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Blood, Vol. 94 No. 1 (July 1), 1999:
pp. 74-86
Genetic Evidence for an Additional Factor Required for
Erythropoietin-Induced Signal Transduction
By
Sarah L. Gaffen,
Stephen Y. Lai,
Gregory D. Longmore,
Kathleen D. Liu, and
Mark A. Goldsmith
From the Gladstone Institute of Virology and Immunology, San
Francisco, CA; the Department of Medicine, School of Medicine,
University of California, San Francisco; and the Departments of
Medicine and Cell Biology, Washington University School of Medicine, St
Louis, MO.
 |
ABSTRACT |
Erythropoietin (EPO) and its receptor (EPOR) are required for the
development of mature erythrocytes. After binding of ligand, the EPOR
activates a variety of signaling pathways that ultimately control
cellular proliferation, survival, and specific gene expression. Although erythroid progenitors appear to be the principal
EPO-responsive cell type in vivo due to the restricted expression of
the EPOR, many growth factor-dependent cell lines expressing the EPOR
can respond to EPO by activating many or all of these pathways. In the
present study, we have identified a cellular context (the interleukin-2
[IL-2]-dependent HT-2 line) in which the EPO stimulation of the EPOR
fails to support cellular proliferation, STAT-5 induction, or MAPK
activation, despite efficient phosphorylation of the EPOR and JAK2 and
inhibition of apoptosis after withdrawal of IL-2. Interestingly, when
we fused HT-2 cells expressing the EPOR with Ba/F3 cells in a
complementation assay, the resulting hybridomas proliferated and
potently activated STAT-5 and MAPK in response to EPO. These data
indicate that an unidentified cellular factor is needed to mediate
signaling by the EPOR. Moreover, Ba/F3 cells apparently express this
factor(s) and somatic fusions can, therefore, confer EPO-responsiveness
to HT-2 cells that lack this factor.
© 1999 by The American Society of Hematology.
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INTRODUCTION |
MAMMALIAN BLOOD CELL differentiation
depends on cellular proliferation and crucial cell-fate decisions that
are coordinated by a family of hematopoietic cytokines, particularly
erythropoietin (EPO). EPO, a 34-kD cytokine, is produced by the kidney
under conditions of low oxygen tension and acts on erythroid
progenitors via a specific erythropoietin receptor (EPOR).1
EPO plays a critical role in mammalian hematopoiesis, because it is
required for the development of mature erythrocytes, and animals
deficient in this cytokine or its receptor die early in fetal
development due to severe anemia.2,3 In addition, EPO is
used extensively in the clinic to treat anemia associated with EPO
deficiency without serious side effects. Numerous signaling molecules
and pathways have been linked to the EPOR, the combined effects of
which generally include cellular survival and/or proliferation,
differentiation, and specific gene expression, ultimately leading to
erythroid maturation and expansion of the red blood cell
compartment.4
The EPOR belongs to the cytokine receptor superfamily, members of which
are characterized by conserved structural motifs in the extracellular
domain including four conserved cysteine residues and a
membrane-proximal WSXWS motif.5 Like other members of this
family, the cytoplasmic tail of the EPOR contains no known enzymatic
activity, but instead associates with various second messenger
molecules to effect signal transduction. EPO stimulation is thought to
trigger homodimerization of EPOR subunits, initiating a cascade of
signal transduction events. Interestingly, a mutation within the EPOR
extracellular region of cysteine for arginine-129 results in
constitutive dimerization through a disulfide bridge between receptor
subunits that causes ligand-independent signaling.6-9 In
addition, synthetic peptide dimers that cause dimerization of the EPOR
function similarly to EPO, and crystal structures of the EPOR
co-complexed with these peptides show that the receptor adapts a
symmetrical twofold dimer configuration.10 Homodimerization of the receptor results in the tyrosine phosphorylation of a variety of
intracellular substrates, including the EPOR itself.11-13
Tyrosine phosphorylation of the receptor and many of its associated
signaling molecules is mediated by the Janus kinase JAK2, which
associates constitutively with EPOR monomeric subunits and is activated
upon ligand binding by autophosphorylation.14 One
consequence of JAK2 activation is the recruitment of the
SH2-domain-containing transcription factors STAT-5A and
STAT-5B15,16 This recruitment leads to tyrosine
phosphorylation of STAT-5A/5B and dimerization via reciprocal
SH2-phosphotyrosyl interactions, followed by rapid nuclear import of
STAT-5 dimers, resulting in transcription of EPO-responsive genes such
as oncostatin-M17 and the signaling inhibitor
CIS.18 Studies using dominant negative STAT-5 mutants or
EPOR mutants defective in STAT-5-association domains have linked the
activation of STAT-5 to cellular proliferation and/or differentiation in certain cultured cell lines.19-22 However, mice
deficient in STAT-5A, STAT-5B, or both exhibit normal
erythropoiesis,23-25 indicating that STAT-5 is not
essential for EPOR function in vivo. Tyrosine phosphorylation of the
receptor also leads to the recruitment of other SH2-containing proteins
such as Shc, PI3K, SHIP, Vav, and PLC , and the tyrosine phosphatases
SHP-1 and SHP-2.4,26 Therefore, the EPOR activates a
striking array of signaling pathways that are characteristic of the
cytokine receptor superfamily and act in concert to coordinate the
physiological effects of this cytokine.
The study of cytokine signaling pathways has been aided greatly by the
availability of cytokine-dependent cell lines. For example, in the
interleukin-3 (IL-3)-dependent cell lines 32D (pro-myeloid) and Ba/F3
(pro-B), ectopic expression of the EPOR allows growth in EPO in place
of IL-3. Similarly, transfection of the IL-2R chain into these cell
lines renders them responsive to IL-2.27 These cell lines
have been valuable in molecular mapping studies of the EPOR, allowing
specific signaling events to be linked to particular subdomains of the
EPOR cytoplasmic tail. In addition, such studies suggest that there is
tremendous overlap in the array of signals used by various cytokine
receptors. For example, in the 32D and Ba/F3 cell lines, the EPOR,
IL-2R, and IL-3R complexes all activate JAKs, STAT-5A/B, the MAPK
cascade, c-fos gene expression/DNA binding, bcl-2 and
bcl-xL gene expression, and inhibit apoptosis. The
present report describes a cell line in which signaling through the
EPOR is markedly but selectively impaired, while signaling through the
IL-2R is intact. Although the EPOR can activate JAK2 and exert a potent
anti-apoptotic effect in this cellular background, it fails to activate
STAT-5 or MAPK, or to support EPO-dependent proliferation, despite the
ability of the IL-2R to engage these pathways efficiently. Genetic
complementation by somatic fusion to permissive cells rescues
signaling, indicating that a missing factor(s) can be provided in
trans to restore full EPOR function. Thus, this cell line
provides a valuable system in which to identify previously unrecognized
components of the EPOR signaling pathway.
