Recombinant human thrombopoietin (TPO) stimulates erythropoiesis by inhibiting erythroid progenitor...

Recombinant human thrombopoietin (TPO) stimulateserythropoiesis by inhibiting erythroid progenitor cell apoptosis

MARIUSZ Z. RATAJCZ AK, JANINA RATAJC ZAK, WOYTEK MARLICZ,† CHARLES H. PLETCHER, JR, BOGDAN MACHAL INSKI,JO NN I MO ORE, HSIAO-LING HUNG AND AL AN M. GEWIRT Z* Departments of Pathology and Laboratory Medicine, and*Internal Medicine, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania, U.S.A.

Received 14 November 1996; accepted for publication 14 April 1997

Summary. Thrombopoietin (TPO) has been reported tostimulate erythropoiesis, but the stimulatory mechanismhas not been defined. To address this issue, we performedserum-free cell-culture experiments with recombinanthuman TPO and purified human progenitor cells. Wefound that TPO alone was able to stimulate the mega-karyocyte colony formation in serum-free cultures, buterythroid colonies were never observed. Only in the presenceof EPO (erythropoietin) þIL-3 was TPO able to stimulate asmall increase (,25%) in erythroid colony formation.Accordingly, we hypothesized that TPO might have aneffect on erythroid progenitor cell viability, rather than adirect stimulatory effect. To test this idea, CD34þ cells werecultured for 7 d in serum-free methylcellulose in thepresence or absence of TPO, after which time KL+ EPO wasadded to the cultures. Cells which were pre-cultured for 7 d

in the presence of TPO gave rise to approximately 6 timesas many burst forming unit-erythroid (BFU-E) coloniesas cells which were pre-cultured in the absence ofTPO. Further, when primitive CD34þ, Kitþ MNC werecultured for 3–7 d under serum-free conditions in thepresence or absence of TPO, significantly fewer cells culturedin the presence of TPO displayed apoptotic changes whencompared to cells cultured in the absence of TPO. Takentogether, these results suggest that TPO has little directstimulatory effect on erythroid progenitor cells, but mightindirectly enhance erythropoiesis by preventing very earlyerythroid progenitor cells from undergoing apoptotic celldeath.

Keywords: thrombopoietin, erythropoiesis, erythroid pro-genitor cell apoptosis.

Erythrocytes and megakaryocytes have been postulated toderive from a common bi-potent progenitor cell (McDonald &Sullivan, 1993). Originally supported by indirect evidence,such as the simultaneous expression of erythroid (glyco-phorin A) and megakaryocyte (glycoprotein IIb/IIIa)lineage markers on primitive haemopoietic cells (Rowleyet al, 1992), more direct proof of the existence of acommon progenitor has recently been provided by cellcloning studies (Debili et al, 1996). Given the closeness oftheir origins, it may not be surprising that two of themajor regulators of erythroid and megakaryocyte develop-ment, EPO and TPO, have been shown to have a highdegree of amino terminal amino acid sequence homology(Lok et al, 1994). In this same regard, EPO has mild,but consistently observed, trophic effects on in vitro

megakaryocyte development (An et al, 1994; Dessypriset al, 1987; Ishibashi et al, 1987; Tsukada et al, 1992),whereas TPO has been reported to stimulate erythroid cellgrowth in a variety of in vitro and in vivo systems (Fibbe et al,1995; Kaushansky et al, 1995; Kobayashi et al, 1995).

Nevertheless, since it is difficult to discern direct causeand effect with an in vivo model, and since the culturestudies cited above were not carried out under serum-freeconditions, it remains uncertain if TPO exerts a directstimulatory effect on erythroid progenitor cells, and evenless clear whether such stimulation is of physiologicalsignificance. The aim of this study was to address theseissues by examining TPO’s effects on growth of purifiednormal human haemopoietic progenitor cells in serum-free cultures plated at normal and low density. Whenexamined under such conditions we found that TPO alone,or in combination with other cytokines, possessed littleerythroid colony stimulating ability. Rather, TPO appeared toprevent primitive erythroid progenitor cells from undergoingapoptosis. Such an effect could explain the apparent

British Journal of Haematology, 1997, 98, 8–17

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† On leave from Department of Cell Pathology, Pommeranian Schoolof Medicine, Szczecin, Poland.

