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Prepublished online as a Blood First Edition Paper on June 7, 2002; DOI 10.1182/blood-2002-03-0686.
HEMATOPOIESIS
From the Institut National de la Santé et de la
Recherche Médicale (INSERM) U362 and the Centre National de la
Recherche Scientifique (CNRS, Unité Mixte de Recherche 1599),
Institut Gustave Roussy, Institut Fédératif de Recherche
54, Villejuif, France; CNRS, Unité Mixte de Recherche 8603, Université René Descartes (Paris V), Institut
Fédératif de Recherche Necker, and INSERM U129, Institut
Cochin de Génétique Moléculaire, Paris, France; and
Service des Maladies du Sang, Hôpital Claude Huriez, Lille,
France.
Platelets are formed from mature megakaryocytes (MKs)
and arise from the development of long and thin cytoplasmic extensions called proplatelets. After platelet release, the senescent MKs (nucleus
surrounded by some cytoplasm) undergo cell death by apoptosis. To
explore the precise role of apoptosis in proplatelet formation, we grew
human MKs from CD34+ cells and assessed the possible role
of caspases. Proteolytic maturation of procaspase-3 and procaspase-9
was detected by immunoblots in maturing MKs as well as in
proplatelet-bearing MKs and senescent MKs. Cleavage of caspase
substrates such as gelsolin or poly adenosine diphosphate (ADP)-ribose
polymerase (PARP) was also detected. Interestingly, activated forms of
caspase-3 were detected in maturing MKs, before proplatelet formation,
with a punctuate cytoplasmic distribution, whereas a diffuse staining
pattern was seen in senescent and apoptotic MKs. This localized
activation of caspase-3 was associated with a mitochondrial membrane
permeabilization as assessed by the release of cytochrome c, suggesting
an activation of the intrinsic pathway. Moreover, these MKs with
localized activated caspase-3 had no detectable DNA fragmentation. In
contrast, when apoptosis was induced by staurosporine, diffuse caspase
activation was seen; these MKs had signs of DNA fragmentation,
and no proplatelet formation occurred. The pan-caspase inhibitor
z-VAD.fmk as well as more specific inhibitors of caspase-3
and caspase-9 blocked proplatelet formation, whereas an inhibitor of
calpeptin had no effect. Overexpression of Bcl-2 also inhibited
proplatelet formation in maturing MKs. Thus, localized caspase
activation is causal to proplatelet formation. We conclude that
proplatelet formation is regulated by a caspase activation limited to
only some cellular compartments.
(Blood. 2002;100:1310-1317) Megakaryocytopoiesis is a unique process, which
leads to platelet production that has 2 unique
characteristics.1 First, the megakaryocyte (MK), the
direct platelet precursor, is a polyploid cell.2 This
polyploidization occurs by a process called endomitosis, corresponding
to nuclear endoreplication without cytokinesis.3,4 At the
end of polyploidization, MKs complete their cytoplasmic maturation to
finally shed platelets. Second, platelets are anucleate cells formed by
the fragmentation of the MK cytoplasm.5 Until recently,
platelet formation was poorly understood. It was first believed that
demarcation membranes, internal membranes of the MK, determined
platelet territories corresponding to future platelets, which would be
liberated via cytoplasmic fragmentation.6,7 However,
Radley and colleagues5,8 demonstrated that demarcation membranes were internal invaginations of the cytoplasmic membrane and
rather served as a reservoir of cytoplasmic membrane permitting the
extension of long pseudopods. Such cytoplasmic extensions contain all
the platelet organelles including mitochondria and have been termed
"proplatelets." Using in vitro cultures in presence of
thrombopoietin (TPO), it has now been clearly demonstrated that
platelets are released from proplatelets,9-11 confirming a
hypothesis originally formulated by Becker and De Bruyn.12 Proplatelet formation requires profound changes in the organization of
the cytoskeleton,10,11,13,14 which may be regulated by the
nuclear factor-erythroid 2 (NF-E2) transcription
factor.15,16
In vivo platelet formation does not occur directly in the marrow. MKs
either entirely migrate into the circulation17 or extrude
proplatelets into bone marrow sinusoids; these extensions are
subsequently fragmented into platelets. After release of platelets, the
MK nucleus, its envelope, and its adjacent cytoplasm usually remain in
the marrow and are subsequently phagocytosed by
macrophages.18 These denuded MKs have been called
senescent MKs, and Zauli et al19 have suggested that
senescent MKs correspond to apoptotic cells. However, the precise in
vitro relationship between MK apoptosis and the proplatelet formation
process is elusive. Intriguingly, in transgenic mice overexpressing
Bcl-2 under the control of a hematopoietic cell-specific promoter, a
2-fold reduction in platelet numbers was found, although MK numbers
remained unchanged.20 A similar observation was obtained
in mice with a homozygous deletion of the proapoptotic gene
Bim.21 Moreover, terminal differentiation associated with loss of the nucleus (as it occurs in lens cells and
keratinocytes and during erythropoiesis) might be regulated by
caspases, a class of proteases usually activated during
apoptosis.22-26 Caspase activation may result from the
ligation of plasma membrane receptors such as CD95 (the extrinsic
pathway) or from the activation of a cytosolic caspase activation
complex, the apoptosome (the intrinsic pathway).27 The
apoptosome is activated when cytochrome c (Cyt-c) is released from
mitochondria and gains access to cytosolic apoptotic
protease-activating factor 1 (Apaf-1), which in turn activates
procaspase-9. Therefore, the intrinsic pathway involves a critical
Bcl-2-controlled step of mitochondrial membrane permeabilization (MMP).28
In this paper, we report that proplatelet formation is the consequence
of a caspase-dependent mechanism that appears to be localized in the cytoplasm.
