From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
Metabolic Adaptation During Erythropoietin-Mediated Terminal Differentiation
of Mouse Erythroid Cells
By Hyun Dju Kim, Mark J. Koury, Sang Joon Lee, Jeong Hyok Im, and Stephen T. Sawyer
Metabolic development was examined in erythroid precursor
cells, which were isolated from the spleens of mice infected
with the anemia-inducing strain of Friend virus ( N A cells).
FVA cells undergo differentiation invitro from the proerythroblast stage through the reticulocyte stage over a 48-hour
period in the presence of erythropoietin. Concomitant with
marked decreases in cellular size and energy demand, metabolic capacities of both glycolysis and oxygen consumption
diminish after 48 hours in culture by 7- and 18-fold, respectively. Because the oxidative capacity decreases more than
glycolytic ability does, the metabolic machinery increasingly
shifts toward anaerobic metabolism. During the 48-hour
period of differentiation, the 2,3-diphosphoglyceric acid (DPG)
content per cell and 2,3-DPG mutase activity per cell increased eightfold and threefold, respectively. Freshly har-
vested FVA cells have adenosine triphosphate (ATP) levels of
7.23 f 2.52 pmol/lO’O cells or 3.76 -c 1.31 pmol/mL cell water
which are 12- or 2.3-fold higher, respectively, than the ATP
levels of mature red blood cells. In the course of FVA cell
differentiation, ATP content per cell decreases by fourfold,
but ATP concentration in cell water remains unchanged
because of a corresponding decrease in cellular size and
water content during differentiation. These studies show
that in the face of dramatic decreases in cell size and cellular
energy demand, terminally differentiating erythroid cells
maintain a constant ATP level by undergoing an involution of
their glycolytic machinery as well as by losing their aerobic
metabolic capacity.
o 1991by TheAmerican Society of Hematology.
T
nal differentiation of erythroid cells. In addition, we report
the metabolic data for mature erythrocytes from mice
because (1) the mature erythrocyte is the final product of
erythroid differentiation, and (2) much metabolic data has
not been previously reported for mouse erythrocytes. Compared with mature erythrocytes, freshly harvested FVA
cells show a very large glycolytic as well as an oxidative
capacity. Over the 48-hour period of differentiation with
Epo, both metabolic capacities decrease by an order of
magnitude. However, because the oxidative capacity decreases more than the glycolytic ability, the metabolic
reliance progressively shifts toward anaerobic metabolism
during differentiation. Despite the large changes in both
glycolysis and aerobic respiration during terminal differentiation of FVA cells, adenosine triphosphate (ATP) concentrations in cell water remain remarkably constant. These
findings show that, in the face of dramatic decreases in cell
size and cellular energy demand during terminal differentiation, the metabolic machinery of erythroid cells undergoes
a complex involution but maintains stable energy levels.
HE INVESTIGATION of metabolic changes which
occur in mammalian erythroid cells during their terminal stages of differentiation has long been hampered by the
lack of an appropriate in vitro culture model. However, the
recent availability of genetically engineered erythropoietin
(Epo) and the development of an erythroid precursor cell
system using the anemia-inducing strain of Friend erythroleukemia virus permits procurement of a large, relatively
homogenous population of proerythroblasts that will terminally differentiate in vitro in response to Epo. Erythroid
cells, which are obtained from the spleens of mice infected
with the anemia-inducing strain of Friend erythroleukemia
virus (FVA cells),’,*are either in or just before the proerythA routine isolation using two
roblast stage of de~elopment.~
spleens yields 5 x lo8FVA cells. FVA cells cultured in the
presence of Epo differentiate into basophilic erythroblasts
by 24 hours and are orthochromatic erythroblasts and
reticulocytes by 48 hours: while those cultured in the
absence of Epo fail to differentiate and soon die.4
A sequence of specific events has been documented in
FVA cells during the 48 hours of Epo-mediated
differentiation.’ RNA synthesis increases and DNA and
protein syntheses remain stable for the first 24 hours of
culture and then all decrease ~ h a r p l yThe
. ~ plasma membrane undergoes many structural alterations. Some prominent erythrocytic proteins, such as the globins,’X6 show
progressive increases in synthesis throughout the 48-hour
period.’ The anion transporter is accumulated in the
plasma membrane and the developing membrane skeleton
has progressive incorporation of spectrin and actin.’ Functionally, the cell maintains a high rate of iron accumulation
through complex variations in the surface numbers and
recycling pattern of transferrin receptors.’ While all of
these differentiation-related events are proceeding, FVA
cells are dividing with a significant decrease in individual
cell size and ultimately extrude their nuclei.’
