Binding of erythropoietin (EPO) to its receptor (EPOR) on erythroid cells induces the activation of numerous signal transduction pathways, including the mitogen-activated protein kinase Jun-N-terminal kinase (JNK). In an effort to understand the regulation of EPO-induced proliferation and JNK activation, we have examined the role of potential autocrine factors in the proliferation of the murine erythroleukemia cell line HCD57. We report here that treatment of these cells with EPO induced the expression and secretion of tumor necrosis factor alpha (TNF-α). EPO-dependent proliferation was reduced by the addition of neutralizing antibodies to TNF-α, and exogenously added TNF-α induced proliferation of HCD57 cells. EPO also could induce TNF-α expression in BAF3 and DA3 myeloid cells ectopically expressing EPOR. Addition of TNF-α activated JNK in HCD57 cells, and the activity of JNK was partially inhibited by addition of a TNF-α neutralizing antibody. Primary human and murine erythroid progenitors expressed TNF-α in either an EPO-dependent or constitutive manner. However, TNF-α had an inhibitory effect on both immature primary human and murine cells, suggestive that the proliferative effects of TNF-α may be limited to erythroleukemic cells. This study suggests a novel role for autocrine TNF-α expression in the proliferation of erythroleukemia cells that is distinct from the effect of TNF-α in normal erythropoiesis.

Erythropoietin (EPO) is the glycoprotein hormone necessary for the production of mature erythroid cells. Erythroid cells at the colony-forming units–erythroid (CFU-E) to proerythroblast stage of differentiation respond to EPO with proliferation, survival, and terminal differentiation. EPO affects these diverse cellular events via association with its receptor (EPOR) and subsequent activation of numerous signal transduction pathways, which then direct the appropriate cellular response. These pathways include Janus kinase/signal transducers and activators of transcription (JAK2/STAT5),1-4 PI3-kinase,5,6 the adaptor protein SHC,7-9 Src homology 2 domain–containing inositol 5-phosphatase (SHIP),10,11 and the MAP kinase pathways Jun N-terminal kinase (JNK)12,13 extracellular signal related-kinase (ERK)14,15 and p38.16,17 The ERK pathway is primarily associated with the proliferation of erythroid cell lines.12,18,19 The kinetics of the activation of these signals upon activation of the EPO receptor differ, however. In the EPO-dependent murine erythroleukemia cell line HCD57, the JAK2/STAT5, ERK, and PI3-kinase pathways are activated maximally within 5 minutes of EPO binding to its receptor, followed by a decrease in the signal to a lower basal level that is maintained as long as EPO is present. JNK and p38, by contrast, are activated 1 to 4 hours after EPO addition and reach maximum activation 24 hours after EPO addition.12 Whereas much is known about the early signals generated from EPO binding to its receptor, these long-lasting signals that result from EPO treatment are not well understood. One possible mechanism may be that the initial hormone triggers the autocrine or paracrine release of a new factor that maintains the cells.

Tumor necrosis factor alpha (TNF-α) is a cytokine produced by a variety of cell types, including macrophages, monocytes, lymphoid cells, and fibroblasts, usually in response to inflammation or infection.20 The TNF-α signal is mediated via 2 distinct receptors, TNF-α receptor-1 (TNFR1 p55) and TNF-α receptor 2 (TNFR2 p75).21 The extracellular domains of these receptors are closely related to those of CD30, CD40, CD27, and Fas, which are all members of the TNF-α superfamily. Although TNF-α is usually considered an inflammatory cytokine, inducing fever, shock, and apoptosis, TNF-α also has been shown to promote proliferation of human leukemia cells22 and differentiation of macrophages23 in vitro. TNF-α has been shown to stimulate proliferation of both lymphoid22 and nonlymphoid24 cells, as well as some cancers, including chronic lymphoid leukemia25,26 and ovarian cancer.27 This stimulation may be direct activation of the TNF-α receptor or secondary effects resulting from TNF-α–dependent secretion of an intermediate factor such as granulocyte-macrophage colony-stimulating factor.28 TNF-α has been shown to inhibit erythropoiesis,29-31 although it has been demonstrated that the inhibitory effects of TNF-α were likely mediated by β-interferon produced by macrophages in response to TNF-α and not due to direct actions of TNF-α.32,33TNF-α also has been reported to promote proliferation of CD34+ human hematopoietic cells.34 The direct action of TNF-α on erythroid cells therefore remains unclear.

Our laboratory uses HCD57 cells as a model cell system for the study of erythroid proliferation, survival, and apoptosis. These cells depend on EPO for survival and proliferation but do not differentiate in the presence of EPO. In an effort to elucidate the long-term EPO-induced signals of these erythroleukemia cells, we searched for potential autocrine factors that might promote proliferation of these cells. In this report, we will show that whereas TNF-α appears to inhibit both human and murine primary cell erythropoiesis, EPO can induce the production and secretion of TNF-α to promote proliferation in HCD57 cells, and TNF-α may mediate this proliferative signal by activation of JNK.

Reagents

Murine tumor necrosis factor-α (TNF-α), MTT reagent (3-(4,5-dimethyl-2-thiozol)-2,5-diphenyl-2H-tetrazolium bromide), and inhibitors PD98059, SB203850, U0126, and LY294002 were purchased from Calbiochem (La Jolla, CA). Recombinant stem cell factor was purchased from Intergen (Purchase, NY). Phosphospecific antibodies against JNKs (Thr183/Tyr185), ERKs (Thr/Tyr204), protein kinase B/AKT (Ser473), c-jun (Ser 63/Ser73), and p38 (Thr 180/Tyr182) were obtained from Cell Signaling Technologies (Beverly, MA). The goat polyclonal antibody recognizing both phosphorylated and nonphosphorylated forms of JNK1 (C-17) was obtained from Santa Cruz Biotechnologies (Santa Cruz, CA). The neutralizing goat anti–mouse TNF-α antibody and isotype control were obtained from R&D Systems (Minneapolis, MN).

Cell culture

Murine HCD57 cells were cultured in Iscove modified Dulbecco medium (IMDM) (Invitrogen, Carlsbad, CA), 25% fetal calf serum (FCS) (Hyclone, Logan, UT), and 10 μg/mL gentamicin (Invitrogen) at 37°C in a 5% CO2 environment and maintained in 1 U/mL EPO media (EPOGEN, Amgen, Thousand Oaks, CA). Murine DA3-EPOR and BAF3-EPOR cells were cultured in RPMI media, 10% FCS, 10 μg/mL gentamicin, and 100 μg/mL geneticin (Invitrogen). Normal human colony-forming cells were purified in the laboratory of Dr Amittha Wickrema at the University of Illinois at Chicago. The human erythroid progenitors highly enriched for CFU-E were purified by a previously published method.35 Human CD34+cells were isolated as previously published,36 and 6.7 × 104 CD34+ cells were cultured in triplicate in IMDM, 30% FCS, 2 U/mL EPO in the presence or absence of 0.2, 2.0, or 20 μg neutralizing rabbit anti–human TNF-α antibody (Calbiochem) per milliliter or 2 μg/mL rabbit IgG negative control for 7 days at 37°C in a moist CO2 environment. For isolation of murine bone marrow cells, TNF-α homozygous (−/−) and control (wild type) mice (C57BL/6 × 129 genetic background) (Jackson Laboratories, Bar Harbor, ME) were humanely killed at 6 to 8 weeks of age by CO2 asphyxiation, and femurs were removed. Bone marrow was extracted in 5 mL of IMDM, 10% FCS medium, using a 23-gauge needle. Cells were enumerated with trypan blue and plated at the desired density.

