vectors (Promega Corporation, Madison, WI) containing complementary DNA (cDNA) inserts from murine erythroprotein (mEpo) (nucleotide
[nt] 194-579, RefSeq NM_007942) or murine EPAS1 (mEPAS1) (nt
1378-1890, RefSeq NM_010137) using a SP6/T7 Maxiscript Kit (Ambion,
Woodland Hills, TX) and 35S-uridine 5⬘-triphosphate (Amersham Biosciences, Piscataway, NJ).
Microscopy and image acquisition
We reviewed all histological preparations on a Leica Laborlux S photomicroscope (Leica Microsystems, Wetzlar, Germany) equipped with brightfield and incidental dark-field illumination. We imaged paraffin-embedded
or frozen tissue sections without oil or water immersion using 20 ⫻ or 40 ⫻
Leica objectives with plan-achromatic optics of 0.45 and 1.00 numerical
aperture, respectively. We photographed images using this same Leica
photomicroscope and an Optronics DEI-750 analog CCD color camera
(Optronics, Goleta, CA) interfaced with a Scion CG-7 frame-digitizerequipped (Scion Corporation, Frederick, MD) Macintosh G4 desktop
computer (Apple Computer, Cupertino, CA). We captured images using
Image J v1.23 acquisition and analysis software (Scion Corporation,
Frederick, MD & NIH, Bethesda, MD). We processed and adjusted the
images with Adobe Photoshop 7.0 (Adobe Systems, San Jose, CA).
Samples were imaged under identical light conditions. Posthoc image
adjustments of brightness/contrast or magnification, if performed, were
made with identical modification of parameters for EPAS1⫺/⫺ and
EPAS1⫹/⫹ samples.
Hematopoietic and serum studies
For spun hematocrits, we collected blood using tail-vein sampling with
heparinized capillary tubes. For complete blood cell counts, we collected
blood samples from anesthetized mice via retro-orbital punctures just prior
to euthanasia and performed automated cell counts (Dallas Children’s
Hospital Pathology Laboratory, Dallas, TX). Serum was prepared from
blood and frozen at ⫺80°C until use. We processed samples using
automated analyses for routine metabolites or electrolytes (Dallas Children’s Hospital).
Molecular studies
We obtained kidney samples from freshly euthanized mice matched for
genotype, gender, age, litter, and treatment group. For each sample, we
generated total RNA followed by first-strand cDNA preparations. Quantitative real-time reverse transcription–polymerase chain reaction (rtRT-PCR)
reactions identified changes in gene expression of candidate genes with
SYBR-green based rtRT-PCR measured in an ABI Prism 7700 sequence
detection system instrument and software (Applied Biosystems, Foster
City, CA). The genes examined included erythropoietin (Epo) as well as a
housekeeping gene for the internal standard (␤-actin) with primers designed to span introns and validated prior to use as described earlier.9,46
Gene expression for EPAS1⫺/⫺ or EPAS1⫹/⫺ samples were compared with
results obtained from EPAS1⫹/⫹ samples using the threshold cycle method
normalized to ␤-actin (User Bulletin Number 2; Applied Biosystems). The
data for each genotype are the overall mean of 4 individual subject means
with each subject mean generated from duplicate rtRT-PCR data points.
HIF-2␣ REGULATES MURINE ERYTHROPOIETIN
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Western blot analyses
For total kidney protein extracts, freshly harvested whole kidney was
minced in ice-cold harvesting buffer (50 mM Tris, pH 7.4; 500 mM NaCl; 2
mM EDTA [ethylenediaminetetraacetic acid], pH 7.9; 0.5% Nonidet-P40;
100 ␮M CoCl2; 1 mM dithiothreitol [DTT]) supplemented with protease
inhibitor cocktail (Complete Mini; Roche, Indianapolis, IN; cat no.
1 836 153). The tissues then were extracted with 10:1 volume/weight
ice-cold homogenization buffer (25 mM Tris, pH 7.4; 1 mM EDTA, pH 7.9;
10% glycerol; 100 ␮M CoCl2; 1 mM DTT) with protease inhibitor cocktail.
Protein concentrations were determined by bicinchoninic acid (BCA)
protein assay (Pierce Biotechnology, Rockford, IL; cat no. 1 856 210) with
bovine serum albumin as standard.
