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,
Victor J. Yuste,1,
Gaël Roué,1,
Sandrine Barbier,1
Patricia Sancho,1
Clémence Virely,1
Manuel Rubio,2
Sylvie Baudet,3
Josep E. Esquerda,4
Hélène Merle-Béral,3
Marika Sarfati,2 and
Santos A. Susin1*
Apoptose et Système Immunitaire, CNRS-URA 1961, Institut Pasteur, 25 rue du Dr. Roux. 75015 Paris, France,1 Centre de Recherche du CHUM, Hôpital Notre-Dame, Laboratoire d'Immunorégulation, 1560 Sherbrooke St. East, Montréal, QC H2L 4M1, Canada,2 Service d'Hématologie Biologique, Groupe Hospitalier Pitie-Salpêtrière, Paris, France,3 Unitat de Neurobiologia Cellular, Departament de Ciencies Mediques Basiques, Facultat de Medicina, Lleida, Spain4
Received 13 November 2006/ Returned for modification 14 February 2007/ Accepted 24 July 2007
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ABSTRACT |
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 Top ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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CD47 (integrin-associated protein) is a widely expressed member of the immunoglobulin (Ig) superfamily, which functions both as a receptor for thrombospondin (TSP) and as a ligand for the transmembrane signal regulatory proteins SIRP-
and -
. These molecules regulate various biological phenomena in the immune system, including platelet activation, leukocyte migration, and macrophage multinucleation (9). Importantly, they are involved in the negative regulation of the inflammatory response both in vitro and in vivo. For instance, CD47/TSP interaction negatively regulates antigen-presenting-cell and T-cell function in human cells (45). Moreover, TSP null mice display persistent inflammation in several organs (15), and CD47 null mice have impaired responses to bacterial pathogens (47) and defects in dendritic cell (DC) migration (30, 83). Most relevant to the present work, CD47 ligation, by TSP or immobilized CD47 monoclonal antibody (MAb), induces a form of caspase-independent cell death, which seems to be different from classical type I PCD (43, 44, 50, 53, 54, 58, 69).
Aberrant regulation of cell growth has traditionally been viewed as the major mechanism for tumor formation. However, it is clear that cellular changes leading to inhibition of apoptosis or PCD play an essential role in tumor development (90). The elucidation of the apoptotic pathways is thus an important area of study that may provide insight into the causes of drug resistance and facilitate the development of novel anticancer therapies. B-cell chronic lymphocytic leukemia (CLL) is the most common hematological malignancy in Western countries. Characterized by a progressive expansion of apparently quiescent B cells, CLL generally follows an indolent course. Despite the development of new chemotherapeutic agents that utilize the caspase-dependent pathway to provoke apoptosis, CLL is not considered curable (42). Future goals in CLL research are the identification of new factors sustaining the life span of the malignant B cells and the subsequent development of therapeutic agents that interfere with these molecules to provoke cell death. For this reason, the study of the molecular basis of alternative PCD pathways can provide new means of improving the current therapeutic strategies employed in the treatment of CLL.
The aim of the present work is to investigate the morphological, biochemical, and molecular mechanisms characterizing CD47-mediated PCD in normal and leukemic cells. Our approach shows that CD47 ligation induces a caspase-independent type III PCD process characterized by chymotrypsin-like serine protease activation, striking mitochondrial inner membrane alterations, loss of mitochondrial transmembrane potential (
m), production of reactive oxygen species (ROS), and outer leaflet exposure of phosphatidylserine (PS) in the plasma membrane. Importantly, our results lead us to identify a key mediator of type III PCD, dynamin-related protein 1 (Drp1) (86).
The unraveling of the mechanisms regulating CD47-mediated caspase-independent type III PCD should facilitate the understanding of alternate cell death pathways that take part in the control of immune cell homeostasis. In addition, induction of CD47-mediated caspase-independent PCD in CLL cells may be the basis for the development of novel anticancer therapies.
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MATERIALS AND METHODS |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Cell death induction and inhibition.
To induce CD47-mediated cell death, cells were cultured at different times with soluble TSP (20 µg/ml; Calbiochem) or on precoated plates with CD47 MAb (5 µg/ml; clone B6H12). Alternatively, cells were treated for 20 h with hydrocortisone (HC) (0.5 mM) or brefeldin A-cycloheximide (5 µg/ml and 10 µg/ml, respectively); for 16 h with etoposide (5 µM), thapsigargin (10 µM), hydroxychloroquine (10 µM), dexamethasone (DEX) (1 µM), or Taxol (5 µM); for 6 h with staurosporine (STP) (1 µM); or for 3 h with H2O2 (10 mM). For protease inhibition assays, Q-VD.OPh (QVD) (10 µM), z-VAD.fmk, z-DEVD.fmk, z-VDVAD.fmk, z-VEID.fmk, z-LEHD.fmk, or z-IETD.fmk (50 µM); tosylsulfonyl phenylalanyl chloromethyl ketone (TPCK) (1 to 20 µM); N-p-tosyl-L-lysine chloromethyl ketone (TLCK) (20 µM); MG101 (50 µM); N-acetyl-Leu-Leu-methional (50 µM); z-FA.fmk (100 µM); leupeptin (100 µM); or the proteasome inhibitors MG132, lactacystin, and NLVS (4-hydroxy-5-iodo-3-nitrophenylacetyl-Leu-Leu-leucinal-vinyl sulfone) (50 µM) (Merck Biosciences) were added 30 min before induction of cell death. Intracellular chymotrypsin-like, trypsin-like, and cathepsin activities were measured using Suc-Leu-Leu-Val-Tyr-7-amino-4-methylcoumarin (AMC), Boc-Leu-Arg-Arg-AMC, and Z-Arg-Arg-AMC (100 µM) (Calbiochem), respectively. Actinomycin D was used at 10 µM, cycloheximide at 100 µM, and bafilomycin A at 500 nM.
