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Sensitivity, Resistance to Therapy and Apoptosis

Study of apoptosis-related responses of leukemic blast cells to in vitro anthracycline treatment


Anthracyclines trigger an apoptotic cell death but their molecular targets are not totally explored. We investigated the apoptotic response of blast cells and lymphocytes from medullary samples of 31 de novo acute leukemia. Mononuclear cells were treated in vitroby therapeutic concentrations of either daunorubicin (DNR) or idarubicin (IDA) for 1 h, washed and cultured for 18 h. A multivariate analysis using flow cytometry and a CD45 gating on lymphocytes and blast cells was performed. DNR and IDA induced a Fas enhancement on both leukemic and normal cells. In blast cells the DEVDases were activated and the caspase 3 was cleaved in relation to phosphatidyl serine exposure, showing a caspase-dependent pathway in anthracycline-induced apoptosis. Apoptotic percentages were always higher for blast cells than for lymphocytes, confirming that anthracycline toxicity mainly affected tumor cells. Moreover, drug-induced apoptosis was not related to spontaneous apoptosis, suggesting that variations in response intensities were due to individual variations of sensitivity rather than to programmed life span time. The apoptotic response of P- glycoprotein-expressing blast cells was not significant, giving biological argument for the poor prognosis of multidrug resistance leukemia. Finally, Fas induction and anthracycline-induced apoptosis on blast cells were significantly higher when a complete remission was achieved, thus shedding light on potential new prognostic factors in acute leukemia.


Fas (APO-1, CD95) is a tumor necrosis factor family membrane protein which triggers programmed cell death when engaged by anti-Fas antibody or Fas-ligand.1 The death pathway initiated by Fas activation involves a series of death-induced molecules.1 Fas-associating protein with death domain (FADD) or MORT-1 is recruited to Fas upon its engagement.23 FADD then binds FADD-like ICE (FLICE) or MORT-1-associated CED-3 homologue.45 The association with the Fas death-inducing signaling complex activates FLICE,6 followed by activation of the caspase proteolytic cascade and apoptosis.7 Fas is found on immunity cells (lymphocytes, NK cells, monocytes) but also on numerous tumor cells such as tumor cell lines or leukemic blast cells.89 Its expression is decreased during tumor cell enhancement or during the cell acquisition of the drug resistance phenotype.1011

Several cytotoxic drugs (cisplatin, doxorubicin, bleomycin, mitomycin, methotrexate) as well as ionizing radiation can sensitize many tumor cells to Fas-induced apoptosis.1213141516 This sensitization is mediated by a transcriptional and a post- transcriptional regulation of the fas gene, which leads to receptor accumulation on the drug-treated membrane cells. The increase in Fas on tumor cells after low-dose chemotherapy may be used as a therapeutic target. Because a synergy between Fas-ligand and cytotoxic drugs was observed in vitro, the Fas increase could enhance tumor cell elimination by the immune cells.17 These observations supported the rationale for a new therapeutic strategy which could combine chemotherapy and immunotherapy for cancer treatment.1418

For many years, the anthracyclines have been used in leukemia treatment and it is known that anthracyclines induce an apoptotic cell death in a wide range of cultured cells.192021 Therapeutic daunorubicin (DNR) concentrations are about 1–2 μM.22 At these doses, DNR induces apoptosis in U937 and HL60 leukemic cell lines.2324 The Fas apoptotic pathway could at least partially play a role,1213141516 involving both death receptor/ligand accumulation on the cell surface17252627 and activation of the death effectors such as caspases in the cytoplasm.28 Expression of the mdr-1 gene leads to the multidrug resistance phenotype, which is one of the major cause of failure in cancer therapy.2930 This gene encodes for a 170 kDa ATP binding glycoprotein (PGP) which acts as a pump to exclude therapeutic agents from the cell,31 thus decreasing the intracellular concentration of the drug and the apoptotic cell response.32 However, the mechanisms mediating the anthracycline-induced killing of leukemic blast cells are not definitively known and a possible correlation between the apoptotic potential of blast cells and the clinical outcome of patients has never been explored.