Nuclear extracts were prepared by centrifugation of the whole kidney
extracts at 800g for 5 minutes at 4°C. The nuclear pellets were gently
resuspended with ice-cold X-wash buffer (0.25 M sucrose; 20 mM Tris, pH
7.8; 1.5 mM MgCl2; 8.5% glycerol; 5% Triton X-100; 100 ␮M CoCl2; 1
mM DTT) in presence of protease cocktail. After centrifugation at 1500g
for 10 minutes at 4°C, the pellets were washed with ice-cold X-wash buffer
minus Triton X-100 and then centrifuged. The pellets were homogenized in
ice-cold lysis buffer (20 mM Tris, pH 7.8; 450 mM NaCl; 5 mM EDTA, pH
7.9; 10 mM EGTA [ethylene glycol-bis(beta-aminoethyl ether)-N,N,N⬘,N⬘tetraacetic acid]; 25% glycerol; 100 ␮M CoCl2; 1 mM DTT) in presence of
protease inhibitor cocktail and incubated for 1 hour at 4°C. Nuclear debris
was pelleted by centrifugation at 13 000g at 4°C for 30 minutes and the
supernatants collected as nuclear extract.
For Western blot analyses of HIF proteins, nuclear extract protein
preparations (HIF-1␣, 50 ␮g per lane; HIF-2␣, actin: 15 ␮g per lane) were
separated by 6% sodium dodecyl sulfate (SDS)–polyacrylamide gels and
transferred to polyvinylidene difluoride (PVDF) membrane (Amersham
Biosciences, Buckinghamshire, United Kingdom). The blots were blocked
in Blotto-Tween solution (5% nonfat dry milk, 0.1% Tween in PBS) for 1
hour and then incubated overnight in Blotto-Tween with a mouse monoclonal antibody for HIF-1␣ (1:500; Novus, Littleton, CO), HIF-2␣ (1:1000;
Novus), or ␤-actin (1:2000; Sigma, St Louis, MO). Peroxidase-coupled
AffiniPure rabbit anti–mouse immunoglobulin G (1:10 000; Jackson Immunoresearch Laboratories, West Grove, PA) was used in secondary incubations for 1 hour followed by the detection of reactive bands with a
chemiluminescence detection system (ECL) plus kit (Amersham Biosciences, Piscataway, NJ).
For serum erythropoietin protein evaluation, serum was prepared from
blood collected from the inferior vena cava of anesthetized mice. Total
serum protein (150 ␮g per lane) was separated by a 10% SDS–
polyacrylamide gel electrophoresis (PAGE) gel and transferred to PVDF
membranes. The blots were blocked in Blotto-Tween solution (5% nonfat
dry milk, 0.05% Tween in PBS) for 1 hour and then incubated in
Blotto-Tween for 2 hours with a rabbit anti–human erythropoietin (H-162)
polyclonal antibody (1:200; Santa Cruz Biotechnology, Santa Cruz, CA).
The blots were then incubated with horseradish peroxidase (HRP)–
conjugated goat anti–rabbit immunoglobulin (Ig) G (1:5000; Santa Cruz
Biotechnology) for 30 minutes. Labeled protein was visualized by using an
ECL (Amersham Biosciences, Piscataway, NJ) and quantified by densitometry. For control of loading, the blots were first incubated in stripping buffer
(62.5 mM Tris-HCl, 100 mM ␤-mercaptoethanol, 2% SDS) at 50°C for 30
minutes, and then incubated with an HRP-conjugated sheep anti–mouse
IgG (1:1000; Amersham Biosciences) for 1 hour prior to detection by
chemiluminescence. The bands corresponding to the light chain of normal
mouse IgG were used as an internal reference to serum erythropoietin.
Statistics
We reported data as the mean with standard error of the mean (SEM).
Statistical analyses were performed using Excel (Microsoft, Redmond,
WA) or JMP Statistics Software (SAS, Cary, NC). Comparisons were by
Student t tests for paired data, analysis of variance for 3-way comparisons,
and chi-squared or the exact binomial test for proportions.
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SCORTEGAGNA et al
BLOOD, 15 APRIL 2005 䡠 VOLUME 105, NUMBER 8
Results
Mice normally exhibit developmental aspects during the first
month of life, including anemia of prematurity and a lack of
response to exogenous erythropoietin administration.1 For this
reason, we evaluated mice older than 1 month of age to assess for
abnormalities in global hematopoiesis attributable to defects in
erythropoietin synthesis rather than to altered developmental
sequelae. Spun hematocrit measurements of 2-month-old EPAS1⫺/⫺
mice confirmed a reduced red blood cell mass (Figure 1A) as also
seen with 1-month-old EPAS1⫺/⫺ mice.47 We noted increased
kidney–to–body-weight ratios in 2-month-old EPAS1⫺/⫺ mice,
indicating that reduced renal mass was not a cause of the anemia
(Figure 1B). Serum creatinine levels were not significantly different between EPAS1⫺/⫺ and EPAS1⫹/⫹ mice, indicating preservation
of gross renal function (data not shown).