Flow cytometry.
We used 40 nM DiOC6(3) for m quantification, 2 µM hydroethidine (Invitrogen) for the measurement of ROS generation, 100 nM LysoTracker Red (Invitrogen) for the quantification of lysosomal stability, annexin V-allophycocyanin (APC) (BD Biosciences) for the assessment of PS exposure, and propidium iodide for cell viability analysis. Chymotrypsin-like serine protease cytofluorometric detection was performed with a SerPase kit from Imgenex. Determination of Bax and Bak activation was performed as described previously (5), with MAbs designed to recognize the active form of Bax (MAb 6A7; BD Biosciences) or Bak (MAb Ab-1, Calbiochem), respectively. Data analysis was carried out in a FACScalibur (BD Biosciences) on the total cell population (10,000 cells).
Determination of ATP content. Cells treated as indicated were lysed, and the total ATP content was assessed with a luciferin-luciferase kit from Sigma. Luminescence was measured in a Berthold LB96V MicroLumat Plus. The ATP content is expressed relative to cell protein in arbitrary units.
DNA electrophoresis. Oligonucleosomal DNA fragmentation was detected by agarose gel electrophoresis as described elsewhere (65).
Caspase activity. Cells treated as indicated were lysed in caspase assay buffer containing 40 mM HEPES-NaOH (pH 7.2), 300 mM NaCl, 20 mM dithiothreitol, 10 mM EDTA, 2% Nonidet P-40, 20% sucrose, and 100 µM of Ac-DEVD-AFC. Caspase activity was read in a Fluoroskan Ascent fluorimeter (Thermo Labsystems).
Quantitative real-time reverse transcription-PCR. Total RNA from control or CLL cells was extracted with Trizol reagent (Invitrogen) according to standard procedures. Samples were examined in an ABI Prism 7000 sequence detector system with TaqMan Assays-on-Demand Gene Expression Products (Applied Biosystems). Data were analyzed using the comparative threshold cycle method according to the manufacturer's protocol. The amount of mRNA measured in CLL cells was normalized according to an endogenous reference (human 18S rRNA housekeeping gene) and relative to a calibrator (B cells from control donors).
Cell transfection and RNA interference assays. Jurkat cells were stably transfected either with pcDNA3.1 control vector (Jk-Neo) or with human Bcl-2, human Bcl-XL (Jk-Bcl-2 and Jk-Bcl-XL; inserts provided by J.L. Fernández-Luna, University Hospital of Santander, Spain), human Bcl-2 targeted to the ER (Jk-Bcl-2-ER; insert supplied by C. W. Distelhorst, University Hospital of Cleveland), or human Mcl-1 (Jk-Mcl-1; cDNA provided by I. Marzo, University of Zaragoza, Spain). For transient overexpression, Jurkat cells were transfected in a Nucleofector system (program S-18, kit V; Amaxa) with human Drp-1 (Jk-Drp1) and human Drp1 mutated in the GTPase domain (Jk-Drp1K38A and Jk-Drp1K679A inserts supplied by A.M. Van der Bliek, David Geffen School of Medicine at UCLA, and C. Blackstone, NINDS-NIH). For downregulation assays, Jurkat cells were similarly transfected with small interfering RNA (siRNA) double-stranded oligonucleotides designed against human Bax (5'-GGTGCCGGAACTGATCAGA-3'), Bak (5'-CCGACGCTATGACTCAGAG-3'), Bim (5'-TTACCAAGCAGCCGAAGAC-3'), Drp1 (Drp1a, 5'-GGTGCCTGTAGGTGATCAA-3'; Drp1b, 5'-TCCGTGATGAGTATGCTTT-3' [41]; Drp1c, 5'-CAGTATCAGTCTCTTCTAA-3') or hFis1 (hFis1a, 5'-GCGGACAAGGTACAATGAT-3'; hFis1b, 5'-AGGCATCGTGCTGCTCGAG-3' [76]). As a control, we used an irrelevant oligonucleotide (5'-GCGATAAGTCGTGTCTTAC-3') or an siRNA oligonucleotide against lamin A (5'-CTGGACTTCCAGAAGAACA-3'). At 24 h after transfection, live cells were selected in a standard Ficoll gradient before cell death induction.