Apart from one study that has examined glycolytic
enzymes during terminal erythroid differentiation using
regenerating murine erythroid cells,8 little is known about
the energetics of terminally differentiating erythroid cells.
We report here the metabolic changes accompanying termiBlood, Vol77, No 2 (January 15), 1991: pp387-392
MATERIALS AND METHODS
Cell procurement and culture. FVA cells were isolated and
cultured as previously described.*Briefly, splenic cells were obtained from CD,F, or BALBlc mice that had been infected 2 weeks
earlier with 1 x lo4 spleen focus-forming units’ of the anemia~
~
~~
From the Department of Pharmacology, University of MissouriColumbia; and the Department of Medicine, Vanderbilt University,
Nashville, TN.
Submitted January 2,1990; accepted September 25, 1990.
Supported in part by National Institutes of Health Grants DK33456,
Dx31513, and DK39781.
Address reprint requests to Hyun Dju Kim, PhD, Depament of
Pharmacology, University of Missouri-Columbia, Columbia, MO
65212.
The publication costs of this article were defrayed in part by page
charge payment. This article must therefore be hereby marked
“advertisement” in accordance with 18 U.S.C.section I734 solely to
indicate this fact.
1991 by The American Society of Hematology.
0006-4971l91/7702-O010$3.0010
387
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
388
inducing strain of Friend leukemia virus. These cells were separated by velocity sedimentation at unit gravity and those cells
sedimenting at 6 mm or more per hour were collected and pooled
as the starting (0 hour) cells. Erythrocytes and late-stage erythroblasts are essentially absent from the 0-hour cell population, which
consists mainly of proerythroblasts. The EVA cells were cultured at
37°C in humidified air plus 5% CO, at 1 X 106 cells/mL in Iscove’s
modified Dulbecco’s medium (IMDM) supplemented with 30%
(vol/vol) fetal bovine serum (FBS), 1% deionized bovine serum
albumin (BSA), 0.1 mmol/L a-thioglycerol, and 0.15 U/mL of
recombinant human Epo (AmGen, Thousand Oaks, CA). The cells
were collected from culture at various times of incubation, washed
once in IMDM or incubation medium, and then used for the
various assays as described below.
Blood was either taken from heart puncture of ether-anesthetized adult mice or collected into heparinized containers after
severing the jugular vein. Red blood cells (RBCs) were washed
several times by alternate suspension and centrifugation. The buffy
coat was removed in each cycle by aspiration.
Determination of cellular water content and dry weight. Cells
were suspended in 1.5-mL microfuge tubes in 1 mL of IMDM
containing 500,000 cpm of ‘Wabeled BSA. Cell number was
determined by counting a IO-pL aliquot of the suspension in a
hematocytometer (at least 200 cells were counted for every sample
and counts were made in triplicate) and then the cells were
pelleted by centrifugation and the supematant medium was removed. The tube containing the wet pellet was immediately
weighed and then desiccated in a 15-pm Hg vacuum at 25°C for 24
hours. The tube containing the dried pellet was weighed and
125
I-albumin in the trapped extracellular water was measured in a y
counter. The water content was determined by the difference in
weight between the wet pellet and dry pellet minus the trapped
extracellular water. The dry pellet was directly weighed after the
‘=I content had been measured.