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MATERIALS AND METHODS |
Cell culture, preparation of cell lines, and cytokine stimulations.
HT-2 cells (American Type Culture Collection [ATCC], Rockville, MD)
were maintained in RPMI 1640 medium (Life Technologies, Gaithersburg,
MD), 10% fetal bovine serum (FBS), 2 mmol/L glutamine, 0.05 mmol/L
2-mercaptoethanol and 1 nmol/L recombinant IL-2 (generously supplied by
Chiron Corp, Emeryville, CA). HT-2.EPOR.3 cells were maintained in this
media with 1 mg/mL G418 (Life Technologies). HT-2EPO cells were
grown in this media with EPO (5 U/mL, a gift of Ortho Biotech, Raritan,
NJ) in place of IL-2. HT-2 transfectant cell lines were prepared as
described.28,29 Briefly, cells were electroporated at 300 V/950 µF and cultured without selective media for 24 hours. They were
transferred to selective media and monoclonal lines obtained by
limiting dilution. 32D and Ba/F3 cells were cultured in RPMI 1640, 10%
FBS, 2 mmol/L glutamine, 0.05 mmol/L 2-mercaptoethanol, and 10%
WEHI-3B conditioned medium (CM) as a source of IL-3. 371.2 cells are
32D cells that stably express a transgene encoding IRS-1,30
and were also maintained in this media. HCD57 cells were cultured in
Iscove's medium (Life Technologies), 20% FBS, 2 mmol/L glutamine, and
1 U/mL EPO. For cytokine stimulations, cells were washed twice in
phosphate-buffered saline (PBS), stripped with a 30-second incubation
in 10 mmol/L sodium citrate/140 mmol/L NaCl, and incubated in
serum- and growth factor-free RPMI 1640 medium with 1% bovine serum
albumin (BSA fraction V; Sigma, St Louis, MO) for 2 to 4 hours
(immunoprecipitations and nuclear extracts) or in growth factor-free
medium for 12 to 15 hours (mRNA preparations) or 40 hours
(apoptosis assays). Stimulations were for indicated time periods with
recombinant human IL-2 (10 nmol/L), EPO (0.01 to 100 U/mL), or IL-3
(10% WEHI-3B-CM).
Proliferation assays.
Conventional 24- to 48-hour [3H]thymidine incorporation
assays and transient proliferation assays were performed as described previously.31 For the transient proliferation assay,
107 cells were electroporated with 20 µg DNA at 300 V/950
µF and incubated in 1 nmol/L IL-2 for 24 hours. Cells were washed
twice with PBS and incubated in 96-well plates with IL-2 (10 nmol/L), EPO (50 U/mL), or no cytokine. [3H]thymidine
incorporation was measured between days 6 and 16. Data are presented as
averages of triplicate samples expressed as a percentage of
[3H]thymidine incorporation of cells treated with 10 nmol/L IL-2 (HT-2 cells) or 10%WEHI-3B-CM (32D and 371.2 cells).
Apoptosis assays.
Cells, 2 to 5 × 106, were incubated for 40 hours in
medium without cytokines, or stimulated with EPO (50 U/mL) or IL-2 (10 nmol/L). Cells were harvested, washed with PBS/2.5 mmol/L CaCl/2% FBS
(Wash Buffer), and incubated with a 1:1,000 dilution of Annexin-GFP (generously provided by Dr Joel Ernst, University of California, San
Francisco32) on ice for 10 minutes. Cells were washed twice with Wash Buffer and resuspended in 1 mL Wash Buffer and 1 µL of 50 µg/mL Propidium Iodide (Pharmingen, San Diego, CA). Samples were
analyzed on a Becton Dickinson FACScan using the Cellquest computer program.
MAPK assay.
Cells, 10 to 20 × 106, were rested for 6 hours and
stimulated with no cytokines, IL-2 (10 nmol/L), EPO (50 U/mL), or
murine IL-4 (10 ng/mL). In vitro MAPK assays were performed using an MAPK assay kit (New England Biolabs, Beverly, MA). In short, cells were
lysed after stimulation with indicated cytokines. Erk1/Erk2 MAPK was
immunoprecipitated using an antiphospho-MAPK antibody. Kinase assays
were performed using an Elk1-glutathione S-transferase fusion
protein as a substrate with subsequent immunoblotting for phosphorylated Elk1.
Immunoprecipitations, nuclear extracts, and electrophoretic mobility
shift assays (EMSAs).
For immunoprecipitations of JAK kinases, EPOR, and STAT-5, 20 to 60 × 106 cells were rested as described above and lysed
in 1% Nonidet P-40 (Calbiochem, La Jolla, CA), 20 nmol/L Tris-HCl pH
8.0, 150 mmol/L NaCl, 50 mmol/L NaF, 100 mmol/L sodium orthovanadate, 1 mmol/L phenylmethylsulfonyl fluoride, 10 µg/mL leupeptin, 10 µg aprotinin, 1 µg/mL pepstatin A. Immunoprecipitations were performed with 5 µg of the indicated JAK antibodies (Upstate Biotechnology Inc,
Lake Placid, NY), 3 µL of a C-terminal, polyclonal rabbit anti-EPOR
antisera,33 or 4 µL of anti-STAT-5A and STAT-5B
antisera34 and protein A-Sepharose (Boehringer Mannheim,
Indianapolis, IN). Immunoprecipitations were separated on 8.75% (JAK
and STAT-5) or 10% (EPOR) sodium dodecyl sulfate-polyacrylamide gel
electrophoresis (SDS-PAGE) and transferred to nitrocellulose. Membranes
were probed with 1:1,000 dilution of anti-phosphotyrosine 4G10
antibodies (Upstate Biotechnology Inc). Membranes were stripped per
manufacturer's instructions and reprobed with anti-JAK1, -JAK2, and
-JAK3 antisera (data not shown) or with anti-STAT-5A or anti-EPOR
antisera (not shown) to verify equivalent loading. For nuclear
extractions, 30 × 106 cells were rested in serum-free
medium containing 1% BSA for 2 to 4 hours and stimulated with
indicated cytokines for 15 minutes. Cells were lysed and nuclear
extractions were prepared. DNA binding assays were performed
using 1 × 105 cpm of
32P-radiolabeled FcRI oligonucleotide encoding an STAT
response element and 10 µg of nuclear extract/reaction as described
previously.35,36
RNA preparation and Northern blot analysis.