In the present work, we investigated the apoptotic response of in vitro anthracycline-treated leukemic blast cells and normal lymphocytes isolated from medullary samples of 31 patients suffering from de novo acute leukemia (AL). Some mechanisms of cellular drug resistance (PGP expression, apoptotic deficiency) and others of cell death (Fas expression, DEVDase activation, caspase 3 activation, phosphatidyl serine exposure) were assayed. A possible correlation between these parameters and the clinical outcome of patients was then examined. We clearly show that both anthracyclines (DNR and idarubicin (IDA)) induce apoptosis in leukemic blast cells and lymphocytes. This death was accompanied by a Fas expression enhancement on both lymphocytes and blast cells. Caspase 3 was cleaved and consequently activated and the caspase 3 cleavage was correlated with phosphatidyl serine exposure. When PGP was expressed on the cell membrane, the apoptotic potential of blast cells was strongly decreased, giving biological argument for the poor prognosis of PGP-positive acute leukemia. The apoptotic response intensity of blast cells was significantly associated with the clinical outcome of patients, thus shedding light on potential new prognostic factors in acute leukemia.

Patients and methods

The study included 30 patients with previously untreated de novo AL who were treated in our institution between November 1998 and August 1999. Another patient who relapsed 1 year after the end of chemotherapy was included. One patient was not followed up for his clinical evolution because he was treated in another institution.

All the patients gave informed consent. Their disease was classified according to the recommendations of the French–American–British (FAB) committee.33 Twenty-four acute myeloid leukemia (AML) and seven acute non-myeloid leukemia (ANML) were found. According to the type of AL, the induction chemotherapy varied but all patients received an anthracycline, either DNR or IDA. On September 1999, 15 patients had survived in complete remission (SCR) and 10 patients died after treatment failure (DTF). The remaining were not classified because no post-mortem biopsies were done or because there was not enough distance from the first chemotherapy induction. Complete remission was defined as the presence of fewer than 5% blast cells in a cellular BM smear, and the absence of circulating blast cells and extramedullary leukemic cell infiltration. The clinical and hematological characteristics are summarized in Table 1.

Table 1  Clinical and hematological characteristics of patients

BM collection and preparation

Before induction chemotherapy, BM aspirates were collected for diagnosis and 1 ml was sampled on sodium heparinate tubes (Roche, France). Bone marrow mononuclear cells (BM-MNC) were isolated by centrifuging half diluted BM over Lymphoprep (Nyegeaard, Oslo, Norway) for 40 min at 400 g. BM-MNC were washed in RPMI 1640 (Gibco-BRL, Eragny, France) and resuspended in culture medium (CM) consisting of RPMI 1640 medium supplemented with 10% fetal calf serum (FCS) and 1 mM L-glutamine (Gibco-BRL), 10 mM Hepes (Gibco-BRL), 100 units/ml penicillin (Gibco-BRL), 50 μg/ml streptomycin (Gibco-BRL). Finally, the cell concentration was adjusted to 1 × 106 cells/ml.

Cell incubation with daunorubicin or idarubicin

The cell suspension was distributed in three tubes. One was incubated without anthracyclines (NT) for 1 h at 37°C, the second was incubated with 1 μM daunorubicin (DNR, RPR-Bellon, Neuilly, France) for 1 h at 37°C and the third was incubated with 0.2 μM idarubicin (IDA, Pharmacia, Saint Quentin en Yvelines, France) for 1 h at 37°C. Then, BM-MNC were washed, resuspended in CM and cultured for 18 h in humidified 95% O2 and 5% CO2 atmosphere at 37°C.

Cellular response to anthracycline exposure

Flow cytometry (FCM) analysis:

Fas and PGP expression, apoptosis percentage, and caspase 3 activation were assayed by FCM with an EPICS XL cytometer (Coulter, Margency, France). The CD45 differential expression allowed a selective gating of lymphocytes and blast cells34 and the cell response to anthracycline exposure was thus assessed simultaneously in these two cellular populations.

After centrifugation (5 min, 400 g), 5 × 105 cells from NT, DNR and IDA suspensions were resuspended in 100 μl phosphate buffer saline (PBS, Eurobio, France) supplemented with 10% FCS (PBS-FCS) and labeled with 5 μl of PC5-conjugated anti-CD45 antibodies (Coulter-Immunotech), 5 μl of FITC-conjugated anti-Fas antibodies (clone UB-2, Immunotech, France), and 5 μl of PE-conjugated anti-PGP antibodies (clone UIC2, Immunotech, France). Negative controls were established with corresponding FITC or PE-conjugated-isotypic irrelevant IgG and PC5-conjugated anti-CD45 antibodies. Erythrocytes were lyzed with multi-Q-Prep (Coulter-Immunotech) according to the manufacturer's instructions and analysis was performed by FCM.

