Apoptosis in refractory anaemia with ringed sideroblasts is initiated at the stem cell level and associated with increased activation of caspases


Eva Hellström Lindberg MD, PhD, Department of Medicine, Division of Haematology, Karolinska Institutet at Huddinge University Hospital, 141 86 Huddinge, Sweden. E-mail: Eva.Hellstrom-Lindberg@medhs.ki.se


Treatment with granulocyte colony-stimulating factor plus erythropoietin may improve haemoglobin levels in patients with ringsideroblastic anaemia (RARS) and reduce bone marrow apoptosis. We studied bone marrow from 10 RARS patients, two of whom were also investigated after successful treatment. Mononuclear, erythroid and CD34+ cells were analysed with regard to proliferation, apoptosis, clonogenic capacity and oncoprotein expression, in the presence or absence of Fas-agonist, Fas-blocking antibody 2 and caspase-3 inhibitor. During culture, RARS bone marrow cells showed higher spontaneous apoptosis (P < 0·05) and caspase activity (P < 0·05)) than bone marrow cells from healthy donors. Eight out of nine patients had reduced growth of erythroid colony-forming units (CFU-E) (< 10% of control) and granulocyte–macrophage CFU (CFU-GM) (< 50% of control) from CD34+ cells. Fas ligation increased apoptosis and decreased colony growth equally in RARS and controls, but caused significantly more caspase activation in RARS (P < 0·01). Fas-blocking antibody showed no significant inhibitory effect on spontaneous apoptosis or ineffective haematopoiesis, as measured using phosphatidylserine exposure, the terminal deoxynucleotide transferase-mediated dUTP-biotin nick-end labelling technique, caspase activity or clonogenic growth. Caspase inhibition reduced apoptosis, increased proliferation and enhanced erythroid colony growth from CD34+ cells in RARS, but showed no effect on normal cells. CFU-E improved > 1000% after successful treatment. Thus, erythroid apoptosis in RARS is initiated at the CD34+ level and growth factor treatment may improve stem cell function. Enhanced caspase activation at the stem cell level, albeit not mediated through endogenous activation of the Fas receptor, contributes to the erythroid apoptosis in RARS.

The myelodysplastic syndromes (MDS) are characterized by cytopenia, ineffective haematopoiesis and a risk of progression to acute myeloid leukaemia (AML) (Greenberg, 1994). Refractory anaemia with ringed sideroblasts (RARS) is a subtype of MDS in which excess iron accumulates in the mitochondria of the erythroid precursors. The mitochondria localize in a ring around the nucleus, thus giving the erythroid progenitor the classic ringsideroblastic appearance (Björkman, 1956; Gattermann, 1999). The main clinical problems for this subgroup of MDS are severe anaemia, chronic transfusion need and the subsequent risk of iron overload (Bennett et al, 1982; Greenberg et al, 1997).

Recent studies have shown an increased incidence of apoptotic precursors in myelodysplastic bone marrow samples. However, the exact association between bone marrow cell apoptosis, ineffective haematopoiesis and peripheral blood cytopenia, or the cause of the increased apoptosis, has not been clarified (Clark & Lampert, 1990; Raza et al, 1995; Hellström-Lindberg et al, 1997). Several hypotheses have been suggested based on abnormal in vitro findings in myelodysplastic cells. Upregulated surface expression of the Fas receptor (CD95/APO-1) on myelodysplastic bone marrow cells, including CD34+ cells, has been shown, as well as an increased expression of the Fas ligand (Bouscary et al, 1997; Gersuk et al, 1998; Kitagawa et al, 1998; Lepelley et al, 1998; Stahnke et al, 1998). Fas stimulation may induce functional apoptosis in selected patients (Bouscary et al, 1997; Lepelley et al, 1998) and there are also studies showing increased growth of granulocyte–macrophage colony-forming units (CFU-GM) by adding antagonistic Fas and tumour necrosis factor-α (TNF-α) antibodies to long-term bone marrow cultures (Gersuk et al, 1998; Gupta et al, 1999). Next, it has been proposed that inflammatory cytokines, such as TNF-α and transforming growth factor-β (TGF-β), play a key role in the apoptotic process in MDS (Shetty et al, 1996; Mundle et al, 1999a), while other studies have described increased levels of intracellular proteases (caspases) in myelodysplastic bone marrow (Mundle et al, 1996, 1999b). Finally, the expression of oncoproteins involved in cell death or survival is altered in MDS. Two separate studies have reported that patients with ‘low-risk’ MDS show a higher relative expression of pro-apoptotic vs. anti-apoptotic oncoproteins (Rajapaksa et al, 1996; Parker et al, 1998).

Caspases are cysteine proteases that cleave preferentially after Asp residues (Nicholson & Thornberry, 1997). Caspases are present as proforms in the cell and can be activated by proteolytic cleavage, which is thought to initiate a cascade of proteolytic events in which activated caspases activate other downstream caspases. The caspases can be divided in different groups based on their principal enzymatic function. Initiator caspases, such as caspase-8 and caspase-9, induce activation of other caspases and, therefore, play a role in upstream events of the enzymatic cascade. Others, such as caspase-3, caspase-6 and caspase-7, have effector functions and play a role in downstream events by cleaving so-called ‘death substrates’ in the cell.

