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Mol Cell Biol. Feb 2003; 23(4): 1278–1291.
PMCID: PMC141136

Erythroid Differentiation Sensitizes K562 Leukemia Cells to TRAIL-Induced Apoptosis by Downregulation of c-FLIP


Regulation of the apoptotic threshold is of great importance in the homeostasis of both differentiating and fully developed organ systems. Triggering differentiation has been employed as a strategy to inhibit cell proliferation and accelerate apoptosis in malignant cells, in which the apoptotic threshold is often characteristically elevated. To better understand the mechanisms underlying differentiation-mediated regulation of apoptosis, we have studied death receptor responses during erythroid differentiation of K562 erythroleukemia cells, which normally are highly resistant to tumor necrosis factor (TNF) alpha-, FasL-, and TRAIL-induced apoptosis. However, upon hemin-mediated erythroid differentiation, K562 cells specifically lost their resistance to TNF-related apoptosis-inducing ligand (TRAIL), which efficiently killed the differentiating cells independently of mitochondrial apoptotic signaling. Concomitantly with the increased sensitivity, the expression of both c-FLIP splicing variants, c-FLIPL and c-FLIPS, was downregulated, resulting in an altered caspase 8 recruitment and cleavage in the death-inducing signaling complex (DISC). Stable overexpression of both c-FLIPL and c-FLIPS rescued the cells from TRAIL-mediated apoptosis with isoform-specific effects on DISC-recruited caspase 8. Our results show that c-FLIPL and c-FLIPS potently control TRAIL responses, both by distinct regulatory features, and further imply that the differentiation state of malignant cells determines their sensitivity to death receptor signals.

Programmed cell death or apoptosis is an active form of cellular destruction, which in concert with cell division controls immune responses, embryonic development, and tissue homeostasis. The most specific way to induce apoptosis is stimulation of the different types of death receptors, for example, tumor necrosis factor (TNF) receptor I (for review see reference 66), Fas/CD95 (28), or the TNF-related apoptosis-inducing ligand (TRAIL) receptors DR4 and DR5 (also known as TRAIL-R1 and -R2) (50, 51, 56). Stimulation of death receptors with their specific ligands FasL, TNF alpha (TNF-α), and TRAIL causes in responsive cells oligomerization or aggregation of the receptor and activation of the cytoplasmic caspase machinery, resulting in disruption of normal cellular and nuclear morphology followed by DNA fragmentation and finally apoptosis (for review see reference 59).

The TRAIL (67) receptors are members of the TNF receptor family and are widely expressed on the surface of many different cell types. The resistance of cells to TRAIL seems to be tightly regulated in a cell-type- and differentiation-dependent manner. TRAIL induces apoptosis in several cancer cell lines, whereas most nontransformed cells are resistant to TRAIL-mediated apoptosis (52, 67). Therefore, TRAIL has been regarded as a potential anticancer agent. Although the TRAIL receptors have been established as bona fide death receptors, with similar assembly of FADD and caspase 8 to the death-inducing signaling complex (DISC) (6), as has been shown for Fas, the mechanisms regulating the resistance against TRAIL-mediated apoptosis are not equally well established. To fully understand the physiological roles of TRAIL and to enable optimal exploitation of its anticancer properties, it is important to gain detailed knowledge of the molecular mechanisms and signaling pathways conferring TRAIL receptor sensitivity.

Apoptotic responses are regulated at several different levels, for example, at the activated receptor, along the mitochondrial pathway, and at the level of caspase activation, enabling a finely tuned regulation by antiapoptotic and proapoptotic signals and proteins. FLICE inhibitory protein (FLIP) was first found in viruses (v-FLIP) (61), and subsequently, protein homologs in vertebrate cells were identified as c-FLIP (27), also named as Casper (57), I-FLICE (25), FLAME-1 (58), CASH (18), CLARP (26), MRIT (19), and usurpin (54). Cellular FLIP exists as two alternatively spliced isoforms: c-FLIPL is homologous to caspase 8, except in lacking critical amino acids for proteolytic caspase activity, whereas c-FLIPS consists of only two death effector domains (DED) (see Fig. Fig.5A).5A). While there are various results on the role of c-FLIP in apoptotic signaling, the majority of studies have established c-FLIP as a potent suppressor of apoptotic signals especially induced by the Fas receptor (for review see reference 37). The variation in the obtained results from different cellular and apoptosis models suggests that c-FLIP is far from the only regulator of death receptor signaling and that c-FLIP-independent regulation exists. c-FLIP is recruited to the activated death receptor via FADD, thereby either preventing the recruitment of procaspase 8 to the DISC or inhibiting the proximity-induced activation of caspase 8 (38, 55). c-FLIP expression has been reported to fluctuate in a cell-type-specific manner and in response to various stimuli: transcriptionally through the NF-κB pathway (36, 47) and at the protein level via altered rates of proteasomal degradation (15, 34), which makes it a versatile inhibitor of apoptotic responses mediated by death receptors. Therefore, it is not surprising that upregulation of c-FLIP has been shown in several cancer cell lines that are resistant to death ligand-induced apoptosis. For example, high levels of c-FLIPL have been reported in melanoma cells (27) and elevated expression of c-FLIP has also been linked to the escape of tumors from immune surveillance (14, 46).

FIG. 5.
Sensitization corresponds to downregulation of c-FLIP. (A) A schematic comparison of c-FLIP isoforms and procaspase 8 (modified from reference 37). (B and C) Western blot analysis of c-FLIPL and c-FLIPS, respectively, in K562 cells treated with 30 μM ...