Given its key role in definitive erythropoiesis, we were interested in assessing erythropoietin mRNA levels in EPAS1⫺/⫺ mice.
Despite the anemia, we found markedly reduced renal erythropoietin mRNA levels in 2-month-old EPAS1⫺/⫺ mice in comparison to
EPAS1⫹/⫹ mice maintained at room air (RA) (Figure 1C). Treatment of EPAS1⫺/⫺ mice with hypoxia (short-term intermittent
hypoxia; STIH) resulted in a relative increase in erythropoietin
mRNA levels compared with RA, but remained markedly depressed in the absolute level of erythropoietin mRNA expression
compared with even EPAS1⫹/⫹ mice maintained at RA (Figure 1C).
We next performed histologic and in situ hybridization
studies to evaluate for any pathologic damage and to localize
sites of EPAS1 expression. There were no overt histologic
abnormalities in kidneys from 2-month-old EPAS1⫺/⫺ mice
(Figure 2A) versus EPAS1⫹/⫹ mice (Figure 2B). Activity
staining for nuclear-localized ␤-galactosidase, a surrogate marker
for EPAS1 in this knockout,50 revealed strong expression
predominantly in vascular endothelial cells, but also in the renal
interstitial cell compartment; ␤-galactosidase staining was not
evident in renal tubular cells of 2-month-old EPAS1⫺/⫺ (Figure
2C) or EPAS1⫹/⫺ mice (data not shown). The in situ hybridization analyses revealed prominent expression of EPAS1 mRNA in
vascular endothelial cells (Figure 2D) as well as in a subset of
the renal interstitial cell population (Figure 2E) of EPAS1⫹/⫹
Figure 1. Anemia of EPAS1ⴚ/ⴚ mice accompanied by depressed erythropoietin
mRNA levels. (A) Hematocrit levels for EPAS1⫹/⫹ (䡺) or EPAS1⫺/⫺ (f) mice (n ⫽ 11
per genotype). Statistically meaningful difference in means between the hematocrits
of EPAS1⫹/⫹ and EPAS1⫺/⫺ mice is indicated with an asterisk (1-tailed t test with
P ⬍ .001). (B) Kidney weight–to–body weight ratios (KW/BW) for EPAS1⫹/⫹ (䡺) or
EPAS1⫺/⫺ (f) mice (n ⫽ 7 per genotype). Statistically meaningful difference between
the KW/BW of EPAS1⫹/⫹ and EPAS1⫺/⫺ mice is indicated with an asterisk (1-tailed t
test with P ⬍ .005). (C) Real-time RT-PCR quantification of erythropoietin (Epo)
mRNA levels in EPAS1⫹/⫹ (䡺) or EPAS1⫺/⫺ (f) kidney for mice maintained at room
air (RA) or after exposure to short-term intermittent hypoxia (STIH) (n ⫽ 4 per
genotype per treatment group). Statistically meaningful differences in means between the erythropoietin mRNA levels of EPAS1⫹/⫹ and EPAS1⫺/⫺ mice maintained
at RA (1-tailed t test with P ⬍ .001) or after STIH exposure (1-tailed t test with
P ⬍ .001) are indicated with an asterisk. Values are expressed as the mean plus or
minus standard error of the mean.
Figure 2. EPAS1 is expressed in renal interstitial cells. Representative kidney
sections from hematoxylin and eosin–stained (A) EPAS1⫺/⫺ (–/–) or (B) EPAS1⫹/⫹
(⫹/⫹) kidney (⫻40 fields) and (C) ␤-galactosidase–stained EPAS1⫺/⫺ (–/–) kidney
(⫻40 field). Dark-field (top) and bright-field images (bottom) of in situ studies
performed on kidney samples from EPAS1⫹/⫹ mice after exposure to short-term
continuous hypoxia (STCH) with probes specific for (D) and (E) EPAS1 or (F) Epo
mRNA. The arrows indicate areas of prominent signal in vascular endothelial cells
(VECs), juxtaglomerular complex cells (JGCs), or interstitial cells (ICs) within the
kidney. The black bars in panel B and the white bar in panel C correspond to 100 ␮m;
the black bar in panel F corresponds to 200 ␮m.
mice. Expression of Epo mRNA was characterized by strong
expression in a subset of the renal interstitial cells (Figure 2F).