Protein extractions and immunoblotting. Mitochondrial and cytosolic fractions were obtained with the help of a kit from Pierce. Cell fractions and whole protein extracts from B lymphocytes or Jurkat cells were lysed in 20 mM Tris-HCl (pH 7.4), 150 mM NaCl, 1% Triton X-100, and 1 mM EDTA. Protein content was determined with the Bio-Rad DC kit, and 15 to 80 µg of protein was loaded on a sodium dodecyl sulfate-polyacrylamide gel. After blotting, polyvinylidene difluoride filters were probed with anti-human caspase 9 (Cell Signaling), anti-activated caspase 3 (BD Biosciences), anti-Bcl-2 (BD Biosciences), anti-BclXL (BD Biosciences), anti-Mcl-1 (BD Biosciences), anti-Bax (BD Biosciences), anti-Bak (BD Biosciences), anti-Bim (BD Biosciences), anti-AIF, anti-cytochrome c, anti-Smac/DIABLO (ProScience), anti-EndoG or anti-Omi/HtrA2 (Alexis), anti-Cox IV (Invitrogen), anti-ß-tubulin or anti-DRP1/DLP1 (BD Biosciences), or anti-hFis1 (Alexis) or with antibodies against the mitochondrial respiratory chain (MRC) complex I subunits p39 (clone 20C11; Invitrogen) and p30 (clone 3F9, Invitrogen). All were detected with anti-mouse or anti-rabbit IgG-horseradish peroxidase conjugated according to standard procedures.
Recombinant proteins.
N-terminal His-tagged Drp1, Drp1K38A, Drp1K679A, and Drp1(1-335) human recombinant proteins were produced from a Novagen pET28b expression vector and purified from Escherichia coli strain BL21 on a nickel-nitrilotriacetic acid affinity matrix column. The retaining extract, which contains the desired recombinant protein, was further purified onto a gel filtration chromatographic column (Superdex 200; Amersham). The eluted protein (
95% purity) was stored in 50 mM HEPES (pH 7.9), 100 mM NaCl, 1 mM dithiothreitol, and 10% glycerol until use. Bax recombinant protein was from Abnova.
Cell-free system with isolated mitochondria.
Mitochondria were isolated as previously described (79). Assessment of m was carried out by incubating 100 µg of mitochondria with 500 nM of the indicated recombinant protein in 80 nM rhodamine 123 (Invitrogen) and scoring by flow cytometry. For the mitochondrial swelling assay, 100 µg of mitochondria was incubated with recombinant Drp1, Drp1K38A, or Drp1K679A proteins (500 nM) and the A520 was recorded in an Ultrospec 3300 spectrophotometer (Amersham). For detection of cytochrome c release, mitochondria were incubated for 30 min at room temperature (RT) with the different recombinant proteins and then mitochondria were removed by centrifugation and supernatants and pellets were analyzed by immunoblotting.
Native gel electrophoresis. In situ detection of MRC complex I activity by native polyacrylamide gel electrophoresis was done as described previously (37, 72). Briefly, purified mitochondria were incubated with each recombinant protein for 30 min at RT and then loaded onto a 5 to 15% native polyacrylamide gel. Immediately after electrophoresis, the gel was incubated in 0.1 M Tris-HCl (pH 7.4), 1 mM NADH, and 2 mM nitroblue tetrazolium. The reaction was stopped with water after appearance of the band.
Oxygen consumption assessment. Mitochondrial respiration was measured in isolated mitochondria and digitonin-permeabilized cells using a Clark oxygen electrode (Oxygraph; Hansatech) as described previously (64). Substrate-driven respiration rates were measured as described previously (21) and expressed as nmol of O2/min/mg of proteins. Complex I substrates and inhibitor were added at the following final concentrations: 5 mM malate, 5 mM glutamate, and 2 µM rotenone.
Cell growth measurements. Cell growth was analyzed using a Quantos cell proliferation assay kit (Stratagene) according to the manufacturer's instructions.
Detection of ROS in isolated mitochondria. Measurement of ROS in purified mitochondria was carried out by incubating 100 µg of mitochondria with 500 nM of the indicated recombinant protein in buffer containing 100 µM of the luminol analog L-012 (Wako) (17). Chemiluminescence was counted in a Berthold LB96V MicroLumat Plus.
Immunofluorescence and imaging. Cells labeled with 5 µM CellTracker Green (Invitrogen) and 1 µM Hoechst 33342 were subjected to Hoffman modulation contrast (HMC) or fluorescent microscopic assessment. Images were visualized in a Nikon Eclipse TE-2000-U fluorescence microscope with a Plan Apo 60x/1.4 objective, acquired with a Nikon Digital DXM 1200 camera, and analyzed using the Nikon ACT-1 2.2 software. May Grünwald-Giemsa (MGG) staining solutions were from Ral. In immunofluorescence experiments, cells were fixed with 4% paraformaldehyde and stained for the detection of activated Bax (MAb 6A7; BD Biosciences); activated Bak (MAb Ab-1; Calbiochem); calnexin, KDEL, and Hsp60 (Stressgen), DRP1/DLP1 (BD Biosciences); and cytochrome c, Smac/DIABLO, Omi/HtrA2, AIF, and EndoG. All were mounted with Vectashield and detected by anti-mouse or anti-rabbit IgG conjugated with Alexa Fluor (Invitrogen) according to standard procedures. The quantification of different parameters by fluorescence microscopy was performed in blind testing by at least two investigators, on 100 cells for each data point, and was repeated at least four times in independent experiments. Images were visualized at RT in an Apotome-equipped Zeiss Axioplan (Axiovert 200 M; Zeiss) with an Apochromat 100x/1.4 objective, acquired with a charge-coupled device Roper Scientific Coolsnap HQ camera, and analyzed using the Axiovision 4.4 software. MGG and Bax activation images were visualized at RT in a Nikon Eclipse TE-2000-U fluorescence microscope as described above.