Cytoplasmic RNA was prepared from 1 to 2 × 107 cells
using an RNeasy kit (Qiagen, Valencia, CA) and quantified by UV
spectrophotometry. Denaturing 1.4% agarose gels were run with 10 to 15 µg RNA/lane, blotted to Zeta Probe membranes (BioRad, Hercules,
CA), and hybridized with 32P-labeled c-fos,
bcl-2, and GAPD probes as described
previously.34,37 The 435-nucleotide bcl-x probe
(derived from the 5' end of the bcl-x cDNA that
recognizes both bcl-xS and bcl-xL transcripts) was from ATCC.38
Somatic fusions.
A protocol to create fusion cell lines was adapted from Goldsmith et
al.39 1.5 × 106 HT-2.EPOR.3 and 1.5 × 106 Ba/F3 cells were washed in RPMI 1640, mixed in
a 1:1 ratio (or 3 × 106 of either cell type alone) in
flat-bottomed 12-well plates, centrifuged for 10 minutes at 1,200 rpm
in an RT6000B table top centrifuge (Sorvall, Newtown, CT), and depleted
of media by gentle aspiration. 0.3 mL of RPMI 1640 prewarmed to
37°C containing various concentrations (40% to 65%) of
polyethylene glycol (PEG) 4000 (ATCC) diluted in RPMI 1640 was overlaid
slowly onto the cells at room temperature. After 3 minutes, 1.5 mL RPMI
1640 was slowly added to wells and removed by gentle aspiration. Wells
were washed four to five times in RPMI 1640 and resuspended in complete
media containing 50 U/mL EPO, transferred to fresh 6-well plates, and
cultured at 37°C.
125I-EPO binding assays.
Cells were washed in RPMI 1640 supplemented with 10% FBS, resuspended
at a concentration of 6 × 107 cells/mL, and incubated
at room temperature for 30 minutes and pelleted. Aliquots of 3 × 106 cells in triplicate were incubated with a range of
concentrations of iodinated EPO at 4°C overnight in binding buffer
(RPMI/10% FBS/50 mmol/L HEPES, pH 7.2) in the absence or presence of
100-fold excess unlabeled EPO. After overnight incubation, cells were
determined to be greater than 90% viable by trypan blue exclusion
analysis. Free EPO was separated from cell-bound EPO by centrifugation
through 100% FBS cushion and the cell pellets were counted in a counter. The data were plotted according to the method of Scatchard.
Dissociation constants (kd) and cell-surface receptor
numbers were then determined through Scatchard analyses.
| |
RESULTS |
The EPOR inhibits apoptosis in HT-2 cells.
To examine the function of the EPOR in an IL-2-responsive cell, stable
HT-2 cell lines were generated that expressed the EPOR (designated
HT-2.EPOR.329), and mRNA expression was confirmed by
Northern blotting (Fig 1A). It has been
shown that the EPOR serves to inhibit apoptosis in erythroid
progenitors,40,41 and several cytokines inhibit apoptosis
in HT-2 cells.42 Therefore, we examined the ability of the
EPOR to inhibit apoptosis induced by IL-2 withdrawal by incubating
cells for 40 hours without cytokine, with IL-2 or with EPO. Cells were
then stained with propidium iodide (PI) and Annexin V coupled to green
fluorescent protein (Annexin-GFP)32 and analyzed by flow
cytometry. First, PI staining was used to measure cell death.
Incubation of HT-2 or HT-2.EPOR.3 cells without cytokines resulted in
significant cell death, because only 25.2% or 14.2% of HT-2 or
HT-2.EPOR.3 cells excluded PI, respectively, in a typical experiment
(Fig 1B). The slight differences in cell death among cell lines
represent experimental variation, because they were not consistently
observed over multiple experiments. However, signaling through the EPOR
dramatically and reproducibly protected HT-2.EPOR.3 cells from death in
the presence of EPO, as 62.1% of cells excluded PI, compared with
27.3% of the parental HT-2 cells lacking the EPOR. Finally, IL-2
signaling prevented cell death in both cell lines (94.8% of HT-2 and
83.2% of HT-2.EPOR.3 cells were PI-negative). Thus, stimulation by EPO
conferred cell survival on EPOR-expressing HT-2 cells, although
the protection was not as complete as that observed when cells were
incubated with IL-2.
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| Fig 1.
(A) HT-2.EPOR cells express EPOR mRNA. Ten micrograms of
cytoplasmic RNA from HT-2.EPOR.3 and HT-2 cells was separated by
denaturing agarose gel electrophoresis, transferred to Zeta Probe
membranes and probed with a 32P-labeled 1.2-kb DNA fragment
corresponding to the EPOR extracellular domain. (B and C) The EPOR
protects HT-2 cells from cell death and apoptosis. HT-2 and HT-2.EPOR.3
cells were incubated for 40 hours with no cytokine (None), 20 U/mL or
indicated doses of EPO (Epo), or 10 nmol/L IL-2 (IL-2).
Cells were costained with PI and Annexin-GFP and analyzed by flow
cytometry.
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To determine whether the EPOR was inhibiting apoptosis per se, the
PI-negative populations were examined for staining by Annexin-GFP, which binds to phosphatidylserine and serves as a marker of
apoptosis43,44 (Fig 1B). Virtually all HT-2 and HT-2.EPOR.3
cells incubated without cytokine that were still alive (ie,
PI-negative) bound to Annexin-GFP (96.7% of HT-2 and 92.2% of
HT-2.EPOR.3 cells), indicating that they were apoptotic. Similarly,
93.1% of PI-negative HT-2 cells (that did not express the EPOR) were
Annexin-GFP-positive in the presence of EPO, indicating that the
entire culture was undergoing apoptosis. In contrast, HT-2.EPOR.3 cells
showed significant, EPO-dependent protection from apoptosis, because
only 10.2% of the cells stained positively with Annexin-GFP in the
presence of EPO. As expected, IL-2 treatment strongly protected both
cell lines from apoptosis (6.2% of HT-2 and 3.7% of HT-2.EPOR.3 cells were Annexin-GFP-positive). In addition, the anti-apoptotic effect of
EPO occurred in a dose-responsive fashion, with detectable inhibition
of apoptosis and a corresponding increase in percentage of live cells
evident at concentrations of EPO as low as 0.2 U/mL (Fig 1C). These
data show that the EPOR prevents HT-2.EPOR.3 cell death upon cytokine
withdrawal by delivering a potent anti-apoptotic signal.