The mean fluorescence intensity (MFI) was measured in lymphocyte and blast cell populations of NT, DNR and IDA samples and the specific mean fluorescence intensity (MFIsp) relating Fas or PGP expression was calculated by subtracting negative control values. Fas or PGP expressions were considered as positive when the MFIsp was found positive. Such an interpretation was allowed by gating on the homogeneous population.

BM-MNC (5 × 105) from NT, DNR and IDA suspensions were incubated in 100 μl culture medium containing 5 μl of PC5-conjugated anti-CD45 antibodies (Coulter-Immunotech) for 20 min. The FITC-annexin V kit (Coulter-Immunotech) was used after modification of the manufacturer's specifications: binding buffer (500 μl) containing 5 μl FITC-annexin V (Coulter-Immunotech) was added and further incubated for 20 min on ice. The addition of propidium iodide which is usually proposed to stain permeabilized dead cells was omitted because the cells were further fixed, permeabilized and stained with a PE-conjugated antibody whose fluorescence spectrum overlaps the propidium iodide spectrum. The cells were then pelleted, resuspended in 1 ml Permeafix (Ortho Diagnostic Systems, Roissy, France) and incubated for 40 min at 20°C. The suspension was then centrifuged and the pellet was washed twice with washing buffer (0.2 mM EDTA, 5% FCS in phosphate buffer saline (PBS)). Labeling was performed by adding to the cells 100 μl of washing buffer containing 5 μl of polyclonal PE-conjugated anti-active caspase 3 antibodies (Pharmingen-Becton Dickinson, Le Pont de Claix, France). The samples were gently stirred for 1 h, and washed in washing buffer. A negative control sample incubated with PE- conjugated IgG was run in parallel. Then, FCM analysis was performed at 525 nm for annexin binding, 575 nm for cleaved caspase 3 labeling and 675 nm for CD45 expression. The FITC-annexin V binding cells were considered as apoptotic and the negative cells as viable. From the same analysis, positive and negative cell populations for the active caspase 3 were individualized.

Fluorimetric analysis:

After CD45 labeling as described above, blast cells were sorted from NT, DNR, and IDA samples with an ELITE cell sorter (Coulter). The cell flow was set to analyze 2000 events/s, and cells in the windows of interest were sorted in sterile tubes containing 0.5 ml PBS. Sorted cells (5 × 105) were washed in PBS and processed for DEVDase activity as described below.

DEVDase activity was evaluated as previously described.2435 Briefly, 3 × 105 cells were washed in PBS, then suspended in 50 μl of a pH 7.4 permeabilizing buffer containing 10 mM HEPES, 5 mM dithiothreitol, 0.02% Saponine, 1 mM PMSF, 10 μg/ml Pepstatin A, 10 μg/ml Leupeptin, 72 μM fluorogenic substrate Ac-DEVD aminomethylcoumarin (UBI, Euromedex, Souffelweyersheim, France). Cells were incubated for 10 min at 37°C and then centrifuged for 3 min at 14000 r.p.m. The supernatant was diluted in 1 ml dH2O. Fluorescence was measured using a Jobin and Yvon spectrofluorometer (λexc = 380 nm, λem = 480 nm). A blank was performed without addition of cells in the substrate buffer and its value was subtracted from all the measurements. A control without any drug treatment was processed for each experiment. The relative enzyme activity was expressed as the ratio of treated sample activity to untreated control activity or as fluorescence units per 105 sorted blast cells.

Statistical analysis

The paired Student's t-test and the Chi-square test were used to analyze the data.


Fas expressed by blast cells and lymphocytes in all the AL

Because the Fas apoptotic pathway is involved in anthracycline-induced apoptosis,17 Fas expression was assessed on the membranes of blast and lymphocyte cells. Anthracycline-treated and control cells were labeled with an anti-CD45 and an anti-Fas antibody as described in the Patients and methods section and analyzed by FCM after blast cell or lymphocyte CD45 gating (Figure 1a). Almost all the AL were found positive for Fas expression on both blast and lymphocyte cells (Figure 1d). Moreover, whatever the gating on blast or lymphocyte cells, histogram analysis after anti-Fas labeling (Figure 1c, left) was clearly right-shifted in comparison with the isotypic control (Figure 1b, left), so the whole blast cell and lymphocyte populations expressed Fas.