Determination of 0, consumption. 0, consumption was measured using an Instech (Nordham, PA) polarographic oxygen
electrode mounted in a water-jacketed 0.6-mL chamber at 37°C.
The electrode was calibrated using air-saturated water at 37°C
(total 0.64 pmoles of 0, in the chamber) and then sodium
dithionite was added to the chamber to determine the electrode
reading in the absence of 0,. The output from the electrode was
recorded on a chart recorder and the rate of oxygen consumption
was determined from the slope on the chart in units of micromoles
of 0, per minute. FVA cells taken immediately after isolation or
cells cultured for either 24 hours or 48 hours in the presence of Epo
were washed once in incubation medium (140 mmol/L NaCl, 5
mmol/L KCl, 1 mmol/L CaCl,, 1 mmol/L MgCl,;-’ 5 m m o m
glucose, 0.1% BSA, 1 mmol/L Na phosphate, and 10 mmol/L
Tris-HEPES p H 7.4) and then resuspended in incubation medium,
which was air saturated at 37°C. At least 5 X lo7 cells were
introduced into the chamber and 0, consumption was measured.
0, consumption varied linearly with time and cell number. Mature
blood erythrocytes prepared in the same manner had no detectable
0, consumption and KCN totally blocked 0, consumption in all
cells tested.
Determination of glucose consumption, lactate production, 5 3 diphosphoglyceric (DPG) mutase activily and content of 2,3-DPG and
ATP. FVA cells or RBCs were incubated in the same medium,
which was used for the determination of 0, consumption described
above, but containing 6 to 20 pCimL of D-[Z’H]-glucose. At
various times, samples were taken, from which perchloric acid
extracts were prepared.
Glucose consumption rate was measured according to the
procedure of Neely et all0 as modified by Kim.” Briefly, the
production of 3H,0 from [t-’H]-glucose catalyzed by phosphoglu-
KIM ET AL
cose isomerase reaction was used as the measure of glycolytic flux.
Approximately 0.2 to 0.4 mL perchloric acid (PCA) extract was
adsorbed onto a 0.7 x 1.5-cm Dowex-1 (1 X 4 - 200) borate
column and eluted with water. ’H,O in the eluate was determined
by liquid scintillation spectroscopy, and glucose consumption rate
was calculated from the ’H,O production and the specific activity of
glucose.
Lactate in PCA extracts was measured by the method of
Lundholm et al.” 2,3-DPG mutase activity was determined by the
method of Beutler” on FVA cell lysates, which were prepared by
the method of Nijhof et a1.8
PCA extracts of FVA cells and mature RBCs were used to
measure 2,3-DPG content employing a modification of the fluorometric method of Keitt.14
ATP in neutralized PCA extract was determined by high
performance liquid chromatography (HPLC) technique according
to the procedure published elsewhere.”
RESULTS
Differentiating erythroblasts in the presence of Epo
undergo remarkable reductions in cell size. Figure 1 shows
measurements of intracellular water (femtoliters per cell),
dry weight (picogramsper cell), and water content (percentage of total cell weight) of FVA cells in culture. Intracellular water content of 192 -C 8 =/cell in freshly harvested
erythroblasts is decreased by approximately 17% during the
20c
d
I“
15C
[L
4I
3YlOC
dd
V
4
50
z
A o
70
60
-I
I
50
(3-
Izz 4c
f\
> x30
(L
0
20
10
B c
n
L
24
34
45
TIME IN CULTURE, h
RBC
Fig 1. Changes in water content in FVA cells in culture with Epo.
The splenic erythroblasts were cultured as described in Materials and
Methods and samples were taken at times indicatedfor the determination of (A) intracellular water (femtolkers per cell); (B) dry weight
(picograms per cell); and (C) water content (percent). Results are
given as the mean -c standard deviation from five t o nine determinations.