We next addressed whether a haploinsufficiency state of EPAS1
would alter renal erythropoietin gene expression. Renal erythropoietin expression was not significantly different between EPAS1⫹/⫺
and EPAS1⫹/⫹ mice under normoxic (ie, RA) conditions (Figure
3A). However, after exposure to short-term intermittent hypoxia
(STIH) or short-term continuous hypoxia (STCH) treatment,
induction of Epo gene expression in EPAS1⫹/⫺ mice was impaired
relative to EPAS1⫹/⫹ mice (Figure 3A). Exposure of EPAS1⫹/⫹ or
EPAS1⫹/⫺ mice to STIH or STCH conditions induced activation of
the HIF pathway to a comparable degree as defined by stabilization
of HIF-1␣ protein (Figure 3B). With respect to HIF-2␣, EPAS1⫹/⫺
mice had reduced HIF-2␣ protein levels relative to EPAS1⫹/⫹
mice under RA, STIH, or STCH conditions (Figure 3B) although
there were no significant differences within each genotype for
the 3 conditions.
The differences in renal erythropoietin mRNA levels between
EPAS1⫹/⫺ and EPAS1⫹/⫹ mice translated into functional deficits as
well (Figure 3C). Baseline (RA) hematocrit values were mildly
decreased in EPAS1⫹/⫺ as compared with EPAS1⫹/⫹ mice. Hematocrit determinations performed shortly after initiation of STIH or
STCH trended toward lower values for EPAS1⫹/⫺ as compared
with EPAS1⫹/⫹ mice (Figure 3C). Following prolonged or longterm continuous hypoxia exposure (LTCH), there remained a small,
but significant, difference in the hematocrits of EPAS1⫹/⫺ versus
EPAS1⫹/⫹ mice (Figure 3D).
The molecular characterization and hematocrit results suggest
that a haploinsufficiency state for EPAS1 has measurable effects on
short-term (ie, Epo gene expression) as well as long-term (ie,
erythropoiesis) responses to hypoxic exposure. Since there is a
temporal delay between changes in de novo erythropoietin gene
expression and protein levels, we examined serum erythropoietin
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BLOOD, 15 APRIL 2005 䡠 VOLUME 105, NUMBER 8
Figure 3. Erythropoietin levels in EPAS1ⴙ/ⴚ mice are not induced by hypoxia.
(A) Renal erythropoietin mRNA levels for EPAS1⫹/⫹ (䡺) or EPAS1⫹/⫺ (u) mice
exposed to room air (RA), short-term intermittent hypoxia (STIH), or short-term
continuous hypoxia (STCH) (n ⫽ 4 per genotype per treatment group). Statistically
meaningful differences in means between erythropoietin mRNA levels from EPAS1⫹/⫹
or EPAS1⫹/⫺ mice after STIH exposure (one-tailed t test with P ⫽ .07) or STCH
exposure (one-tailed t test with P ⫽ .06) are indicated with an asterisk. (B) HIF-1␣,
HIF-2␣, or actin protein levels as assessed by Western blot analyses of nuclear
extracts from representative EPAS1⫹/⫹ (⫹/⫹) or EPAS1⫹/⫺ (⫹/–) mice maintained
under RA, STIH, or STCH conditions. A nonspecific (NS) band seen with use of the
HIF-2␣ monoclonal antibody is indicated for comparison between samples. (C)
Hematocrit levels for EPAS1⫹/⫹ (䡺) or EPAS1⫹/⫺ (u) mice maintained at RA (n ⫽ 21
per genotype), STIH (n ⫽ 22 per genotype), or STCH (n ⫽ 13 per genotype).
Statistically meaningful difference in means between the hematocrit levels of
EPAS1⫹/⫹ or EPAS1⫹/⫺ mice maintained at RA is indicated with an asterisk
(one-tailed t test with P ⫽ .01). (D) Hematocrit levels for EPAS1⫹/⫹ (䡺) or EPAS1⫹/⫺
(u) mice maintained at RA or after long-term continuous hypoxia (LTCH) exposure
(n ⫽ 6 per genotype per treatment group). Statistically meaningful difference in
means between the hematocrit levels of EPAS1⫹/⫹ or EPAS1⫹/⫺ mice maintained at
LTCH is indicated with an asterisk (one-tailed t test with P ⫽ .003). Values are
expressed as the mean plus or minus standard error of the mean. (E) Erythropoietin
protein levels as assessed by Western blot analyses of serum from representative
EPAS1⫹/⫹ (⫹/⫹) or EPAS1⫹/⫺ (⫹/–) mice maintained under room air (RA) or
medium-term continuous hypoxia (MTCH) conditions. The same membrane was
reprobed for mouse serum IgG as a comparison between samples.