RNAse protection analysis

For RNAse protection analysis of HCD57, DA3-EPOR, and BAF3-EPOR cells, 5 × 106 cells per time point were deprived of EPO for 18 hours in the following manner: cells were washed 3 times in 10 mL media per 5 × 106 cells in IMDM media with no serum or growth factors and incubated in complete media without EPO for 18 hours prior to stimulation with either 1 U/mL EPO, 10 ng/mL stem cell factor (SCF), or 10 ng/mL TNF-α for the times indicated in the figure legends. For the inhibitor studies, the cells were pretreated with the indicated concentrations of inhibitor for 2 hours or 0.1% dimethyl sulfoxide (DMSO) vehicle control for 2 hours prior to addition of EPO for 4 hours. For the human cells, following isolation of purified CFU-E as previously described,352 × 107 cells were washed 3 times to eliminate EPO. The cells were cultured in the same serum-free media without EPO, and 1 × 107 cells were collected at 1 and 6 hours after EPO withdrawal. Cells were harvested, and total RNA was isolated using the RNeasy RNA isolation kit (Qiagen, Valencia, CA). The radioactive RNA probe was transcribed from the mCK-3 (Figure 1A-C), mCK-3b (Figure 1D), or hCK-3 (for the human CFU-E) template sets (BD Pharmingen, San Diego, CA) using 32P-UTP and an in vitro transcription kit (BD Pharmingen) RNAse protection was carried out using the Riboquant RNase Protection kit (BD Pharmingen) using 10 micrograms (μg) of total RNA for the mouse RNA and 5 μg for the human CFU-E RNA, and 5.9 × 105 cpm of probe per sample. Protected fragments were resolved on a 6% polyacrylamide, 7M urea gel and visualized by autoradiography for 24 hours at −80°C with an intensifying screen.

Fig. 1.

EPO induces the expression of TNF-α in hematopoietic cell lines.

All panels represent RNAse protection analysis of total RNA isolated from the cell lines indicated. Arrows on left indicate the presence of protected fragments for lymphotoxin β (LTβ), TNF-α, interleukin-6 (IL-6), TGF-β2 (TGF-β), interferon gamma (IFN-γ), interferon-β (IFN-β), macrophage inhibitory factor-1 (MIF-1), and housekeeping genes L32 and glyceraldehyde phosphate dehydrogenase (GADPH). P indicates undigested probe. (A) HCD57 cells were deprived of EPO for 18 hours and then stimulated with nothing (lane 1) or EPO (lanes 2-7) for the times indicated. Mouse control RNA (lane 8) and yeast RNA (lane 9) were used as positive and negative controls for the RNAse protection, respectively. Lines on right of panel indicate location of undigested probe; arrows on left indicate protected fragments indicative of expression of factors and housekeeping genes. (B) HCD57 cells were deprived of EPO overnight and then stimulated with nothing (lane 1), EPO (lanes 2-4), SCF (lanes 5, 6), or TNF-α (lanes 7, 8) for the times indicated. C indicates positive control RNA; Y, yeast RNA. (C) HCD57 (lanes 1, 2), DA3-EPOR (lanes 3-6), or BAF3-EPOR (lanes 7-10) cells were cultured either continuously in EPO (c, lanes 3, 7) or in 10 ng/mL IL-3 overnight (lanes 4, 8), or deprived of EPO overnight and then stimulated with nothing (lanes 1, 5, 9) or EPO (lanes 2, 6, 10) for 4 hours. Y indicates yeast RNA (lane 11). (D) HCD57 cells were deprived of EPO for 18 hours and then pretreated with DMSO vehicle (lane 2), 5 and 50 μM PD98059 (lanes 4, 5), 1 or 10 μM U0126 (lanes 6, 7), 5 or 50 μM LY294002 (lanes 8, 9), or 2 and 20 μM SB203580 (lanes 10, 11) for 2 hours prior to addition of EPO for 4 hours (lanes 2-11). C indicates positive control RNA; Y, yeast RNA; P, undigested probe.

Fig. 1.

EPO induces the expression of TNF-α in hematopoietic cell lines.

All panels represent RNAse protection analysis of total RNA isolated from the cell lines indicated. Arrows on left indicate the presence of protected fragments for lymphotoxin β (LTβ), TNF-α, interleukin-6 (IL-6), TGF-β2 (TGF-β), interferon gamma (IFN-γ), interferon-β (IFN-β), macrophage inhibitory factor-1 (MIF-1), and housekeeping genes L32 and glyceraldehyde phosphate dehydrogenase (GADPH). P indicates undigested probe. (A) HCD57 cells were deprived of EPO for 18 hours and then stimulated with nothing (lane 1) or EPO (lanes 2-7) for the times indicated. Mouse control RNA (lane 8) and yeast RNA (lane 9) were used as positive and negative controls for the RNAse protection, respectively. Lines on right of panel indicate location of undigested probe; arrows on left indicate protected fragments indicative of expression of factors and housekeeping genes. (B) HCD57 cells were deprived of EPO overnight and then stimulated with nothing (lane 1), EPO (lanes 2-4), SCF (lanes 5, 6), or TNF-α (lanes 7, 8) for the times indicated. C indicates positive control RNA; Y, yeast RNA. (C) HCD57 (lanes 1, 2), DA3-EPOR (lanes 3-6), or BAF3-EPOR (lanes 7-10) cells were cultured either continuously in EPO (c, lanes 3, 7) or in 10 ng/mL IL-3 overnight (lanes 4, 8), or deprived of EPO overnight and then stimulated with nothing (lanes 1, 5, 9) or EPO (lanes 2, 6, 10) for 4 hours. Y indicates yeast RNA (lane 11). (D) HCD57 cells were deprived of EPO for 18 hours and then pretreated with DMSO vehicle (lane 2), 5 and 50 μM PD98059 (lanes 4, 5), 1 or 10 μM U0126 (lanes 6, 7), 5 or 50 μM LY294002 (lanes 8, 9), or 2 and 20 μM SB203580 (lanes 10, 11) for 2 hours prior to addition of EPO for 4 hours (lanes 2-11). C indicates positive control RNA; Y, yeast RNA; P, undigested probe.