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
ERYTHROPOIETIN-DEPENDENTMETABOLISM
389
Table 1. Glycolytic Capacity of FVA Cells and Mature RBCs
Culture
Time (h)
Lactate
Production
Glucose
Consumption
Ratio of
Lactate Produced/
Glucose Consumed
(~mol/lO'oceHsx h)
45.84f 10.06 (n = 7) 128.83f 43.13
23.43 f 2.32 (n = 3) 68.80 f 15.43
6.60 f 2.58 (n = 7) 17.34 f 5.81
Mature RBC 0.69 f 0.1 1 (n = 12) 1.77? 0.37
0
24
48
2.84 f 0.92
2.94 f 0.66
3.11 f 1.96
2.58 f 0.43
Results are given as mean f standard deviation with the number of
determinations (n) given in parentheses.
first 24 hours in culture followed by a larger decrease to
27% of the original value after 45 hours in culture. The dry
weight undergoes a similar decrease. The volume of a single
mouse RBC was approximately 40 Wcell, which is slightly
less than the values reported in the literature of 50 Wcell."
Having determined water content of FVA cells, the concentration of metabolites can be expressed in terms of their
concentrations in cell water.
Concomitant with morphologic changes, glycolyticrate of
FVA cells decreases dramatically during differentiation.
The results are summarized in Table 1. We found that
glycolytic capacity of freshly harvested FVA cells is more
than 66-fold higher than that of mature RBCs when
expressed in terms of the rate per cell. After 2 days in
culture, glucose consumption rate is decreased to approximately 14% of the original cell capacity but is still ninefold
higher than that of mature cells. Because FVA cells become
highly unstable after a 48-hour culture,'6 no determination
was made on cultures later than 48 hours. Unexpectedly,
FVA cells produced more lactate than can be accounted for
by the glucose consumption.
Figure 2 shows a time course of glucose consumption,
and the ratio of lactate produced to glucose consumed by
adult mouse RBCs. In the course of the isolation of RBCs,
glucose was temporarily withheld from the cells. Upon
re-exposure to glucose at 37"C, glucose consumption commences at a low rate during the first 15 minutes followed by
n
I
W
a significantly higher rate that then remains linear through
a 3.5-hour incubation. The initial metabolic adjustment to
glycolytic carbon flow is reflected by the dramatic changes
in the ratios of lactate produced to glucose consumed seen
in the early phase of the incubation. After 1-hour incubation, ratios reach a steady state but remain slightly larger
than 2.0.
Table 2 compares aerobic and anaerobic metabolism in
differentiating FVA cells. In addition to having robust
glycolytic ability, freshly harvested FVA cells are endowed
with a high capacity for oxygen consumption, as expected.
In experiments performed with glucose as the sole exogenous substrate, it can be shown that approximately 65% of
total glucose metabolism takes place by the aerobic pathway. The oxidative capacity of FVA cells also decrease
rapidly in parallel with the time course of glycolytic reduction in that by 48 hours in culture, only 6% of the original
cellular oxidative capacity remains. However, the relative
reliance on anaerobic metabolism increases with differentiation such that by 48 hours in culture, only 44% of glucose
metabolism occurs by the aerobic pathway. As in other
mammalian RBCs, mature RBCs of mice are unable to
consume oxygen.
Table 3 shows 2,3-DPG levels and 2,3-DPG mutase
activities of erythroid cells undergoing differentiation in the
presence of Epo. 2,3-DPG in the splenic erythroblasts is
barely detectable, as reported earlier." However, in the
course of a 48-hour culture 2,3-DPG content per cell
increases by approximately eightfold, despite the decrease
in the metabolic rate. When expressed in terms of its
concentration in cell water, 2,3-DPG increases by more
than 30-fold. 2,3-DPG content per cell or 2,3-DPG concentration in cell water of FVA cells in culture for 48 hours
amounts to 23% or 12% of 2,3-DPG levels found in mature
RBCs, respectively. The increase in 2,3-DPG levels occurs
concomitant with an increase in 2,3-DPG mutase activity in
the FVA cells. 2,3-DPG mutase activity per milliliter of cell
water increased almost 12-fold over the 48 hours of their
differentiation in vitro. However, the activity of the enzyme
per milliliter of cell water achieved only 18%of the activity
of mature RBCs (38% for activity per cell).