To investigate the mechanisms by which differentiation processes affect apoptotic responses, we studied the regulation of death receptor-mediated apoptosis during erythroid differentiation of K562 human erythroleukemia cells, which are capable of differentiating along the erythroid lineage following treatment with hemin, a ferric chloride salt of heme (2, 42). In this study, we show that K562 cells, which are normally resistant to death receptor-mediated apoptosis, were potently sensitized to TRAIL upon hemin-induced differentiation. The differentiation-mediated sensitization occurs also in other differentiating malignant cells, since erythroid differentiation of HEL (human erythroleukemia) cells and granulocytic differentiation of HL-60 (human promyelocytic leukemia) cells increased the sensitivity to TRAIL. The TRAIL-mediated apoptosis in the sensitized K562 cells occurred by direct caspase activation without any apparent involvement of the mitochondrial apoptotic signaling pathway. Analysis of components in the DISC revealed that antiapoptotic protein c-FLIP was downregulated concomitantly with the observed sensitization and that this downregulation was reflected in the composition of the TRAIL receptor DISC, leading to a differential cleavage of the major initiator caspase, caspase 8. Stable overexpression of both c-FLIP splicing variants efficiently rescued the cells from TRAIL-induced apoptosis after hemin sensitization, by interfering with caspase 8 cleavage and recruitment to the TRAIL receptor DISC, underlining the importance of c-FLIP as an efficient modulator of death receptor responses.


Cell culture and treatments.

K562, HEL, HL-60, and Jurkat (acute T-cell leukemia) cells were cultured in a humidified 5% CO2 atmosphere at 37°C in RPMI 1640 supplemented with 10% fetal calf serum, antibiotics (penicillin and streptomycin), and 2 mM l-glutamine. For K562 cells, hemin (Sigma) was added to a final concentration of 30 μM and tetradecanoyl phorbol acetate (TPA) to 20 nM. For HEL cells, hemin was added to a final concentration of 60 μM and HL-60 cells were treated with 1% dimethyl sulfoxide (DMSO). Apoptosis in K562, HEL, and Jurkat cells was induced by adding 100 ng of FLAG-tagged TRAIL (Alexis)/ml together with 2 μg of cross-linking M2 anti-FLAG antibody (Sigma)/ml. For HL-60 cells 50 ng of TRAIL/ml and 1 μg of M2/ml were used. Caspase 8 inhibitor (Caspase 8 inhibitor II; Calbiochem) was used at a final concentration of 10 μM.

SDS-polyacrylamide gel electrophoresis and Western blotting.

For Western blot analysis cells were harvested by centrifugation and washed once with phosphate-buffered saline (PBS). Cells were lysed in lysis buffer (30 mM Tris, pH 7.5, 150 mM NaCl, 1% Triton X-100, 10% glycerol, 1 mM phenylmethylsulfonyl fluoride, and Complete mini protease inhibitor cocktail [Roche]) and were centrifuged to remove cell debris. The protein concentration was determined by the Bradford method, and each lysate containing 30 to 50 μg of protein was loaded and resolved on a sodium dodecyl sulfate (SDS)-polyacrylamide gel and transferred to nitrocellulose membrane (Protran nitrocellulose; Schleicher & Schuell) by using a semidry transfer apparatus (Bio-Rad). Western blotting was performed using antibodies against poly(ADP-ribose) polymerase (PARP) (Sigma), caspase 8 (C15 caspase 8 antibody [a kind gift from Peter Krammer, German Cancer Research Center, Heidelberg, Germany]), DR4 (Santa Cruz), DR5 (Santa Cruz, Alexis), TNF-R1 (Santa Cruz), Bcl-XL (Santa Cruz), c-FLIP (Alexis, NF6 FLIP antibody; kindly provided by Peter Krammer), FADD (Transduction Laboratories), and Hsc70 (StressGen). Horseradish peroxidase-conjugated secondary antibodies were purchased from Promega, Amersham, and Southern Biotechnology. The bands were visualized using the enhanced chemiluminescence method (ECL; Amersham).

Microscopic analysis of cellular morphology.

After 16 h of hemin treatment, 5 × 105 cells were treated with TRAIL together with M2 antibody and sealed into an incubation chamber. After 3 h, cellular morphology was analyzed by viewing live cells under a Leica DMRB microscope using differential interference contrast (DIC) illumination.

Caspase 3 activity assay.

After hemin and TRAIL treatments, cells were harvested with centrifugation and washed once with PBS. Cells were lysed in lysis buffer (30 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1% Triton X-100, and 10% glycerol) and were centrifuged to remove cell debris. Caspase 3 activity was assayed from lysed samples consisting of 3 × 104 cells with the LANCE homogenous time-resolved fluorescence quench caspase 3 assay according to the manufacturer's protocol (Wallac Oy, PerkinElmer Life Sciences).

Analysis of mitochondrial membrane potential by confocal microscopy.

To measure mitochondrial membrane potential, hemin-treated and untreated cells were equilibrated with 50 nM tetramethyl rhodamine methyl ester (TMRM; Molecular Probes) in RPMI 1640 buffered with 25 mM HEPES (pH 7.2) for 1 h at 37°C in darkness. Subsequently, TRAIL and M2 antibody were added to the equilibration medium. TMRM fluorescence and transmitted light images at given time points were collected with a Leica TCS SP confocal microscope with a 63× 1.4-numerical-aperture oil immersion planapochromat objective. Red fluorescence of TMRM was imaged by using 568-nm excitation light from an argon/krypton laser, and emitted light was collected through 575 to 705 nm.

Analysis of cytochrome c immunofluorescence by confocal microscope.

