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Address correspondence to S.B. Brown, Centre for Inflammation Research, Medical School, University of Edinburgh, Teviot Place, Edinburgh EH8 9AG, UK. Tel.: 44-131-6511606. Fax: 44-131-6511607. E-mail: simon.brown{at}ed.ac.uk
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Abstract |
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Key Words: mitochondria; Fas; apoptosis; caspases; thrombopoiesis
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Introduction |
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For some time, we have been interested in the relationship between MK apoptosis and platelet production. In particular, we were intrigued by the possible role of caspases in both processes, as we had previously shown that mature blood platelets were committed to a caspase-independent program of cell death (Brown et al., 2000). Thus, when deprived of plasma as a source of survival factors, cultured platelets exhibited morphological condensation accompanied by exposure of phosphatidylserine (PS), maintenance of plasma membrane integrity, and specific recognition and engulfment by monocyte-derived macrophages (MDMs) using the class-A scavenger receptor (Brown et al., 2000). Although these events were strongly reminiscent of constitutive apoptosis in cultured leukocytes, a key difference was that platelet death was not inhibited by broad-spectrum caspase inhibitors, nor was there evidence of activation by cleavage of effector caspase-3 (Brown et al., 2000), which is found in abundance in platelets.
The current study supports the hypothesis that MKs produce platelets by a novel compartmentalized form of caspase-directed apoptosis (De Botton et al., 2002), but importantly extends those studies to show that such proplatelet MKs yield functional platelets that retain inner mitochondrial membrane potential (M) and membrane PS asymmetry. By contrast, the MK cell body exhibited typical nuclear morphological features of apoptosis. Further evidence of compartmentalized progenitor cell death was the absence from viable platelets of caspase-9 present in MKs, accounting for the caspase-independent nature of constitutive platelet death.
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Results |
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Platelets generated by MK apoptosis are not ingested by macrophages
To confirm the morphological and functional evidence that platelets produced by MEG-01 MKs undergoing apoptosis were viable, we investigated whether MDMs would selectively clear nonfunctional platelets and MK fragments. Incubating MK culture supernatants with MDMs resulted in the selective clearance of all PS-positive bodies, leaving a population of functional platelets demonstrating agonist-induced shape change (Fig. 3 D).
Mitochondrial permeability transition is not observed in proplatelet MK extensions and occurs only as mature platelets die
The foregoing data strongly implied that a compartmentalized form of apoptosis in proplatelet MKs gave rise to viable platelets and an apoptotic remnant body. Because mitochondrial permeability transition is a prominent feature of caspase-mediated apoptosis, we investigated proplatelet MK M using JC-1, a mitochondrial dye that fluoresces orange in respiring mitochondria that maintain
M (Petit et al., 1995; Salvioli et al., 1997). Importantly, we found that mitochondria, localized to platelet-sized nodes along the cytoplasmic extensions of proplatelet MKs, had not undergone permeability transition despite double staining with Hoechst 33342 showing clear morphological evidence of nuclear condensation and fragmentation within the main cell body (Fig. 5, A and B). In addition, confocal microscopy revealed that mitochondria remaining within the cell body were polarized to the MK edge with the remaining proplatelet "bridge" still attached (Fig. 5 C). Furthermore, viable MK culture-derived platelets that were allowed to adhere and spread on glass also showed no evidence of mitochondrial permeability transition (Fig. 5 D). This was only observed when mature human blood platelets were cultured in the absence of plasma-derived survival factors for 16 h to allow constitutive death (Brown et al., 2000), or when fresh blood platelets were treated with the respiratory chain uncoupler mCCCP (Fig. 6 A).
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Unlike MKs, freshly isolated platelets lack caspase-9, which is required for caspase-3 activation in platelet lysates
We were intrigued by mitochondrial permeability transition and cytochrome c release in senescent platelets because our previous work had clearly shown that constitutive death in such platelets was caspase-independent, with no evidence of the caspase-3 activation normally seen in caspase-mediated apoptosis (Brown et al., 2000). Together, these data implied that freshly isolated mature blood platelets lacked either APAF-1 or caspase-9, key components of the apoptosome that cleaves caspase-3 to its active p12/17 form (Liu et al., 1996; Li et al., 1997).