Electron microscopy. Cells were fixed with 2% glutaraldehyde in phosphate buffer (pH 7.4) for 2 h at RT, washed, and postfixed in 2% OsO4 before being embedded in Durcupan. Analysis was performed with a transmission electron microscope (Carl Zeiss MicroImaging), on ultrathin sections stained with uranyl acetate and lead citrate.
Statistical analysis. The significance of differences between experimental data included in Fig. 4A and B was determined using the Student t test for unpaired observations.
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RESULTS |
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Next, we characterized the biochemical events of CD47-mediated type III PCD. A detailed kinetic analysis confirmed that TSP and CD47 MAb-treated primary B cells underwent PS exposure, cell viability loss, m disruption, ROS production, and lysosomal permeabilization (Fig. 1E). Of note, pretreatment of cells with agents that block lysosomal function, such as bafilomycin A, did not prevent the PS exposure/cell viability loss associated with CD47 MAb treatment (see Fig. S1 in the supplemental material). This fact strongly suggests that lysosomal permeabilization is a secondary event in this type of PCD. CD47-mediated PCD is a rapid and time-dependent process, with significant alterations observed as soon as 30 min after treatment and a maximum effect at 6 h (Fig. 1E). In contrast, classical HC-induced type I PCD is a slower process, starting at 6 h and plateauing at 20 h. It is important to remark that CD47-mediated PCD does not require new transcription or translation. Indeed, CD47-mediated PCD is not modulated by actinomycin D or cycloheximide (see Fig. S2 in the supplemental material). Despite the aforementioned dysfunctions, CD47 MAb-treated cells did not show the biochemical hallmark of nuclear apoptosis, namely, oligonucleosomal DNA fragmentation, observed after HC treatment (Fig. 1F). Interestingly, the mitochondrial alterations observed after CD47 triggering (e.g., m loss and ROS generation) were coupled with a drop in cellular ATP levels (Fig. 1G). The cellular ATP pool diminished in a time-dependent manner with kinetics similar to those of CD47-mediated death, confirming that CD47 triggering provokes an effect on the levels of cellular ATP.
Next, we sought to determine the involvement of caspases in CD47-mediated PCD. First, we found that CD47-mediated PCD was not affected by the pan-caspase inhibitors QVD.OPh and z-VAD.fmk or by inhibitors of caspases 2, 3, 6, 7, 8, 9, and 10. As depicted in Fig. 1H, as opposed to the effect observed in the HC-mediated caspase-dependent PCD process, neither broad nor specific caspase inhibitors elicited a change in the mitochondrial damage or the PS exposure induced by CD47 ligation. Additional evidence that CD47 ligation does not induce caspase activity was seen using a fluorogenic substrate-based assay (Fig. 1I). This probe revealed a caspase 3/7 activity associated with HC-treatment and a residual activity, similar to that measured in control cells, in CD47 MAb-treated lymphocytes. Finally, we assessed the activation of two key players in caspase-dependent PCD: caspase 9 and caspase 3. These proteases are synthesized as inactive proenzymes of 49 and 32 kDa, respectively. After a caspase-dependent apoptotic insult, such as HC, caspases 9 and 3 are cleaved to yield active subunits of 37/39 and 17 kDa, respectively. Immunoblot analysis demonstrated that, in contrast to HC, caspase 9 or caspase 3 did not become activated even 6 h after CD47 MAb treatment (Fig. 1J). Overall, these results demonstrated that CD47-mediated PCD is a caspase-independent process.
Together, our results indicate that the cell damage induced by CD47 ligation is defined exclusively by cytoplasmic alterations. The morphological and biochemical features of CD47-mediated PCD represent the hallmarks of caspase-independent type III PCD.
The chymotrypsin-like family of serine proteases controls CD47-mediated killing.
The absence of caspase activation led us to evaluate the possible effect of a specific family of proteases on the regulation of CD47-mediated cell death. Among the protease inhibitors tested, including inhibitors of trypsin-like proteases; calpains I and II; cathepsins B, D, and L; proteasome; and other general inhibitors of serine or cysteine proteases, only the chymotrypsin-like serine protease inhibitor TPCK maintained the cellular viability and mitochondrial function of CLL cells (Fig. 2A and B). As a control in this pharmacological approach, we verified that trypsin-like proteases, calpains, cathepsins, or proteasome inhibitors precluded other previously described types of PCD (1, 22, 23, 29). Cytosolic extracts from CD47 MAb-treated cells, which contained a chymotrypsin-like TPCK-inhibitable protease activity but were devoid of either trypsin-like or cathepsin protease activity, supported our pharmacological conclusion (Fig. 2C). These results were further corroborated in normal and leukemic B cells by fluorescence microscopy (Fig. 2D) and flow cytometry (Fig. 2E) with the help of a fluorochrome-labeled TPCK analog that covalently binds to the active site of this subtype of serine proteases. In untreated cells, this analog yields negative results. However, after CD47 MAb ligation, B cells displayed a positive labeling, which is specifically inhibited by TPCK and not TLCK. Overall, these results demonstrate the involvement of chymotrypsin-like proteases in type III PCD. In addition, given that the inhibition of this type of protease suppresses the mitochondrial alterations characterizing CD47-mediated PCD (e.g., m loss), our data imply a hierarchical relationship between chymotrypsin-like serine proteases and mitochondria.