For immunofluorescence analysis, K562 cells were centrifuged on glass coverslips, washed with PBS, and fixed with 3% paraformaldehyde. Subsequently, cells were permeabilized with 0.5% Triton X-100-PBS for 10 min at room temperature. After blocking with normal goat serum (GS), samples were incubated with mouse anti-cytochrome c antibody (clone 6H2.B4; BD Pharmingen) (1:150 in PBS-0.01% Triton X-100 with 1.5% GS) for 2 h in a humidified dark chamber at 37°C. After three washes with PBS-0.01% Triton-X-100, samples were incubated with Alexa 488-conjugated goat anti-mouse immunoglobulin G (1:150 in PBS-0.01% Triton X-100 with 1.5% GS; Molecular Probes) for 45 min in a dark chamber. After three washes with PBS-0.01% Triton X-100, the nuclei were counterstained with 4′,6′-diamidino-2-phenylindole (DAPI) (0.1 μg/ml) and coverslips were mounted on microscope slides in 80% glycerol in PBS. Cytochrome c release and nuclear morphology of the cells were imaged by a Leica TCS SP MP confocal microscope with a 63× 1.4-numerical-aperture oil immersion planapochromat objective. Alexa 488 fluorescence was excited by using a 488-nm excitation line from an argon/krypton laser, and the emission window was set at 492 to 560 nm. DAPI fluorescence was imaged by using multiphoton excitation at 780 nm from a Ti-Sapphire (Tsunami) laser. Emission was recorded at 400 to 490 nm.

Surface expression analysis of DR4 and DR5.

Cells (n = 0.5 × 106) were treated with hemin for 16 h and then washed two times with PBS. After washing, cells were blocked for 30 min with 1% bovine serum albumin in PBS. Cells were then incubated with DR4- or DR5-specific antibodies (5 μg/ml; Alexis) in 1% bovine serum albumin in PBS for 30 min followed by washing with PBS. Finally, cells were incubated with Alexa 488-conjugated goat anti-mouse immunoglobulin G (Molecular Probes) for 30 min. After washes, cells were analyzed on a FACScan flow cytometer. Samples without primary antibody were used as negative controls.

Biotin labeling of cell surface proteins.

After treatments, 3 × 106 cells were washed once with PBS and were incubated 15 min at room temperature with 0.5 μg of EZ-Link Sulfo-NHS-LC-Biotin (Pierce)/ml in PBS. After incubation, cells were lysed with lysis buffer supplemented with protease inhibitors and the lysate was cleared by centrifugation. Lysates were incubated for 2 h at 4°C with streptavidin-coated agarose beads (Pierce), and the beads were washed three times with lysis buffer. The beads were boiled in 3× Laemmli sample buffer, and samples were resolved by SDS-8% polyacrylamide gel electrophoresis. Western blotting was performed as described above.

RNase protection assay.

K562 cells (n = 5 × 106) were treated as indicated, and total RNA was isolated with the Trizol (Gibco-BRL) method according to the manufacturer's recommendations. The amount of various apoptosis-related transcripts and the internal controls L32 and glyceraldehyde-3-phosphate dehydrogenase was analyzed by using the hAPO-3b multiprobe template set (Pharmingen). Probe synthesis, hybridization, and RNase treatments were done with the RiboQuant Multi-Probe RNase Protection Assay System (Pharmingen) following the manufacturer's instructions. Protected RNA was resolved on a 5% urea polyacrylamide gel and quantified with a phosphorimager (Fuji).

Fluorescence-activated cell sorter analysis of cellular caspase 3 activity.

After treatments, cells were washed once with ice-cold PBS and the caspase 3 activity was analyzed with the phycoerythrin (PE)-conjugated monoclonal active caspase 3 antibody apoptosis kit 1 (BD Pharmingen) according to the manufacturer's protocol. Briefly, cells were fixed and permeabilized with Cytofix/Cytoperm solution followed by two washes with Perm/Wash buffer. PE-conjugated anti-active caspase 3 antibody was diluted in the Perm/Wash buffer and incubated with the fixed cells. After one wash with Perm/Wash buffer, cells were analyzed for PE fluorescence by flow cytometry.

Plasmid constructs, stable cell lines, and transfections.

For transfections, 5 × 106 K562 cells were centrifuged and resuspended in 0.4 ml of OptiMEM (Gibco-BRL) and 30 μg of plasmid DNA encoding FLAG-tagged c-FLIPL or c-FLIPS (a kind gift from Jürg Tschopp, Institute of Biochemistry, University of Lausanne, Lausanne, Switzerland) was added. Cells were subjected to a single electric pulse (975 μF, 200 V) in 0.4-cm gap electroporation cuvettes (BTX) using a Bio-Rad Gene Pulser electroporator, followed by dilution to 5 × 105 cells/ml in RPMI 1640 with 10% fetal calf serum and antibiotics. For making stable cell lines, neomycin-resistant cells were selected by G418 (500 μg/ml; Life Technologies, Inc.) for 2 weeks. The resistant pool was serially diluted on a 96-well plate in the presence of G418 selection. Single-cell clones were upscaled and screened for c-FLIP expression by Western blotting.

TRAIL receptor immunoprecipitation and DISC analysis.