3 .O
Table 2. Aerobic and Anaerobic Metabolism in
DifferentiatingR I A Cells
5 .O
0
w w
mnr
0 3
u m
3 2
J
O
0 m u
Time
(h)
F
U \
U
W
O
+ w
u u
-13
u o
u o
_lu
a
1.o
1.0
0
1
2
3
4
0 .o
HOURS
Fig 2. Glucose consumption and ratio of lactate produced to
glucose consumed by mouse RBCs. Blood from four to six mice was
pooled and RBCs were isolated as described in Materials and Methods. Glucose consumption rate was measured by the production of
'H,O from [2-H]-glucose in PCA extracts of a 10% cell suspension.
Results are given as the mean ? standard deviation from five
determinations.
0
24
48
RBC
0,
Consumed*
Glucose
Converted
to C0,t
Glucose
Converted
to Lactate*
(Fmol/lO'o
cells x h)
703 86
117.2
300 f 17
50.0
39 * 8
6.6
0
0
Anaerobic
Metabolism
(% of total)
64.1
34.4
8.4
0.9
35
40
56
100
'0, consumption measured with oxygen electrode as described in
Materials and Methods. Data are mean f standard deviation of three
separate experiments.
tGlucose conversion calculated from 0, consumption (CO, produced
by hexosemonophosphate shunt is not included), assuming no other
energy source.
*Glucose conversion calculated from lactate levels in Table 1,
assuming no other energy source.
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
390
KIM ET AL
Table 3. 2,3-DPG Levels and 2.3-DPG Mutase Activity in R I A Cells During Culture With Epo
2.3-DPG Mutase Activity*
2.3-DPG
Erythroblasts
Culture (h)
prnol/lO'o Cells
pmolImL Cell Water
pmolllO'OCellslh
pmol/mL Cell Waterlh
0
24
48
0.09
0.46 f .08
0.75 k .08
3.2 5 0.3
0.05
0.29 5 ,005
1.50 ? .02
13.33 k 1.25
3.48 k 2.48
4.85 2 0.44
10.65 2 0.86
28.18 -t 6.47
1.81 2 1.29
3.11 f 0.28
21.30 2 1.72t
117.4 5 26.9t
-
RBC
Mean f standard deviation, n = 3.
*Activity expressed as rate of micromoles of 1.3-DPG converted to 2,3-DPG.
tn = 4.
ATP levels in differentiating FVA cells are shown in Fig
3. Freshly harvested FVA cells have ATP levels of 7.23 f
2.52 Fmol/lO1' cells or 3.76 f 1.31 p.mol/mL cell water. By
comparison, mature RBCs have ATP content of 0.58 f 0.12
p,moI/10" cells (n = 16). Thus, in terms of ATP content per
cell, FVA cells have 12-fold higher level of ATP than do
mouse RBCs, but only twofold higher when expressed in
terms of ATP concentration in cell water. Although ATP
content per cell decreases by fourfold after 48 hours in
culture, ATP concentrations in cell water do not decrease.
This apparent discrepancy between ATP content and
concentration stems from the different water content of
FVA cells shown in culture as shown in Fig 1. After 48
hours in culture, FVA cells that have now differentiated to
the reticulocyte stage possess ATP concentrations that are
twofold to threefold higher than mature RBCs. Thus, there
is a significant decrease in ATP concentration in cell water
in differentiating erythroid cells that seems to occur between the reticulocyte and erythrocyte stages.