EPOR alone fails to support proliferation in HT-2 cells.
The ability of the EPOR to mediate growth signaling in the HT-2 cell
background was evaluated by measuring incorporation of [3H]thymidine in cells incubated in either IL-2 or EPO.
As a positive control, HT-2 cells expressing chimeric receptors in
which the cytoplasmic domain of the EPOR was replaced with the
cytoplasmic tails of either the IL-2R or c chains (EPO and
EPO) responded vigorously to EPO after 24 hours of stimulation
(Fig 2A28,29). In contrast,
parental HT-2 cells failed to incorporate significant levels of
[3H]thymidine in response to EPO. Strikingly, HT-2.EPOR.3
cells did not incorporate high levels of [3H]thymidine,
although there was a small but reproducible signal that peaked at
approximately 8% of IL-2 levels in very high doses of EPO (Fig 2A).
However, EPO failed entirely to sustain growth of HT-2.EPOR.3 cells in
the absence of IL-2 (data not shown). Thus, EPO triggers a minimal but
transient proliferative response in HT-2 cells that is greatly reduced
compared to that induced by IL-2. To confirm that the EPOR cDNA used in
these experiments encodes a functional receptor that supports cell
growth in an permissive cellular background, the EPOR was expressed
stably in the IL-3-dependent myeloid cell line, 32D.45 In
contrast to HT-2 cells, 32D.EPOR cells exhibited a vigorous
proliferative dose response to EPO, while the parental 32D line lacking
the EPOR failed to proliferate (Fig 2B). Together with the
anti-apoptotic effect described earlier, these data show that
inhibition of apoptosis and induction of proliferation are separable
signaling events.
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| Fig 2.
(A) The EPOR fails to support proliferation signaling in
HT-2 cells. HT-2  ), HT-2.EPOR.3 ( ), and HT-2EPO () cells
were incubated with no cytokine, various doses of EPO, or 10 nmol/L
IL-2 and incorporation of [3H]thymidine was measured at
48 hours. Data are presented as percent of [3H]thymidine
incorporation relative to IL-2. (B) The EPOR supports proliferation
signaling in 32D cells. 32D () and 32D.EPOR () cells were
incubated with no cytokine, various doses of EPO, or 10%WEHI-CM
(IL-3). Data are presented as percent of [3H]thymidine
incorporation relative to IL-3. (C) The EPOR can pair with EPO to
support EPO-dependent proliferation signaling in HT-2 cells.
HT-2.EPOR.3 cells were transfected with 20 µg of pCMV4 vector with no
insert (control, ) or pCMV4.EPO (EPO, ) and
[3H]thymidine incorporation measured on days 6-16, as
indicated.
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Finally, to verify that the HT-2.EPOR.3 cells express potentially
functional EPO receptors, the EPO chimeric receptor was transiently
transfected into HT-2.EPOR.3 cells, and [3H]thymidine
incorporation was measured over a 16-day period. We have previously
reported that, although the EPO receptor alone is not sufficient to
support growth signaling, it can signal proliferation when paired with
either the EPO chimera or with a truncated EPOR retaining the
JAK2-binding domain [EPOR(1-321)29]. EPO also
exhibited a characteristically potent proliferative response to EPO
when paired with the full-length EPOR in HT-2.EPOR.3 cells, whereas a
control vector alone failed to induce [3H]thymidine
incorporation (Fig 2C). Thus, these experiments show that the HT-2 cell
line is restricted in its ability to respond to cytokines: while the
IL-2R (or EPO/EPO chimeras) delivers effective growth signals to
these cells, the EPOR is insufficient to promote proliferation or
long-term survival in EPO.
The EPOR fails to activate STAT factors in HT-2 cells.
A dominant pathway activated by cytokine receptors is the JAK-STAT
pathway, and EPOR-derived proliferation signaling has been linked to
JAK2 and STAT-5 activation in some settings.14,15,20,21 Therefore, to determine the basis of the signaling block in HT-2.EPOR.3 cells, the ability of EPO to activate JAK2 and STAT-5 was examined. HT-2.EPOR.3 cells were starved of IL-2 and incubated for 10 minutes with no cytokine, EPO, or IL-2 (in these and subsequent experiments, supra-physiological doses of EPO were used to achieve maximal stimulation of the EPO receptors expressed on target cells). Cellular lysates were prepared and immunoprecipitated with antibodies to JAK1,
JAK2, or JAK3. Immunoprecipitates were separated by SDS-PAGE, transferred to nitrocellulose membranes, and probed with
antiphosphotyrosine antibodies (Fig 3A). As
expected, when HT-2.EPOR.3 cells were stimulated with IL-2, JAK1 and
JAK3, but not JAK2, became phosphorylated on tyrosine. In contrast,
when the same cells were stimulated with EPO, JAK2 alone was found to
be phosphorylated (Fig 3A, lane 5). Therefore, JAK2 activation is
clearly not the block to EPO signaling in this context.
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| Fig 3.
EPO signaling mediates phosphorylation of JAK2 and the
EPOR but not STAT-5 in HT-2 cells. (A) HT-2.EPOR.3 cells were rested
and stimulated with no cytokine (U), 50 U/mL EPO (E), or 10 nmol/L IL-2
(2) and lysates were immunoprecipitated with 5 µg of antibodies to
JAK1 (lanes 1 through 3), JAK2 (lanes 4 through 6), or JAK3 (lanes 7 through 9). Immunoprecipitates were separated on 8.75% SDS-PAGE,
transferred to nitrocellulose, and probed with antiphosphotyrosine
antibodies. Migration of molecular-weight markers is indicated. The
presence of a small amount of phosphorylated JAK3 in lanes 7 and 8 is
probably background phosphorylation resulting from incomplete IL-2
deprivation of the cells before restimulation with the indicated
cytokines. (B) HT-2.EPOR.3 and 371.2 cells were rested and stimulated
for 15 minutes with no cytokine (U), 50 U/mL EPO (E), 10 nmol/L IL-2
(2), or 10% WEHI-CM (IL-3) (3) and nuclear extracts were prepared.