K562 cells (4 × 107/sample) were left untreated or pretreated with 30 μM hemin for 16 h at a cell density of 4 × 105/ml. To stimulate TRAIL receptors, cells were pelleted (500 × g, 7 min) and were resuspended in 1 ml of prewarmed RPMI medium, and thereafter 1 μg of FLAG-tagged TRAIL (Alexis) and 2 μg of anti-FLAG monoclonal M2 antibody (Sigma) were added to the cell suspension. Cells were incubated in a 37°C water bath for 20 min, and the reaction was stopped by adding 10 ml of ice-cold PBS to the cell suspension. Cells were pelleted, washed with ice-cold PBS, and lysed in 1 ml of lysis buffer (20 mM Tris-HCl, pH 7.4, 150 mM NaCl, 10% glycerol, 0.2% Nonidet P-40, 0.1% sodium deoxycholate, and Complete protease inhibitor cocktail [Roche]) for 30 min on ice. The cell debris was removed by centrifugation at 15,000 × g for 15 min at 4°C. The amount of protein was determined by Bradford assay, and an equal amount of protein from each sample was precleared with 50 μl of Sepharose-CL-4B for 2 h at 4°C. One microgram of TRAIL and 1 μg of M2 antibody were added to unstimulated control lysates. Immunoprecipitation was carried out with 15 μl of protein G beads (Amersham) for 2.5 h at 4°C. Beads were washed six times in 1 ml of lysis buffer, finally resuspended in 3× Laemmli sample buffer, and boiled for 3 min. About one-third of immunoprecipitation samples and 20 to 50 μg of protein from cell lysates were analyzed by SDS-12.5 or 10% polyacrylamide gel electrophoresis. Western blotting was performed as described above.


K562 cells lose their TRAIL resistance during erythroid differentiation.

The K562 erythroleukemia cell line is normally relatively resistant to apoptosis induced by FasL (39), TNF-α (29), and TRAIL (13). Since several studies indicate that death receptor responses are strictly regulated upon erythroid differentiation (11, 69), we examined whether induction of K562 cells to erythroid differentiation would affect their sensitivity to death receptor-mediated apoptosis. Pretreatment with hemin had no effect on the sensitivity to FasL or TNF-α (data not shown), whereas TRAIL potently induced apoptosis in differentiating K562 cells, as illustrated by views of live K562 cells under DIC illumination (Fig. (Fig.1A).1A). TRAIL treatment alone did not have any apparent effect on proliferation or differentiation status of K562 cells (data not shown). Enhanced cleavage of procaspase 8 to intermediate cleaved fragment (p43/41) and active p18 caspase subunit was observed after a 2-h TRAIL treatment in differentiating K562 cells, whereas only a modest cleavage of procaspase 8 to the intermediate fragment was detectable in nondifferentiating cells (Fig. (Fig.1B).1B). Caspase 3, in turn, was only activated in differentiating TRAIL-stimulated cells with kinetics similar to those of caspase 8 when the enzymatic activity of caspase 3 was analyzed by measuring DEVDase activity, i.e., the proteolytic activity toward a synthetic substrate containing the consensus sequence DEVD (Fig. (Fig.1C),1C), or by detecting the cleavage of its physiological substrate PARP to the 85-kDa fragment (Fig. (Fig.1D).1D). It is notable that hemin itself did not induce any caspase 8 or caspase 3 activity.

FIG. 1.
Hemin-mediated differentiation sensitizes K562 cells to TRAIL-induced apoptosis. (A) K562 cells pretreated for 16 h with or without hemin were treated or left untreated for 3 h with 100 ng of TRAIL/ml together with 2 μg of M2 cross-linking antibody/ml. ...

Sensitization to TRAIL is not unique for erythroid differentiation of K562 cells but occurs also upon erythroid differentiation of HEL cells and granulocytic differentiation of HL-60 cells.

To study whether the observed sensitization was specific to erythroid differentiation or differentiation of malignant cells in general, we tested the TRAIL sensitivity of two additional cell lines with well-established ability to differentiate in response to chemical stimuli (9, 44). Nondifferentiating HEL cells displayed modest sensitivity to TRAIL. But when the cells were induced to differentiate along erythroid lineage with hemin (24 h), their sensitivity towards TRAIL was doubled as quantified with fluorescence-activated cell sorter analysis of caspase 3-positive cells (Fig. (Fig.2A).2A). Another tested differentiation process was DMSO-induced granulocytic differentiation of HL-60 cells, which has been earlier shown to moderately sensitize the cells to Fas- and TNF-mediated apoptosis during early differentiation (63). As shown in Fig. Fig.2B,2B, 24 h of DMSO-induced granulocytic differentiation led to a modest sensitization of HL-60 cells to TRAIL-induced apoptosis. Since K562 cells are also capable of differentiating along the megakaryocytic lineage in response to the phorbol ester TPA, we tested whether differentiation in general is sufficient to render K562 cells sensitive to TRAIL. However, megakaryocytic K562 cells displayed no increased sensitivity towards TRAIL after a 24-h TPA treatment, whereas hemin strongly sensitized the cells (Fig. (Fig.2C).2C). In conclusion, it appears that induction of differentiation in several malignant cell lines, along at least two distinct lineages, markedly increases the sensitivity towards TRAIL-induced apoptosis. The sensitization is, however, not a universal differentiation-related phenomenon, as demonstrated by the lack of sensitization during megakaryocytic differentiation of K562 cells.

FIG. 2.
Erythroid differentiation of HEL cells and granulocytic differentiation of HL-60 cells, but not megakaryocytic differentiation of K562 cells, sensitizes to TRAIL. (A) Analysis of apoptosis in HEL cells after combined hemin (60 μM, 24 h) and TRAIL ...

K562 cells retain their mitochondrial membrane potential while undergoing TRAIL-induced apoptosis.