DISCUSSION
Little is known about the development of the metabolic
machinery during erythroid differentiation. Because of the
lack of an appropriate model for studying metabolism in
differentiating erythroprogenitor cells, investigations of
metabolic development have largely been limited to the
transition from the reticulocyte to the erythrocyte stage.
The availability of FVA cells derived from mice affords a
unique opportunity to delineate the metabolic changes
accompanying erythroid differentiation. The FVA cells
12.0
1
1
5 .O
4 .O
3 .O
2.0
1.o
0 .o
HOURS OF CULTURE
Fig 3. ATP levels of FVA cells in culture with Epo. Results are given
as the mean ~ f -standard deviation of FVA cells at 0 hour (n = 4). FVA
cells at 48 hours; (n = 6). and RBC (n = 16).
which accumulate in the spleens of mice infected with the
anemia-inducing strain of Friend erythroleukemia virus are
recognized as colony-forming units-erythroid (CFU-E) and
proerythroblasts. That these FVA cells would require a
high energy supply from metabolism is evident from their
extensive protein synthesis and attendant cell division.
Indeed, FVA cells were found to have an extremely active
glycolytic and oxidative metabolism relative to mature
erythrocytes.
With respect to glycolytic capacity, we found that both
FVA and mature RBCs display higher ratios of lactate
produced to glucose consumed than the anticipated ratio of
2.0. To enhance the sensitivity of glucose consumption
measurements, we have used the measurement of 3H,0
production from D-[2-3H]-glucosecatalyzed by phosphoglucose isomerase. A potential complication of this method is
the breakdown of glycogen, which we did not measure,
resulting in the dilution of the specific activity of glucosedphosphate pool. If FVA cells have a glycogen store, which is
degraded in glucose media, this could have conceivably
caused a slight underestimation of glycolytic rate in FVA
cells. On the other hand, the glycolytic rate measured in
mature RBCs remains highly reliable because mature
mammalian RBCs are practically devoid of glycogen.18
From a comparative point of view, mouse erythrocytes have
a glycolytic capacity that falls into the middle of the range
varying from nonglycolytic pig RBCs to the most prolific
glucose users, rat RBCs.19 Unlike lactate production, glucose consumption exhibits a lag period of a few minutes in
which glucose is consumed at a much reduced rate (Fig 2).
As a result, ratios of lactate produced to glucose consumed
are high in the early phase of incubation but decrease
gradually, approaching metabolic steady state after a prolonged period of incubation. Although the reason for high
glycolyticratios observed in mouse erythroid cells (Table 1)
is not entirely clear, it should be recalled that glycolytic
rates were measured after 1-hour incubation. Thus, it is
conceivable that a prolonged incubation may lead to lower
ratios of lactate produced to glucose consumed. Apparently, endogenous substrates and phosphorylated glycolytic
intermediates are depleted in the presence of glucose at
least during early phase of incubation. This finding raises a
possibility that more than glucose is ordinarily used in vivo
by mouse cells as has been postulated for human RBCs."