protein levels for an intermediate-length continuous hypoxia
protocol, medium-term continuous hypoxia (MTCH). The results
obtained revealed reduced serum erythropoietin protein levels in
EPAS1⫹/⫺ versus EPAS1⫹/⫹ mice (Figure 3E), consistent with the
observed changes in renal erythropoietin gene expression found at
earlier time points in the continuous hypoxia protocol (ie, STCH).
To determine if the hematopoietic defect in EPAS1⫺/⫺ mice
would respond to erythropoietin replacement therapy, we administered exogenous erythropoietin to EPAS1⫺/⫺ mice. Erythropoietin
treatment using low-dose (500 U/kg body weight) erythropoietin
dosing of EPAS1⫺/⫺ mice from 30 to 50 days of age resulted in a
time-dependent increase in hematocrit levels close to that of
normal (ie, untreated EPAS1⫹/⫹) mice (Figure 4A). All EPAS1⫺/⫺
mice universally responded to 20 days of low-dose erythropoietin
treatment, although EPAS1⫺/⫺ mice that began treatment with the
most depressed hematocrit levels were slightly below average
wild-type levels after 20 days of erythropoietin treatment.
To assess if supra-physiologic levels (5000 U/kg) of erythropoietin would further stimulate red blood cell production, we increased the erythropoietin dosage administered to EPAS1⫺/⫺ mice
from low- to high-dose levels from 50 to 60 days of age. After 10
days of high-dose erythropoietin treatment, hematocrit levels of
erythropoietin-treated EPAS1⫺/⫺ mice were slightly higher on
average than untreated EPAS1⫹/⫹ mice (Figure 4A). Continued
treatment of EPAS1⫺/⫺ mice with high-dose erythropoietin for an
additional 10 days resulted in erythrocytosis, similar to that
observed in wild-type or heterozygous control mice treated in an
identical manner (data not shown), indicating that erythropoietin
resistance or a decreased erythroid progenitor reserve is not a cause
of the hematopoietic defect observed in untreated EPAS1⫺/⫺ mice.
HIF-2␣ REGULATES MURINE ERYTHROPOIETIN
3137
Figure 4. Pancytopenia of EPAS1ⴚ/ⴚ mice responds to erythropoietin treatment. (A) Hematocrit levels for EPAS1⫹/⫹ (䡺) compared with EPAS1⫺/⫺ (f)
littermate mice treated with erythropoietin (n ⫽ 4 per genotype per time point). The
erythropoietin dosage was either low-dose (500 U/kg) for days 0 to 20 or high-dose
(5000 U/kg) for days 21 to 30 of treatment. Statistically meaningful differences in
means between the hematocrits of EPAS1⫹/⫹ or EPAS1⫺/⫺ mice prior to initiation of
erythropoietin treatment (one-tailed t test with P ⫽ .002) or after 10 days of low-dose
erythropoietin treatment (one-tailed t test with P ⫽ .02) are indicated by an asterisk.
No further differences between groups of mice were noted for all other time points
(one-tailed t test with P ⬎ .1). (B) Complete blood cell count analyses for untreated
EPAS1⫹/⫹ (䡺) or EPAS1⫺/⫺ (f) mice (n ⫽ 6 per genotype). Statistically meaningful
differences in means between values from EPAS1⫹/⫹ or EPAS1⫺/⫺ mice for white
blood cells (WBCs; one-tailed t test with P ⫽ .0001), red blood cells (RBCs;
one-tailed t test with P ⫽ .003), platelets (PLTs; one-tailed t test with P ⫽ .0018), or
reticulocytes (RETs; one-tailed t test with P ⫽ .049) are indicated by an asterisk. (C)
Complete blood cell count analyses for EPAS1⫹/⫹ (䡺) or EPAS1⫺/⫺ (f) littermate
mice after erythropoietin treatment (n ⫽ 4 per genotype). Statistically meaningful
differences in means between values of EPAS1⫹/⫹ or EPAS1⫺/⫺ mice for platelets
(one-tailed t test with P ⫽ .0001) or reticulocytes (one-tailed t test with P ⫽ .0005) are
indicated by an asterisk. The units correspond to 103/␮L for WBCs, 106/␮L for RBCs,
105/␮L for PLTs, and 105/␮L for RETs. Values are expressed as the mean plus or
minus standard error of the mean.