Figure 1

 Percentage of AL or AML expressing Fas or PGP on lymphocytes and blast cells. MNC were labeled with a PC5-anti-CD45 antibody, a FITC-anti-Fas, and a PE-anti-PGP antibody. FCM analysis was performed. Bivariate CD45-SSc, Fas-PGP and the Fas- or PGP-fluorescence histograms are shown. (a) Gating based upon the differential CD45 expression allowed the simultaneous study of Fas and PGP on lymphocytes and blast cells. CD45 fluorescence is on x axis and side scatter (SSc) on the y axis. Examples of histograms gated on blast cells are represented respectively for isotypic controls (b) and positive labeling (c). The FITC (Fas-related) fluorescence histograms are on the left, the PE (PGP-related) histograms on the right and the bivariate (FITC/PE) histograms in the middle of the Figure. The mean fluorescence intensity of isotypic labeling was subtracted from the corresponding mean fluorescence intensity after Fas or PGP labeling and constituted the specific mean fluorescence intensity (MFIsp). A cell population was considered to express Fas or PGP when its MFIsp was positive (b, c). The frequencies (in percentages) of acute leukemias (AL) expressing Fas (d) or PGP (e) on their blast cells and/or on their lymphocytes are shown.

PGP more frequently found on lymphocytes than on blast cells

PGP expression is able to confer multidrug resistance phenotype36 and so is likely to modify the cellular response to anthracyclines. As was done for Fas expression, PGP expression was assessed by FCM on blast and lymphocytes cells as described in Patients and methods. Interestingly, 29% of our AML were positive for PGP expression on lymphocytes although only 13% expressed PGP on blast cells (Figure 1e). Thus, studying PGP expression on total MNC could lead to overestimating the percentage of PGP-positive AL. Therefore, a specific analysis of blast cells, as individualized by low CD45 expression, appeared necessary. Isotypic control (Figure 1b, right) and PGP-labeled (Figure 1c, right) histograms were almost directly superimposed and only a small population was positive after PGP labeling, so only a part of blast and lymphocyte cell populations expressed PGP in some of the AL. Bivariate histogram analysis (Figure 1b and c, middle) showed that PGP-positive cells expressed Fas as well as PGP-negative cells.

IDA induces Fas expression on leukemic cells

The variation in Fas expression on anthracycline-treated cells was measured by FCM. The results showed that IDA, but not DNR, led to a significant increase in Fas expression on blast cells of total studied AL (Figure 2a) and on PGP negative blast cells of AML (Figure 2b) and ANML (Figure 2c). Fas was not induced when PGP was expressed (Figure 2b), suggesting that Fas expression may be related to the intracellular anthracycline accumulation. Induction of the expression of Fas on anthracycline-treated tumor cells could sensitize these cells to apoptosis.17 As caspase activation is considered to be a common feature in apoptosis,37 it was interesting to verify if the effect of anthracyclines was accompanied by activation of caspases. DEVDase activity was therefore measured on sorted blast cells.

Figure 2

 Fas expression on leukemic blast cells after anthracycline exposure. MNC were treated by either DNR (1 μM) or IDA (0.2 μM) or non-treated (NT) and labeled for CD45, Fas and PGP as described in Patients and methods. FCM analysis was performed. For each sample, the Fas expression on blast cells was calculated as the MFIsp and the mean Fas expression was plotted for the AL (a), for the AML expressing or not the PGP on blast cells (b), and for the ANML (c). The s.d. are indicated. * to n*: P < 5.10−2 to P < 5.10−(n+1) for the paired Student's t-test.

Anthracyclines activate caspase 3 and induce DEVDase activity in leukemic blast cells

Blast cells were isolated by flow cell sorting and the DEVDase activity was measured by fluorimetry in anthracycline-treated and control samples as described in Patients and methods. Both DNR and IDA led to a significant activation of DEVDase in AL blast cells (Figure 3a). The increase in blast cells DEVDase values after anthracycline treatment was nearly two-fold compared to the control (Figure 3a) and the DEVDases were similarly activated after both DNR and IDA treatment. Interestingly, when PGP was expressed on AML blast cells, no DEVDase activation occurred (Figure 3b). As for Fas expression, the DEVDases activation may be related to the intracellular anthracycline accumulation.