Correspondence: Dr Alan M. Gewirtz, Rm 514, Stellar-ChanceLaboratories, University of Pennsylvania School of Medicine, 422Curie Blvd, Philadelphia, PA 19104, U.S.A.

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erythroid stimulatory activity that has been attributed to thiscytokine.

MATERIALS AND METHODS

Progenitor cell isolation and enrichment. Normal light-density bone marrow mononuclear cells (MNC) wereobtained from eight consenting donors and depleted ofadherent cells and T lymphocytes (A¹T¹ MNC) as previouslydescribed (Ratajczak et al, 1995, 1992). CD34þ cells wereenriched from the A¹T¹MNC population by immunoselec-tion with anti-CD34 murine monoclonal antibody (anti-HPC1; Becton Dickinson, Mountainview, Calif.) and mag-netic beads according to the manufacturer’s protocol (Dynal,Oslo, Norway). Briefly, 2 × 107 A¹T¹MNC were incubated in1.5 ml Eppendorf tubes for 1 h in 1 ml of Iscove DMEM(Gibco, U.S.A.) þ 5% Bovine Calf Serum (BCS) (Hyclone,U.S.A.) with 50 ml of anti-CD34 antibodies (anti-HPC1) at48C on a rotating rack. The cells were washed three times inIscove DMEM þ 5% BCS and then resuspended in 1 ml ofIscove DMEM þ 5% BCS. 150 ml of coated immunomagneticbeads (Dynal, Oslo, Norway) were added and the cells wereincubated again for 45 min at 48C on a rotating rack. Cellswere subsequently transferred to 5 ml polystyrene plastictubes and collected for 2 min using a magnetic rack (Dynal,Oslo, Norway). This magnetic purification procedure wasrepeated three times. After the third separation, cells wereresuspended in Iscove DMEM þ 10% BCS. Cell viability was>98%. Final purity was >90%.

In some cases the c-kit receptor bright (Kit-R bright) subsetof CD34þ cells was isolated by fluorescence-activated cellsorting (FACS). Briefly, 2 × 107 human A¹T¹MNC weresuspended in PBS supplemented with 5% bovine calf serum(BCS) and labelled for 30 min at 48C with anti-Kit Rmonoclonal antibodies (SR-1) (1:1000) (generously pro-vided by Dr V. Broudy, University of Washington, Seattle,Wash.). Cells were washed three times in ice-cold PBSsupplemented with 5% BCS and then incubated with Cy5conjugated goat anti-mouse monoclonal antibody (JacksonImmuno Research Labs, West Grove, Pa.) (1:100) for 30 minat 48C. After incubation, cells were washed again three times inice-cold PBS supplemented with 5% BCS and then finallyincubated with anti-CD34 monoclonal antibody directlyconjugated with phycoerythrin (PE anti-HPCA-2, BectonDickinson, San Jose, Calif.; 20 ml/106 cells) for 30 min at 48C.After final incubation, cells were washed three times in ice-coldPBS supplemented with 5% BCS and then subjected to FACSusing a FACS Star Plus II (Becton Dickinson, San Jose, Calif.).

Serum-free cell cultures. 104 CD34þ or 105 A¹T¹ MNCwere cloned in 1 ml of medium containing 0.8% methyl-cellulose (Methocel MC, Fluka, Switzerland) in Iscove DMEMsupplemented with 1% delipidated, deionized, and charcoaltreated BSA (Sigma, St Louis, U.S.A.), 270 mg/ml ironsaturated transferrin (Sigma, St Louis, U.S.A.), insulin (20mg/ml), 5.6 mg/ml cholesterol (Sigma, St Louis, U.S.A.), and2 mmol/l L-glutamine in 3.5 cm plastic petri dishes andincubated (378C, 95% humidity, 5% CO2).

In some experiments, recombinant human (rH) TPO(R & D, Minneapolis, Minn.) was added to cultures on day

0 at concentrations of 10 ng/ml or 50 ng/ml. After 7 d, rHKL (100 ng/ml; R & D, Minneapolis, Minn.) þ rH EPO (5 U/ml, Amgen Inc., Thousand Oaks, Calif.), or rH EPO þ rH IL-3(20 U/ml, Genetics Institute, Cambridge, Mass.) was added tothe cultures by diluting the growth factors in 50 ml of IMEMand then overlaying the methylcellulose with the solution.Effects on BFU-E colony formation were observed over thenext 14 d. Colonies were counted with an inverted micro-scope on day 14.