Close modal

Enzyme immunoassay

Triplicate samples of 1 × 105 HCD57 cells were cultured in 1 U/mL EPO or 10 nanograms (ng) SCF for 24, 48, 72, or 96 hours or with no growth factor for 96 hours as indicated in the figures. Culture media was filtered through a 0.45-mM filter and subjected to an enzyme immunoassay (EIA) using the TNF-α EIA kit from BD Pharmingen. TNF-α was quantitated against a standard curve of known concentrations of TNF-α.

MTT assay

HCD57 cells (1 × 105 in triplicate) were deprived of EPO as described above for 18 hours and then incubated with no additional growth factor, 1 U/mL EPO, 1 U/mL EPO with 0.01, 0.1, or 1.0 μg/mL anti–TNF-α neutralizing antibody, or 1, 10, 100, and 1000 ng/mL TNF-α alone for 48 hours. MTT in 1 × phosphate-buffered saline (PBS) was added to a final concentration of 5-μg/mL, and the cells were incubated for 4 hours at 37°C. The cells were then lysed with an equal volume of 0.2 N HCl in isopropanol, and the absorbance was read at 540 nM with a 630-nM reference filter.

Western blot analysis and in vitro kinase assay

For each time point, 5 × 106 cells were used. For Western blot analysis of JNK phosphorylation, the cells were washed to deprive them of EPO for 18 hours as indicated above. The cells were then stimulated with 1 U/mL EPO or 100, 10, or 1 ng/mL TNF-α for 2 hours at 37°C. The cells were lysed in 1 × sample buffer (0.05 M Tris [tris(hydroxymethyl)aminomethane], pH = 8, 2% sodium dodecyl sulfate, 0.1% bromophenol blue, 10% glycerol, 10% β-mercaptoethanol) and sonicated for 10 seconds each to shear the genomic DNA. Equal volumes (40 microliters [μL]) of sample were electrophoresed on an 8.5% sodium dodecyl sulfate–polyacrylamide gel (SDS-PAGE) and subjected to Western blot analysis with the phospho-specific antibodies JNK, ERK, and AKT, as previously described.12 Specific reactive proteins were detected using enhanced chemiluminescence (Amersham Biosciences, Piscataway, NJ). The blot was stripped as previously described37 and reprobed with an antibody to JNK-1 to ensure equal loading of proteins. For the in vitro kinase assays, 5 × 106 cells per sample were incubated in EPO in the absence or presence of anti–TNF-α antibody or IgG control antibody for 18 hours at 37°C. TNF-α (100 ng/mL) was added to one sample containing anti–TNF-α (Figure 4B, lane 5) 2 hours prior to cell harvesting. Total cell extracts were immunoprecipitated as previously described with anti–JNK-135 and subjected to an in vitro kinase assay according to Cell Signaling Technologies' protocol for the SAPK/JNK in vitro kinase assay. Then 20 μL of the assay was electrophoresed on a 10% acrylamide SDS-PAGE gel and subjected to Western blot analysis using a phospho-cJun–specific antibody (1:1000 dilution) overnight at 4°C. Following exposure of the phospho-cJun, the blot was stripped and then probed with the anti-JNK1 antibody to ensure equal loading of proteins.

Flow cytometry analysis of CD34+ cells

Following incubation of the human CD34+ cells in the presence or absence of neutralizing TNF-α antibodies, the cells were collected, washed once in 1 × fluorescence-activated cell-sorter scanner (FACS) buffer (1 × PBS/5% FCS/0.1% sodium azide), resuspended in 10 μg/mL AB24G2 antibody (BD Pharmingen) to block FCγRII receptors, and incubated for 10 minutes at 4°C. Phycoerythrin (PE)–labeled anti–glycophorin A monoclonal antibody (clone GA-R2, BD Pharmingen) or PE-labeled mouse IgG isotype control (clone 27-35, BD Pharmingen) was then added to a final concentration of 10 μg/mL and incubated for 30 minutes at 4°C. The cells were washed twice with FACS buffer and resuspended in FACS buffer, and gycophorin-A–positive cells were detected using a FACSscan flow cytometer (Becton Dickinson, Franklin Lakes, NJ) gated on an FL-2 channel.

Colony-forming cell assays

To assess murine CFU-E and burst-forming unit–erythroid (BFU-E) colony formation, 6 × 105 total murine bone marrow cells were added to 3 mL methylcellulose media (Methocult M3334, Stem Cell Technologies, Vancouver, BC, Canada) containing 3 units EPO/mL but no other cytokines in the presence or absence of 1 or 10 ng/mL TNF-α. Of these cells, 1.1 mL were plated in duplicate onto 30-mM plates and cultured at 37°C in a moist CO2 environment. CFU-Es were counted 2 days after the start of the experiment, and BFU-Es were counted 8 days after the start of the experiment.

As an initial screen to detect possible autocrine secretion of growth factors in HCD57 cells, we tested for the presence of likely candidate cytokines using RNAse protection analysis (RPA) templates that detect the mRNA expression of numerous cytokines. RPA of total RNA isolated from HCD57 cells cultured in the absence or presence of EPO revealed that TNF-α mRNA was expressed in the presence of EPO but expression was greatly reduced when EPO was removed (Figure1A). EPO induced the expression of TNF-α within 15 minutes of EPO addition (Figure 1B, lane 2). TNF-α expression reached a maximum 4 hours after EPO addition and was maintained over further incubation for 24 or 48 hours (Figure 1A, lanes 2-4). Other growth factors also were expressed (lymphotoxin B, interferon-γ, and TGF-β), but their expression was EPO independent. SCF also induced the expression of TNF-α, but this induction was weaker than the EPO-induced TNF-α expression. TNF-α did not affect its own expression (Figure 1B, lanes 7 and 8). To determine if the ability of EPO to activate TNF-α expression was unique to HCD57 cells, TNF-α expression was next tested in 2 cell lines ectopically expressing EPOR: DA3-EPOR and BAF3-EPOR cells. These cell lines are normally dependent on interleukin-3 (IL-3) for proliferation and survival, and EPOR can replace the IL-3 receptor in their proliferative and antiapoptotic properties.38 39 Both DA3-EPOR and BAF3-EPOR cells expressed TNF-α mRNA in response to EPO treatment (Figure 1C, lanes 6 and 10), indicating that EPOR has the capacity to signal TNF-α mRNA expression in other cells. IL-3 induced expression of TNF-α in DA3-EPOR cells (Figure 1C, lane 4) but not in BAF3-EPOR cells (Figure 1C, lane 8). DA3-EPOR and BAF3-EPOR cells also expressed interferon-β and IL-6, in addition to TGF-β and IFN-γ (Figure1C). Other human EPO-dependent cell lines tested (UT-7-EPO and TF-1) expressed TNF-α but did not express it in an EPO-dependent manner (data not shown).