As do many malignant cells, K562 cells display strong resistance towards chemotherapeutic agents (45). This is probably due to efficient protection against the mitochondrial apoptotic pathway, mediated by mutations in the Apaf-1 protein (30) as well as high expression of Bcl-XL, maintained by constitutive activity of the Bcr-Abl tyrosine kinase (24). Since TRAIL induced a strikingly rapid apoptosis in differentiating K562 cells (Fig. (Fig.1),1), we were prompted to study whether it could be mediated by removal of molecular protection mechanisms acting at the mitochondrial apoptotic signaling pathway. Although a prolonged treatment of K562 cells with hemin has been shown to lead to downregulation of Bcl-XL (5), we were not able to detect any downregulation in Bcl-XL levels after 16 and 24 h of hemin treatment (Fig. (Fig.3A),3A), when a strong sensitization to TRAIL was observed. To further examine the role of the mitochondrial apoptotic pathway in K562 cells undergoing TRAIL-induced apoptosis, mitochondrial membrane potential was analyzed with confocal microscopy. For this purpose, live cells undergoing TRAIL-induced apoptosis were loaded with the red fluorescent TMRM dye that accumulates to polarized mitochondria. We used Jurkat cells as a positive control, where TRAIL induced complete mitochondrial depolarization, as indicated by a disappearance of mitochondrial TMRM fluorescence in cells displaying an apoptotic morphology (Fig. (Fig.3B).3B). Surprisingly, there was very little or no loss in TMRM fluorescence even in late apoptotic K562 cells, with numerous characteristic membrane blebs, indicating that mitochondria remained polarized during TRAIL-induced apoptosis.

FIG. 3.
Hemin-treated K562 cells retain their mitochondrial membrane potential during TRAIL-induced apoptosis. (A) Western blot analysis of Bcl-XL in K562 cells treated with hemin for indicated time periods. Hsc70 was blotted to show equal loading. (B) K562 cells ...

To exclude the possibility of cytochrome c release without loss of mitochondrial membrane potential, we analyzed the subcellular localization of cytochrome c in K562 cells undergoing TRAIL-induced apoptosis. Confocal microscopy showed that TRAIL alone does not cause release of cytochrome c from mitochondria, since cytochrome c was located in distinct structures that correspond to mitochondria (Fig. (Fig.3C).3C). Interestingly, a closer examination of samples treated with hemin and TRAIL revealed cells in which most of the cytochrome c-specific fluorescence was structured, but some distinct parts showed a diffuse fluorescent pattern, likely to represent an apoptotic bleb (Fig. (Fig.3C;3C; Hemin+TRAIL, Max. projection). In these cells the nucleus was fragmented, indicating the presence of prominent effector caspase activities. Taken together, the late and partial loss of mitochondrial integrity upon TRAIL-induced apoptosis suggests that differentiating K562 cells retain their mitochondrial protection and undergo apoptosis by direct caspase activation, which is characteristic for cells classified as type I cells, i.e., without requirement for mitochondrial signal amplification to execute apoptosis. This result is in agreement with the high levels of Bcl-XL in the sensitive differentiating K562 cells (Fig. (Fig.3A3A).

Differentiation-mediated sensitization is not accompanied by upregulation of death receptors DR4 and DR5.

Since many chemotherapeutic agents have been shown to mediate TRAIL sensitization via upregulation of the surface expression of TRAIL receptors, we examined whether erythroid differentiation leads to an elevated surface expression of DR4 and DR5. For this purpose, we immunolabeled DR4 and DR5 receptors with monoclonal antibodies, together with fluorescent secondary antibodies, and analyzed the surface expression of the DRs by flow cytometry. No major differences were found in expression of either DR4 or DR5 after 16 h of hemin treatment (Fig. (Fig.4A).4A). Although it has been earlier shown that K562 cells do not express decoy receptors for TRAIL (13), we wanted to confirm that downregulation of TRAIL decoy receptors does not play a role in this sensitization. TRAIL-binding receptors were saturated on the cell surface with FLAG-tagged TRAIL, after which they were immunolabeled with FLAG-specific antibody and fluorescent secondary antibody. The flow cytometry analysis did not show any changes in TRAIL-binding ability (Fig. (Fig.4A),4A), which together with the data from DR4 and DR5 expression analysis indicates that overall expression of TRAIL receptors was not changed during hemin treatment. The flow cytometry analysis was complicated by the increase in autofluorescence after hemin treatment, which can be seen from the secondary antibody controls (Fig. (Fig.4A).4A). This was most probably due to the increased synthesis of fluorescent porphyrins during differentiation (43). To overcome the background fluorescence in hemin-treated samples, the surface proteins of unpermeabilized cells were labeled with biotin and purified with streptavidin-coated agarose. Based on Western blot analysis of purified cell surface proteins, there was no increase in the surface expression of DR4 or DR5 during hemin-induced differentiation (Fig. (Fig.4B),4B), excluding the possibility that elevated surface expression of TRAIL receptors might contribute to the observed sensitization.

FIG. 4.
Surface expression of DR4 and DR5 is not increased during erythroid differentiation of K562 cells. (A) For surface analysis of TRAIL receptors, K562 cells were treated for 16 h with 30 μM hemin or left untreated. After treatments the cells were ...

Downregulation of c-FLIP corresponds to the differentiation-mediated sensitization of K562 cells.