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m loss provoked by the PCD inductor etoposide (a topoisomerase II inhibitor) (Fig. 3B). When targeted to the ER, overexpressed Bcl-2 also prevented apoptosis induced by the ER calcium-mobilizing agent thapsigargin (Fig. 3B), confirming previous work suggesting that Bcl-2 maintains Ca2+ homeostasis within the ER, thereby inhibiting apoptosis induction by thapsigargin (32). The death action of the microtubule-interfering reagent Taxol, which provokes the cytosolic-mitochondrial redistribution of the proapoptotic protein Bim (81), was significantly inhibited by the downregulation of this protein (Fig. 3B). Thus, our findings reveal that, in contrast to apoptosis caused by etoposide, thapsigargin, or Taxol, the mitochondrial dysfunctions characterizing CD47-mediated PCD are not controlled by the most representative members of the Bcl-2 family of proteins. An additional set of experiments validated these findings in CLL cells. In contrast to HC-treated cells, CD47 MAb-treated cells showed no activation of the proapoptotic proteins Bax and Bak (Fig. 3C and D). Moreover, this form of cell death proceeded without the release of the mitochondrial proapoptotic proteins AIF, cytochrome c, Smac/DIABLO, EndoG, and Omi/HtrA2 from mitochondria to cytosol (Fig. 3E and F).
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m dissipation. Indeed, the lack of implication of these Bcl-2 family members could explain the absence of release of proapoptotic proteins from mitochondria in this mode of PCD. Drp1 redistributes from cytosol to mitochondria after CD47 ligation. We next searched for a key element integrating the morphological and biochemical hallmarks of CD47-induced PCD. Accumulating evidence suggests that dynamin-related proteins, a family of proteins incriminated in mitochondrial remodeling (2, 10, 18, 46), establish a link between the mitochondrial dysfunctions cited above and PCD. Remodeling of mitochondria is controlled by the balance between fission and fusion and, more specifically, by the balance between Drp1 (33, 38, 74, 75, 86), Opa1 (13, 26, 34, 55), mitofusin 1 (Mfn1) (12, 67), and mitofusin 2 (Mfn2) (39). In this context, the striking mitochondrial inner membrane morphological changes observed in CD47-mediated PCD (Fig. 1C) led us to investigate the implication of these proteins in type III PCD. We thus quantified Drp1, Opa1, Mfn1, and Mfn2 mRNA expression in B cells from control donors and in B lymphocytes from 30 CLL patients described in Table S1 in the supplemental material. This analysis indicates that in most of the CLL patients tested, Drp1 and Mfn1, but not Opa1 or Mfn2, mRNA expression was high compared to that in B cells from control donors (Fig. 4A). Importantly, the Drp1 mRNA and protein expression strongly correlated with the susceptibility of B cells to CD47-induced PCD (Fig. 4A and B). Indeed, in patients 1 to 28 an elevated Drp1 expression level corresponds with a greater degree of responsiveness to CD47-mediated PCD, while patients 29 and 30, like subjects with normal B cells, presented both lower Drp1 expression and a poor cell death response to TSP and CD47 MAb (Fig. 4A and B; see Table S1 in the supplemental material). Interestingly, when comparing CD47-mediated PCD and HC-induced apoptosis in this panel of patients, CD47-mediated PCD represented a more efficient pathway to induce death. In fact, this comparative analysis distinguished three different types of response in CLL cells (Fig. 4B). In B cells from 19 patients (group 1), the response to TSP or CD47 MAb was comparable to that to HC treatment. CD47 ligation appeared to be a more efficient means of inducing PCD in B cells from nine patients (group 2), while in B cells from patients 29 and 30 (group 3) we observed a lower degree of death, similar to the level found in B lymphocytes from control donors. Overall, these results led us to further investigate the role of Drp1 in CD47-mediated PCD.
When redistributed from cytoplasm to mitochondria, Drp1 plays an essential role in the morphological changes observed during type I apoptosis (4, 7, 25, 40, 60, 66, 76). We showed that, after CD47 triggering, Drp1 translocated from cytoplasm to mitochondria in normal and leukemic cells in a time-dependent manner with kinetics similar to those for the CD47-mediated cell death response (Fig. 4C and D). Interestingly, we also showed that Drp1 redeployment strictly correlated with the response of CLL cells to CD47 triggering. In fact, mitochondrial relocalization of Drp1 was observed in a similar degree in B cells from CLL patients displaying a greater degree of responsiveness to CD47 ligation (e.g., patients 1 and 27) but to a lesser extent in B cells from a patient displaying poor cell death response to TSP or CD47 MAb (patient 30) (Fig. 4E). In contrast, Drp1 did not redistribute to mitochondria in HC-induced caspase-dependent apoptosis, suggesting that HC-mediated apoptosis utilizes a different molecular pathway than CD47-mediated PCD (Fig. 4E). These data strongly suggest that the localization of Drp1 in mitochondria is a central event in CD47-mediated PCD.