EMSAs were performed with 10 µg of nuclear extracts and a
32P-labeled oligonucleotide corresponding to the FcRI
STAT-response element.46 Supershifts were performed by
preincubating nuclear extracts with anti-STAT-5A and -5B antisera
(lane 4) or preimmune rabbit sera (lane 5) for 45 minutes before the
EMSA reaction. (C) HT-2.EPOR.3 cells were rested and stimulated with no
cytokine (U), EPO (E), or IL-2 (2) and lysates were immunoprecipitated
with antibodies to STAT-5A and -5B. Immunoprecipitates were separated
on 8.75% SDS-PAGE, transferred to nitrocellulose membranes, and probed
with antiphosphotyrosine antibodies. The membrane was stripped and
reprobed with anti-STAT-5A and -5B antibodies. (D) 5 × 106 HT-2.EPOR.3 cells/sample or 2 × 106
371.2.EPOR cells/sample were rested and incubated with no cytokine (U)
or 50 U/mL EPO (E) and lysates were immunoprecipitated with anti-EPOR
antibodies. Immunoprecipitates were separated by 10% SDS-PAGE,
transferred to nitrocellulose, and probed with antiphosphotyrosine
antibodies.
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We next examined the ability of EPO to activate STAT-5 in HT-2.EPOR.3
cells. Cells were starved of cytokine and stimulated for 15 minutes
with EPO, and nuclear lysates were prepared and subjected to EMSA using
a generic STAT-response element derived from the FcRI gene that
recognizes a variety of STAT factors.46 As previously
reported,36 IL-2 stimulation of HT-2.EPOR.3 cells resulted
in the appearance of a specific retarded nucleoprotein complex (Fig 3B,
lane 3) that could be supershifted with antibodies to STAT-5A and
STAT-5B (Fig 3B, lane 4). In contrast, EPO stimulation of HT-2.EPOR.3
cells failed to cause significant retardation of the probe (Fig 3B,
lane 2). Notably, the STAT-response element used in this experiment can
also bind STAT-1 and STAT-3,42 which have been linked to
the EPOR47; therefore, the EPOR fails to induce any known
STAT factors in this cellular context. Furthermore, EPO stimulation of
371.2.EPOR cells (derived from 32D cells30) induced STAT-5,
verifying that the EPOR cDNA is fully functional in permissive cells
(Fig 3B, lane 7).
Because STAT-5 DNA binding activity was not induced by the EPOR, we
next examined whether STAT-5 was inducibly phosphorylated on tyrosine
after receptor stimulation. HT-2.EPOR cells were starved of IL-2 and
incubated with no cytokine, EPO, or IL-2 for 15 minutes. Cell lysates
were immunoprecipitated with antibodies to STAT-5A and -5B, separated
by SDS-PAGE, transferred to nitrocellulose and probed with
antiphosphotyrosine antibodies (Fig 3C). Although IL-2 induced strong
phosphorylation of STAT-5A and -5B in HT-2 cells, EPO failed to induce
detectable phosphorylation. Thus, the initial step of STAT-5 activation
by phosphorylation on tyrosine fails to occur in HT-2.EPOR.3 cells.
This lack of STAT-5 activation could be explained if the EPOR itself
were not phosphorylated after receptor engagement and JAK2 activation,
because it would then be unable to provide docking sites for signaling
intermediates such as STAT-5. To determine whether the EPOR was
phosphorylated in HT-2.EPOR.3 cells, cells were stimulated with EPO for
25 minutes, and cell lysates were immunoprecipitated with anti-EPOR
antibodies.33 Lysates were separated by SDS-PAGE,
transferred to nitrocellulose, and probed with antiphosphotyrosine
antibodies (Fig 3D). In both HT-2EPOR.3 cells and control 371.2.EPOR
cells, EPO stimulation resulted in tyrosine phosphorylation of the EPOR
(Fig 3D, lanes 2 and 4), indicating that the defect in signaling is not
attributable to impairment of this proximal step.
The EPOR fails to activate the MAPK cascade or specific gene
expression in HT-2 cells.
To characterize other signaling pathways known to be induced by the
EPOR in permissive cells, the ability of EPO to induce phosphorylation
of MAPK on serine residues was examined. Cells were starved and
stimulated with cytokine, and lysates were prepared and
immunoprecipitated with antibodies to phosphorylated MAPK. The
immunoprecipitates were then subjected to an in vitro kinase reaction,
separated by SDS-PAGE, transferred to nitrocellulose membranes, and
probed with antibodies to a phosphorylated substrate, Elk1. EPO
stimulation of a positive control cell line HCD57 (a murine,
EPO-dependent erythroleukemia cell line) induced MAPK phosphorylation
of its substrate (Fig 4, lane 1). However,
in HT-2.EPOR.3 cells, EPO stimulation failed to activate MAPK, while IL-2 stimulation resulted in efficient MAPK induction (Fig 4, lanes 4 and 6).
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| Fig 4.
Absence of MAPK induction by the EPOR in HT-2 cells.
HCD57 cells (lanes 1 and 2) or HT-2.EPOR.3 cells (lanes 3 through 6)
were starved and stimulated with no cytokine (U), 50 U/mL EPO (E), 10 ng/mL murine IL-4 (4), or 10 nmol/L IL-2 (2). Total cell lysates were
prepared and immunoprecipitated with anti-phosphoMAPK antibody.
Immunoprecipitates were subjected to in vitro kinase assays with an
Elk1-glutathione S-transferase fusion protein, separated by
10% SDS-PAGE, transferred to nitrocellulose, and probed with
anti-phospho-Elk1 antibodies. Arrow indicates phosphorylated Elk1.
|
|
Another common consequence of MAPK activation is the induction of
specific gene expression, including the proto-oncogene
c-fos.48 To assay c-fos gene expression,
cells were starved of cytokine and stimulated with EPO or IL-2 for the
indicated time periods. Cytoplasmic RNA was prepared from these cells,
separated on denaturing agarose gels, transferred to membranes, and
probed for c-fos or GAPD
(Fig 5A). Parental HT-2 cells lacking the
EPOR failed to induce c-fos gene expression (Fig 5A, lanes 1 through 4), while IL-2 stimulation resulted in marked c-fos
mRNA induction which peaked at 30 to 60 minutes, as described
previously (Fig 5A, lanes 5 through 834). As expected, EPO
stimulation of HT-2EPO cells also induced c-fos mRNA (Fig
5A, lanes 14 through 17), but EPO stimulation of HT-2.EPOR.3 cells
failed to induce detectable c-fos gene expression (Fig 5A,
lanes 9 through 13). As c-fos gene expression is considered
typically to be a consequence of MAPK signaling, this result is
consistent with the lack of MAPK activation by EPO in HT-2.EPOR.3 cells
(Fig 4).
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| Fig 5.