With respect to oxidative metabolism, it is of interest to
note that relative partition between aerobic and anaerobic
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
ERYTHROPOIETIN-DEPENDENT METABOLISM
391
metabolism shows a major reliance on oxidative metabolism
(Table 2). In the course of FVA cell differentiation, both
glycolytic and oxidative capacities decrease rapidly. The
rapid decline in glycolytic ability seen in differentiating
FVA cells is consistent with the previously observed marked
decreases in the activities of several glycolytic enzymes
including hexokinase, phosphofructokinase, aldolase, enolase, pyruvate kinase, and G-6-P dehydrogenase during the
terminal differentiation of regenerating CFU-E.' While the
relative reliance of FVA cells on aerobic metabolism
decreases with time during differentiation, dependency on
anaerobic metabolism increases. By a 48-hour culture,
when most FVA cells have differentiated to the reticulocyte
stage, anaerobic metabolism accounts for 56% of glucose
utilization. This finding is in good agreement with previous
findings on rabbit reticulocytes in which approximately 55%
of glucose carbon flow takes place through anaerobic
metabolism.z0
During the reductions in cell size and diminished energy
need in erythroid differentiation not all metabolic parameters decrease. The activity of 2,3-DPG mutase and the level
of 2,3-DPG, which are at the level of detection in freshly
harvested FVA cells, increase with culture. By a 48-hour
culture, 2,3-DPG content per cell or its concentration in
cell water amounts t o 23% or 12% of 2,3-DPG levels found
in mature RBCs, respectively. Similar results showing an
increase in 2,3-DPG levels were reported in Friend leuke-
mic cells that were induced to differentiate by dimethyl
sulfoxide (DMSO)?'322
In keeping with this observation, the
biosynthesis of 2,3-DPG mutase is initiated only after the
induction of Friend leukemic cell differentiation by DMSO,'*
and increases markedly during differentiation of regenerating CFU-E.' During rabbit reticulocyte maturation, 2,3DPG is reported to either increaseB or not change.% We
interpret the increases in 2,3-DPG levels and 2,3-DPG
mutase activity toward the very high levels found in RBCs
as showing that differentiating FVA cells are not simply
restricting their energy-related metabolism, but are rather
undergoing specific developmental changes that will ultimately result in the metabolic machinery characteristic of
mature erythrocytes.
Of particular interest pertaining to energetics of FVA
cells is the finding that ATP content per cell decreases but
not ATP concentrations in cell water during Epo-dependent differentiation. Apparently, throughout this developmental program in which the metabolic machinery is being
remodeled, the energy demands of the cells are met by a
steady-state maintenance of ATP concentration as the cells
decrease in size.
ACKNOWLEDGMENT
The unstinting technical assistance of Jane Burnett and Dr
Sarvandaman Rana is gratefully acknowledged.The authors thank
Judy Richey and Kristin Nelson for typing of the manuscript.
REFERENCES
1. Koury MJ, Sawyer ST, Bondurant M C Splenic erythroblasts
in anemia-inducing Friend disease: A source of cells for studies of
erythropoietin-mediated differentiation. J Cell Physiol 121:526,
1984
2. Sawyer ST, Koury MJ, Bondurant M C Large-scale procurement of erythropoietin-responsive erythroid cells: Assay for biological activity of erythropoietin. Methods Enzymol 147340,1987
3. Koury ST, Koury MJ, Bondurant MC: Morphologicalchanges
in erythroblasts during erythropoietin-inducedterminal differentiation in vitro. Exp Hematol16:758,1988
4. Koury MJ, Bondurant M C The maintenance by erythropoietin of viability, proliferation and maturation of murine erythroid
precursor cells. J Cell Physioll3765,1988
5. Koury MJ, Bondurant MC, Atkinson JB: Erythropoietin
control of terminal erythroid differentiation: Maintenance of cell
viability, production of hemoglobin, and development of the
erythrocyte membrane. Blood Cells 13:217,1987
6. Bondurant MC, Lind RL, Koury MJ, Ferguson ME: Control
of globin gene transcription by erythropoietin in erythroblastsfrom
Friend virus-infected mice. Mol Cell Biol5:675, 1985
7. Sawyer ST, Krantz SB: Transferrin receptor number, synthesis, and endocytosis during erythropoietin-induced maturation of
Friend virus-infected erythroid cells. J Biol Chem 261:9187, 1986
8. Nijhof W, Wierenga PK, Staal GEJ, Jansen G: Changes in
activities and isozyme patterns of glycolytic enzymes during erythroid differentiationin vitro. Blood 64:607,1984
9. Axelrad AA, Steeves RA:Assay for Friend leukemia virus: A
rapid quantitative method based on enumeration of macroscopic
spleen foci in mice. Virology 24:513,1964
10. Neely JR, Denton RM, England PS, Randle PS: The effects
of increased heart work on the tricarboxylate cycle and its interactions with glycolysis in the perfused rat heart. Biochem J 128:149,
1972
11. Kim HD: Is adenosine a second metabolic substrate for
human red blood cells? Biochim Biophys Acta 1036:113,1990
12. Lundholm L, Mohme-Lundholm M, Vamos N: Lactic acid
assay with L(+) lactic acid dehydrogenase from rabbit muscle.