The pathways upstream of EPO-induced TNF-α expression were next explored by treatment of HCD57 with inhibitors of known signal transduction pathways activated by EPO. Treatment with the PI3-kinase family inhibitor LY294002 inhibited EPO-induced TNF-α expression in HCD57 cells (Figure 1D, lanes 8 and 9); the map kinase kinase (MEK) inhibitors PD98059 and U0126 partially inhibited TNF-α expression in HCD57 cells (Figure 1D, lanes 4-7). The p38 inhibitor SB203580 had no significant effect on EPO-induced TNF-α activity at any concentration tested (Figure 1D, lanes 10 and 11). The inhibitors had no significant effects on the expression of other cytokines expressed, such as interferon-γ Therefore, EPO-induced expression of TNF-α is mediated in part by the activation of a PI3-kinase–related pathway or a non–PI3-kinase pathway inhibited by LY294002.

The ability of EPO to induce TNF-α protein secretion was then tested using an enzyme immunoassay. HCD57 cells secreted TNF-α into the media in response to EPO (Figure 2). SCF, which can promote proliferation but cannot promote survival of HCD57 cells, induced secretion of TNF-α within 48 hours of SCF addition (Figure 2, lane 8), but this amount did not increase further (Figure 2, lane 9).

Fig. 2.

EPO induces the secretion of TNF-α by HCD57 cells.

TNF-α enzyme immunoassay (EIA) of media harvested from HCD57 cells treated with no growth factors (lane 1), 1 U/mL EPO (lanes 2-5), or 10 ng/mL SCF (lanes 6, 7) for the number of hours indicated. TNF-α levels are measured in pg/mL compared with a standard curve using known quantities of TNF-α.

Fig. 2.

EPO induces the secretion of TNF-α by HCD57 cells.

TNF-α enzyme immunoassay (EIA) of media harvested from HCD57 cells treated with no growth factors (lane 1), 1 U/mL EPO (lanes 2-5), or 10 ng/mL SCF (lanes 6, 7) for the number of hours indicated. TNF-α levels are measured in pg/mL compared with a standard curve using known quantities of TNF-α.

Close modal

The role of TNF-α as an autocrine factor for proliferation in HCD57 cells was tested by assessing the ability of TNF-α to stimulate proliferation of these cells using a standard MTT dye reduction assay. Addition of exogenous TNF-α was able to induce proliferation in HCD57 cells in a dose-dependent manner. Likewise, treatment of HCD57 with neutralizing antibodies to TNF-α inhibited EPO-induced proliferation in a dose-dependent manner, and the inhibition could be partially reversed by the addition of excess TNF-α to the media (Figure 3B, lane 6).

Fig. 3.

TNF-α induces proliferation of HCD57 cells, and a neutralizing antibody to TNF-α inhibits EPO-induced proliferation of these cells.

Proliferation was measured using the MTT dye reduction assay as indicated in “Materials and methods.” (A) HCD57 (lanes 1-5) cells were deprived of EPO for 18 hours prior to addition of 1 (lane 2), 10 (lane 3), 100 (lane 4), or 1000 (lane 5) ng/mL TNF-α for 48 hours. (B) HCD57 cells were deprived of EPO overnight and then treated with EPO in the presence (lanes 3-6) or absence (lane 2) of neutralizing anti–TNF-α antibody for 48 hours. Indicated is μg/mL neutralizing antibody added. Excess TNF-α (10 ng/mL) was added to counteract the effect of the neutralizing antibody (lane 6).

Fig. 3.

TNF-α induces proliferation of HCD57 cells, and a neutralizing antibody to TNF-α inhibits EPO-induced proliferation of these cells.

Proliferation was measured using the MTT dye reduction assay as indicated in “Materials and methods.” (A) HCD57 (lanes 1-5) cells were deprived of EPO for 18 hours prior to addition of 1 (lane 2), 10 (lane 3), 100 (lane 4), or 1000 (lane 5) ng/mL TNF-α for 48 hours. (B) HCD57 cells were deprived of EPO overnight and then treated with EPO in the presence (lanes 3-6) or absence (lane 2) of neutralizing anti–TNF-α antibody for 48 hours. Indicated is μg/mL neutralizing antibody added. Excess TNF-α (10 ng/mL) was added to counteract the effect of the neutralizing antibody (lane 6).

Close modal

TNF-α has been shown previously to activate JNK and ERK, 2 kinases which have been shown to be important in the proliferation of erythroid cells.12 We therefore explored the ability of these kinases to be activated by TNF-α in HCD57 cells. We previously have reported that TNF-α induced the activation of JNK within 2 hours of cytokine addition in HCD57 cells.12 Treatment of HCD57 cells for 2 hours with increasing amounts of TNF-α resulted in a dose-dependent increase in phosphorylation of JNK, thus confirming and extending our previously published results (Figure4A). No phosphorylation of ERK or AKT in response to TNF-α treatment was detected. Furthermore, treatment of HCD57 cells with the TNF-α neutralizing antibody inhibited JNK activity (Figure 4B, lanes 3 and 4). This effect could be reversed by the addition of exogenous TNF-α (Figure 4B, lane 2) and was not seen with a rat IgG control antibody (Figure 4B, lane 5). Therefore, TNF-α may transduce its signal in erythroid cells via activation and activity of JNK.

Fig. 4.

TNF-α activates JNK in HCD57 cells.

(A) Western blot analysis of HCD57 cells deprived of EPO for 18 hours prior to treatment. Cells were treated with nothing (lane 1), 100, 10, or 1 ng/mL TNF-α (lanes 2-4), or EPO (1 U/mL) (lane 5), and whole cell lysates were probed for anti–phospho-JNK1/2 (top panel), anti–phospho AKT and anti–phospho ERK1/2 (middle panel), and anti-JNK1 (bottom panel). (B) In vitro kinase assay of JNK1 immunoprecipitates using glutathione-S–transferase (GST)–cJun as a substrate from HCD57 cells treated with EPO in either the absence (lane 1) or presence of 0.1 or 1.0 μg/mL anti–TNF-α antibody (lanes 3, 4), 1.0 μg/mL anti–TNF-α antibody plus 100 ng TNF-α (lane 2), or goat IgG control (lane 5). Shown are 2 separate experiments to indicate reproducibility of the result. Phosphorylated GST-cJun (top panel) and total JNK1 (bottom panel) are indicated.

Fig. 4.

TNF-α activates JNK in HCD57 cells.

(A) Western blot analysis of HCD57 cells deprived of EPO for 18 hours prior to treatment. Cells were treated with nothing (lane 1), 100, 10, or 1 ng/mL TNF-α (lanes 2-4), or EPO (1 U/mL) (lane 5), and whole cell lysates were probed for anti–phospho-JNK1/2 (top panel), anti–phospho AKT and anti–phospho ERK1/2 (middle panel), and anti-JNK1 (bottom panel). (B) In vitro kinase assay of JNK1 immunoprecipitates using glutathione-S–transferase (GST)–cJun as a substrate from HCD57 cells treated with EPO in either the absence (lane 1) or presence of 0.1 or 1.0 μg/mL anti–TNF-α antibody (lanes 3, 4), 1.0 μg/mL anti–TNF-α antibody plus 100 ng TNF-α (lane 2), or goat IgG control (lane 5). Shown are 2 separate experiments to indicate reproducibility of the result. Phosphorylated GST-cJun (top panel) and total JNK1 (bottom panel) are indicated.