(A) The EPOR fails to induce the c-fos or
bcl-2 proto-oncogene mRNA in HT-2 cells. HT-2, HT-2.EPOR.3, and
HT-2EPO cells were incubated in media without growth factors for
15 hours and stimulated with no cytokine (U, lanes 1, 5, and 14) 50 U/mL EPO (lanes 2 through 4, 9 through 13, 15 through 17) or 10 nmol/L
IL-2 (lanes 6 through 8) for the indicated time periods. Total cellular
RNA was prepared, separated on a denaturing agarose/formaldehyde gel,
and probed with 32P-labeled c-fos, bcl-2, or
GAPD cDNA probes. (B) The EPOR fails to induce bcl-x
proto-oncogene mRNA in HT-2 cells. HT-2.EPOR.3 and 371.2.EPOR cells
were starved as described in (A) and stimulated with no cytokine (lane
1), EPO (lanes 2 through 4), or IL-2 (lanes 5 through 7) for the
indicated time periods. RNA from 371.2.EPOR cells serves as a positive
control (lane 8). RNA was prepared and blotted as in (A) and probed
with a 32P-labeled bcl-x cDNA probe derived from
the 5' end of the gene.
|
|
We likewise examined whether the anti-apoptotic gene bcl-2 is
induced by EPO in HT-2.EPOR.3 cells. However, despite the potent anti-apoptotic effect of EPOR seen in Fig 1, bcl-2 mRNA was
only weakly induced upon EPO stimulation (Fig 5A, lane 13), although IL-2 induced a strong upregulation of bcl-2 transcripts in HT-2 cells (Fig 5A, lane 8) and EPO induced bcl-2 mRNA in
HT-2.EPO cells (Fig 5A, lane 17). This experiment suggests that
the anti-apoptotic effect of EPOR probably occurs via a pathway
independent of Bcl-2. Recent evidence has also implicated the
Bcl-2 family member Bcl-xL in anti-apoptotic effects
mediated by the EPOR.49,50 However, we found by analysis of
RNA that HT-2.EPOR.3 cells do not express detectable levels of
bcl-xL or bcl-xS mRNA, nor is
bcl-xL mRNA induced by either IL-2 or EPO (Fig 5B).
Somatic hybrids between HT-2 and Ba/F3 cells rescue EPO signaling.
To determine whether the defect in EPOR signaling in HT-2 cells could
be overcome by genetic complementation, we generated a series of stable
hybridoma cell lines between HT-2.EPOR.3 and Ba/F3, a cell line lacking
EPOR expression. Cells were cocultured, fused with various
concentrations of PEG, and selected for growth in 50 U/mL EPO. Notably,
when the fusion protocol was performed with either HT-2.EPOR.3 or Ba/F3
cells alone, no cell lines whatsoever survived selection in EPO.
However, EPO-dependent fusion lines were readily generated when
HT-2.EPOR.3 and Ba/F3 cells were mixed in the fusion reaction. Two
polyclonal fusion lines were chosen for further analysis, designated
HB40 and HB55 (for HT-2-Ba/F3 fused at 40% or 55%
PEG). Both HB40 and HB55 cells proliferated vigorously in response to
EPO, while Ba/F3 and HT-2.EPOR.3 cells showed little or no response
(Fig 6A). The abilities of HB40 and HB55 to
proliferate were directly dependent on the dose of EPO, and these cells
responded to physiological doses of EPO (Fig 6B).
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| Fig 6.
Somatic fusions between HT-2.EPOR.3 and Ba/F3 cells
restore EPO-dependent signaling. (A) Indicated cell lines were
incubated without cytokine (U), 50 U/mL EPO or 10%
WEHI-CM (as a source of IL-3), and [3H]thymidine
incorporation measured after 48 hours. ( ), HB40; ( ), HB55; ( ),
Ba/F3; ( ), HT-2.EPOR.3. (B) Indicated cell lines were incubated
without cytokines or with various doses of EPO, and
[3H]thymidine incorporation was measured after 48 hours.
(), HB40; ( ), HB55. (C) HT-2.EPOR.3, HB40, and HB55
cells were rested and stimulated for 15 minutes with no cytokine (U),
50 U/mL EPO (E), or 10 nmol/L IL-2 (2). Nuclear extracts were subjected
to EMSA with the FcRI probe. Note that all lanes were derived from
the same gel, but in the photograph, irrelevant intervening lanes were
omitted between lanes 3 and 4. (D) HT-2.EPOR.3, HB40, and HB55 cells
were rested and stimulated with no cytokine (U), EPO (E), or IL-2 (2)
and lysates prepared and immunoprecipitated with anti-STAT-5A and -5B
antibodies. Lysates were separated on 8.75% SDS-PAGE, transferred to
nitrocellulose membranes, and probed with antiphosphotyrosine
antibodies. Membranes were stripped and reprobed with anti-STAT-5A and
-5B antibodies to confirm equivalent loading. (E) HT-2.EPOR.3, HB40,
and HB55 cells were starved and stimulated with no cytokine (lanes 1, 5, and 9) or 50 U/mL EPO (lanes 2 through 4, 6 through 8, and 10 through 12) as described in the legend to Fig 5. RNA was separated on a
denaturing agarose/formeldehyde gel, transferred to Zeta Probe
membranes, and probed with 32P-labeled c-fos or
GAPD cDNA probes.
|
|
Moreover, 125I-EPO binding analyses showed that the fusion
cell lines expressed approximately the same number of surface EPO receptors with similar affinities as the parental HT-2.EPOR.3 cells
(Table 1). Both the HT-2.EPOR.3 cells and
the HB40 and HB55 cells expressed 200 to 250 cell-surface EPO
receptors, with affinities in the range of 1,500 to 1,800 pmol/L,
whereas the parental HT-2 cells did not express detectable receptors
for EPO (Table 1). These results indicate that restoration of signaling in the fusion cell lines is not simply caused by an increase in receptor number.
We next examined STAT-5 activation by the fusion cell lines. In
contrast to HT-2.EPOR.3 cells, HB40 and HB55 cells activated STAT-5 DNA
binding activity vigorously in response to EPO (Fig 6C). In addition,
phosphorylation of STAT-5 proteins in response to EPO was also restored
in the fusion lines (Fig 6D). Thus, Ba/F3 cells are able to complement
the EPOR signaling defect of HT-2 cells. Furthermore, rescuing the
cells' ability to grow in EPO correlates with concurrent restoration
of STAT-5 activation, which suggests that the missing component(s)
functions upstream of both STAT-5 activation and proliferation.
Finally, to determine whether the MAPK/c-Fos pathway is likewise
rescued in the fusion cell lines, we measured induction of c-fos mRNA. Cells were starved of cytokines, stimulated for various times with EPO, and mRNA and Northern blots were prepared and probed
with 32P-labeled c-fos and GAPD probes.