Cytokines may be administered to patients with MDS in order to improve granulocyte counts and haemoglobin levels (Cazzola et al, 1998). We showed that the combination of granulocyte colony-stimulating factor (G-CSF) and erythropoietin (EPO) improved the anaemia in 38% of a cohort of 80 low-risk patients with MDS (Hellström-Lindberg et al, 1998). The best and most durable responses were observed in patients with RARS and a pronounced in vivo synergy between G-CSF and EPO was also observed in this group. Erythroid improvement was associated with a decrease in the number of apoptotic bone marrow precursors (Hellström-Lindberg et al, 1997). In contrast, other clinical studies have suggested a role for T-cell suppression in MDS, but the favourable results and the in vitro finding of activated T cells have almost exclusively been observed in refractory anaemias (RA) (Molldrem et al, 1997, 1998; Jonasova et al, 1998).

Thus, there is accumulating evidence that the presence of a significant number of ringsideroblasts implies a specific pathogenesis in MDS, and that RA and RARS should be considered as different diseases. Most in vitro investigations comprise a sample of various MDS subcategories and the number of patients in each subgroup is often too small to allow specific conclusions.

The present study investigated the effects of inhibition and stimulation of the Fas system, as well as of caspase inhibition on bone marrow cells from patients with RARS. The aim of the study was to define the starting point for the erythroid apoptosis, describe the main pathways for the increased apoptosis and investigate bone marrow function after successful clinical treatment with G-CSF + EPO. A homogeneous and well-characterized patient group and the use of several parallel methods for apoptosis detection, in addition to functional assays performed on different subsets of bone marrow cells, have enabled us to draw new and valuable conclusions concerning apoptotic mechanisms in RARS.


Patients and controls Ten patients with RARS with a mean age of 74 ± 13 years were included in the study (Table I). Bone marrow from two patients was examined twice: before and after 20 weeks from the start of treatment with G-CSF + EPO. Normal bone marrow samples were obtained from 10 patients undergoing thorax surgery and from three healthy volunteers. Six of these were used only for the analysis of spontaneous apoptosis in suspension cultures, while seven subjects also served as controls for the specific induction experiments described below. Relatively age-matched controls (mean age 58 ± 20 years) were used, as an age-related decrease in clonogenic capacity has been described (Marley et al, 1999). Informed consent was obtained from both patients and controls, and the study followed the guidelines of the local ethical committee of Karolinska Institutet.

Table I.  Baseline data of 11 patients with RARS.
  Duration ofBlood values Ongoing
PatientAge/Sexdisease (months)Hb value (g/dl)WBC (gran.) (× 109/l)Platelet count (× 109/l)Karotypetreatment
  1. *Regular transfusions of packed red blood cells.
    †G-CSF was according to the protocol dose to obtain a granulocyte count of 6–10 × 109/l.

  2. Two patients (2 and 3) were included in the ongoing Scandinavian trial on G-CSF + EPO and analysed twice, before and after 20 weeks of successful treatment. WBC, white blood cells (granulocytes).

RARS 185/Female 12 9·25·7 (4·2)25246 XXTransfusions*
RARS 2a81/Female  211·25·5 (3·6)26846 XXTransfusions
RARS 2b  13·014·3 (12·0)363 G-CSF + EPO
RARS 3a82/Female  6 9·56·2 (4·1)33646 XXTransfusions
RARS 3b  12·524·6 (20·1)356 G-CSF + EPO
RARS 462/Female 1810·06·9 (4·6)29846 XXNo treatment
RARS 585/Female 54 9·25·5 (3·5)42146 XX, 20q-Transfusions
RARS 659/Male 88 8·31·6 (1·2)10346 XYTransfusions
RARS 774/Male154 9·34·7 (1·8)30746 XYNo treatment
RARS 881/Male 6411·04·4 (2·5)21846 XYNo treatment
RARS 947/Female  4 8·55·2 (2·8)26346 XXTransfusions
RARS 1075/Female  4 7·67·2 (3·2)16646 XXTransfusions

Serological and other reagents Inhibition of the Fas receptor was achieved using Fas receptor-blocking antibodies [f(ab)′2] that were generously supplied by Professor Peter Krammer (Dhein et al, 1995). F(ab)′2 were used at a concentration of 1 μg/ml and the biological effects were confirmed in the Jurkat T-cell line. Agonistic stimulation of the Fas receptor was achieved using the antibody CH-11 (Medical Biological Laboratories) at a concentration of 1 μg/ml. Caspase-3-like activity was blocked by the inhibitory peptide DEVD-fmk (Enzyme Systems Products) at a concentration of 10 μmol/l [final dimethyl sulphoxide (DMSO) concentration not exceeding 2‰]. Antibodies against caspase-3 (generous gift from Dr Donald Nicholson), c-myc (Oncogen Research Products, Cambridge, MA, USA), Bcl-2 (Dako, Glostrup, Denmark) and caspase-8 (generous gift from Professor P. Krammer) were used at the concentrations 1:5000, 1:100, 1:100 and 1:20 respectively.