In other experiments, KL (100 ng/ml)þIL-3 (50 U/ml) þ

IL-11 (50 U/ml) was added to cultures of CD34þ MNC alone,or in combination with TPO (100 ng/ml). Effects on BFU-Ecolony formation were observed over the next 14 d. Colonieswere counted with an inverted microscope on day 14. Aftercounting, the methycellulose was solubilized, the colonieswere disaggregated, and individual cells were analysed forexpression of glycophorin A (GPA), CD33 or plateletglycoprotein IIb/IIIa as described below.

Finally, experiments were carried out in which FACSisolated Kit-Rþ, CD34þ MNC were plated in serum-freemedium in 96-well plates at a concentration of 100 cells/well (0.2 ml). The cells were stimulated with EPO þ IL-3, andEPO þ KL in the presence or absence of TPO (50 ng/ml).Erythroid colonies were scored 14 d later using an invertedmicroscope.

Immunochemical phenotyping of BFU-E colonies. At days 4,6 or 9, cells growing in haemopoietic colonies were isolatedby diluting the methylcellulose with PBS. Liquefied cultureswere washed again with PBS and cells isolated from thecultures were stained with murine monoclonal anti-GPA-Aantibody (1:2000) (kind gift from Dr Steven Spitalnik,University of Pennsylvania, Philadelphia), anti-IIb/IIIa anti-body (1:100) (Immunotech, Marseille, France) or CD33antigen antibody (20 ml/106 cells) (Becton Dickinson,Mountainview, Calif.) for 30 min. After washing, cellsincubated in primary antibody, were then incubated withPE-conjugated sheep anti-mouse Ab (1:100) (Sigma,St Louis, Mo.). Expression of GPA-A, IIb/IIIa and CD33 wasevaluated by FACS.

Isolation and mRNA phenotyping of haemopoietic colony andmarrow cells. Colonies growing under serum-free conditionswere plucked from cultures on days 4–9 using a Pasteurpipette with the aid of an inverted microscope. Approxi-mately 20 colonies from each culture dish were utilized foreach mRNA extraction. Extracted colonies were suspendedin 10 ml of Iscove DMEM and incubated for 1 h at 378C todissolve the methylcellulose. Cells were then washed twice inPBS, and RNA extracted as previously reported (Ratajczak etal, 1995, 1992). Briefly, the cells were lysed in 200 ml ofRNAzol (Tel-Test Inc., Friendswood, Texas) þ22 ml of chloro-form. The aqueous phase was collected and subsequentlymixed with 1 vol of isopropanol. RNA was precipitatedovernight at ¹208C. The RNA pellet was washed in 75%ethanol and resuspended in 3 × autoclaved H2O. mRNA wassimilarly extracted from isolated CD34þA¹T¹ marrow MNC.

RT-PCR was carried out as previously reported (Ratajczaket al, 1995). mRNA (0.5 mg) derived from the various cellularsources was reverse transcribed with 500 U of Moloneymurine leukaemia virus reverse transcriptase (MoMLV-RT)

and 50 pmol of an ODN primer complementary to 30 end ofthe following sequence of TPO receptor (c-mpl) (nts 691–709) (50-CTG CTG TCA GAG CTG AAG C-30). The resultingcDNA fragments were amplified using 5 U of Thermusaquaticus (Taq) polymerase and primers specific for 30 endof TPO receptors (50-ACA TTT GAG GAC CTC ACT TGC-30)(nts 691–709) and of the 50 end of TPO receptors (50-ACATTT GAG GAC CTC ACT TGC-30) (nts 130–151). The Kitprimers employed have been reported (Ratajczak et al, 1992).