Induction of c-fos mRNA by EPO was indeed evident in these cell
lines (Fig 6E, lanes 7 and 11). Thus, all aspects of EPOR signaling
measured were reconstituted in the fusion cell lines HB40 and HB55.
| |
DISCUSSION |
This study shows that the EPOR fails to mediate STAT-5 and MAPK
activation or to deliver proliferative signals in the IL-2-dependent T-cell line, HT-2, despite the presence of potentially functional cell-surface EPO receptors that can mediate JAK2 and EPOR
phosphorylation and inhibit apoptosis. The signaling defect(s) can be
complemented by somatic hybridization between HT-2.EPOR cells and
IL-3-dependent Ba/F3 cells, suggesting that a missing factor required
for signaling in HT-2 cells can be provided in trans by Ba/F3
cells. These results delineate a dichotomy between the signaling
processes that drive anti-apoptosis and those that induce
proliferation. In addition, they provide a valuable platform for
identifying new components of the EPOR signaling cascade, which may
also yield further insights into signaling mechanisms by other cytokine receptors.
Cytokines regulate a myriad of physiological processes within their
target cells, including proliferation, survival, and differentiation. Other studies have suggested that the mechanisms that promote cell
survival by inhibiting apoptosis are distinct from those that trigger
cellular proliferation.51-54 The HT-2.EPOR.3 cell line
described here supports this separation of function, as the EPOR
delivers a strong anti-apoptotic signal while failing to induce
proliferation. The immature erythroid cell line J2E-NR exhibits similar
characteristics with respect to proliferation; while the EPOR fails to
induce growth signaling or differentiation in these cells, it does
serve to enhance cell viability.55 However, in many other
cytokine-dependent cells, the EPOR supports proliferative signaling
(eg, 32D cells, Fig 2B). These results may indicate that certain cell
types need only an anti-apoptotic signal to transit the cell cycle and
proliferate, while HT-2 cells require an additional impetus to
proliferation that apparently can be provided by the IL-2R but not by
diverse cytokine receptors such as the EPOR (this study),
IL-7R,56 IL-9R,42 IL-4R,57 growth hormone receptor (GHR), thrombopoietin receptor (c-mpl), or granulocyte colony-stimulating factor receptor (G-CSFR) (data not shown). Moreover, HT-2 cells provide a valuable system to define regions of
the EPOR that deliver anti-apoptotic signals versus those that mediate
proliferation. Although there is evidence that STAT-5 may regulate the
anti-apoptotic signals of the EPOR in some cell backgrounds,58 the EPOR fails to activate any known STAT
factors in HT-2 cells, thus arguing that STAT-5 is not involved in this particular anti-apoptotic pathway. Likewise, despite evidence that
induction of the AP1 transcription factor (which includes the
c-Fos proto-oncogene) by the MAPK pathway has been linked to
protection from apoptosis,59 it is clear that MAPK
activation is not required for inhibition of apoptosis in the HT-2 cell
background, because this pathway is not engaged by the EPOR. In 32D
cells, the EPOR is known to induce Bcl-2 and Bcl-xL to
effect apoptosis inhibition.60 However, neither of these
genes is significantly induced by the EPOR in HT-2 cells (Fig 5).
Therefore, the molecular mechanism underlying EPOR-mediated apoptotic
inhibition in this context is distinct from recognized pathways and
remains to be elucidated.
In vivo, some developmental events appear to require cytokines mainly
to prevent apoptosis, while others appear to need additional proliferative signals.61 Although the EPOR does exert
anti-apoptotic effects 40,41 (Fig 1), prevention of
apoptosis in erythroid progenitors is probably not sufficient to
eliminate their dependence on EPO for maturation.62
Specifically, in mice expressing a bcl-2 transgene under the
control of an erythroid promoter, growth of erythroid progenitors in
vitro is impaired in the absence of EPO. While apoptosis of colony
forming units-erythroid (CFU-E) is delayed in these transgenic mice, no
spontaneously differentiating colonies are found in the absence of
EPO.62 It will be interesting to see the results of
crossing these mice with EPO / or
EPOR/ mice2,3 to determine
whether differentiation in vivo is equally affected. However, using a
bcl-2 transgene to inhibit apoptosis in erythroid progenitors
may not mimic native anti-apoptotic signaling by the EPOR, because our
work shows that the EPOR is able to prevent apoptosis without inducing
bcl-2 mRNA significantly (Figs 1 and 5A), and because Bcl-2 is
apparently not expressed in erythroid progenitors.63
Another in vivo system illustrating the importance of apoptotic
inhibition is mice lacking the IL-7R chain (or its partner, c).64-66 IL-7R/ mice are
characterized by profound immunodeficiency, lacking mature B and T
cells.67 When these mice were crossed with animals that
overexpress the human bcl-2 gene, T-cell development was rescued, but not B-cell development.64-66 Thus, the
IL-7-derived signal necessary for T-cell maturation can be replaced
solely by inhibiting apoptosis, but B-cell differentiation requires
additional signals from the IL-7R. Similarly, the EPO-EPOR system
appears to require such additional signals for erythroid development.
The dominant paradigm of the JAK-STAT pathway indicates that STAT
factors are recruited to the receptor complex following phosphorylation
of receptor tyrosine residues. In the case of the EPOR, this model is
supported by the severe diminution in STAT-5 activation by EPO receptor
mutants that lack cytoplasmic tyrosine residues.20,68,69
However, the block in JAK-STAT signaling in this study shows that there
is likely to be an additional component of this pathway that has not
been previously recognized. Although very proximal signals are intact
in this pathway (eg, phosphorylation of JAK2 and the EPOR, Figs 3A and
3D), phosphorylation of STAT-5 is impaired (Fig 3C) despite the
presence of potential phosphotyrosine docking sites on the EPOR in
these cells. It is interesting to note that the DNA binding complexes
induced by EPO in 371.2 cells migrate slightly faster than does the
IL-2-induced STAT-5 complex in HT-2 cells or the EPO-induced STAT-5
complex in HB40 and HB55 cells. We have not yet determined the
biochemical basis for this difference.