Bone marrow samples and suspension cultures Bone marrow aspirate (5–10 ml) was obtained from the posterior crista iliaca (patients and healthy volonteers) or sternum (thorax patients) and subjected to Lymphoprep (NYCOMED) density gradient centrifugation to isolate mononuclear cells (MNCs). After washing twice in phosphate-buffered saline (PBS), the cells were resuspended in Roswell Park Memorial Institute (RPMI)-1640 medium supplemented with 10% fetal calf serum (FCS; Gibco BRL). Cells were then seeded at a concentration of 5 × 105 cells/ml in tissue culture flasks and incubated at 37°C and 0·5% CO2. MNCs were cultured in the presence of FCS only (control), antagonistic Fas antibody (f(ab)′2), agonistic Fas antibody (clone CH-11) or caspase-3 inhibitor (DEVD-fmk). Cell cultures were checked for viability (trypan blue exclusion), cell number, [3H]-thymidine incorporation and apoptosis on d 0, 1, 2, 3 and 4.

CD34 cell separation and colony assay When MNC numbers were high enough to allow a CD34 separation (eight out of nine patients and both patients analysed after treatment), 63–300 × 106 cells were allocated for this purpose (17–70 × 106 cells in the control samples). The cells were separated for CD34 positivity using the Mini Macs system (Miltenyi Biotec). Aliquots of CD34+ cells were incubated overnight according to positions 1–4. The following day, 10 000 CD34+ cells/ml were sown in triplicate from each of positions 1–4 on MethoCult 4434 medium (StemCell Technologies), containing fetal bovine serum, rh stem cell factor (SCF), rh granulocyte–macrophage CSF (GM-CSF), bovine serum albumin (BSA), methylcellulose in Iscove's modified Dulbecco's medium (IMDM), 2-mercaptoethanol, rh erythropoietin and l-glutamine, and Petri dishes 1008 (35 × 10 mm), and cultured for 14 d. Erythroid colonies (CFU-E), granulocytic colonies (CFU-G) and monocytoid colonies (CFU-M) colonies were counted and a mean value was calculated for each position. In the final report, the sum of CFU-G + CFU-M is presented as CFU-GM.

[3H]-thymidine incorporation assay Aliquots containing 1 × 105 cells in 200 μl of medium were sown in triplicates from each of positions 1–4 on a 96-well test plates (Nunc), with 37 kBq (20 μl of 1·85 MBq/ml) [3H]-thymidine at 37°C. Labelling was initiated at 0, 24, 48 and 72 h after the start of cell culture and [3H]-labelled DNA was measured harvesting the aliquots after 24 h labelling with a Combi Cell Harvester (Skatron, Norway), absorbing the cells on pieces of filter that were put into a scintillation fluid (2 ml of OptiScint ‘HiSafe’, Wallac Scintillation Products). The radioactivity was measured with a liquid scintillation counter (Wallac 1409).

TUNEL staining The commercial in situ apoptosis detection kit (ApopTag, Oncor) was used for assessment of terminal deoxynucleotidyl transferase (TdT)-mediated dUTP-biotin nick end labelling (TUNEL) positivity. Cell suspensions were fixed in 4% neutral buffered formalin for 10 min. Cytospins were made with 100 000 cells per slide. The cytospins were digested in Equilibration Buffer (for 1 min at room temperature). Dioxigenin-dUTP was catalytically added to DNA using TdT enzyme (incubation, 1 h at 37°C) and visualization of the reaction was carried out by incubation with anti-dioxigenin antibody conjugated to fluoroscein isothiocyanate (FITC) (30 min at room temperature). Counterstaining was performed using propidium iodide (0·6 μg/ml DABCO anti-fade). The percentage of cells with FITC-positive nuclei (TUNEL-positive cells) was based on differential counts of 200 nucleated cells.

Flow cytometry Flow cytometric analysis of fresh and cultured mononuclear bone marrow cells in suspension was performed on a FACScan cytometer, using the CellQuest software (Becton-Dickinson, San Jose, CA, USA). Flow cytometer performance was checked by calibration using the FACSComp program and Calibrate beads (Becton-Dickinson). Appropriate dilutions of the following fluorochrome-conjugated (FITC, F; Phycoerythrin, PE) antibodies were used in a standard, direct immunofluorescent staining protocol: CD45-F/F/14-PE (Dako A/S, Glostrup, Denmark), Glycophorin A-PE (Dako), CD95-PE (Dako), CD34-PE (B-D) and isotype controls (IgG1-F/IgG2a-PE; Dako). Exposure of phosphatidylserine on the cell surface was detected by Annexin V-FITC (APOPTEST, NeXins Research BV, Hoeven, The Netherlands), according to the manufacturer's instructions. Mononuclear cells were gated on forward-vs.-side-scatter dot plots. Leucocytes were gated on CD45-vs.-side-scatter gates. Erythoid cells were gated on glycophorin A (GPA)-vs.-side-scatter. Late apoptotic/necrotic cells were defined as cells that were permeable to propidium iodide and early apoptotic cells as Annexin V-positive, propidium iodide-negative cells.

Western blot analysis After incubation for 0 h, 18 h, 42 h and 66 h, 0·5 × 106– 1 × 106 cells were collected from each of the four different treated MNC fractions. The cells were washed twice in PBS and frozen as a cell pellet at −80°C. Cells from two patients and two donors were resuspended in 80 μl of proteinase inhibitor cocktail (Complete Mini, Roche Diagnostics Scandinavia AB, Bromma, Sweden, 1 tablet in 10 ml PBS) before freezing. If the inhibitor cocktail was not added before freezing, the cells were resuspended with 80 μl of this inhibitor immediately after they were taken out of the freezer to prevent enzymatic activity during the thawing. All cells were thawed on ice. 8 μl of 5x Laemmli's loading buffer was added and the samples were boiled for 6 min. These samples were resolved on 15% SDS polyacrylamide gels, as described earlier (Fadeel et al, 1999a).