Two lines of evidence demonstrated that platelets (but not MKs) lacked caspase-9. First, Western blot analysis of blood platelets confirmed that APAF-1 was present and at comparable levels to Jurkat T cells (Fig. 7 A). However, although caspase-9 was readily identified in Jurkat T cells, migrating with an apparent molecular size of 48 kD, caspase-9 could not be detected in blood platelets. Importantly, the MK cell lines MEG-01 and SET-2 expressed caspase-9, and at levels equivalent to Jurkat T cells using both a polyclonal (Fig. 7 B) and a monoclonal antibody (unpublished data), indicating exclusion of the enzyme during platelet formation.
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Discussion |
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Our findings differ importantly from the probable role of caspases in generation of the erythrocyte, another anucleate cell (Zermati et al., 2001). Rather than production of multiple daughter cells from a single progenitor (Stenberg and Levin 1989; Cramer et al., 1997; Italiano et al., 1999), the data suggest that viable erythrocytes transiently activate caspases, resulting in cleavage of a subset of structural proteins that may help lead to the enucleation of the erythroblast (Zermati et al., 2001) to produce a single erythrocyte (Gregory and Eaves, 1978). Nevertheless, it seems clear that evolution has achieved adaptations of the basic program of cell death by apoptosis to produce anucleate cells of critical importance in blood. However, such adaptations do not yield immortal cells; viable platelets arising from compartmentalized MK death are committed to a short life span and later death (Brown et al., 2000), in keeping with nucleated blood cells such as neutrophils (Savill et al., 1989).
Our data also reinforce the concept that plasma membrane changes of apoptosis may be dissociated from the caspase-directed program of nuclear condensation and fragmentation (Knepper-Nicolai et al., 1998; Harper et al., 2001). Clearly, further work will be required to define the mechanisms by which caspase activation in one part of the cell can lead to nuclear changes, whereas mitochondria in a different area of the cell retain their M, even when cell death is induced by Fas ligation. Nevertheless, because it is believed that there is specific delivery of progenitor cell cytoplasmic components into the extensions of proplatelet MKs (Italiano et al., 1999), it is possible that there is specific exclusion of death pathway components upstream of mitochondria analogous to the exclusion of caspase-9 suggested by our data. Unfortunately, we could not directly test this possibility because platelet yields from MK cultures were insufficient for blotting studies, and the delicate nature of proplatelet-bearing MKs, which tended to lose their processes on manipulation, excluded direct immunofluorescent localization of caspase-9. However, despite our biochemical and functional evidence that platelets lack caspase-9, others have reported its presence (Wolf et al., 1999), even though our data would suggest this should have resulted in caspase-3 cleavage, which we were unable to detect. Indeed, the possibility that we might have artifactually "missed" caspase-9 activity, because calcium-dependent calpains can indirectly block caspase-3 activation by inactivating caspase-9 (Wolf et al., 1999; Chua et al., 2000; Lankiewicz et al., 2000), appeared most unlikely in our studies given the lack of Ca2+, and presence of EGTA, EDTA, and calpain inhibitors within the lysis buffer. Moreover, it should be noted that we were particularly careful to ensure that our platelet preparations were free of low grade contamination by leukocytes, which may have served as an artifactual source of caspase-9 (Webb et al., 2000; Bantel et al., 2001).