Close modal

The EPO-induced expression of TNF-α in these cell lines led us to investigate whether primary erythroid cells might express TNF-α. We therefore investigated the ability of EPO to induce TNF-α in both human colony-forming cells (CFU-E)35 and CD34+human erythroid progenitors,36 and in primary murine erythroid progenitors purified from the spleens of mice infected with the anemia strain of the Friend spleen focus-forming virus (FVA cells).40 RNAse protection analysis of CFU-Es cultured in the presence of EPO revealed expression of TNF-α (Figure 5, lane 3). When EPO was removed, TNF-α expression decreased, whereas the expression of TGF-β increased (Figure 5, lane 1). CD34+ human cells and FVA cells also expressed TNF-α in the absence of EPO (data not shown). Addition of EPO to either the CD34+ or the FVA cells, however, failed to further induce TNF-α expression (data not shown).

Fig. 5.

TNF-α is expressed in primary human colony-forming cells.

RNAse protection analysis of human CFU-E cultured in the presence of EPO (lane 3) or deprived of EPO for 1.5 and 6 hours (lanes 1, 2). Cytokines expressed are indicated by arrows.

Fig. 5.

TNF-α is expressed in primary human colony-forming cells.

RNAse protection analysis of human CFU-E cultured in the presence of EPO (lane 3) or deprived of EPO for 1.5 and 6 hours (lanes 1, 2). Cytokines expressed are indicated by arrows.

Close modal

Because TNF-α traditionally has been considered an inhibitor of erythropoiesis,41 we wished to investigate the effect of TNF-α on primary cell erythropoiesis in both human and murine erythroid cells. First, human CD34+ cells were cultured in EPO for 7 days with neutralizing antibodies to TNF-α, and the number of mature erythroid cells was determined by immunostaining against the erythroid-specific protein glycophorin A (Figure6A). During the experiment, approximately 10% of total cells were found to express glycophorin A after culture (data not shown). The presence of neutralizing TNF-α antibody resulted in a dose-dependent increase in the number of glycophorin A–positive cells, indicating that TNF-α was inhibiting erythroid cell proliferation in this system. To investigate the effect of TNF-α on murine erythropoiesis, bone marrow cells were isolated from wild-type and TNF-α−/− mice, and these cells were assayed for CFU-Es and BFU-Es in either the absence or presence of TNF-α in semisolid media. TNF-α had no significant effect on the number of CFU-Es in these experiments (data not shown). Furthermore, the addition of TNF-α had no significant effect on the number of BFU-Es in TNF-α wild-type mice (Figure 6B, lanes 2 and 3). However, the addition of TNF-α to TNF-α–deficient bone marrow cells inhibited the formation of BFU-Es in a dose-dependent manner (Figure 6B, lanes 5 and 6). Taken together, these results suggest that TNF-α has an inhibitory effect on both human and murine normal erythropoiesis in vitro.

Fig. 6.

Effects of TNF-α on human and murine primary erythroid cells.

(A) Human CD34+ cells were cultured in EPO or in EPO with 0.2, 2.0, or 20.0 μg/mL neutralizing TNF-α antibody for 7 days and assessed for glycophorin A expression by flow cytometry. Increasing amounts of antibody resulted in an increase in the number of glycophorin A–positive cells (lanes 2-4), whereas the addition of control IgG had no effect (lane 5). (B) Total bone marrow isolated from TNF-α wild-type (WT) (lanes 1-3) and TNF-α−/− (lanes 4-6) mice was incubated in EPO alone (lanes 1, 4) or EPO with 1 or 10 ng/mL TNF-α (lanes 2, 3, 5, 6) in semisolid media. The number of BFU-Es was counted 8 days after the start of the experiment and is expressed as number of BFU-Es detected per 30-mM plate.

Fig. 6.

Effects of TNF-α on human and murine primary erythroid cells.

(A) Human CD34+ cells were cultured in EPO or in EPO with 0.2, 2.0, or 20.0 μg/mL neutralizing TNF-α antibody for 7 days and assessed for glycophorin A expression by flow cytometry. Increasing amounts of antibody resulted in an increase in the number of glycophorin A–positive cells (lanes 2-4), whereas the addition of control IgG had no effect (lane 5). (B) Total bone marrow isolated from TNF-α wild-type (WT) (lanes 1-3) and TNF-α−/− (lanes 4-6) mice was incubated in EPO alone (lanes 1, 4) or EPO with 1 or 10 ng/mL TNF-α (lanes 2, 3, 5, 6) in semisolid media. The number of BFU-Es was counted 8 days after the start of the experiment and is expressed as number of BFU-Es detected per 30-mM plate.

Close modal

TNF-α has been shown to be a potent inhibitor of hematopoiesis.30,41,42 TNF-α inhibition of erythropoiesis has been demonstrated in normal hematopoietic progenitors,43,44 and TNF-α expression and suppression of erythropoiesis has been associated with a number of hematopoietic disorders such as Fanconi anemia,45 myelodysplastic disease,46 aplastic anemia,47 and anemia due to chronic disease.32,48 49 The finding that HCD57 cells could not only express and secrete TNF-α in response to EPO, but also respond to TNF-α with enhanced proliferation is therefore intriguing. We also detected EPO-inducible TNF-α expression in DA3-EPOR and BAF3-EPOR cells, indicating that the ability of EPOR to transduce a TNF-α–inducing signal is not a unique property of HCD57 cells. The EPO-dependent expression of TNF-α may be mediated by PI3-kinase activation, since treatment with the PI3-kinase inhibitor LY294002 greatly inhibited TNF-α expression in both HCD57 cells. Treatment with the MEK inhibitors PD90859 and U0126 also partially inhibited TNF-α expression in HCD57 cells, suggesting a partial contribution of the ERK/MAP kinase pathway to TNF-α expression as well.

TNF-α activated the JNK pathway in a dose-dependent manner in HCD57 cells (Figure 4). The fact that proliferation, JNK activation, and JNK activity were only partially inhibited by the neutralizing antibody to TNF-α suggests that additional EPO-dependent pathways also contribute to these processes. Alternatively, there may be internal TNF-α that is not secreted, so that the neutralizing antibody would have no effects on these internal TNF-α–activated events.