Caspase in vitro assay The measurement of IETD-AMC and DEVD-AMC cleavage was performed in a fluorometric assay, modified from Nicholson et al (1995), as estimations of caspase-3-like activity. Briefly, 1 × 106 cells from each of the four different fractions were taken, washed twice in PBS and frozen as a dry cell pellet at −80°C before and after 4 h of incubation. For the assay, the cells were kept on ice and resuspended in 50 μl of PBS containing calpain inhibitor I (1:1000, Roche Diagnostics Scandinavia AB, Bromma, Sweden). The appropriate peptide substrate, DEVD-AMC (Peptide Institute, Osaka, Japan), was combined in a standard reaction buffer containing 100 mmol/l HEPES, 10% sucrose, 5 mmol/l dithiothreitol (DTT), 10−4% octylphenoxy polyethoxy ethanol (NP-40) and 0·1% 3-[(3-cholamidopropyl) dimethylammonio]-1-propane sulphonate (CHAPS), pH 7·25, and added to the cell lysates on a microtitre plate. Cleavage of the fluorogenic peptide substrate was monitored by AMC liberation in a Fluoroscan plate reader (Labsystems, Stockholm, Sweden) using 355 nm excitation and 460 nm emission wavelengths. Fluorescence units were converted to pmol of AMC using a standard curve generated with free AMC. Data from duplicate samples were then analysed by linear regression.

Statistical analysis Results are given as mean values ± SD and also as median values + range. For comparison of related and unrelated samples, paired and unpaired t-tests were used respectively. Simple linear regression was used to analyse correlation between variables.


Fas expression on myelodysplastic and normal erythroid cells

Cell yield enabled a flowcytometric analysis of Fas expression on erythroid (GPA+) MNCs in five patients and four controls (Table II). The mean expression of CD95 on normal GPA+ cells was 12·8 ± 8% with no clear increase during culture. RARS patients showed a heterogeneous pattern. The three transfusion-dependent patients showed higher Fas positivity (42–69%), while the two with stable haemoglobin levels showed lower expression. There was no increase of Fas expression in the transfused patients, probably because the Fas-positive erythroid cells were deleted by apoptosis. In contrast, Fas positivity increased over time in the patients with stable haemoglobin, indicating that these erythroid cells remained in culture. Analysis of the f(ab)′2-treated cultures verified the binding of the fragments to the receptor. The binding of the CD95-PE antibody was 10% of that in the FCS cultures on d 1 and 21–57% on d 3. The agonistic CH-11 also reduced Fas expression, but not to the same extent (Table II). Caspase inhibition showed no major impact on Fas expression in any of the cultures.

Table II.  Fas (CD 95) expression on myelodysplastic and normal erythroid cells.
  Day 1Day 2/3
PatientDay 0FCSf(ab)′2CH-11DEVDFCSf(ab)′2CH-11DEVD
  • *

    Few cells.

  •   Flow cytometric data on MNCs from five RARS and four control subjects. The percentage of Fas (CD95)-positive erythroid (glycophorin A+) cells is shown, with the percentage of erythroid cells of total MNCs within brackets.

RARS 2a34% (29)35%3·6%11·4%34·5%14·9% 8·6%NDND
RARS 3b17·3% (52)29·4%1·4%24·9% 4·1%*47·6%15·7%27·8%52%
RARS 647% (29) 9%1·4% 2·7%12%18·4% 4·7% 6%14·7%
RARS 7 7·1% (69) 3·9%0·4% 1·7% 5·9%38·8% 8·2%14·6%14·9%
Controls12·8% 7·4%0·73% 1·2% 7·4%15·9% 8% 8%20·6%
Mean ± SD±7·9% (22)±3·6±0·25±0·42±4·9±5·2±3·0±1·6±5·7

Caspase inhibition increases proliferation in RARS cultures

Viability in the MNC suspension cultures was > 90% in all analysed samples during the first 48 h and > 70% during the last 48 h. [3H]-TdR was measured during all 24-h periods during the cultures, but results are presented for period 2 (24–48 h, Fig 1) and period 5 (96–120 h). Spontaneous proliferation in the FCS cultures did not differ between patients and controls during any of the four measured periods (P > 0·50). Fas-induced changes of proliferation showed a heterogeneous pattern. Fas ligation induced an increase in two of the five controls during periods 2 (200–220%) and 5 (260–450%), while the remaining three controls showed minor decreases. Two patients with relatively mild disease showed a Fas-induced increase of [3H]-TdR (Fig 1), while proliferation in response to CH-11 was unchanged or decreased in the remaining patients. Interestingly, both patients analysed after successful treatment had switched from a Fas-induced decrease to an increase in proliferation. The presence of Fas-blocking fragments did not significantly affect [3H]-TdR in either RARS or controls. In the presence of the caspase inhibitor, DEVD-fmk, there was an increase in mean [3H]-TdR in RARS during period 1 (mean increase 128 ± 41%, P < 0·05), while no significant change was observed in the controls.

Figure 1.