There are two opposing hypotheses to explain the lack of EPOR signaling
in the HT-2 cell line. The most likely explanation is that the cells
are missing a factor(s) necessary to mediate signaling. Supporting
evidence comes from the somatic fusions with Ba/F3 cells, which can
complement the lack of function in HT-2.EPOR.3 cells. The resulting
fusion cell lines rescue not only proliferation signaling, but also
STAT-5 activation and proto-oncogene induction. Although the nature of
this factor(s) can only be determined by isolating a gene from an
EPO-responsive cell line that can rescue the proliferative defect, some
possibilities include an additional EPOR subunit or an adapter molecule
that might couple STAT-5 to JAK2 or to the EPOR. Indeed, there have
been hints that the EPOR complex might contain an additional
subunit70-73; in this scenario, the missing receptor would
be expressed in most cell lines but would be absent in HT-2 cells. We
are confident that the missing factor is not the adapter Shc, because
IL-2 signaling through Shc that leads to MAPK and c-fos mRNA
induction proceeds normally in this cell line (Figs 4 and 5, and refs
34, 74, and 75). On the other hand, it is formally possible that HT-2 cells express an overabundance of a suppressor factor that inhibits the
EPOR. Although it is known that the tyrosine phosphatases SHP-1 and
SHP-2 are involved in EPOR signaling,76-78 mRNA levels of
SHP-1 and SHP-2 in HT-2 cells are comparable to those in EPO-responsive cell lines such as HCD57 and Ba/F3 (data not shown). Furthermore, it is
unlikely that the cytokine inhibitors of the SOCS/CIS
family79 are responsible, because: (1) the STAT
downregulation that occurs upon SOCS activation is linked to a decrease
in JAK activity, while JAK2 activation by the EPOR is retained in HT-2
cells (Fig 3); (2) none of the SOCS family members identified so far
are specific to the EPOR,80,81 and IL-2, IL-7, and IL-9 can
activate STAT-5 vigorously in HT-2 cells36,42,57; and (3)
the EPOR can pair with EPO to activate JAK2 and STAT-529
and is thus unlikely to carry any other dominant suppressor molecule(s)
that inhibits proximal signaling within this hybrid receptor complex. Therefore, the available evidence supports the hypothesis that HT-2
cells are missing a necessary signaling component that links the EPOR
to STAT-5, MAPK, and proliferation signaling.
An issue of considerable interest is whether cytokine signaling differs
depending on cell context. Although there are dozens of cytokines, each
with a specific receptor, the downstream events that follow receptor
ligation are remarkably similar among members of the cytokine receptor
superfamily; eg, both the IL-2R and EPOR activate STAT-5, c-Fos, Bcl-2,
MAPK, IRS-2, and PI3K.27 Do the target cells of different
cytokines "interpret" these signals differently, or is the basis
for this dichotomy simply their anatomical location? To address this
question in the context of homodimeric receptors such as the EPOR,
recent work using an in vivo differentiation system showed that the
cytoplasmic tail of the EPOR can be substituted with the cytoplasmic
domains of diverse cytokine receptors (G-CSFR, c-mpl, and GHR) yet
exhibit similar signaling outcomes with regard to erythropoiesis, thus
favoring the notion that cytokine signals are generally
replaceable.82 Similarly, prolactin and its receptor can
substitute for the EPOR in driving differentiation of erythrocytes in
vitro.83 In an analogous study, the cytoplasmic tail of the IL-2R chain was found to substitute functionally for the IL-7R chain in driving proliferation of B cells in an ex vivo culture system.84 However, the data presented here suggest that the EPOR cannot replace the IL-2R, despite the apparent similarity in the
intracellular signaling molecules engaged by these receptors.
Although the restoration of STAT-5 activation in the HT-2-Ba/F3
hybridomas implies a link between STAT-5 function and proliferation, STAT-5 activation is clearly not sufficient to drive proliferation in
the HT-2 cell background. First, it has been previously shown that
IL-7R signaling fails to support proliferation in HT-2 cells, despite a
vigorous induction of STAT-5 DNA binding.56 Second, the
IL-9R activates STAT-5 (and STAT-1 and STAT-3) quite well in HT-2 cells
and also mediates an anti-apoptotic response, but does not permit
proliferation.42 Finally, the IL-4R also induces JAK-STAT
activation (in this case, STAT-6) without driving proliferation of HT-2
cells.57 At present, it is unclear precisely how IL-2 signaling differs from that of the other c-utilizing cytokine receptors and the EPOR to induce proliferation in HT-2 cells.
The failure of EPOR to signal in a T-cell background has also been
described for sublines of the IL-2-dependent T-cell line CTLL
transfected with the EPOR.85 In this case, the cells'
nonresponsiveness was attributed to the failure of JAK2 to associate
with the EPOR. This mechanism contrasts with our results in HT-2.EPOR.3
cells in which JAK2 is potently activated in response to EPO, while STAT-5 activation is impaired. Because IL-2 functions as a growth factor for mature, peripheral T cells such as HT-2 and CTLL, it is
tempting to speculate that the lack of permissivity to signaling in the
HT-2 and CTLL lines reflects their biologic origins. It would be
interesting to determine whether EPOR-mediated signal transduction can
occur in normal T cells.
In conclusion, the present report describes a unique cell context in
which signaling by the EPOR is impaired, despite the ability of a
highly related cytokine (IL-2) to trigger the same signaling pathways.
This system will serve as a valuable tool for identifying essential
components that link the EPOR to the JAK-STAT, MAPK pathways, and
proliferative machinery.
| |
ACKNOWLEDGMENT |
The authors acknowledge the valuable assistance of John Carroll, Neile
Shea, and Heather Livesay in the preparation of this manuscript, and Dr
N. Abraham for critical reading. Annexin-GFP was the kind gift of Dr
Joel Ernst, and 371.2 cells were provided by Dr Morris White (Joslin
Diabetes Center, Harvard University). IL-2 was a gift from
Chiron Corporation, and EPO was a gift from Ortho Biotech.
| |
FOOTNOTES |
Submitted August 24, 1998; accepted February 23, 1999.
Supported by the J. David Gladstone Institutes, and National Institutes
of Health (NIH) Grants No. GM54351 to M.A.G. and CA75315 to G.D.L.
S.L.G. was supported by the Bank of America-Giannini Foundation and the
NIH Immunology Training Grant at the University of California, San
Francisco. S.Y.L. was in the NIH Medical Scientist Training Program
(MSTP) and the Biomedical Sciences Program at the University of
California, San Francisco (UCSF), and K.D.L. is in the NIH MSTP and
Program in Biological Sciences at UCSF.
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
1734 solely to indicate this fact.
Address reprint requests to Mark A. Goldsmith, MD, PhD, Gladstone
Institute of Virology and Immunology, PO Box 419100, San Francisco, CA
94141; e-mail: mgoldsmith{at}gladstone.ucsf.edu.
| |
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