Our finding that TNF-α had the capacity to induce proliferation of erythroleukemia cells led us to investigate the expression of TNF-α in primary erythroid cells and the effect of TNF-α on erythropoiesis. We detected TNF-α expression in both human and murine primary erythroid cells. This expression, however, was not reliably shown to be EPO dependent. The expression of TNF-α in FVA primary proerythroblasts may be constitutive due to activation of signaling pathways by Friend virus infection, because the virus activates the MAP kinase pathway but not other EPO-dependent signals.50TNF-α expression also has recently been reported in CD34+hematopoietic cells and BFU-E cells that we now confirm in CD34+ human cells.51 The decrease in TNF-α expression upon EPO withdrawal from the human CFU-E for 6 hours suggests that EPO-dependent TNF-α expression may occur in these cells. We cannot rule out that the decrease in TNF-α expression was due to a general down-regulation of transcription that occurs when the cells undergo apoptosis due to EPO withdrawal; however, the increase in TGF-β expression upon EPO withdrawal (Figure 5, lane 1) suggests that not all messages are down-regulated upon EPO withdrawal. This result strongly suggests that the TNF-α in the purified primary cells does not arise from contaminating nonerythroid cells but from the erythroid cells themselves.

Our results in both human CD34+ cells treated with neutralizing antibodies to TNF-α and in murine bone marrow cells isolated from TNF-α–deficient mice treated with TNF-α indicate that in these systems, TNF-α inhibits either the proliferation or differentiation and/or induces apoptosis of maturing erythroid cells. Therefore, the EPO-dependent TNF-α secretion and proliferation from HCD57 cells may be a result of the transformation of the cells and may not be an inherent property of primary erythroid progenitors. This still does not explain, however, how HCD57 cells can proliferate in response to TNF-α and activate JNK in response to TNF-α, whereas differentiation of erythroid progenitors is inhibited by TNF-α. TNF-α has been reported to induce apoptosis through sustained activation of JNK.52 However, in many systems, TNF-α–induced JNK activation does not induce apoptosis and has even been reported to be cytoprotective.53 It also has been demonstrated that TNF-α can be a synergistic inducer of proliferation in immature CD34+/CD38 cells34 and may induce proliferation of multipotent hematopoietic progenitors while inhibiting the development of committed progenitors.54 HCD57 erythroleukemia cells are arrested at an early stage in erythroid development, as evidenced by the fact that forced expression of the activator protein-1 (AP1) transcription factor JunB induced the expression of some mature erythroid markers such as β-globin and spectrin-α and required at least 48 hours before these markers were seen.55 It is possible, therefore, that committed erythroid cells must reach a certain stage at which they are insensitive to inhibition by TNF-α. Alternatively, there may be a loss of a proapoptotic signal usually stimulated by JNK that is absent in HCD57 cells. There also may be a gain of an antiapoptotic TNF-α–induced signal (such as NF-κB56,57) or an antiapoptotic signal unrelated to TNF-α (such as Bcl-x(L)58) that suppresses TNF-α–induced apoptosis. Therefore, it might be possible to render these cells sensitive to TNF-α–induced killing or reduced proliferation by identifying and suppressing these pathways.

TNF-α also has been shown to induce the proliferation of myeloid leukemia cells lines,22 but this effect has not been demonstrated for an erythroleukemia cell line. It is possible that the HCD57 cell line has acquired, as a part of its leukemic phenotype, the characteristics of other myeloid lineages; however, the expression of mature erythroid proteins by the induction of JunB or hemin indicate that it has retained the characteristics of an immature erythroid cell.55,59 The acquisition of EPO-dependent TNF-α expression may benefit erythroleukemic cells by gaining the ability to induce its own proliferation and/or by killing cells in the bone marrow, spleen, or blood that might compete with the leukemic cells for resources. Inhibition of this TNF-α autocrine loop may therefore provide a means to specifically inhibit the leukemic cell proliferation. It is interesting that TGF-β is very strongly expressed in HCD57, DA3-EPOR, and BAF3-EPOR (Figure 1), in human primary colony-forming cells (Figure 6), and in FVA cells. Recent studies have indicated that TGF-β may drive the differentiation of erythroid progenitors and EPO-dependent erythroid cell lines; autocrine TGF-β may therefore play a role in this process.60 61 

In conclusion, this study demonstrates that some erythroid cell lines have the capacity to proliferate in response to TNF-α and that EPO-activated EPOR has the capacity to induce the synthesis and secretion of TNF-α in some cells but not others. TNF-α appears to induce proliferation by modulation of the JNK pathway. Inhibition of this TNF-α–dependent modulation of JNK slowed the proliferation of these leukemic cells. The identification of this autocrine loop suggests the possibility that anti–TNF-α strategies may be useful in inhibiting the proliferation or survival of these leukemias in a clinical setting. The study also suggests the possible existence of a negative feedback of TNF-α expressed by mouse and human CFU-Es or proerythroblasts that act on immature erythroid progenitors to suppress maturation of these cells.

The authors would like to thank Dr Maurice Bondurant for his assistance with RNAse protection analysis of primary murine erythroid progenitors. D.L.B is a research scientist at the National Cancer Institute of Canada.

Prepublished online as Blood First Edition Paper, August 29, 2002; DOI 10.1182/blood-2001-11-0084.

Supported by grants R01DK39781 (S.T.S.), R01HL65906 (S.T.S.), RO1 AI43433 (J.J.R.), RO1CA91839 (J.J.R.), R01CA88906 (P.D.), and R01DK52825 (P.D.) from the National Institutes of Health; grant 9804806U from the American Heart Association (S.M.J.-H.); Intramural Research Grant (IRG)-100036 from the American Cancer Society (S.M.J.-H.); grant 98-0148 (P.D.) from the Department of Defense; the Department of Veterans Affairs (E.N.D.); and Canadian Institute for Health Research (D.L.B.).

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.