EPAS1⫺/⫺ mice at baseline exhibit marked abnormalities in all the
major hematopoietic lineages (Figure 4B). After 30 days of
erythropoietin treatment, we observed correction of all major blood
lineages in the peripheral blood of 2-month-old EPAS1⫺/⫺ mice,
revealing an effect of erythropoietin on global hematopoiesis
(Figure 4C).
Erythropoietin treatment of EPAS1⫺/⫺ mice alleviated other
aspects of the EPAS1-null state. The hypocellular bone marrow of
EPAS1⫺/⫺ mice (Figure 5A) was corrected in erythropoietin-treated
EPAS1⫺/⫺ mice (Figure 5B), resulting in bone marrow similar in
Figure 5. Erythropoietin treatment alters other aspects of the EPAS1ⴚ/ⴚ
phenotype. Giemsa staining of bone marrow from representative (A) untreated
(-Epo) EPAS1⫺/⫺ (–/–), (B) erythropoietin-treated (⫹Epo) EPAS1⫺/⫺ (–/–), or (C)
EPAS1⫹/⫹ (⫹/⫹) mice (all images are ⫻20 fields, equivalent size). The black bar in
panel C corresponds to 100 ␮m. Representative oil red O (ORO) staining of liver from
(D) untreated (-Epo) EPAS1⫺/⫺ (–/–), (E) erythropoietin-treated (⫹Epo) EPAS1⫺/⫺
(–/–), or (F) EPAS1⫹/⫹ (⫹/⫹) mice (all images are ⫻20 fields, equivalent size). The
black bar in panel F corresponds to 100 ␮m.
From www.bloodjournal.org by guest on August 3, 2017. For personal use only.
3138
SCORTEGAGNA et al
appearance to that of EPAS1⫹/⫹ mice (Figure 5C). The liver–body
weight ratios of EPAS1⫺/⫺ mice improved after erythropoietin
treatment compared with untreated EPAS1⫺/⫺ mice (data not
shown). The lipid deposits found in EPAS1⫺/⫺ livers (Figure 5D)
reversed after erythropoietin treatment (Figure 5E) so as to be
indistinguishable from EPAS1⫹/⫹ mice (Figure 5F).
The biochemical abnormalities normally seen in EPAS1⫺/⫺
mice also were affected by erythropoietin treatment. We noted
improvements in serum glucose (no significant differences between
groups) and lactate levels for erythropoietin-treated EPAS1⫺/⫺
mice, although lactate levels were still mildly elevated relative to
EPAS1⫹/⫹ mice (2.0 ⫾ 0.5 versus 6.4 ⫾ 0.3, EPAS1⫹/⫹ versus
erythropoietin-treated EPAS1⫺/⫺ mice). The treatment protocol as
performed did not alleviate all aspects of the EPAS1⫺/⫺ phenotype;
the retinopathy and cardiac hypertrophy observed in EPAS1⫺/⫺
mice did not differ significantly after erythropoietin treatment (data
not shown). Hence, other aspects of the EPAS1⫺/⫺ phenotype likely
have additional molecular etiologies besides erythropoietin deficiency or are multifactorial in nature.
Discussion
We have determined that HIF-2␣ is a major regulator of renal
erythropoietin gene expression in vivo. A reduction in EPAS1 gene
dosage has profound effects for EPAS1⫺/⫺ mice and intermediate
effects for EPAS1⫹/⫺ mice on erythropoiesis. EPAS1⫺/⫺ mice
exhibit a moderate-to-severe anemia under ambient oxygen conditions whereas EPAS1⫹/⫺ mice have a mild anemia under these same
conditions. Molecular characterization revealed a marked depression of renal erythropoietin gene expression for EPAS1⫺/⫺ mice
maintained at room air and no significant differences in renal
erythropoietin gene expression for EPAS1⫹/⫺ mice under these
same conditions.
Although renal erythropoietin mRNA levels in EPAS1⫹/⫺ mice
were not significantly different from those of EPAS1⫹/⫹ mice under
ambient oxygen conditions, we observed a slight reduction in
resting hematocrits in EPAS1⫹/⫺ mice compared with EPAS1⫹/⫹
mice. Since erythropoietin mRNA levels are near the limits of
detection for RA samples even by real-time RT-PCR, we cannot
exclude a minor reduction in erythropoietin levels as being
responsible for this difference in EPAS1⫹/⫺ mice compared with
EPAS1⫹/⫹ mice. We also cannot exclude an effect of EPAS1
haploinsufficiency on paracrine erythropoietin production in the
bone marrow and resultant effects on definitive hematopoiesis as
a result.