Figure 3

 DEVDases and caspase 3 activation in anthracycline-treated cells. MNC were exposed to anthracyclines as previously described. (a, b) After blast cell sorting based upon their low CD45 expression, DEVDase activity was measured as described in Patients and methods using a fluorogenic substrate and spectrofluorimetry. The mean relative DEVDase activation (activity of the DNR or IDA treated sample/activity of the untreated control) was plotted for the AL (a), and for the AML expressing or not the PGP on blast cells (b) and the s.d. are indicated. The cleavage of caspase 3, which led to the activation of this enzyme, was assayed by FCM after MNC labeling with a PE-anti-cleaved-caspase 3 antibody as described in Patients and methods (c). Neg indicates the cells negative for active caspase 3 (uncleaved form) and Pos the cells positive for the active caspase 3 (cleaved form). (d) The mean percentage of blast and lymphocytes cells containing active caspase 3 was plotted for the 10 AL studied and the s.d. were indicated.

Because caspase 3 is the main effector of the caspase family and belongs to the DEVDase family, the specific activation of this enzyme was assayed by FCM after anthracycline treatment. During apoptosis, the enzyme is activated by the cleavage of the procaspase 3 zymogen. This activation was evaluated by FCM both in lymphocytes and blast cells after CD45 gating using a specific anti-cleaved-caspase 3 antibody as described in Patients and methods (Figure 3c). Anthracycline (DNR and IDA) treatment triggered caspase 3 activation in the lymphocytes and blast cells of 10 AL (Figure 3d). The percentage of positive cleaved caspase 3 lymphocytes was significantly lower than the percentage of blast cells in control samples (P < 5.10−4, Figure 3d), suggesting that the life span was higher for lymphocytes. Moreover, this effect was strictly reproducible for all anthracycline treated samples, thus suggesting a greater specificity of anthracycline for tumor cells.

The last event of the ‘roads to ruin’38 is apoptosis by itself. We previously described that caspase activation is an early event of apoptosis, occurring before morphological changes.24 Consequently, apoptosis following caspase activation in lymphocytes and blast cells was quantified after anthracycline treatment in lymphocytes and blast cells by the binding of FITC-annexin V.

Anthracyclines trigger caspase 3-mediated apoptosis in normal and leukemic cells

The percentage of apoptotic lymphocytes and blast cells was quantified by FCM after annexin V labeling and CD45 gating as described in Patients and methods. DNR and IDA induced apoptosis both in lymphocytes and in blast cells of total AL (Figure 4b). When PGP was expressed on AML blast cells, no significant apoptosis was measured beyond spontaneous apoptosis (Figure 4c). This observation reinforces the idea that anthracycline-induced apoptosis is related to intracellular drug accumulation and gave supplementary evidence for the deleterious role of PGP in AL.

Figure 4

 Anthracycline-induced caspase-dependent apoptosis in leukemic blast cells. Bone marrow mononuclear cells were either untreated or treated with DNR or IDA. The cells were then labeled with PC5-CD45, FITC-annexin V, PE-anti-cleaved caspase 3 as described in Patients and methods. Blast cells and lymphocytes were gated as described in on the basis of their CD45 expression. (a) Flow cytometric analysis of untreated and anthracycline-treated blast cells after annexin V and anti-activated caspase 3 double labeling. The corresponding monovariate analysis are shown on the extreme left and extreme right parts of the Figure. The mean percentages of apoptotic (annexin V positive) blast cells were plotted for the total AL (b). The AML were divided in AML with either PGP expressing or PGP-negative blast cells (c). The mean and the s.d. of N patients are shown. (d) For 10 AL, the percentage of apoptotic (annexin V-positive) lymphocytes or blast cells and the percentage of activated-caspase 3-positive lymphocytes or blast cells were measured in anthracycline-treated and control samples as described in (a). The apoptotic values (x axis) and the activated-caspase 3 values (y axis) were plotted and the equations and correlation coefficients obtained from linear regressions were indicated.