The late and partial appearance of mitochondrial changes upon apoptosis led us to study events associated with DISC signaling. One of the antiapoptotic proteins acting at the level of activated death receptors is c-FLIP, which is a DED-containing antiapoptotic protein that is homologous to caspase 8, except for its lack of amino acids critical for the protease activity (Fig. (Fig.5A).5A). c-FLIP exists as two alternatively spliced isoforms, which are thought to act by inhibiting recruitment and proximity-induced activation of caspase 8 in the DISC. To examine the c-FLIP protein levels, K562 cells were treated with hemin for different time periods and the cell lysates were analyzed by Western blotting with an anti-c-FLIP antibody. As shown in Fig. 5B and 5C, both c-FLIPL and c-FLIPS were potently downregulated in response to hemin-induced differentiation. Interestingly, the kinetics of c-FLIPS downregulation were different from those of c-FLIPL, since c-FLIPS remained at the control level until 10 h of hemin treatment when it rapidly disappeared, whereas downregulation of c-FLIPL was more gradual. To study the onset of apoptotic sensitivity upon erythroid differentiation, K562 cells were pretreated with hemin for indicated time periods, followed by a 3-h TRAIL treatment to induce apoptosis. The TRAIL resistance gradually decreased after 8 h of hemin treatment (Fig. (Fig.5D),5D), which corresponds to the overall downregulation of c-FLIPL and c-FLIPS (Fig. (Fig.5B5B and C).

c-FLIP expression has been proposed to be regulated by mechanisms involving changes in transcription and mRNA stability (for review see reference 37). Therefore, we examined the c-FLIP mRNA levels upon hemin treatment using RNase protection assay with a set of probes specific to apoptosis-related transcripts. As shown in Fig. Fig.6,6, during a 16-h hemin treatment, c-FLIP mRNA was downregulated to 60% of the control levels when normalized against the L32 housekeeping gene. Although clearly reproducible, the downregulation was relatively modest in comparison to the protein levels, suggesting that additional downregulation is also likely to occur at the protein level. Surprisingly, the mRNA levels of caspase 8 were upregulated 1.9-fold, although no increase in the protein levels could be detected.

FIG. 6.
c-FLIP mRNA is modestly downregulated in hemin-treated K562 cells. RNase protection assay analysis of various apoptosis-related genes, including c-FLIP, in cells left untreated (C) or treated with hemin for 16 h (H16). Each signal was normalized by using ...

Downregulation of c-FLIP affects procaspase 8 cleavage in the DISC.

To study whether the observed downregulation of c-FLIP was reflected at the level of the DISC, we performed TRAIL receptor DISC immunoprecipitation experiments (Fig. (Fig.7A).7A). The successful immunoprecipitation was controlled by the presence of DR5 in the immunoprecipitated samples. FADD and caspase 8 were recruited to the TRAIL receptor DISC, and the activation of procaspase 8 in the immunoprecipitated DISC was demonstrated by the presence of intermediate cleaved p43/41 fragments. Interestingly, the intermediate p43/41 fragments of caspase 8 were detected in the samples treated with TRAIL alone (displaying no apoptosis), whereas, in differentiating K562 cells much less, if any, p43/41 fragments were present in the DISC. The absence of intermediate fragments of caspase 8 in DISC devoid of c-FLIP most probably reflects rapid processing and diffusion of activated caspase 8 fragments followed by replacement with procaspase 8. In addition, TRAIL-induced procaspase 8 recruitment was consistently ~1.5 fold stronger in hemin-treated cells. As expected, c-FLIPL and c-FLIPS were readily recruited to the DISC in TRAIL-stimulated cells, whereas, in hemin-treated cells, no c-FLIP was found in the DISC. It is also worth noticing that all c-FLIPL recruited to the DISC was cleaved to the p43 fragment. The immunoprecipitation experiments also supported the assumption that K562 cells are so-called type I cells, in which a large amount of DISC is formed upon TRAIL receptor stimulation.

FIG. 7.
Downregulation of c-FLIP alters the composition of TRAIL receptor (TRAIL-R) DISC. (A) For TRAIL receptor DISC analysis, K562 cells treated with hemin (16 h) or left untreated were stimulated with TRAIL and M2 for 20 min and then lysed. For unstimulated ...

To further specify that caspase 8 was the principal initiator caspase mediating the TRAIL-induced apoptotic signaling in the K562 cell model, we used 10 μM caspase 8-specific inhibitor Z-IETD-fmk and measured the number of apoptotic cells by flow cytometry after labeling with caspase 3-specific PE-conjugated antibody (Fig. (Fig.7B).7B). Inhibition of caspase 8 activity efficiently protected hemin-treated cells from TRAIL-induced apoptosis, reducing the amount of apoptosis from 36% to less than 10%. This result indicates that caspase 8 acts as the major initiator caspase in K562 cells upon TRAIL receptor stimulation.

Stable overexpression of c-FLIP prevents hemin-mediated sensitization.

In order to study the effect of c-FLIP on TRAIL-induced apoptotic signaling, we generated stable cell lines overexpressing different amounts of FLAG-tagged c-FLIPL (1F6, 2E10, and 2G11) and c-FLIPS (1E5 and 2E11) (Fig. (Fig.8A).8A). To determine, whether expression of ectopic c-FLIP could rescue K562 cells from hemin-mediated sensitization to TRAIL, c-FLIPL and c-FLIPS cell lines were subjected to hemin-mediated differentiation followed by TRAIL receptor stimulation. In addition to parental K562 cells, a neomycin-resistant pool containing an empty plasmid was used as a mock control. As shown in Fig. Fig.8B,8B, all c-FLIPL-overexpressing cell lines were efficiently protected from apoptotic TRAIL signaling, showing only 4 to 8% of apoptosis after hemin-induced sensitization, whereas the parental K562 and the mock-transfected cells displayed 25 to 30% apoptosis. Thus, the efficiency of the c-FLIPL protection was comparable with the effect of caspase 8 inhibitor Z-IETD-fmk. Interestingly, 1F6 displaying the lowest expression level was the most resistant cell line, suggesting that the apoptotic machinery in K562 cells becomes saturated with a relatively modest amount of c-FLIPL and that an excess may have a proapoptotic effect, as has been shown by others using different cell systems (27, 57). Similar experiments with the cell lines stably overexpressing c-FLIPS revealed that the antiapoptotic capacity of ectopic c-FLIPS was even stronger than that of c-FLIPL. No trace of elevated apoptosis could be detected in c-FLIPS-overexpressing cells after the combined treatments with hemin and TRAIL (Fig. (Fig.8C8C).