Proliferation in mononuclear cells from patients and controls: effects of Fas antibodies and caspase inhibition. Proliferation as measured by [3H]-TdR analysis (cpm/min) in mononuclear bone marrow cells from patients with RARS (A) and healthy controls (B). The figure shows the effects of incubation with FCS (control culture), f(ab)′2 (Fas-blocking antibody), CH-11 (Fas-triggering antibody) and DEVD-fmk (caspase-3 inhibitor). The analysis covers the period 24–48 h after start of incubation. One patient (patient 1), who also had severe rheumatoid arthritis and had just recovered from pneumonia, showed a clear stimulation of proliferation after Fas inhibition. She spontaneously improved and stabilized her haemoglobin level around 10·0 g/dl 4 weeks after the sampling, which implies that her period of severe anaemia was also influenced by these other conditions.

Caspase, but not Fas, inhibition reduces apoptosis in RARS cultures

TUNEL results are presented in Figs 2 and 3. Mean TUNEL positivity on d 0 was somewhat higher in RARS than in normal cells (3·6 ± 4·0% vs. 1·2 ± 0·8%, P = 0·08). The RARS MNCs underwent significantly more spontaneous apoptosis in suspension cultures than normal MNCs (Fig 2). Day 4 TUNEL positivity was 12·9 ± 6·4% in RARS vs. 6·3 ± 3·5% in the controls, P = 0·01. Fas ligation induced apoptosis in all cultures (controls 203 ± 99%, RARS 223 ± 78%), with no difference between the groups. Presence of Fas-blocking fragments, f(ab)′2, showed no effect on apoptosis in either RARS or controls (106 ± 27% vs. 138 ± 37%, NS) on either d 4 or on any other day (data not shown). Caspase inhibition resulted in different patterns in controls and RARS. While the control cultures showed an increase in the number of apoptotic cells (167 ± 115%), RARS cultures showed a decrease (79 ± 36%, P < 0·05).

Figure 2.

Spontaneous apoptosis in mononuclear cells from patients and controls. Apoptosis as measured by TUNEL analysis (% positive cells) in mononuclear bone marrow cells from patients with RARS (A) and healthy controls (B). TUNEL positivity was analysed just after separation (d 0) and after 4 d in culture with FCS.

Figure 3.

Apoptosis in mononuclear cells from patients and controls: effects of Fas antibodies and caspase inhibition. Apoptosis as measured by TUNEL analysis (% positive cells) in mononuclear bone marrow cells from patients with RARS (A) and healthy controls (B). The figure shows the effects on d 4 of incubation with FCS (control culture), f(ab)′2 (Fas-blocking antibody), CH-11 (Fas-triggering antibody) and DEVD-fmk (caspase-3 inhibitor).

Phosphatidylserine exposure parallells nuclear apoptotic changes

Flowcytometric analyses were performed on MNCs from five RARS before treatment, in both patients analysed after treatment and in five of the controls (Fig 4). Phosphatidylserine (PS) exposure on MNC cells incubated with FCS, f(ab)′2 and CH-11 parallelled nuclear apoptotic changes analysed using the TUNEL technique (Fig 5). Basal PS exposure, shortly after separation, was somewhat higher in untreated RARS than in controls (11·5 ± 9·7 vs. 2·5 ± 1·1, P = 0·07), a tendency that was maintained during culture. RARS cultures were more sensitive to Fas stimulation than control cultures (PS on d 3, 25·1 ± 14·1 vs. 9·1 ± 4·8, P < 0·05). Significant changes in PS expression were not induced by f(ab)′2 or DEVD-fmk.

Figure 4.

Phosphatidylserine surface expression during cell culture in one RARS patient and one healthy control: effects of Fas antibodies and caspase inhibition. Annexin V staining showing phosphatidylserine surface expression on mononuclear bone marrow cells after separation and after 3 d in culture. The figure shows one representative patient with RARS (left, light grey areas) and one control (right, dark areas). (A) After separation, d 0. (B) FCS. (C) f(ab)′2 (Fas-blocking antibody). (D) CH-11 (Fas-triggering antibody). (E) DEVD-fmk (caspase-3 inhibitor).

Figure 5.

Correlation between two methods for detection of apoptosis: phosphatidylserine surface expression and TUNEL positivity. Fas-induced phosphatidylserine surface expression (PS), as detected by Annexin V, corresponded well with Fas-induced TUNEL positivity. Figure shows results of induction with CH 11; x-axis, TUNEL positivity (%), d 4; y-axis, Annexin V (%), d 3. P = 0·14.

Caspase inhibition results in enhanced erythroid colony growth from RARS CD34+ cells

Details regarding the CD34+ separation are given in Table III. The mean yield (%) of CD34+ cells was 0·76 ± 0·36% from RARS MNCs and 0·54 ± 0·32% from control MNCs. After two consecutive separations, the percentage of CD34+ cells exceeded 95%. Basal CFU-E growth was significantly reduced in RARS cultures compared with control cultures (39 ± 42 vs. 246 ± 168, P < 0·01). A less pronounced reduction was observed in CFU-GM growth (72 ± 72 vs. 138 ± 75, P = 0·13). Normal erythroid and myeloid growth was observed only in one patient, with mild disease. Fas stimulation reduced erythroid growth to approximately two thirds of the FCS control in both RARS and controls. Fas-blocking fragments did not induce significant changes in either erythroid or myeloid colony growth, even if single patients with RARS showed an increase in the number of colonies. No difference was observed between RARS and control in this respect (P-value 0·52). Caspase inhibition did not affect myeloid growth in either control or RARS cultures (81 ± 57% vs. 99·8 ± 43% of the FCS control respectively, NS). DEVD-fmk enhanced CFU-E between 58% and 320% in four of the RARS cultures. The overall increase in CFU-E after caspase inhibition was significant for RARS (162 ± 99%, P < 0·05), while control cultures remained unaffected (96 ± 41%, NS).