As noted for caspase 3 activation, there were fewer apoptotic lymphocytes than blast cells in both untreated and anthracycline-treated samples (data not shown). This again suggested that lymphocytes could be more resistant to cytostatic agents and have a longer in vitro life span than blast cells. Both DNR and IDA induced similar percentages of apoptotic cells.

Interestingly, caspase 3 cleavage and annexin V binding were strongly related in blast cells and in lymphocytes (Figure 4d). Moreover, bivariate analysis showed that activated caspase 3 positive cells were also positive for annexin V binding (Figure 4a). These data strongly support the hypothesis that anthracyclines induce apoptosis through the caspase pathway. Moreover, drug-induced apoptosis was not related to spontaneous apoptosis in either blast cells or lymphocytes (data not shown) suggesting that anthracyclines induce apoptosis by a specific mechanism, independently of the cell life span.

Apoptotic potential of blast cells higher for SCR patients

The apoptotic potential of blast cells was compared with the patients clinical evolution. Fas induction on blast cells was significantly higher for SCR patients compared to DTF patients (Figure 5a). Moreover, a statistical analysis using the chi-square test showed that a Fas-induction higher than 50% on blast cells was more frequently observed for SCR patients (χ2 < 0.05). Similarly, blast cell apoptosis was greater for SCR patients (Figure 5b). This difference was significant for the Student's t-test only after IDA treatment. An IDA-induced-apoptosis higher than 20% was more frequently found in SCR patients (χ2 < 0.05). Finally, no correlation was found between leukocytosis on the day of diagnosis and the apoptotic potential of blast cells (data not shown). Leukocytosis reflecting a ratio between apoptosis and proliferation thus appeared independent of the potentiality of blast cells to undergo apoptosis under anthracycline treatment.

Figure 5

 Clinical evolution of patients in relation with the in vitro apoptotic potential of blast cells. Survival in complete remission (SCR) and death after therapeutic failure (DTF) were compared to the in vitro blast cell response after anthracycline exposure. MNC were treated by either DNR or IDA, and the Fas expression and the percentage of apoptosis were evaluated by FCM on blast cells as described above. A control without anthracyclines was processed for each experiment. The rate of increase in Fas expression on blast cells (a) and the percentage of apoptotic blast cells (b) were plotted for SCR and DTF groups and the s.d. are indicated.


The use of anthracyclines heralded a new era in the treatment of leukemia.39 The fact that anthracyclines had a pro-apoptotic effect20 opened up new research fields to investigate the molecular mechanisms triggering tumor cell death. Better understanding of the anthracycline apoptotic effect may lead to new therapeutic agents and to better biological support for drug evaluation.

In this work, we studied the in vitro apoptotic response of lymphocytes and blast cells from medullary samples of 31 patients suffering from de novo AL. MNC were exposed for 1 h to anthracyclines reproducing the in vivo pharmacokinetics of these drugs.2240 Because anthracyclines activate the Fas apoptotic pathway on cell lines,1213141516 Fas expression was assessed on lymphocytes and blast cells. We found that Fas was expressed on both tumor blast cells and lymphocyte cells. This confirms on clinical samples previous work that showed a CD95 expression on various tumor cell lines.9 The use of FCM allowed the detection of Fas in different cellular populations based on a differential CD45 expression,34 thus reflecting the true expression on each cell type and avoiding discrepancies due to contamination by other cell populations.

The expression of Fas on all leukemic blast cells theoretically would enhance the apoptotic response. Indeed, it has been shown that cytotoxic agents such as cisplatin, doxorubicin, and camptothecin sensitize tumor HT29 cells to Fas-mediated apoptosis.17 This sensitization is based upon an upregulation of the fas gene, which leads to an overexpression of the receptor at the cell surface. Since we found such an increase on leukemic blast cells after IDA exposure, this reinforces the idea that IDA, which exhibit a greater cytotoxicity than DNR,41 could be more active on leukemic cells. Fas induction was also found on lymphocytes after IDA and DNR treatment (data not shown), suggesting the activation of similar mechanisms in both normal and tumor cells after chemotherapy. Interestingly, the induction of Fas was not significant when PGP was expressed on blast cells suggesting that Fas expression might be related to intracellular anthracycline accumulation.