Future studies should also examine whether retention of M and plasma membrane asymmetry (denying recognition by phagocytes) in those parts of the dying MKs destined to become platelets reflect special expression of anti-apoptotic members of the Bcl-2 family. Such proteins have been implicated in platelet production because knockout of the pro-apoptotic Bcl-2relative Bim results in reduced circulating platelet counts, whereas other circulating blood cells are increased (Bouillet et al., 1999), with identical effects being observed when the balance of Bcl-2 family members is similarly shifted toward survival of hemopoietic cells by targeted overexpression of Bcl-2 (Ogilvy et al., 1999). Although such observations provide indirect support for MK apoptosis being involved in platelet production, such data require careful interpretation in the light of intriguing observations suggesting special patterns of antiapoptotic Bcl-XL expression in MKs producing platelets (Sanz et al., 2001). Thus, although Bcl-XL was greatly up-regulated (Terui et al., 1998; Sanz et al., 2001) as MKs differentiated from CD34+ progenitors and was detectable in proplatelet fragments and mature platelets, potentially explaining the maintenance of
M, Bcl-XL was apparently absent from senescent/apoptotic MK cell bodies (Sanz et al., 2001). A further degree of complexity is found with overexpression of Bcl-2 in CD34+ progenitor cells cultured in vitro where proplatelet extensions were inhibited (De Botton et al., 2002). This further suggests that Bcl-2 members may be differentially targeted to different populations of mitochondria, where Bcl-2 is known to be absent from mature blood platelets (Brown et al., 2000; Sanz et al., 2001). Such data may support sorting of cytoplasmic proteins during formation of proplatelet extensions by MKs and emphasize the future need to overcome technical and logistical problems to follow location of Bcl-XL, caspase-9, and other relevant proteins in proplatelet forming MKs in real time.
Nevertheless, our findings contrast with those of De Botton et al. (2002) in assigning relative importance to cytochrome c and the intrinsic cell death program in the initiation of proplatelet formation. Mitochondrial staining with the M-sensitive dye JC-1 suggested that during platelet formation the mitochondria retained an active electron-transport chain and were actively sorted to the proplatelets, consistent with the high energy demands that are placed on platelet formation (Watanabe et al., 1990) and the subsequent utilization of oxidative phosphorylation by platelets throughout their life span (Doery et al., 1970; Akahori et al., 1995; Parker and Gralnick, 1997). Given the intensity of the JC-1 stain, we also failed to observe mitochondria with altered fluorescence properties, indicative of a loss in
M even when distinctive chromosomal margination of the attached MK body was apparent. The ability of mitochondria to resist participation in an apoptotic program that results in proplatelet formation would also be assisted by the high levels of reported Bcl-XL (Terui et al., 1998; Sanz et al., 2001), thus able to inhibit an intrinsic (but not extrinsic) cell death program. This is in keeping with further results where the enforced release of cytochrome c by antagonism of anti-apoptotic Bcl-XL with "BH3 mimetics" resulted in the loss of
M and formation of platelet-like progeny that were dysfunctional (unpublished data). Therefore, it is unlikely that cytochrome c would be released into the cytosol of MKs destined to extend processes and form functional platelets. Furthermore, to our knowledge there is no precedent for the long-term maintenance of
M after release of mitochondrial cytochrome c.
Our findings further support the idea that abnormalities of circulating platelet numbers, important in disorders of hemostasis and in thrombotic diseases such as stroke and myocardial infarction, could reflect abnormalities in control of MK apoptosis. Indeed, in keeping with our findings, mice deficient in components of the Fas death pathway exhibit thrombocytopenia (Rieux-Laucat et al., 1995; Le Deist et al., 1996), which has largely been attributed to autoimmune thrombocytopenia. Indeed, one might speculate that pharmacological manipulation of MK apoptosis, and the Fas pathway in particular, might provide a novel therapeutic strategy for the management and control of thrombostasis.