Acta Physiol Scand 58:243,1961
13. Beutler E: Red Cell Metabolism (ed 2). Philadelphia, PA
Grune and Stratton, 1975, p 54
14. Keitt AS: Reduced nicotinamide adenine dinucleotidelinked analysis of 2,3-diphosphoglycericacid: Spectrophotometric
and fluorometricprocedures. J Lab Clin Med 77:470,1971
15. Wintrobe MM, Lee GR, Boggs DR, Bithell TC, Athens JW,
Foerster J (eds): Clinical Hematology (ed 7). Philadelphia, PA,
Lea and Febiger, 1974, p 1807
16. Koury MJ, Bondurant MC, Rana SS: Changes in erythroid
membrane proteins during erythropoietin-mediatedterminal differentiation. J Cell Physiol133:438,1987
17. Kim HD, Tsai YS, Lee SJ, Im JH, Koury MJ, Sawyer S T
Metabolic development in erythropoietin-dependent maturation
of erythroid cells, in Brewer GJ (ed): The Red Cell 7th Ann Arbor
Conference, Progress in Clinical and Biological Research, ~01319.
New York, NY,Liss, 1989, p 491
18. Moses SW, Bashan N, Gutman A Glycogen metabolism in
the normal red blood cell. Blood 40:836,1972
19. Kim HD: Postnatal changes in energy metabolism of mammalian red blood cells, in Agar NS, Board PG (eds): Red Blood
Cells of Domestic Mammals. New York, NY, Elsevier Science,
1983, p 339
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
392
20. Siems W, Muller M, Dumdey R, Holzhutter HG, Rathmann
J, Rapoport SM: Quantification of pathways of glucose utilization
and balance of energy etabolism of rabbit reticulocytes. Eur J
Biochem 124567,1982
21. Yeoh GCT Levels of 2,3-diphosphoglycerate in Friend
leukaemic cells. Nature 285:108,1980
22. Narita H, Yanagawa S, Sasaki R, Chiba H: Induction of
KIM ET AL
2,3-bisphosphoglycerate synthase in Friend leukemia cells. Biochem Biophys Res Commun 103:90,1981
23. Ohyama H, Minakami S: Studies on erythrocyte clygoclysis.
V. Change of the glycolytic intermediate pattern of reticulocytes
during maturation. J Biochem 61:103,1976
24. Bartlett GR: Phosphate compounds of rat erythrocytes and
reticulocytes. Biochem Biophys Res Commun 701055,1976
From www.bloodjournal.org by guest on June 18, 2017. For personal use only.
1991 77: 387-392
Metabolic adaptation during erythropoietin-mediated terminal
differentiation of mouse erythroid cells
HD Kim, MJ Koury, SJ Lee, JH Im and ST Sawyer
Updated information and services can be found at:
http://www.bloodjournal.org/content/77/2/387.full.html
Articles on similar topics can be found in the following Blood collections
Information about reproducing this article in parts or in its entirety may be found online at:
http://www.bloodjournal.org/site/misc/rights.xhtml#repub_requests
Information about ordering reprints may be found online at:
http://www.bloodjournal.org/site/misc/rights.xhtml#reprints
Information about subscriptions and ASH membership may be found online at:
http://www.bloodjournal.org/site/subscriptions/index.xhtml
Blood (print ISSN 0006-4971, online ISSN 1528-0020), is published weekly by the American
Society of Hematology, 2021 L St, NW, Suite 900, Washington DC 20036.
Copyright 2011 by The American Society of Hematology; all rights reserved.