Table III.  Colony data from CD 34+ cells from nine patients with RARS and mean and median values from five healthy controls.
PatientMNC × 109/% CD34+Controlf(ab)′2CH-11DEVD-fmkControlf(ab)′2CH-11DEVD-fmk
  • *

    Expressed as the mean of three wells.

  •   †Number of cells put on column/% yield of CD34+ after two separations.

  •   ‡This patient had a normal total number of CFU-GM, but showed a disturbed pattern with a dominance of small monocytic colonies.

  • Two patients (2 and 3) were analysed twice, before and after 20 weeks of successful treatment with G-CSF + EPO.

RARS 1100/0·45% 15 28  0 16 23 52  0 37
RARS 2a 80/0·19% 17  7  4  6216197 54206
RARS 2b300/0·48%109 10 10 12 87  7  3 15
RARS 3a100/0·45%  2  0  1  2  5  4  1  7
RARS 3b100/0·60% 18 15  8 13 58 69 27 54
RARS 4 63/1·35%126186 75139150231115187
RARS 6106/0·85%  9 39  8 29 67125 16 60
RARS 7100/1·15%  8  7  0 13 35 27  4 14
RARS 8200/0·70% 83112130131  9 12 19 65
RARS 9124 × 109/0·73% 36 95 49109 39 20 22 16
RARS 10100 × 109/0·90% 51ND 21 93101ND 40 95
Mean patients124 ± 71/0·70 ± 0,35% 38 ± 42 59 ± 66 30 ± 46 56 ± 56 65 ± 71 84 ± 71 26 ± 37 69 ± 75
Median patients100 (range 63–300) 17 (2–126) 34 (0–186)  4 (0–130) 29 (2–139) 39 (5–216) 40 (4–231) 16 (0–115) 37 (7–206)
Mean C124 ± 22/0·54 ± 0·32%186 ± 89243 ± 143121 ± 110165 ± 112151 ± 80146 ± 108 70 ± 68140 ± 114
Median C 25 (range 17–70)200 (59–288)241 (102–387)134 (7–245)161 (55–346)144 (79–238) 87 (81–271) 47 (9–181)127 (5–313)

Bcl-2 expression in RARS

All RARS and control MNCs showed very low amounts of Bcl-2 protein, compared with the Jurkat and P 39 cell lines, as detected by Western blot analysis. No changes were observed after incubation with any of the antibodies (data not shown).

Higher caspase-3 expression in myeloid than in erythroid cells

Expression of pro-caspase-3 was analysed in three patient and two control samples. There was no difference between control and RARS cells and no changes over time. No cleavage of the 32-kDa procaspase-3 into its active 17-kDa subunit could be detected and there were no differences in amount of procaspase-3 under the various culture conditions (Fig 6A and B). As caspase inhibition had different effects on erythroid and myeloid colony growth, we also analysed caspase-3 in separated GPA+ and GPA cells, which were obtained using the Mini Macs system, as described for CD34. Fas-treated Jurkat cells are shown as a positive control. The expression of pro-caspase-3 was significantly higher in non-erythroid (GPA) cells than in GPA+ cells (Fig 6C). In contrast, an additional band of approximately 19 kDa was recognized by the caspase-3 antibody in all bone marrow samples, although it was conspicuously absent in the Jurkat cells (Fig 6A).

Figure 6.

Caspase-3 protein expression in bone marrow cells from patients and donor: effects of Fas antibodies and caspase inhibition and differences between erythroid and non-erythroid cells. Western blot analysis of caspase-3 protein expression of mononuclear bone marrow cells from one patient (A) and one control (B). In the patient sample, cells were analysed just after separation (d 0), and after 18 h (d 1) and 42 h (d 2) of incubation. Fas-treated Jurkat cells with the caspase-3 cleavage product p17 are shown in (B) to compare the different molecular weights. (C) shows caspase-3 protein expression and processing in GPA+ vs. GPA subpopulations.

Fas-induced caspase-3-like activity is increased in RARS

The optimal time-point for measurement of caspase-3-like activity was 4–24 h. No significant difference between RARS and controls in the cleavage of DEVD-AMC in MNCs could be observed immediately after separation (Fig 7). After 4 h and 24 h in FCS culture, RARS cells showed more activity than the controls (P = 0·13 and < 0·05 respectively). Fas stimulation induced more caspase activity in RARS than in controls, both after 4 and 24 h (P = 0·003 and P = 0·01 respectively). F(ab)′2 fragments showed no effect on caspase activity in any of the cultures and caspase inhibition, using DEVD-fmk, significantly reduced activity in all cultures. When caspase activity was studied separately in GPA+ vs. GPA cells, the activity just after separation in the GPA cells was 10 times higher than in the GPA+ cells, and a Fas-induced increase was observed only in GPA cells (data not shown).

Figure 7.