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Materials and methods |
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Cell culture
The human megakaryoblastic cell line MEG-01 (Ogura et al., 1985; European Collection of Cell Cultures) and Jurkat T cells were routinely maintained in RPMI 1640 supplemented with 10% FCS, 2 mM L-glutamine, and 0.1 mg/ml penicillin/streptomycin. The human megakaryoblastic cell line SET-2 (Uozumi et al., 2000; provided by K. Uozumi, Kagoshima University, Japan) was maintained in DME supplemented with 10% FCS, 2 mM L-glutamine, 0.1 mg/ml penicillin/streptomycin, 10 µM 2-mercaptoethanol, and 10% nonessential amino acids. Generation of primary murine MKs is described elsewhere (Drachman et al., 1997; Rojnuckarin and Kaushansky, 2001). In brief, bone marrow was flushed from femurs of Balb-C mice and a single cell suspension was obtained by gentle pipetting. Cells were incubated overnight in StemSpanTM H2000 (StemCell Technologies, Inc.) with 40 ng/ml human recombinant TPO (PeproTech), and all nonadherent cells and media were transferred to fresh wells and incubated for a further 72 h. Enrichment of primary or cell linederived mature MKs was by velocity sedimentation through a discontinuous 1.5%, 3.0% BSA gradient at 1 g, collecting those cells reaching the bottom within 30 min. Purified primary MKs were replated and cultured with 30 ng/ml TPO and 10% normal human plasma. MKs were treated with the following reagents: 100 or 10 µM zVAD-fmk (Bachem), 10 µM zDEVD-fmk, 10 µM zIETD-fmk (Calbiochem); 50 ng/ml anti-Fas agonistic antibody CH.11 (Upstate Biotechnology), 1 µg/ml anti-Fas antagonistic antibody ZB4 (Upstate Biotechnology), 5 ng/ml soluble Fas ligand and enhancer (Qbiogene), 25 ng/ml TNF- (R&D Systems), and 250 ng/ml JO-2 (BD Biosciences). Phagocytic assays with MDMs were as described previously (Brown et al., 2000). Direct protein transfection was achieved using BioPORTER® II (Gene Therapy Systems) as instructed. In brief, CrmA (Kamiya Biomedical) or caspase-8 (CHEMICON International) in PBS was complexed with the BioPORTER® reagent for 5 min before the addition of cells in serum-free media. After 3 h of incubation, an equal volume of 20% serum media was applied, and cells were cultured for a further 8 h before enumeration of proplatelet-bearing MKs.
Ex vivo bone culture
Trabecular bone from femoral heads was isolated from a 58-yr old male patient undergoing hip surgery, machined with high precision to cylindrical cores under sterile conditions, and inserted into the loading chambers of a ZetosTM bone perfusion system (Smith and Jones, 1998; EOBM, Philipps-University, Marburg, Germany). Each core was maintained at 37°C /5% CO2 and perfused with 5 ml DME, recirculated at a rate of 5 ml/h. On day 5, the circulating media was replaced with 5 ml of fresh media containing 50 ng/ml Fas agonistic antibody CH.11 and/or 100 µM zVAD-fmk. The fresh media was then perfused and recirculated for 18 h at 5 ml/h before the eluent was collected and the bone core flushed with 5 ml of PBS. Large cellular material was removed by centrifugation (200 g for 5 min) and remaining supernatant components concentrated by centrifugation (1,500 g for 15 min). Resuspended pellets were stained for GpIIb (Serotec) and GpIIIa (CALTAG Laboratories), and analyzed on a FACScanTM (Becton Dickinson), with reference to a fixed count of 10-µm beads (Polysciences, Inc.) for platelet enumeration.
Flow cytometry
Culture supernatants containing platelets were separated by centrifugation at 300 g for 2 min, and platelets were enumerated by reference to a fixed count of 10-µm polystyrene beads (Polysciences, Inc.). Functional platelets were distinguished from cellular debris by a characteristic increase in side scatter on shape change after 0.5 U/ml thrombin or 3 µM ADP treatment, and an absence of 0.3 µl/ml annexin-V binding (Boehringer) to the shape changed population (Otterdal et al., 2001). All conditions contained 10 µg/ml propidium iodide as a dead cell gate. Transient calcium flux was detected using 10 µm Fluo3-AM (Molecular Probes, Inc.). Agonist-induced activation of the fibrinogen receptor (GpIIb/IIIa) by 3 U/ml thrombin was assessed by increased binding of 2.5 µg/ml Fibrinogen-Alexa Fluor (Molecular Probes, Inc.). Samples were analyzed on a FACScaliburTM system (Becton Dickinson) using CellQuest software. Immunofluorescent labeling of intact platelets for M was performed by adding 5 µl of platelets to 400 µl of PBS w/o Ca2+ containing 10 µg/ml JC-1 (Molecular Probes, Inc.), followed by incubation for 20 min at 37°C, before sampling by flow cytometry. Deliberate uncoupling of the respiratory chain with loss of platelet
M was achieved by direct addition of 10 µg/ml of the protonophore mCCCP.