Drp1 induces mitochondrial damage and controls CD47-mediated PCD.
Next, we established that Drp1 provokes the m collapse observed in CD47-induced PCD. After CD47 MAb triggering, cells transiently transfected with three different siRNAs designed against human Drp1 showed less pronounced m loss, as well as inhibition of PS exposure, than control siRNA-transfected cells (Fig. 5A). Thus, Drp1 gene silencing significantly attenuated CD47 MAb toxicity. This was further confirmed by a cell proliferation assay, which revealed that after CD47 MAb treatment, cell growth was significantly higher in Drp1-downregulated than in control transfected cells (Fig. 5B). This result indicates that Drp1-downregulated cells treated with CD47 MAb survived and, therefore, divided. Strikingly, in contrast to Drp1 gene silencing, overexpression of Drp1 sensitized cells to CD47 MAb (Fig. 5C). However, neither Drp1 downregulation nor Drp1 overexpression modulated apoptosis induced by HC, confirming that, as reported for other systems of type I PCD (57), this cell death pathway was Bax/Bak dependent (Fig. 3C) but Drp1 independent. Interestingly, two independent mutations in conserved lysines predicted to reduce the GTPase activity of Drp1 (K38 and K679) (75, 89) failed to inhibit CD47-mediated mitochondrial damage. Indeed, overexpressed Drp1K38A and Drp1K679A proteins, like Drp1, redistributed to mitochondria after CD47 triggering and sensitized cells to CD47-mediated PCD (Fig. 5C and D). These results indicate that the death function of Drp1 was independent of its GTPase activity in CD47-mediated PCD. In this way, pretreatment of cells with two compounds used to decrease intracellular GTP pools, mycophenolic acid and ribavirin, failed to inhibit CD47-mediated cell death (data not shown). Overall, our data indicate that Drp1 induced the m damage that characterized CD47-mediated type III PCD via a GTPase-independent mechanism.
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m loss, and type III PCD, we tested whether downregulation of the mitochondrial Drp1 binding protein hFis1 (36, 85, 87) regulated CD47-induced PCD. Transient transfection of two different siRNA double-stranded oligonucleotides designed against hFis1 alleviated the presence of Drp1 in mitochondria, along with m loss and PS exposure triggered by CD47 ligation (Fig. 5E). Thus, downregulation of Drp1 (Fig. 5A) or the mitochondrial Drp1 receptor hFis1 prevented CD47-mediated PCD, indicating that the presence of Drp1 in mitochondria is a key element in type III PCD.
We further generated the human Drp1 recombinant protein to investigate whether the mitochondrial alterations observed in CD47-mediated death were directly provoked by the protein itself. In this way, Drp1, Drp1K38A, or Drp1K679A was exposed in a cell-free in vitro system to highly purified mitochondria. After incubation, mitochondrial features were monitored by four independent methods: cytofluorometry to measure m, luminometry to assess ROS generation, spectrophotometry to quantify mitochondrial swelling, and immunoblotting to reveal cytochrome c release from mitochondria. Using these systems, we observed that the three recombinant proteins provoked m dissipation and ROS in isolated mitochondria in the absence of swelling or cytochrome c release (Fig. 6A, B, C, and D). Indeed, Drp1, Drp1K38A, and Drp1K679A caused in vitro the same effects as those observed in TSP- or CD47MAb-treated cells. As a negative control in these experiments, we used an inactive Drp1(1-335) mutant protein, which failed to generate damage in purified mitochondria.
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m dissipation and the high ROS production observed during CD47-mediated PCD.
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m dissipation, ROS production, and ATP loss (Fig. 1E and G and 7C). These data strongly suggest that CD47 ligation provokes disruption of MRC through the massive relocalization of Drp1 in mitochondria. Accordingly, TPCK, a pharmacological agent that inhibits CD47-mediated type III PCD and blocks the relocalization of Drp1 in mitochondria (Fig. 2 and data not shown), restored the normal mitochondrial respiration rate in CD47 MAb-treated cells (Fig. 7C).
Together, these results showed that following CD47 ligation, Drp1 redistributes to mitochondria (Fig. 4C and D), where the protein provokes impairment of MRC activity (Fig. 7), loss of m (Fig. 1C and 6A), production of ROS (Fig. 1C and 6B), ATP loss, and disruption of the mitochondrial structure (Fig. 1C).
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DISCUSSION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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m dissipation, ROS generation, and ATP loss. These mitochondrial alterations (e.g., massive ROS production) may certainly enhance the damage to other organelles, such as the ER, lysosomes, and Golgi apparatus, facilitating the execution of this necrotic cell death pathway.