PGP, which functions as a pump to expel drugs from the cells, was able to modify the cellular response to drug exposure. Moreover, it has been shown that PGP expression on leukemic blast cells at diagnosis or in relapse is strongly correlated to therapeutic failure.42 In our study, PGP was expressed on lymphocytes in 29% of the AML, while it was found on blast cells only in 13%. A recent study using multivariate analysis on blast cells from 352 patients found a comparable incidence.43 Conversely, older studies on whole MNC found elevated frequencies in the expression of PGP, nearly equal to 50%.3642 Taken together, these data underline the interest of FCM analysis after CD45 gating for the exact determination of the percentage of positive PGP-positive AL.

Evidence for PGP expression on lymphocytes has already been demonstrated444546 and we found lymphocytes expressing the PGP in 29% of the AML. The biological relevance of PGP expression on non-tumor cells appears unclear. It could represent an activation phenotype, reflecting a cellular progression toward particular functions. For example, some authors have shown that PGP in leukocytes could function to transport cytokines, cytotoxic effector molecules, or inflammatory mediators.4748 Our FCM histograms analysis showed that PGP was mostly expressed on a fraction of lymphocytes or blast cells. This suggests that the drug sensitivity of PGP-positive AL could be decreased without abolition of the whole cellular response.

Overall, the efficiency of anthracycline treatment is reflected by the ability of tumor cells to undergo the program of apoptosis.1920212349 Caspase 3 is currently described as a common effector of apoptosis required in many models of cell death.7 Caspase 3 is characterized by the DEVDase activity it shared with other members of the caspase family. During the apoptotic process, caspase 3, the main effector of the caspase family, is activated by cleavage.37 We have previously shown that in U937, HL60 and mononuclear cells from leukemic patients, DEVDase activity is evidenced after anthracycline treatment.24 In the present work, we reinforce this observation, showing such an activation in sorted blast cells. DEVDases activation was not significant when PGP was expressed, thus highlighting a potential link with intracellular anthracycline accumulation. Using a specific antibody against activated caspase 3, FCM and multivariate analysis, we demonstrate that DNR and IDA induce cleavage and consequently the activation of caspase 3 in both lymphocytes and blast cells. Therefore, the response in normal and tumor cells is similar. Spontaneous apoptosis was lower in lymphocytes than in blast cells and the rate of caspase 3 activation was significantly higher in blast cells than in lymphocytes, confirming that anthracycline toxicity mainly affected tumor cells. Furthermore, in both blast cells and lymphocytes, caspase 3 activation was highly correlated with phosphatidyl serine exposure, indicating that anthracyclines induced apoptosis through the caspase pathway. When PGP was expressed, both IDA and DNR induced apoptosis to a lesser extent, thus providing biological argument for the poor prognosis of PGP-positive AL.

The clinical evolution of patients was compared to the biological data of our study. First, Fas-induction on anthracycline-exposed blast cells was much higher in SCR patients as compared with DTF patients. A chi-square analysis showed that a Fas-induction higher than 50% of the basal expression was found significantly more frequently for SCR patients. Moreover, anthracycline-induced apoptosis was also greater for SCR patients and an IDA-induced apoptosis higher than 20% of the blast cells was significantly more frequent for SCR patients. These observations shed light on the better biological response of blast cells in the case of SCR and may constitute original short-term prognosis factors.

Finally, our results show that anthracycline could increase susceptibility to Fas-induced apoptosis in both leukemic blast cells and normal lymphocytes. Anthracycline-induced apoptosis was accompanied by a total DEVDase activation and was characterized by caspase 3 cleavage. Susceptibility to chemotherapy was higher in leukemic cells than in lymphocytes and the deleterious role of PGP expression was biologically confirmed in vitro. Interestingly, the biological chemotherapy response of blast cells seems to be associated with patient outcome. Further studies including more patients are now underway to confirm these observations, in order to affirm these new prognostic factors in acute leukemia.


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This work was supported in part by grants from the Ligue Nationale contre le Cancer and from the Association pour la Recherche sur le Cancer.

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Correspondence to F Belloc.

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Belaud-Rotureau, MA., Durrieu, F., Labroille, G. et al. Study of apoptosis-related responses of leukemic blast cells to in vitro anthracycline treatment. Leukemia 14, 1266–1275 (2000).

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  • anthracycline
  • apoptosis
  • leukemia
  • Fas
  • caspase

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