1
Sawyer
ST
Penta
K
Association of JAK2 and STAT5 with erythropoietin receptors: role of receptor phosphorylation in erythropoietin signal transduction.
J Biol Chem.
271
1996
32430
32437
2
Witthuhn
BA
Quelle
FW
Silvennoinen
O
et al
JAK2 associates with the erythropoietin receptor and is tyrosine phosphorylated and activated following stimulation with erythropoietin.
Cell.
74
1993
227
236
3
Miura
O
Nakamura
N
Quelle
FW
Witthuhn
BA
Ihle
JN
Aoki
N
Erythropoietin induces association of the JAK2 protein tyrosine kinase with the erythropoietin receptor in vivo.
Blood.
84
1994
1501
1507
4
Barber
DL
D'Andrea
AD
Erythropoietin and interleukin-2 activate distinct JAK kinase family members.
Mol Cell Biol.
14
1994
6506
6514
5
Damen
JE
Mui
AL
Puil
JL
Pawson
T
Krystal
G
Phosphatidylinositol 3-kinase associates, via its Src homology 2 domains, with the activated erythropoietin receptor.
Blood.
81
1993
3204
3210
6
He
TC
Zhuang
H
Quelle
DE
Wojchowski
DM
Association of the p85 regulatory subunit of phosphatidylinositol 3-kinase with an essential erythropoietin receptor subdomain.
Blood.
82
1993
3530
3538
7
Damen
JE
Liu
L
Cutler
RL
Krystal
G
Erythropoietin stimulates the tyrosine phosphorylation of Shc and its association with Grb2 and a 145-Kd tyrosine phosphorylated protein.
Blood.
82
1993
2296
2303
8
He
TC
Jiang
N
Zhuang
H
Wojchowski
DM
Erythropoietin-induced recruitment of Shc via a receptor phosphotyrosine-independent, Jak2-associated pathway.
J Biol Chem.
270
1995
11055
11061
9
Barber
DL
Corless
CN
Xia
K
Roberts
TM
D'Andrea
AD
Erythropoietin activates Raf1 by an Shc-independent pathway in CTLL-EPO-R cells.
Blood.
89
1997
55
64
10
Damen
JE
Liu
L
Rosten
P
et al
The 145-kDa protein induced to associate with Shc by multiple cytokines is an inositol tetraphosphate and phosphatidylinositol 3,4,5-triphosphate 5-phosphatase.
Proc Natl Acad Sci U S A.
93
1996
1689
1693
11
Boer
AK
Drayer
AL
Vellenga
E
Effects of overexpression of the SH-2–containing inositol phosphatase SHIP on proliferation and apoptosis of erythroid AS-E2 cells.
Leukemia.
15
2001
1750
1757
12
Jacobs-Helber
SM
Ryan
JJ
Sawyer
ST
JNK and p38 are activated by erythropoietin (EPO) but are not induced in apoptosis following EPO withdrawal in EPO-dependent HCD57 cells.
Blood.
96
2000
933
940
13
Nagata
Y
Nishida
E
Todokoro
K
Activation of JNK signaling pathway by erythropoietin, thrombopoietin, and interleukin-3.
Blood.
89
1997
2664
2669
14
Jacobs-Helber
SM
Penta
K
Sun
ZH
Lawson
A
Sawyer
ST
Distinct signaling from stem cell factor and erythropoietin in HCD57 cells.
J Biol Chem.
272
1997
6850
6853
15
Devemy
E
Billat
C
Haye
B
Activation of Raf-1 and mitogen-activated protein kinases by erythropoietin and inositolphosphate-glycan in normal erythroid progenitor cells: involvement of protein kinase C.
Cell Signal.
9
1997
41
46
16
Nagata
Y
Takahashi
N
Davis
RJ
Todokoro
K
Activation of p38 MAP kinase and JNK but not ERK is required for erythropoietin-induced erythroid differentiation.
Blood.
92
1998
1859
1869
17
Nagata
Y
Todokoro
K
Requirement of activation of JNK and p38 for environmental stress-induced erythroid differentiation and apoptosis and of inhibition of ERK for apoptosis.
Blood.
94
1999
853
863
18
Lawson
AE
Bao
H
Wickrema
A
Jacobs-Helber
SM
Sawyer
ST
Phosphatase inhibition promotes antiapoptotic but not proliferative signaling pathways in erythropoietin-dependent HCD57 cells.
Blood.
96
2000
2084
2092
19
Witt
O
Sand
K
Pekrun
A
Butyrate-induced erythroid differentiation of human K562 leukemia cells involves inhibition of ERK and activation of p38 MAP kinase pathways.
Blood.
95
2000
2391
2396
20
Bazzoni
F
Beutler
B
The tumor necrosis factor ligand and receptor families.
N Engl J Med.
334
1996
1717
1725
21
Wallach
D
Varfolomeev
EE
Malinin
NL
Goltsev
YV
Kovalenko
AV
Boldin
MP
Tumor necrosis factor receptor and Fas signaling mechanisms.
Annu Rev Immunol.
17
1999
331
367
22
Liu
RY
Fan
C
Liu
G
Olashaw
NE
Zuckerman
KS
Activation of p38 mitogen-activated protein kinase is required for tumor necrosis factor-alpha -supported proliferation of leukemia and lymphoma cell lines.
J Biol Chem.
275
2000
21086
21093
23
Witsell
AL
Schook
LB
Tumor necrosis factor alpha is an autocrine growth regulator during macrophage differentiation.
Proc Natl Acad Sci U S A.
89
1992
4754
4758
24
Branch
DR
Guilbert
LJ
Autocrine regulation of macrophage proliferation by tumor necrosis factor-alpha.
Exp Hematol.
24
1996
675
681
25
Reittie
JE
Yong
KL
Panayiotidis
P
Hoffbrand
AV
Interleukin-6 inhibits apoptosis and tumour necrosis factor induced proliferation of B-chronic lymphocytic leukaemia.
Leuk Lymphoma.
22
1996
83
90
26
Moberts
R
Hoogerbrugge
H
van Agthoven
T
Lowenberg
B
Touw
I
Proliferative response of highly purified B chronic lymphocytic leukemia cells in serum free culture to interleukin-2 and tumor necrosis factors alpha and beta.
Leuk Res.
13
1989
973
980
27
Marth
C
Zeimet
AG
Herold
M
et al
Different effects of interferons, interleukin-1beta and tumor necrosis factor-alpha in normal (OSE) and malignant human ovarian epithelial cells.
Int J Cancer.
67
1996
826
830
28
Quentmeier
H
Dirks
WG
Fleckenstein
D
Zaborski
M
Drexler
HG
Tumor necrosis factor-alpha-induced proliferation requires synthesis of granulocyte-macrophage colony-stimulating factor.
Exp Hematol.
28
2000
1008
1015
29
Moldawer
LL
Marano
MA
Wei
H
et al
Cachectin/tumor necrosis factor-alpha alters red blood cell kinetics and induces anemia in vivo.
FASEB J.
3
1989
1637
1643
30
Akahane
K
Hosoi
T
Urabe
A
Kawakami
M
Takaku
F
Effects of recombinant human tumor necrosis factor (rhTNF) on normal human and mouse hemopoietic progenitor cells.
Int J Cell Cloning.
5
1987
16
26
31
Ulich
TR
del Castillo
J
Yin
S
Tumor necrosis factor exerts dose-dependent effects on erythropoiesis and myelopoiesis in vivo.
Exp Hematol.
18
1990
311
315
32
Means
RT
Jr
Dessypris
EN
Krantz
SB
Inhibition of human colony-forming-unit erythroid by tumor necrosis factor requires accessory cells.