Nevertheless, type III CD47-mediated cell death shares common features with the other two forms of PCD. Certain biochemical features are conserved, namely, outer leaflet exposure of PS in the plasma membrane, alterations to mitochondria and lysosomes, and production of ROS. In addition, all three forms of PCD are marked by a dependence on input from several cellular compartments, the most important being the mitochondrion. In CD47-mediated type III PCD, four main observations support this last assertion. First, impairment of the MRC, a fall in the m, and the concomitant ROS generation and ATP loss are constant features of this type of PCD. Second, from the available biochemical data, it appears that this kind of cell death does not require the participation of other proapoptotic organelles, such as lysosomes (e.g., through cathepsin B, D, or L activity). Third, inhibitors of the mitochondrial modifications induced by CD47 ligation (e.g., TPCK) abolish CD47-mediated PCD. Finally, downregulation of the Drp1 mitochondrial receptor hFis1 precludes the m loss and PS exposure which are associated with CD47 triggering. Together these data are compatible with the hypothesis that m disruption constitutes a key event in type III PCD. The direct cause-effect relationship between the mitochondrial and cytoplasmic manifestations of type III PCD further underlines the central role of mitochondria in the regulation of this type of cell death.
Control of the Drp1 redistribution mediated by the chymotrypsin-like serine proteases was essential to an understanding of the interplay between the multiple CD47-induced signals. Our results indicate that activity of the chymotrypsin-like family of serine proteases allows Drp1 redistribution from cytosol to mitochondria in CD47-mediated PCD. Thus, as in other types of cell death (3, 84), Drp1 redistributes from cytosol to mitochondria in a specifically controlled manner. Further analysis is necessary to identify the serine protease involved in this kind of cell death. In any case, our results place the chymotrypsin-like family of serine proteases at an early premitochondrial step in caspase-independent type III PCD.
Normally residing in cytoplasm, Drp1 translocates under apoptotic conditions to mitochondria, where it interacts with its mitochondrial partner, hFis1 (36, 76, 85). This mitochondrial translocation is Ca2+ dependent in some types of apoptosis (8, 27). In apparent contradiction to the data reported here, it has been reported that Drp1 and Bax colocalize in mitochondria to induce mitochondrial fission and caspase-dependent cell death (39, 40). As in etoposide or nitric oxide PCD (4, 77), this kind of caspase-dependent PCD relates to the GTPase function of Drp1. In this context, the GTPase-inactive Drp1 mutant Drp1K38A blocks STP- or etoposide-induced mitochondrial fission, m loss, cytochrome c release, and type I apoptosis (39, 77). In HC-mediated caspase-dependent killing, our data indicated that Bax activation was not coupled to Drp1 translocation (Fig. 3C and D). Consequently, HC-mediated PCD is not regulated by overexpression or downregulation of Drp1. In the same way, a recent study confirms that in caspase-dependent PCD induced by actinomycin D or UV irradiation, downregulation of Drp1 or hFis1 does not inhibit Bax-dependent apoptosis (57). Finally, in caspase-independent CD47-mediated type III PCD, the mitochondrial alterations provoked by Drp1 translocation (e.g., m collapse and ROS generation) occur in the absence of Bax activation (Fig. 3C and D), release of mitochondrial apoptogenic proteins (Fig. 3E and F), or mitochondrial fission (J. E. Esquerda and S. A. Susin, unpublished observation). Thus, depending on the form of PCD, it seems that Bax and Drp1 could be coupled or uncoupled in the induction of mitochondrial alterations and cell death.
Interestingly, the mitochondrial dysfunctions monitored in CD47-mediated PCD are independent of the GTPase function of Drp1. In fact, overexpression of wild-type Drp1 or the GTPase-deficient Drp1 mutants Drp1K38A and Drp1 K679A provokes a similar degree of responsiveness to CD47-mediated PCD. This apparent paradox in Drp1 mitochondrial function may reflect the fact that Drp1 is likely to be a bifunctional protein in PCD with a fission, GTP-dependent function and an alternate mitochondrial cell death function. In this way, two recent reports have shown that Drp1 mitochondrial targeting is independent of the GTPase activity of the protein (59) and that Drp1 has other functions in mitochondria that differ from fission regulation (27). A third study has demonstrated that deletion of the hFis1 1-helix inhibits mitochondrial fragmentation but strongly enhances interaction between Drp1 and hFis1. This interaction, which depends on the region from amino acid 61 to 91of hFis1, provokes the same features as those observed in CD47-mediated death: mitochondrial swelling in absence of cytochrome c release (87). These reports and our data strongly favor the hypothesis that massive and specific targeting of Drp1 to the mitochondrial surface and interaction with partners proteins such as hFis1 cause mitochondrial dysfunction. However, other proteins, such as Bax, are needed to induce the release of apoptogenic proteins from mitochondria or to facilitate mitochondrial fission. In STP- or etoposide-mediated PCD, caspase activity or Bax function may enhance Drp1 mitochondrial activity, causing both the release of proapoptotic proteins from mitochondria and the mitochondrial outer membrane modifications that, in cooperation with the GTPase-dependent mechanochemical properties of Drp1, lead to mitochondrial fission. In this context, the lack of implication of caspase and Bax in CD47-mediated PCD explains the absence of both mitochondrial fission and release of apoptogenic proteins.