FIG. 8.
Stable overexpression of both c-FLIP isoforms prevents differentiation-mediated sensitization. (A) Western blot analysis of c-FLIP from parental K562 cells, a mock-transfected cell pool, and cell lines stably overexpressing c-FLIPL (1F6, 2E10, and 2G11) ...

c-FLIP isoforms differentially regulate procaspase 8 cleavage in the TRAIL receptor DISC.

Finally, we examined the mechanism by which the two different splicing variants of c-FLIP regulate TRAIL receptor DISC dynamics, since earlier studies on Fas DISC have shown that c-FLIPL and c-FLIPS are able to differentially regulate caspase 8 cleavage in the DISC (38, 55). In agreement with the earlier studies, our DISC immunoprecipitation experiments employing c-FLIPL and c-FLIPS-overexpressing cells revealed that c-FLIPL was indeed capable of catalyzing the cleavage of procaspase 8 to the intermediate form (Fig. (Fig.9),9), although still being able to prevent apoptotic signaling (Fig. (Fig.8B).8B). Similar to the endogenous c-FLIPL, the ectopic c-FLIPL recruited to the DISC was also completely cleaved to the p43 form (Fig. (Fig.9).9). In contrast to c-FLIPL, c-FLIPS already inhibited the first cleavage efficiently, since only trace amounts of the p43/41 form of caspase 8 were present in the DISC of c-FLIPS-overexpressing cells, demonstrating that the regulatory mechanisms of c-FLIP isoforms in the TRAIL receptor DISC correspond to those reported earlier for Fas (38, 55). Interestingly, the level of overexpressed c-FLIP, especially c-FLIPL, was markedly downregulated upon stimulation with hemin but remained high enough to maintain protection. The downregulation of ectopic c-FLIPL further supports the hypothesis that c-FLIPL is downregulated at the protein level, since we did not observe c-FLIP mRNA downregulation in the c-FLIPL-overexpressing cells (data not shown).

FIG. 9.
DISC analysis of cell lines overexpressing c-FLIP isoforms. G, sample devoid of TRAIL and M2; C, control; H, hemin; T, TRAIL; HT, hemin and TRAIL. Parental K562 cells and lines stably overexpressing c-FLIPL (1F6) or c-FLIPS (1E5) were treated with 30 ...


Regulation of death receptor responses during erythroid differentiation has recently been shown to play an important role in adjusting the rate of erythropoiesis (11, 69). Apart from its significance in differentiation-related processes, this observation may also have ramifications in the regulation of growth in malignant cells, as induction of apoptosis by activation of a differentiation process has been considered a promising strategy in cancer therapy (1; for review see reference 41). Our results imply that a differentiation process has potential to sensitize malignant cells to TRAIL-induced apoptosis. This sensitization is, however, cell and differentiation lineage specific rather than a universal differentiation-related phenomenon. We focused on studying the regulation of death receptor signaling during the erythroid differentiation of K562 leukemia cells. These cells displayed a marked sensitization to TRAIL-mediated apoptosis when induced to differentiate along the erythroid lineage with hemin. Concomitantly with the observed sensitization, expression of the antiapoptotic protein c-FLIP was downregulated, which affected cleavage of the major initiator caspase, caspase 8, in the DISC of the TRAIL receptor. Stable overexpression of both c-FLIP splicing variants efficiently rescued the cells from TRAIL-mediated apoptosis after hemin sensitization by altering caspase 8 processing and its recruitment to the TRAIL receptor DISC.

Several studies have shown that TRAIL is especially efficient in killing malignant cells but not normal cells. The use of TRAIL as an anticancer agent has been studied both in vivo and in vitro (4, 31, 40, 65). Unlike chemotherapy and irradiation, TRAIL has been reported to be capable of inducing apoptosis in several cell lines independently of the mitochondrial pathway and is, therefore, especially promising for elimination of cells that express large amounts of Bcl-2 or Bcl-XL (64). Although previous studies have shown that prolonged erythroid differentiation in K562 cells leads to downregulation of Bcl-XL (5, 60), our results demonstrate that the levels of Bcl-XL remained high within the time frame during which a strong sensitization occurred. In addition, we observed that TRAIL induced no or only a minor loss in the mitochondrial membrane potential and only a partial release of cytochrome c in late apoptotic cells. Furthermore, Bcl-2 overexpression did not protect differentiating K562 cells from TRAIL-induced apoptosis (data not shown). Taken together, these results suggest that the TRAIL sensitivity in K562 cells is mediated by apoptotic activation mechanisms independent of mitochondria.

In addition to the mitochondrial apoptotic signaling pathway, death receptor responses can be regulated at the level of the death receptor complex. It seems that the latter mode of regulation can be adjusted specifically, without affecting the overall sensitivity of the cell to other types of apoptotic stimuli. Several recent studies underline the importance of c-FLIP as a specific gatekeeper, adjusting death receptor responses at the level of DISC (for review see reference 37). For example, c-FLIP−/− embryonic fibroblasts have been shown to be highly sensitive to FasL- or TNF-α-induced apoptosis, showing rapid induction of caspase 8 and caspase 3 activities in response to death receptor stimulation (68). In our study, hemin-mediated differentiation induced a strong overall downregulation of c-FLIP, concomitantly with increasing TRAIL sensitivity. Based on these results, together with those reported by others, c-FLIP appears in death receptor-insensitive cells to be efficiently recruited to the activated receptor, thereby affecting the dynamics of procaspase 8 cleavage.