Microscopy
Images were captured on an inverted microscope (Axiovert S100; Carl Zeiss MicroImaging, Inc.) equipped with a CoolSNAP CCD camera and OpenLab 3.0 image analysis software (ImproVision). Confocal images were captured on a confocal microscope and software system (TCS NT; Leica). Active caspases were detected using CaspaTagTM (Serologicals) as instructed. M was qualitatively determined using JC-1 (Molecular Probes, Inc.) at a final concentration of 5 µg/ml. Nuclear morphology was detected using 2 µg/ml Hoechst 33342. For TEM cells or culture supernatants were fixed with excess 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (EM grade I), followed by embedding within a fibrin plug as described previously (Brown et al., 2000). Plugs were subsequently treated as normal resected tissue and processed with osmium tetroxide, lead citrate, and araldite embedding, followed by 80-nm ultra thin sectioning. Samples were analyzed on an electron microscope (CM12; Philips).
Platelet isolation and culture
Freshly drawn venous blood was obtained from aspirin-free healthy donors, citrated (0.33%; Pharma Hameln), and PRP was prepared by centrifugation (350 g, 20 min). Washed platelets were prepared by diluting PRP with 5 vol of HBSS, pH 6.4, containing EDTA (4 mM final) into a round-bottom capped polystyrene centrifuge tube before centrifugation (280 g for 20 min). After washing, platelets were resuspended in HBSS, pH 6.4, and maintained at 37°C in a closed (capped) tube at 3 x 108 ml. Low levels of contaminating leukocytes were removed from platelet preparations after two successive rounds of centrifugation (220 g for 5 min) in which the supernatant was retained each time.
Cell free apoptosis and cytochrome c immunoprecipitation
Jurkat T cell and platelet lysates were prepared by resuspending cells in lysis buffer (20 mM Hepes, pH 7.5, 10 mM KCl, 1.5 mM MgCl, 1mM EDTA, 1 mM EGTA, 1 mM DTT, 100 µm PMSF, 10 µg/ml leupeptin, and 2 µg/ml aprotinin) and were incubating for 10 min on ice. Platelets were subjected to three cycles of freeze-thaw. Cells were further disrupted by passing through a 25 G needle 10 times. Cell free apoptosis was initiated by addition of 10 µg/ml cytochrome c and 1 mM dATP, and in some experiments, 0.1 U/ml human recombinant caspase-9 (CHEMICON International), followed by incubation at 37°C. At time points indicated aliquots were removed and directly added to x4 SDS Laemmli sample buffer, and immediately boiled for 3 min. For cytochrome c, washed platelets were resuspended in ice-cold 10 mM Hepes buffer (containing 1 mM EDTA, 1 mM 1,10-phenanthroline, 1 mM PMSF, 1 mM benzamidine, 10 µM pepstatin, 10 µM leupeptin, and 10 µM antipain, pH 8.0) and lysed after two rounds of freeze-thaw. The lysed platelets were centrifuged (13,000 g for 10 min) to yield a cytosolic supernatant, whereas the pellet was resuspended in lysis buffer before centrifuging again and dissolving the pellet in 1% TX-100 to release mitochondrial cytochrome c. The soluble fractions were then precleared with a control IgG and Protein G agarose before immunoprecipitating cytochrome c with clone 6H2.B4 (BD Biosciences) and boiling in SDS sample buffer. Cycloheximide (20 µg/ml)-treated Jurkats were used as a positive control and prepared identically.
SDS-PAGE and immunoblotting
SDS-PAGE and immunodetection of transblotted protein to PVDF membranes was performed as described previously (Brown et al., 1997). Western blots were probed with anti-caspase-3 pAb, anti-caspase-9 pAb and mAb (clone B40), anti-APAF-1 pAb, and anti-cytochrome c mAb (clone 7H8.2C12), all purchased from BD Biosciences. Molecular weight markers used were Rainbow markers, SDS-7B, or BenchMark (GIBCO BRL).
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Footnotes |
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Acknowledgments |
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This work was supported in part by the Wellcome Trust (047273, to J. Savill), Chief Scientists Office of the Scottish Executive (CZB/4/8, to S.B. Brown), European Space Agency Microgravity Applications Programme (AO 99-122, to D.B. Jones), the AO ASIF Davos (200J and O2 J7, to D.B. Jones), and a National Kidney Research Foundation Studentship (JS1, to M.C.H. Clarke).
Submitted: 21 October 2002
Revised: 18 December 2002
Accepted: 19 December 2002
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