Cell-free systems have been extremely valuable for the analysis of apoptosis mechanisms (24, 78). Using this in vitro system we demonstrated that, once in mitochondria, Drp1 induced a dramatic decrease in m. Surprisingly, this mitochondrial alteration was not accompanied by mitochondrial swelling or cytochrome c release. In fact, we showed that Drp1 blocked the electron transport activity of MRC and provoked massive ROS generation. In this sense, it is known that electron transport impairment results in m loss, massive ROS production, and lack of ATP generation (31), and that is exactly the picture observed in CD47-mediated PCD. With kinetics similar to those of CD47-mediated Drp1 redistribution, m loss, ROS production, and ATP decrease, we confirmed the dysfunction of MRC in CLL and Jurkat cells subjected to CD47 ligands. Thus, as in caspase-dependent type I PCD (63, 64), the loss of MRC function was implicated in the mitochondrial damage characterizing caspase-independent type III PCD. A more detailed study should be developed to identify whether Drp1 provokes the dysfunction of the MRC electron transport directly, by action on a specific complex (e.g., complex I), or indirectly, by provoking a general MRC disorganization. If the "direct" action applies, important questions need to be resolved. How does the connection between Drp1 (on the outer mitochondrial membrane) and MRC (on the inner mitochondrial membrane) take place? Does Drp1 translocate from the outer to the inner mitochondrial membrane to disrupt mitochondrial electronic transport? Does Drp1 inhibit MRC complex II, III, or IV activity? In the second, "indirect" situation, it seems possible that the presence of Drp1 in mitochondria, which provokes a disorganization of the mitochondrial structure (Fig. 1C), enables a mechanical rupture of the mitochondrial inner membrane. This certainly impairs normal MRC electron transport activity. In our hands, this possibility applied in the pretreatment of purified mitochondria with atractyloside, a ligand of the mitochondrial adenine nucleotide transporter. Atractyloside induced mitochondrial swelling that, in turn, provoked a mechanical rupture of mitochondrial membranes (88). As a consequence, mitochondrial electronic transport was impaired (e.g., as depicted in Fig. 7A). Another example of "indirect" MRC impairment is the pretreatment of mitochondria with the uncoupler mClCCP (carbonyl cyanide m-chlorophenylhydrazone) which also blocks complex I activity (Fig. 7A). In any case, our data fully confirm that in CD47-mediated type III PCD, the action of Drp1 on mitochondria provokes impairment of MRC I activity (Fig. 7), loss of m (Fig. 1C and 6A), production of ROS (Fig. 1C and 6B), ATP loss, and disruption of the mitochondrial structure (Fig. 1C). Of course, it would be of great interest to study whether this novel Drp1 function in type III PCD can be extended to other cell types or PCD pathways (e.g., caspase-dependent type I PCD).
Our experiments, performed with B cells from control donors and a large number of CLL patients, showed that CD47 ligation provoked cell death rapidly, with a higher efficacy in CLL cells than in normal B cells, and more efficiently than HC-mediated apoptosis. Interestingly, sensitivity to CD47-mediated PCD strongly correlated with the expression of Drp1, making Drp1 a potential marker for PCD susceptibility. Our data demonstrate the existence of a biochemical caspase-independent pathway in primary malignant B cells that could be regulated independently of the caspase-dependent pathway. In fact, in CLL, accumulation of B cells reflects their resistance to "classical" caspase-dependent apoptosis. Hence, the study of the molecular basis of alternative cell death pathways can provide new means of improving the current therapeutic strategies employed in the treatment of this disease.
Together with the biochemical mechanisms proposed to mediate CD47-induced PCD in normal and leukemic T cells (43, 44, 50, 62, 69), our observations in primary B cells lead us to conclude that CD47 and its two ligands, TSP and SIRP-, could participate in the elimination of cells during the immune response. In that sense, TSP, secreted by macrophages and DCs at a steady state and during an inflammatory process, may contribute to the maintenance of immune homeostasis. First, TSP/CD47 interaction serves as an autocrine negative regulator of DC maturation and function (20). Second, TSP/CD47 induces caspase-independent PCD in lymphoid cells, sparing the DCs, followed by the engulfment of the dying cells by professional phagocytes (53). Notably, the expression of CD47 on apoptotic cells is required to allow their in vitro phagocytosis by APC (80). Third, TSP establishes "a molecular bridge" between apoptotic cells and phagocytes, facilitating the clearance of dying cells (70).
In conclusion, our work reveals substantial progress in two major areas: (i) in the understanding of the molecular events regulating caspase-independent type III PCD in normal and leukemic cells and (ii) in the development of new approaches to CLL treatment. In fact, the identification of a "downstream" death effector of CD47-mediated PCD, such as Drp1, might pave the way for novel diagnostic and pharmacological strategies for the treatment of this malignancy.
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ACKNOWLEDGMENTS |
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This work was supported by institutional grants from Institut Pasteur and CNRS and by specific grants from Fondation de France, Ligue Contre le Cancer, and Association pour la Recherche sur le Cancer (contract no. 4043) (to S.A.S.), as well as grants from the Spanish Ministry of Education and Science (grant 2005-01535) (to J.E.E.), the Canadian Institute for Health and Research (grant MOP-4490) (to M.S.), and Fonds Recherche France-Canada (to S.A.S. and M.S.). M.B. was supported by Ph.D. fellowships from MENRT and CANAM-Pasteur. V.J.Y. was supported by a Marie Curie fellowship (contract MEIF-2003-501887), S.B. by a Ph.D. fellowship from MENRT, and P.S. by a postdoctoral fellowship from Fondation Manlio Cantarini.
The authors of this paper declare that they have no competing financial interests.
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FOOTNOTES |
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