Detection of apoptosis. CD34þ, Kit-Rþ MNC (105) wereisolated by FACS and then suspended in Iscove DMEMcontaining the serum-free supplement described above. Thecells were cultured in six-well plastic plates in the presence orabsence of TPO (50 ng/ml) for 72 h (378C, 95% humidity, 5%CO2) and then washed once in PBS and fixed in 4% neutralbuffered formalin. After fixation, the cells were sedimentedby gravity onto slides covered with Cell-Tak tissue adhesive

(Collaborative Biomedical Products, Bedford, Mass.).Apoptosis was detected in these cells using an in situapoptosis detection kit: ApopTag (Oncor, Gaithersburg, Md.)according to the manufacturer’s protocol. In this assay,terminal deoxynucleotide transferase (TdT) was used to labelthe fragmented ends of DNA which had undergone apoptosiswith digoxigenin-dUTP. The digoxigenin label was sub-sequently detected with a fluorescently labelled anti-digoxigenin antibody allowing simple, direct enumerationof apoptotic cells with a fluorescence microscope.

Statistical analysis. Arithmetic means and standard devia-tions were calculated on a MacIntosh computer using Instat1.14 (GraphPad, San Diego, Calif.) software. Culture resultsare reported as mean 6 SD of colonies observed in quad-ruplicate plates. Differences in population means werecompared using the Student t-test for unpaired samples. Astatistically significant difference was defined as P < 0.01.

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Fig 1. Phase photomicrograph of TPO (50 ng/ml) stimulated normal human CD34þ marrowcells. Cells stimulated by TPO were cloned for11 d in serum-free methylcellulose cultures.More than 92% of cells isolated from thesedishes were IIb/IIIaþ by FACS analysis.(A) Human megakaryocytes growing inserum-free medium (magnification × 220).(B) Human megakaryocytes growing inserum-free medium (magnification × 660).

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RESULTS

Effect of TPO on erythroid colony cell growth in serum-freeculturesWhen TPO (10 ng/ml or 50 ng/ml) was added on day 0 toserum-free cultures containing CD34þ MNC, no haemo-globinized cells were visible in the cultures after 14 d. Thosecells which were present formed small colonies of translucentcells, many of which appeared to contain mature mega-karyocytes (Fig 1). That colonies formed under theseconditions were predominantly megakaryocytic in naturewas confirmed by immunochemical phenotyping of cells.FACS analysis of the cellular constituents of these culturesrevealed that >90% were IIb/IIIa positive (data not shown).

The addition of TPO in combination with IL-3 (20 U/ml)alone, KL (100 ng/ml) alone, or IL-3 (20 U/ml) þ KL(100 ng/ml) increased the numbers of colonies present,2–5-fold, but again failed to stimulate the appearance ofhaemoglobinized cells (Table I). The absence of cells bearinghaemoglobin was confirmed by benzidine staining of

Table I. Effect of rh TPO on CD34þ MNC cloning efficacy in serum-free culture.

Growth factor(s) added Colonies* 6SD

None 0 6 0TPO (50 ng/ml) 98 6 34†TPO (50 ng/ml) þ IL-3 (20 U/ml) 324 6 114TPO (50 ng/ml) þ KL (100 ng/ml) 227 6 97TPO (50 ng/ml) þ IL-3 (20 U/ml) þ KL (100 ng/ml) 513 6 159

* Colonies per 2 × 104/CD34þ cells. Note that none werehaemoglobinized and that >90% of cells in dishes were IIb/IIIapositive.

† Data from four independent bone marrow donors, each culturedin quadruplicate.

Fig 2. FACS analysis of CD34þ cells grown in serum-free methylcellulose cultures stimulated with TPOþKLþIL-3. Analysis was carried out ondisaggregated colony cells obtained as detailed in the Methods section after 6 d in culture. (A) Flow cytogram of cells isolated from serum-freecultures. (B) and (C) GPA-A and CD33 expression analysis respectively. No cells express GPA-A over background. (D) Expression of glycoproteinIIb/IIIa.

cytocentrifuge preparations of cells grown in the cultures(data not shown). To further confirm that such culturescontained no maturing erythroid elements, FACS basedimmunophenotypic analysis of cells obtained from day 7serum-free cultures stimulated with TPOþIL-3þKL wascarried out (Fig 2). Although many cells expressed IIb/IIIaand/or CD33, no cells expressed glycophorin A. Therefore

TPO appeared to have little or no effect on erythropoietic cellmaturation as a single cytokine, or in combination with IL-3and KL.