Caspase-3-like activity in bone marrow cells from patients and donor: effects of Fas antibodies and caspase inhibition. Caspase-3-like activity in mononuclear bone marrow cells from patients with RARS (1–10) and healthy controls (c1–6). The figure shows the effect of incubation with FCS (control culture, A), f(ab)′2 (Fas-blocking antibody, B), CH-11 (Fas-triggering antibody, C) and DEVD-fmk (caspase-3 inhibitor, D). The figure shows caspase activation after 4 h of incubation. Individual patients/controls are shown by drawing a line between the results of each individual.

Successful treatment with cytokines is associated with improved haematopoietic stem cell function

Both patients analysed before and after treatment showed a complete response to therapy with G-CSF + EPO (Table I). The percentage of myeloblasts, bone marrow cellularity and M/E ratio did not differ significantly before and after treatment. The Fas-induced pattern of proliferation was clearly altered. Both samples after treatment showed a Fas-triggered increase in proliferation, similar to that observed in the two benign RARS cases and in some of the controls (Fig 1). In patient 2, pretreatment proliferation was increased by DEVD-fmk (23 × 103 vs. 41 × 103 cpm), while no change was observed after treatment (17 × 103 vs. 16 × 103 cpm). Caspase-3-like activity at 0 h was lower than in the range of the controls in the samples taken after treatment. Fas-induced activation was lower after treatment (12·42 pmol/min before vs. 3·62 pmol/min after treatment). Endogenous CFU-E were increased in both patients, with 640% and 900% respectively (Table II). CFU-GM were changed in patient 2, in the sense that the increased monocytic growth was reduced, and in patient 3, with an increase in CFU-GM (5–58 colonies). PS exposure on d 0 was unaltered in patient 2, but reduced to normal levels in patient 3, in which Fas-triggered PS exposure was also reduced from 44% to 12%.


In the present study, we chose a relatively homogeneous patient group, RARS, in which there might be a common disease mechanism. The aim was to study different potential mechanisms for the enhanced erythroid apoptosis in this subgroup, as well as the cellular differentiation level for the starting point of this process.

The present data demonstrated that RARS originates in the myeloid stem cell compartment, at least as defined by CD34 positivity. Even patients with erythroid dysplasia only (classified as pure sideroblastic anaemia, PSA, according to Gattermann et al, 1990), showed a CFU-GM growth of less than 10% of control. Evidence for monoclonality has been previously shown in mononuclear cells from patients with RARS (Amenomori et al, 1987; Suzuki et al, 1999). Our results prove that the defect lies at the CD34+ level and affects both erythroid and myeloid differentiation. The reduction of CFU-E reflects the severe anaemia of these patients, but the marked reduction in CFU-GM, in spite of normal leucocyte counts in the majority of patients, remains to be elucidated. One explanation could be mitochondrial DNA mutations affecting cytochrome c, for example, which have recently been described in RARS patients (Gattermann et al, 1997, 1999; Bröker et al, 1998). If RARS patients have mutations affecting respiratory chain processes or iron transport, this could be crucial for erythropoiesis, while the myelopoiesis only needs this system for housekeeping and may manage to maintain effective myelopoiesis in spite of these changes.

The Fas ligand and the caspase pathway are important natural regulators of erythropoiesis. One aim of our study was to characterize the relationship between these two systems for apoptosis and survival. Recent studies have shown that Fas stimulation results in caspase activation, a blockage in erythroid differentiation and a cleavage of GATA-1 (DeMaria et al, 1999a, b). It was also suggested that the Fas ligand, expressed by late erythroid progenitors, inhibits the proliferation of early progenitors, via the Fas receptor, thus creating a natural feedback loop to maintain a steady-state erythropoiesis. A general caspase inhibitor, z-VAD-fmk, was shown to inhibit this reaction. We used z-VAD-fmk in preliminary experiments, but found that viability was reduced after 4 d in culture (data not shown).

We tested the effect of Fas-blocking fragments in several assays, estimating proliferation, clonogenic growth and apoptosis in RARS bone marrow. The Fas-blocking fragments did not show any effect that was not comparable with the effects on normal bone marrow. Thus, even if the Fas system retains a regulatory role in RARS bone marrow, it seems to be equivalent to its role in normal haematopoiesis. Our experiments gave no reason to believe that a continuous overactivation/overstimulation of the Fas receptor to any significant degree participates in the spontaneous erythroid apoptosis in RARS. In a previous study including various subtypes of MDS, there was a correlation between ineffective erythropoiesis and Fas expression (Fontenay-Roupie et al, 1999). However, only three out of nine RARS showed increased Fas expression, which was less than that observed in the other subgroups. Thus, the presence of an increased expression of Fas receptor on the cell surface of MDS cells does not prove that this is the main mechanism for the increased spontaneous apoptosis in MDS. The mechanisms of ineffective erythropoiesis in RA and RARS may be different and it is possible that the Fas system has a more central pathogenetic role in other subtypes of MDS.