J Clin Invest.
86
1990
538
541
33
Means
RT
Jr
Krantz
SB
Inhibition of human erythroid colony-forming units by tumor necrosis factor requires beta interferon.
J Clin Invest.
91
1993
416
419
34
Snoeck
HW
Weekx
S
Moulijn
A
et al
Tumor necrosis factor alpha is a potent synergistic factor for the proliferation of primitive human hematopoietic progenitor cells and induces resistance to transforming growth factor beta but not to interferon gamma.
J Exp Med.
183
1996
705
710
35
Jacobs-Helber
SM
Wickrema
A
Birrer
MJ
Sawyer
ST
AP1 regulation of proliferation and initiation of apoptosis in erythropoietin-dependent erythroid cells.
Mol Cell Biol.
18
1998
3699
3707
36
Angchaisuksiri
P
Grigus
SR
Carlson
PL
Krystal
GW
Dessypris
EN
Secretion of a unique peptide from interleukin-2-stimulated natural killer cells that induces endomitosis in immature human megakaryocytes.
Blood.
99
2002
130
136
37
Penta
K
Sawyer
ST
Erythropoietin induces the tyrosine phosphorylation, nuclear translocation, and DNA binding of STAT1 and STAT5 in erythroid cells.
J Biol Chem.
270
1995
31282
31287
38
Miura
O
Ihle
JN
Subunit structure of the erythropoietin receptor analyzed by 125I-Epo cross-linking in cells expressing wild-type or mutant receptors.
Blood.
81
1993
1739
1744
39
Carroll
M
Zhu
Y
D'Andrea
AD
Erythropoietin-induced cellular differentiation requires prolongation of the G1 phase of the cell cycle.
Proc Natl Acad Sci U S A.
92
1995
2869
2873
40
Bondurant
M
Koury
M
Krantz
SB
Blevins
T
Duncan
DT
Isolation of erythropoietin-sensitive cells from Friend virus-infected marrow cultures: characteristics of the erythropoietin response.
Blood.
61
1983
751
758
41
Johnson
CS
Cook
CA
Furmanski
P
In vivo suppression of erythropoiesis by tumor necrosis factor-alpha (TNF-alpha): reversal with exogenous erythropoietin (EPO).
Exp Hematol.
18
1990
109
113
42
Bonnet
D
Lemoine
FM
Najman
A
Guigon
M
Comparison of the inhibitory effect of AcSDKP, TNF-alpha, TGF-beta, and MIP-1 alpha on marrow-purified CD34+ progenitors.
Exp Hematol.
23
1995
551
556
43
Rusten
LS
Jacobsen
SE
Tumor necrosis factor (TNF)-alpha directly inhibits human erythropoiesis in vitro: role of p55 and p75 TNF receptors.
Blood.
85
1995
989
996
44
Jacobsen
FW
Rothe
M
Rusten
L
et al
Role of the 75-kDa tumor necrosis factor receptor: inhibition of early hematopoiesis.
Proc Natl Acad Sci U S A.
91
1994
10695
10699
45
Otsuki
T
Nagakura
S
Wang
J
Bloom
M
Grompe
M
Liu
JM
Tumor necrosis factor-alpha and CD95 ligation suppress erythropoiesis in Fanconi anemia C gene knockout mice.
J Cell Physiol.
179
1999
79
86
46
Kitagawa
M
Saito
I
Kuwata
T
et al
Overexpression of tumor necrosis factor (TNF)-alpha and interferon (IFN)-gamma by bone marrow cells from patients with myelodysplastic syndromes.
Leukemia.
11
1997
2049
2054
47
Schultz
JC
Shahidi
NT
Detection of tumor necrosis factor-alpha in bone marrow plasma and peripheral blood plasma from patients with aplastic anemia.
Am J Hematol.
45
1994
32
38
48
Allen
DA
Breen
C
Yaqoob
MM
Macdougall
IC
Inhibition of CFU-E colony formation in uremic patients with inflammatory disease: role of IFN-gamma and TNF-alpha.
J Investig Med.
47
1999
204
211
49
Jongen-Lavrencic
M
Peeters
HR
Backx
B
Touw
IP
Vreugdenhil
G
Swaak
AJ
R-h-erythropoietin counteracts the inhibition of in vitro erythropoiesis by tumour necrosis factor alpha in patients with rheumatoid arthritis.
Rheumatol Int.
14
1994
109
113
50
Muszynski
KW
Ohashi
T
Hanson
C
et al
Both the polycythemia and anemia-inducing strains of Friend spleen focus-forming virus induce constitutive activation of Raf-1/mitogen-activated protein kinase signal transduction pathway.
J Virol.
72
1998
919
925
51
Majka
M
Janowska-Wieczorek
A
Ratajczak
J
et al
Numerous growth factors, cytokines, and chemokines are secreted by human CD34+ cells, myeloblasts, erythroblasts, and megakaryoblasts and regulate normal hematopoiesis in an autocrine/paracrine manner.
Blood.
97
2001
3075
3085
52
Guo
YL
Baysal
K
Kang
B
Yang
LJ
Williamson
JR
Correlation between sustained c-Jun N-terminal protein kinase activation and apoptosis induced by tumor necrosis factor-α in rat mesangial cells.
J Biol Chem.
273
1998
4027
4034
53
Sanna
MG
da Silva
CJ
Ducrey
O
et al
IAP suppression of apoptosis involves distinct mechanisms: the TAK1/JNK1 signaling cascade and caspase inhibition.
Mol Cell Biol.
22
2002
1754
1766
54
Rogers
JA
Berman
JW
TNF-alpha inhibits the further development of committed progenitors while stimulating multipotential progenitors in mouse long-term bone marrow cultures.
J Immunol.
153
1994
4694
4703
55
Jacobs-Helber
SM
Abutin
RM
Tian
C
Bondurant
M
Wickrema
A
Sawyer
ST
Role of JunB in erythroid differentiation.
J Biol Chem.
277
2002
4859
4866
56
Wang
CY
Mayo
MW
Baldwin
AS
Jr
TNF- and cancer therapy-induced apoptosis: potentiation by inhibition of NF-kappaB.
Science.
274
1996
784
787
57
Liu
ZG
Hsu
H
Goeddel
DV
Karin
M
Dissection of TNF receptor 1 effector functions: JNK activation is not linked to apoptosis while NF-kappaB activation prevents cell death.
Cell.
87
1996
565
576
58
Manna
SK
Haridas
V
Aggarwal
BB
Bcl-x(L) suppresses TNF-mediated apoptosis and activation of nuclear factor-kappaB, activation protein-1, and c-Jun N-terminal kinase.
J Interferon Cytokine Res.
20
2000
725
735
59
Spivak
JL
Pham
T
Isaacs
M
Hankins
WD
Erythropoietin is both a mitogen and a survival factor.
Blood.
77
1991
1228
1233
60
Zermati
Y
Fichelson
S
Valensi
F
et al
Transforming growth factor inhibits erythropoiesis by blocking proliferation and accelerating differentiation of erythroid progenitors.
Exp Hematol.
28
2000
885
894
61
Zermati
Y
Varet
B
Hermine
O
TGF-beta1 drives and accelerates erythroid differentiation in the epo-dependent UT-7 cell line even in the absence of erythropoietin.
Exp Hematol.
28
2000
256
266

Author notes

Stephen T. Sawyer, Department of Pharmacology/Toxicology, PO Box 980613, Richmond, VA 23298; e-mail:ssawyer@hsc.vcu.edu.

Sign in via your Institution