The hypoxia treatment protocols revealed impaired induction of
erythropoietin gene expression in EPAS1⫺/⫺ as well as EPAS1⫹/⫺
mice. The absolute degree of erythropoietin induction by hypoxia
is dramatically blunted in EPAS1⫺/⫺ mice relative to EPAS1⫹/⫹
mice and mildly decreased in EPAS1⫹/⫺ mice. Blunting of erythropoietin gene expression in EPAS1⫹/⫺ mice occurs despite HIF-1␣
activation as suggested by HIF-1␣ stabilization. In the data
presented, no significant increases in HIF-2␣ protein were observed with nuclear extracts prepared from hypoxia-treated mice;
however, we have noted stabilization of HIF-2␣ protein in some
hypoxia-treated samples similar to previously reported results with
hypoxia-exposed rodents.51 HIF-2␣ protein stabilization may still
occur in localized regions of the kidney and would not be evident in
these experiments using whole kidney extracts. It is also possible
that HIF-2␣ signaling in these experiments involves mechanisms
other than protein stabilization.
BLOOD, 15 APRIL 2005 䡠 VOLUME 105, NUMBER 8
Long-term continuous hypoxia treatment reveals a blunted
erythropoietic response of EPAS1⫹/⫺ mice compared with EPAS1⫹/⫹
mice. Thus, haploinsufficiency for EPAS1, as with HIF-1␣,45 has
long-term effects on erythrocyte steady-state levels under these
conditions. In the kidney, as opposed to most other tissues of the
body, expression of HIF-1␣ and HIF-2␣ protein is mutually
exclusive.52,53 Based on in situ and immunohistochemical studies,
HIF-2␣ is expressed in renal interstitial and endothelial cells51,54
whereas HIF-1␣ is expressed predominantly in the tubular cells.54,55
Using a sensitive surrogate marker for EPAS1 gene expression, we
found EPAS1 gene expression in EPAS1⫺/⫺ mice is coincident with
the interstitial cells and vascular endothelial cells, cell types
previously implicated in erythropoietin production.21,23,25 The in
situ analyses presented in this study support earlier findings
indicating prominent expression of Epo in a subset of renal
interstitial cells. Since the in situ analyses also reveal expression of
EPAS1 in a subset of the renal interstitial cells, it is likely that
HIF-2␣ directly participates in Epo gene expression regulation.
The inability to establish primary erythropoietin-producing
renal interstitial fibroblast or other renal cell lines is indirect
evidence of an in vivo multicellular requirement for renal erythropoietin production. In conjunction with earlier studies examining
the regulation of erythropoietin gene expression by HIF-1␣,9 these
results reveal that in vivo renal erythropoietin production requires
multiple HIF members. It remains to be determined why both
HIF-1␣ and HIF-2␣ are essential for optimal renal erythropoietin
regulation and whether HIF-1␣ and HIF-2␣ alter erythropoietin
gene expression directly or indirectly.
We attribute the depressed erythropoietin gene expression in
EPAS1⫺/⫺ mice to the absence of functional HIF-2␣. It is unlikely
that HIF-1␣ signaling is impaired due to metabolic abnormalities in
EPAS1⫺/⫺ mice since renal Epo mRNA levels remain depressed in
EPAS1⫺/⫺ mice treated with exogenous erythropoietin (data not
shown). HIF-independent mechanisms may contribute to the
reduced basal level of erythropoietin expression seen in EPAS1⫺/⫺
mice under ambient oxygen conditions; however, these mechanisms cannot completely compensate in the complete absence of
HIF-2␣, either under ambient oxygen conditions or after exposure
to hypoxia. HIF-1␣–dependent pathways are likely activated in
EPAS1⫺/⫺ mice exposed to hypoxia, but activation of HIF-1␣–
dependent pathways does not result in efficient induction of Epo
gene expression.