It has recently been suggested that IL-11 may promotefinal erythroid maturation and stimulate haemoglobiniza-tion in EPO receptor (¹/¹) murine fetal liver haemopoieticand embryonic stem (ES) cells (Kieran et al, 1996; Quesniaux

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Fig 3. FACS analysis of CD34þ cells grown in serum-free methylcellulose cultures stimulated with KLþIL-3þIL-11 (A), TPOþKLþIL-3þIL-11(B), or KLþTPO (C). Analysis was carried out on disaggregated colony cells obtained as detailed in the Methods section after 14 d in culture. Flowcytograms of cells isolated from the serum-free cultures are shown in ‘A’ of each experiment (A, B, and C respectively). CD33 and GPA expressionanalysis in relationship to an isotype control antibody are as indicated in each panel.

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et al, 1992). For this reason, we evaluated the ability of TPOto complement erythroid maturation in adult human CD34þ

marrow cells stimulated with KL þ IL-3 þ IL-11 in serum-free cultures. KLþIL-3þIL-11 failed to stimulate erythro-poietic cell development as assessed by formation ofhaemoglobin containing or GPA expressing cells althoughthe expected stimulation of CD33þ myeloid cells wasobserved (Fig 3A). The addition of TPO to the cytokinecombination also failed to promote erythroid cell develop-ment using the same criteria (Fig 3B). To demonstrate that

cells of erythroid phenotype could be detected, Fig 3C showsthat GPA was highly expressed on cells subjected to similaranalysis after culture in EPO and KL.

We then asked whether TPO could augment the effect ofEPO on erythropoiesis in a serum-free culture system (TableII). To address this question, CD34þ cells were cultured inthe presence of EPO alone, EPO þ TPO, EPO þ IL-3 þ TPO,EPO þ KL, and EPO þ KL þTPO. As expected, EPO alone hadno stimulatory effect on BFU-E colony formation (Ratajczaket al, 1992; Wu et al, 1995). The addition of TPO to EPOresulted in the appearance of a small number of small

Table II. Influence of TPO on erythroid colonydevelopment in serum-free culture.

Growth factor(s) added BFU-E* 6SD

Epo† No growthEpo þ TPO‡ 12 6 6Epo þ IL-3§ 119 6 53Epo þ IL-3 þ TPO 163 6 48Epo þ KL¶ 434 6 147Epo þ KL þTPO 419 6 173

* Colonies per 2 × 104/CD34þ cells. Datafrom four independent bone marrow donors,each cultured in quadruplicate.

† EPO concentration ¼ 5 U/ml.‡ TPO concentration ¼ 50 ng/ml.§ IL-3 concentration ¼ 20 U/ml.¶ KL concentration ¼ 100 ng/ml.

Table III. Erythroid colony development byTPO stimulated CD34þ, Kit-Rþ MNC inserum-free culture.

Growth factor(s) added BFU-E* 6SD

Epo† þ IL-3§ 1.2 6 0.7Epo þ IL-3 þ TPO‡ 1.4 6 0.8Epo þ KL¶ 6.8 6 3.4Epo þ KL þTPO 7.3 6 3.6

* Colonies per 1 × 102 CD34þ c-kit-Rþ cells.Data from three independent bone marrowdonors, each cultured in two 96-well plates.

† EPO concentration ¼ 5 U/ml.‡ TPO concentration ¼ 50 ng/ml.§ IL-3 concentration ¼ 20 U/ml.¶ KL concentration ¼ 100 ng/ml.

Fig 4. RT-PCR phenotyping of mRNA for TPO(c-mpl) R (A) and Kit-R (B). RNA was isolatedfrom HEL cells (lane 1), normal human CD34þ

cells (lane 2), day-4 BFU-E colonies (lanes 3and 4), day-6 BFU-E colonies (lanes 5 and 6),and day-9 BFU-E colonies (lanes 7 and 8).Lane 9: negative RT-PCR–H2O reaction.

erythroid bursts (12 6 6 BFU-E) when compared to burstcolony formation in the presence of EPO alone (0 6 0 BFU-E). EPO/TPO synergy with respect to BFU-E colony formationwas again noted when TPO was added to an EPOþIL-3cocktail (163 6 48 BFU-E with TPO versus 119 6 53 BFU-Ewithout TPO; P ¼ 0.02). However, when cultures were moreoptimally stimulated, as with the cytokine combination ofEPOþ KL the burst colony enhancing properties of TPO wereno longer seen (4346147 BFU-E with TPO versus419 6 173 BFU-E without TPO). These results suggest thatTPO has only minor burst-promoting activity.