The multifaceted regulatory role of c-FLIP emerges from the presence of two c-FLIP isoforms. As shown for Fas (38, 55) and in this study for TRAIL receptors c-FLIPL and c-FLIPS regulate distinct steps of procaspase 8 cleavage, since c-FLIPL promotes and c-FLIPS inhibits the first cleavage of caspase 8. This has been further addressed by two recent studies showing that heterodimerization of caspase 8 with c-FLIPL leads to increased proteolytic activity and efficient binding to synthetic substrates (7, 48). While the biochemical basis of the proteolytic activity of the caspase 8-FLIPL heterodimer is only now starting to emerge, the antiapoptotic mechanism of c-FLIPL remains enigmatic. One potential mechanism for c-FLIPL suggested by Micheau and coworkers (48) is targeting of the proteolytic activity of caspase 8 to execute nonapoptotic functions. Another open question is the physiological significance of the presence of two c-FLIP isoforms with differential activities. This diversity is likely to yield both versatility and efficacy to the regulation of death receptor responses and caspase 8 cleavage. In this respect, it has been suggested that the c-FLIP isoforms have a dual role in death receptor signaling: (i) acting as caspase 8 inhibitors and (ii) conferring the nonapoptotic signaling with which death receptors connect. Regarding the inhibitory role, two isoforms give the advantage of a broader range of inhibition, as there is an actively regulated stoichiometric balance between c-FLIPL and c-FLIPS (3, 35). Therefore, the obtained effect extends from that of a single inhibitor, when only one isoform responds to a stimulus, to the synergistic effect between the two c-FLIP isoforms. The synergy has been demonstrated by results showing that the antiapoptotic capacity of coexpressed c-FLIPL and c-FLIPS is more potent than that of a single isoform alone (38). In fact, the results in our study support the assumption of isoform-specific regulation of the expression, as the kinetics of c-FLIPL and c-FLIPS downregulation were different.

The differential activity of the c-FLIP isoforms could also relate to the protective signals mediated by death receptors, independently of their effects on caspase 8. By its ability to recruit important signaling activators to the DISC (e.g., Raf-1, TRAF1, TRAF2, and RIP), c-FLIP has been shown to be involved in death receptor-mediated activation of antiapoptotic signaling pathways, such as NF-κB and mitogen-activated protein kinase (MAPK)/ERK (32). Among the caspase-independent signals activated by death receptors, the MAPK/ERK has especially been demonstrated to have a dominant-inhibiting effect on both Fas (21, 22, 23) signaling and TRAIL (62) signaling, both of which are primarily unaffected by NF-κB protection. Whether downregulation of c-FLIP-mediated activation of MAPK/ERK signaling (and possibly NF-κB) contributes to the regulation of TRAIL sensitivity in the K562 model system remains to be addressed in forthcoming studies.

Great expectations await the employment of TRAIL as an anticancer treatment, since TRAIL has been shown to have potent antitumor effects when administered in vivo to mice and nonhuman primates (4, 65). The recent data on the elevated prevalence of certain types of tumors in TRAIL-deficient mice have further corroborated the importance of TRAIL in the elimination of tumor target cells (10). In the case of normal and malignant hematopoietic cells, TRAIL is emerging as a possible regulator, but a comprehensive understanding is still elusive. For example, immature erythroid cells have been shown to express several death receptors, DR4 and DR5 among others, and the sensitivity of the differentiating erythroblasts to the death receptor stimuli, especially FasL, is carefully controlled during the differentiation process (11, 12). This control is assumed to contribute to negative regulation of erythropoiesis, maintaining the homeostasis of red blood cells. Similar to FasL, TRAIL has been shown to act as a negative regulator of erythropoiesis in adult peripheral blood CD34+ hematopoietic progenitor cells (69). Since TRAIL was shown to selectively kill erythropoietic cells at a certain stage of differentiation, it is plausible that death receptor responses in hematopoietic cells are regulated in a lineage- and differentiation stage-specific manner. However, although TRAIL is able to kill normal differentiating erythroblasts, other reports indicate that it is more potent in selectively killing malignant hematopoietic cells than in killing their normal counterparts (53). For example, in the case of myelodysplastic syndrome, TRAIL preferably targets malignant progenitor cells (70).

This study demonstrates that triggering differentiation in malignant cells modulates the cellular machinery that determines how death receptor signals are routed in a cell and differentiation lineage-specific manner. Therefore, these results support the concept of inducing cell differentiation as a form of anticancer therapy, aiming at maturation of the malignant cell clone and thereby lowering the threshold to apoptosis (for review see reference 41). Although the cytotoxic activity of many conventional chemotherapeutic drugs as well as radiation therapy is often enhanced when combined with TRAIL treatment (8, 16, 17, 33), only a few examples of exploiting TRAIL in combination with differentiating agents are available (20, 49). In the case of normal hematopoietic cells, differentiation leads to increased sensitivity to death receptor stimuli. It is tempting to speculate that combining TRAIL treatments with induction of differentiation could have potential to enhance elimination of malignant cells. However, the full ramifications of the interplay between TRAIL receptor signaling and c-FLIP expression in normal and malignant cells remain to be elucidated in future studies.


The first two authors contributed equally to this work.

We are grateful to Peter Krammer for caspase 8 and c-FLIP antibodies and Jürg Tschopp for c-FLIP constructs. We also thank the members of our laboratories for technical help and constructive criticism during the course of this study and Mika Korkeamäki for help with flow cytometry.

Financial support was obtained from the Academy of Finland, the Sigrid Juselius Foundation, and the Finnish Cancer Organizations. V.H. and M.P. were supported by the Turku Graduate School of Biomedical Sciences.


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