Fas ligation induced apoptosis both in RARS and control cultures but led to significantly more PS exposure and caspase activation in the RARS cultures. Fas expression was detected only in a few percent of the CD34+ cells, which makes it improbable that this would have a significant impact on the induction experiments (data not shown). Fas-induced caspase activity in MDS has also been shown in other studies (Sloand et al, 1998), but our results point out the difference between normal cells and RARS cells. The majority of RARS cultures showed a decrease of proliferation in response to Fas ligation, but two patients with non-severe RARS and both patients responding to cytokine treatment showed a Fas-induced increase in proliferation after treatment. Two recent studies on human fibroblasts and in normal human bone marrow CD34+CD38 cells, respectively, show that Fas stimulation might lead to increased proliferation or cell survival (Aggarwal et al, 1995, Josefsen et al, 1999). We therefore suggest that bone marrow cells from patients with RARS are more susceptible to Fas ligation than normal bone marrow cells, and that the pathogenetic mechanism leading to apoptosis or defective growth in RARS is located downstream of the Fas receptor. This defect may lead to a poorer capacity of proliferation or cell survival in RARS cells in response to Fas stimulation.

Caspase inhibition by DEVD-fmk led to different reactions in RARS and control cultures. It reduced MNC apoptosis and increased proliferation in RARS cells, but not in normal cells. It improved RARS erythroid growth from CD34+ cells, while normal erythroid growth and myeloid growth in all cultures remained unchanged. Thus, a growth-enhancing effect of caspase inhibition was mainly observed on erythroid differentiation from RARS CD34+ cells, which suggests that enhanced caspase activation in RARS is already present in the stem cell population and that erythroid apoptosis is initiated at this level. This hypothesis is further supported by the low protein expression of caspase-3 and the low basal caspase-3-like activity in GPA+ cells, and the fact that Fas-induced caspase-3-like activity was not observed in this cell population (data not shown). Increased caspase activity in MDS bone marrow has been reported, but has not been held responsible for the impaired erythroid differentiation in MDS (Mundle et al, 1999c). Gregoli & Bondurant (1999) showed that EPO deprivation caused erythroid apoptosis in normal cells and that caspase inhibition could delay this apoptotic process. Considering the basically negative results of the Fas inhibition, it is possible that caspase activity is caused not only through Fas-mediated activation, but also through alternative signalling and/or amplification via the mitochondria (Fulda et al, 1998).

Western blot analyses of pro and active caspase-3 revealed the presence of an additional p19 band in normal and RARS cells. Similar results were recently described by Gregoli & Bondurant, 1999). However, in MNC cells deprived of GPA+ cells, this p19 band was absent. The presence of an intermediate p19 band in bone marrow samples, but not in the leukaemic Jurkat T-cell line, suggests that differences in the processing of pro-caspase-3 are cell type-specific. Indeed, a similar possibility was recently discussed by other investigators (Faleiro et al, 1997). Caspase-3-like activity, which was detected by cleavage of fluorogenic substrate at 4 h and decreased at 24 h, was never correlated with the appearance of p17 cleavage product of p32 native pro-caspase-3. One possible explanation is that the sensitivity of the Western blot technique is not high enough to detect this cleavage product. This also related to very low activity of this enzyme in tested samples compared with cell lines.

Normal or RARS MNCs did not express Bcl-2 protein and no changes were observed during culture. This is not an unexpected finding as previous studies have mainly shown Bcl-2 expression in MDS with an increase of bone marrow blasts (Rajapaksa et al, 1996; Parker et al, 1998). It is not possible to draw any specific conclusions about RARS as one of these studies included no patients with RARS and the other grouped all patients with blasts < 10%. However, it is possible that it would be more relevant to study Bcl-XL in stem cells and early progenitors from RARS marrow because this protein has been shown to be more important for erythropoiesis than Bcl-2 (Silva et al, 1998).

Treatment with G-CSF + EPO normalized haemoglobin levels and markedly improved erythroid clonogenic growth. An interesting finding was that Fas-induced proliferation switched from suppression to an increment, thus approaching the pattern observed in two of the controls and in the two RARS patients with mild disease. Moreover, Fas-induced caspase activation was reduced to the same level as in the control samples, and caspase inhibition (which enhanced proliferation before treatment) showed no effect after treatment. These results further support the role of caspase activation in the apoptotic process in RARS. It is also probable that anti-apoptotic treatment with growth factors may reverse this reaction, either by suppressing some steps in the caspase pathway or by selecting less damaged stem cells. It is improbable that the mechanism of action of growth factors is a decrease in Fas expression, as growth factors have been show to increase this expression (Stahnke et al, 1998). Considering that G-CSF significantly improves the treatment response to EPO in RARS and the results showing that EPO deprivation induces caspase activation (Hellström-Lindberg, 1997, 1998; Gregoli & Bondurant, 1999), it is possible that RARS cells are biochemically altered so that they need a supranormal concentration of growth factors to survive.

An unbalanced relation between apoptosis and proliferation is a key feature of many diseases (Fadeel et al, 1999b). The main problem for patients with RARS is the chronic transfusion need caused by ineffective erythropoiesis and excessive erythroid apoptosis. A thorough characterization of the mechanisms of erythoid apoptosis is neccessary to address this problem and improve treatment in RARS. Our results show that enhanced caspase activation, independent of the Fas/Fas ligand interaction contributes to the erythroid apoptosis in RARS. Moreover, the present work provides evidence that erythroid apoptosis in RARS is initiated at the stem cell level.


This work was supported by grants from the Swedish Cancer Society (Cancerfonden, 3829-B98–0310AC and 3689-B99–0510BB). J.S.M. was also supported by a grant from the Cancer Society in Stockholm (98:112).

We thank Dr Nicholson (Canada) and Dr Krammer (Germany) for p17 antibodies and f(ab)′2 blocking antibodies respectively.