Finally, to help exclude potential contributions of ancillarycells present in the cultures, the erythroid colony stimulatingeffects of TPO was examined in a more highly purifiedpopulation of CD34þ/Kit Rþ MNC which were plated at low

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14 Mariusz Z. Ratajczak et alTable IV. Influence of TPO on the survival of human BFU-E in serum-free cultures. CD34þ marrow cells (2 × 104) were cultured in serum-free methylcellulose in the absence (control) or presence of TPO (10or 50 ng/ml). After 7 d EPO (5 U/ml) þ KL (100 ng/ml) was added tothe cultures. BFU-E colonies were scored 14 d later.

TPO TPOControl (10 ng/ml) (50 ng/ml)

BFU-E 72 6 29* 416 6 72† 452 6 49†

* Mean 6 SD of four separate studies, each performed in quad-ruplicate.

† P<0.0001 in comparison to control.

Fig 5. Effect of TPO on apoptotic fate of CD34þ, Kit-Rþ MNC held in serum-free suspension cultures for 72 h. Fragmented DNA, as occurs in cellsundergoing apoptosis, is detected by the presence of green nuclear fluorescence (ApopTag, Oncor, Gaithersburg, Md.). (A) and (B) Correspondingphase and fluorescent photomicrographs of cells cultured with TPO (50 ng/ml). Nuclei were counterstained with propidium iodide. No apoptoticcells are detected in this sample using Apoptag. (C) and (D) Corresponding phase and fluorescent photomicrographs of cells cultured without TPO(50 ng/ml). Note the presence of apoptotic cells which stain positive for green fluorescence.

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density (1 × 102/0.2 ml) in microtitre wells (Table III). Underthese conditions, TPO did not appear to enhance BFU-Ecolony formation over that observed with EPOþIL-3 orEPOþKL.

Assay of BFU-E for Mpl expressionTPO receptor (c-MPL) is weakly expressed by ,2% of CD34þ

cells (Debili et al, 1995). Expression is increased as the cellsprogress along the megakaryocyte maturation pathway andpersists on platelets. If TPO were to exert a direct effect onerythroid cells it is reasonable to assume that such cellswould also express Mpl-R. Since immunochemical methodsmight fail to detect very weakly expressed Mpl protein, weemployed RT-PCR to detect its mRNA. This method is verysensitive so that a negative result would argue strongly thatthe receptor was not expressed by cells of the erythroidlineage. We therefore employed RT-PCR to examine Mplreceptor expression on colony cells developing in serum-freemedium containing cytokines designed to favour growth oferythroid elements. When such colonies were examined atdays 4, 6 and 9 for Mpl receptor expression, none wasdemonstrable by RT-PCR. As positive controls for the RT-PCRreactions, we extracted, and then reverse transcribed, mRNAfrom HEL cells and from normal human CD34þ cells. ThecDNA was amplified with Mpl R primers that were predictedto amplify a 580 nt long fragment. As shown in Fig 4, anappropriately sized fragment was amplified from HEL andCD34þ cells but not from cells present in day 4, 6 or 9 BFU-Ecolonies. The integrity of the isolated mRNA was furtherevaluated by amplification of a cDNA encoding the Kit R. Asis also shown in Fig 4, Kit mRNA could be amplified in all thecells studied. These data strongly suggest that, in contrast tocells of the megakaryocyte lineage, Mpl R is not expressed ondeveloping erythroid cells.