Exogenous erythropoietin has the expected impact on erythropoiesis in EPAS1⫺/⫺ mice, including increased reticulocytosis, and
may also be responsible for increased thrombocytosis.56,57 Surprisingly, erythropoietin treatment also corrected the nonerythroid
deficiencies in EPAS1⫺/⫺ mice. The restoration of all hematopoietic
lineages of EPAS1⫺/⫺ mice after erythropoietin treatment supports
the contention that global hematopoietic maturation is dependent
on erythropoietin. Since both erythropoietin58 as well as the
erythropoietin receptor are expressed in early hematopoietic precursor/progenitor cells,59 erythropoietin may play an important role in
the early maturation process. Erythropoietin is also produced
locally in several tissues, including the bone marrow where it may
have paracrine or autocrine effects.60 The hematopoietic response
of EPAS1⫺/⫺ mice to exogenous erythropoietin may reflect efficient
replenishment of an otherwise absent paracrine source of erythropoietin.61 We cannot exclude an indirect effect of erythropoietin on
global hematopoiesis via erythropoietin-dependent induction of
other cytokines, synergistic interactions of erythropoietin with
other cytokines, or induction by erythropoietin of a downstream
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BLOOD, 15 APRIL 2005 䡠 VOLUME 105, NUMBER 8
HIF-2␣ REGULATES MURINE ERYTHROPOIETIN
signaling pathway that is affected in EPAS1⫺/⫺ mice but not
normally regulated by erythropoietin.62
Several cell types within the bone marrow express HIF-2␣,
including vascular endothelial cells, bone-lining cells, and resident
macrophages.47 Erythropoietin may originate from some of these
cell types. Consistent with the existence of a cytokine-producing
cell type that may originate from within the blood marrow
compartment, aged irradiated mice (2 years after transplantation)
that are recipients of bone marrow stem cells from EPAS1⫺/⫺ mice
exhibit lower hematocrit levels in comparison to irradiated recipient mice that received bone marrow stem cells from EPAS1⫹/⫹
mice (data not shown). Since these 2 groups of mice do not exhibit
differences in their hematocrit levels at younger ages, the long-term
differences in their erythrocyte masses may be due to timedependent turnover of long-lived erythropoietin-producing
EPAS1⫹/⫹ stromal cell types within the bone marrow. In this
scenario, endogenous EPAS1⫹/⫹ stromal cells in transplantrecipient mice that are a paracrine source of erythropoietin in the
bone marrow are eventually replaced by EPAS1⫺/⫺ stromal cells,
originating from the transplanted EPAS1⫺/⫺ stem cells, in which
erythropoietin production is impaired. However, other causes may
also contribute to this age-related phenotype.
The systemic lipid abnormalities otherwise seen in untreated
EPAS1⫺/⫺ mice, including the hepatosteatosis, are also reversed
with erythropoietin treatment of EPAS1⫺/⫺ mice, similar to that
observed after treatment with an antioxidant mimetic.46 The liver is
an extra-hepatic site of erythropoietin production in neonatal mice
3139
and in anemic adult mice. Since HIF members have been implicated in control of erythropoietin expression in extra-renal sites,49,63
it is possible that the pathologic findings observed in the livers of
EPAS1⫺/⫺ mice are in part due to absence of paracrine erythropoietin production. Moreover, other environmental factors such as
oxidative stress modulate erythropoietin gene expression.64-66 Therefore, molecular defects in oxidative stress responses, another aspect
of the EPAS1⫺/⫺-null phenotype,46 also may negatively impact
erythropoietin gene expression.
Erythropoietin, major antioxidant enzymes, and other HIF
target genes may constitute a battery of cellular factors that
alleviate oxidative stress associated with anemia, ischemia/reperfusion,
and hypoxia/reoxygenation. The importance of erythropoietin signaling
in pathologic states is an active source of investigation. Therapeutic
modulators that specifically augment HIF-2␣ activity may thus provide
adjunct or alternative therapies to exogenous erythropoietin treatments.
Future studies of EPAS1-deficient mice will have important translational
implications for disease states involving erythropoietin deficiencies.
Acknowledgments
We thank A. Das for excellent technical assistance, J. Repa for
assistance with real-time RT-PCR, D. Foster and S. McKnight for
generous support, and R. Bruick for critical review of this
manuscript.
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From www.bloodjournal.org by guest on August 3, 2017. For personal use only.
2005 105: 3133-3140
doi:10.1182/blood-2004-05-1695 originally published online
December 30, 2004
HIF-2α regulates murine hematopoietic development in an
erythropoietin-dependent manner
Marzia Scortegagna, Kan Ding, Quiyang Zhang, Yavuz Oktay, Michael J. Bennett, Michael Bennett,
John M. Shelton, James A. Richardson, Orson Moe and Joseph A. Garcia
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