TPO inhibits progenitor cell death in serum-free culturesAs TPO appeared to have little colony stimulating orerythroid cell maturation promoting properties in serum-free cultures, we hypothesized that the reported erythro-trophic effects of TPO might result from expansion of anerythroid progenitor less mature than BFU-E. Alternatively,TPO might inhibit erythroid progenitor cells from undergoingapoptosis. Such inhibition would effectively ‘expand’ the

population and allow the production of more erythrocytesat the end of the developmental cycle. To test this hypo-thesis, CD34þ cells were initially cultured serum-free for1 week in the absence of all growth factors, or in thepresence of TPO alone. EPOþKL was then added to thecultures by layering the cytokines over the cultures whichwere then observed for an additional 14 d (Table IV). In theabsence of TPO, 72 6 29 BFU-E colonies were observed inthe dishes at the end of the experiment. In marked contrast,when cells were exposed to TPO for 7 d prior to the additionof EPOþKL, between 416 6 72 and 452 6 49 BFU-Ecolonies grew in the cultures. It is important to note thatprior to addition of EPOþKL on day 7, TPO’s proliferativeeffects on cells in the cultures were minimal, and would notexplain the 6-fold increase in erythroid colonies noted in theTPO-treated cells. Accordingly, these results strongly suggestthat TPO prevented erythroid progenitor cells from dying inculture.

To directly determine if TPO effected the viability ofcultured progenitor cells, CD34þ, Kit-Rþ marrow cells wereisolated by FACS (Fig 5). The FACS sorted cells were culturedfor 7 d in serum-free medium in the presence or absence ofTPO. As shown in Table V, the number of apoptotic cells wasconsiderably greater in the cells cultured in the absence ofTPO. Accordingly, the addition of TPO to CD34þ, Kit-Rþ cellsappeared to augment the viability of cells in this progenitorenriched population by preventing cells from undergoingapoptosis.

DISCUSSION

Post-chemotherapy and post-transplant thrombocytopeniaremain important clinical problems which, in addition totheir associated morbidity, result in large expenditures forplatelets each year (American Society of AnesthesiologistsTask Force on Blood Component Therapy, 1996; Hassan &Zander, 1996). Although murine (Fibbe et al, 1995;Kobayashi et al, 1995), and primate (Akahori et al, 1996;Farese et al, 1995) studies suggest that TPO may amelioratethrombocytopenia in these settings, TPO’s clinical use wouldbe considerably enhanced if suggestions that it might alsostimulate myeloid and erythroid recovery were substantiated(Sitnicka et al, 1996).

Table V. Effect of TPO (50 ng/ml) on development of apoptotic changes by CD34þ, KIT-Rþ MNC cultured serum free for up to 7 d in suspensionculture. Effects on cell proliferation, viability and apoptosis were determined as noted in the Methods section. Percentages are mean 6 SD oftriplicate cultures.

Day 3 Day 5 Day 7

Day 0 (¹) TPO (þ) TPO (¹) TPO (þ) TPO (¹) TPO (þ) TPO

Cell number 100 000 – – – – 87 000 173 000Viable cells (%) 98 87 6 4 96 6 0.6* 50 6 8 77 6 4† 21 6 2 57 6 9†Apoptotic cells (%) 0 22 6 4 6 6 2† 54 6 5 34 6 3† 91 6 5 55 6 10†

* P < 0.02 compared to cells maintained without TPO.† P < 0.002 compared to cells maintained without TPO.

The present literature regarding TPO is somewhat difficultto evaluate with regard to its effects on non-megakaryocyteprogenitor cells. Mpl ‘knock-out’ mice (Alexander et al,1996; Gurney et al, 1994) appear to have no defects inerythroid or myeloid cell development, although thenumbers of multipotent progenitors present in the knockoutanimals may be reduced (Alexander et al, 1996). Chronicover-expression of retrovirally expressed TPO led to marrowfibrosis with megakaryocyte hyperplasia and thrombocyto-sis, but erythroid hyperplasia and polycythaemia was notobserved (Yan et al, 1995). TPO infusion studies in murineand primate model systems also yielded conflicting results onthe ability of TPO to stimulate non-megakaryocyte progeni-tor cell growth. Infusion of pegylated-TPO into normalprimates was not reported to cause effects on red cell numbersor haemoglobin concentration (Akahori et al, 1996; Harker etal, 1996). In contrast, infusions into normal mice paradoxi-cally resulted in erythroid and lymphoid hypoplasia. In murinetransplant models Molineaux et al (1996) reported no effect onred or white blood cell recovery, and Fibbe et al (1995) reportedenhanced recovery of both lineages when donor cells were pre-treated with TPO. From these reports, one could at leastconclude that TPO is certainly not required for normalerythropoiesis, that supra-physiological levels of TPO do notnecessarily drive erythropoiesis in vivo, and that the effects ofTPO on recovery of non-megakaryocytic lineages after majordamage is uncertain.

Results obtained with in vitro assays, a seemingly less-complicated system, are also ambiguous. Antisenseoligonucleotides targeted to the Mpl receptor perturbmegakaryocyte development but leave erythroid and myeloidcell development unaffected (Methia et al, 1993). Usingtypical serum or plasma containing culture assays, anumber of groups have reported that TPO has little erythroidor myeloid colony stimulating activity on its own, but whenused in conjunction with other cytokines significant colonystimulating synergy results (Kaushansky et al, 1996;Kobayashi et al, 1995; Papayannopoulou et al, 1996).Nevertheless, because these assays systems employed serum,the effects of TPO alone versus a cooperating effect withserum factor(s) are difficult to separate.

To overcome some of the problems alluded to above, weexamined the effect of recombinant human TPO on normalhuman erythroid progenitor cell growth in a serum-freeculture system. Enriched progenitor cells were cloned atusual concentrations (2 × 104/ml) and at more limitingdilutions (1 × 102 cells/0.2ml) in order to detect the effects ofancillary cells which might be present in the cultures. WhenCD34þ cells were exposed to TPO under serum-freeconditions the formation of small cluster-like coloniespositive for platelet glycoproteins IIb/IIIa and CD33 wasstimulated but no cells expressing glycophorin-A or haemo-globin were detected. In the presence of EPO, a few BFU-Ecolonies did appear in culture and in the presence of IL-3 þ EPO a small, but statistically insignificant, increase inBFU-E of >50 cells was also stimulated in comparison to IL-3 þ EPO alone. However, in the presence of more optimalBFU-E stimulation, such as with KLþ EPO, the trophic effectsof TPO on in vitro erythropoiesis were not observed. These

results may be explained at least partially by our inability todetect Mpl receptor expression in colonies developing underconditions designed to favour the growth of erythroidelements. Early in culture, erythroid bursts are undergoingintense proliferative activity, and it is difficult to imagine thatTPO could have significant impact on this process if itsreceptor is not expressed on the putative target cells.

Of course, it is possible that Mpl R is expressed on erythroidcells at levels below the detection limit of the PCR techniquewe employed. We consider this unlikely, in view of the factthat CD34þ cells are known to express Mpl R very weakly(Methia et al, 1993) and we could detect the receptor’spresence on these cells. One might also argue that all thecolonies we plucked for analysis were non-erythroid, but thisis unlikely since EPOþKL in serum-free cultures, at least inour hands, does not stimulate the formation of non-erythroid colonies. Based on these results, we believe thatTPO has little intrinsic burst-promoting activity, and onlyweak colony co-stimulating activity. Importantly however,we do find that TPO inhibits cells in a progenitor enrichedpopulation from undergoing apoptosis and hypothesizethat this may be TPOs major biological effect on non-megakaryocytic cells. TPO has recently been reported toinhibit apoptosis in a human leukaemia cell line (Ritchie et al,1996). Inhibition of cell death might lead to an effectiveexpansion of the progenitor cell compartment as reflected bya greater number of cells being present to respond to theusual stimulatory molecules.

Recently, Kieran et al (1996) reported that TPO can rescueerythroid colony development by fetal liver cells derived fromhomozygous EPO receptor knockout mice. It is worth noting,however, that the fraction of rescued cells was quite small,and that rescue required the presence of either KL, or IL-3 þ IL-11. This effect was not observed on normal adulthuman haemopoietic progenitor cells, suggesting thatcomplementation, or adaptation to an EPO deprivationstate, may be required for such an effect to be observed. Itwas also recently reported that TPO expands primitivehaemopoietic cells in serum-replete conditions (Sitnicka et al,1996) and LTC-IC in a serum-free assay (Petzer et al, 1996).Of interest, it was noted that the progeny of such LTC-IC giverise to predominantly erythroid forming colonies (Petzer et al,1996). This study is intriguing but not inconsistent withour hypothesis that TPO’s primary effect on erythroidprogenitor cells is to inhibit them from undergoing celldeath. Whether this will prove to be of clinical significanceis unclear, but will be answered by clinical trials which arenow in progress.

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