Reagents and antibodies
For indirect immunofluorescence, donkey tetramethyl rhodamine
(TRITC)-labeled antimouse, TRITC-labeled antigoat, or
FITC-labeled antirabbit F(ab') 2 fragments (Jackson ImmunoResearch,
West Grove, PA) were used.
z-VAD-fluoromethylketone (fmk) was obtained from Biomolecular Research
Laboratories (Plymouth Meeting, PA) and staurosporin (STS) from Sigma
Chemical (Saint Quentin Fallavier, France). The calpain inhibitor I
(Ac-Leu-Leu-norleucinal), the z-DEVD.fmk (caspase-3/7 inhibitor), and
the z-LEDH.fmk (caspase-9 inhibitor) were obtained from Alexis (San
Diego, CA).
In vitro generation of megakaryocytic cells
Cell staining protocols Cells were cytospun onto slides at 550 revolutions per minute (rpm) for 4 minutes and processed as previously reported.30 Briefly, cells were subsequently fixed in 2% paraformaldehyde (Serva, Heidelberg, Germany) for 15 minutes at room temperature (RT), permeabilized with 0.2% Triton X100 in phosphate-buffered saline (PBS) at RT for 5 minutes, and blocked with 2% fetal calf serum (FCS) prior to incubation with the anti-caspase-3a and anti-procaspase-3 or anti-VWF Abs. After 3 washes, cells were incubated with the appropriate secondary antibodies. DNA was then labeled with 4'6-diamidino-2-phenylindole (DAPI) in Vectashield (Vector, Burlingame, CA) or Hoechst 33342 (Sigma Chemical) and slides were mounted. In some experiments, cells were stained by PhiPhiLux-G1D2, green fluorescence, a caspase-3 substrate (Alexis), in accordance with the manufacturer's recommendations. Cells were subsequently washed, fixed with 0.5% paraformaldehyde for 15 minutes, treated by 0.1% Triton X-100 for 3 minutes, and incubated with the anti-VWF Ab, which was revealed by a TRITC-conjugated donkey-antimouse Ab. For simultaneous assessment of DNA fragmentation and caspase 3 activation, a double staining was performed using the anti-caspase-3a Ab and the terminal deoxynucleotide transferase-mediated dUTP nick-end labeling (TUNEL) technique. Briefly, after 1 hour's adherence on polylysine-coated slides at 37°C, MKs were fixed with 4% paraformaldehyde at RT. Cells were rehydrated with Tris buffer saline and then permeabilized by 0.2% Triton X-100 for 5 minutes and washed. The anti-caspase-3 staining was performed as above, followed by the TUNEL technique according to the manufacturer's recommendations (Oncogene).Flow cytometric analysis Cells were fixed with 1% paraformaldehyde and permeabilized by 0.1% Triton X-100 treatment (Sigma Chemical). Cells were washed, incubated with the R-PE anti-Bcl-2 mAb, washed again, and analyzed on a FACSort (Becton Dickinson).Conventional and confocal microscopy Conventional examination of samples was performed with a fluorescence microscope equipped with the appropriate filter combinations (Nikon, Tokyo, Japan). Confocal microscopy was performed on a Leica TC-SP (Leica Microsystems, Rueil Malmaison, France) equipped with an Argon Krypton laser and an Argon laser mounted on an inverted Leica DM IFBE microscope with a UV 100 × 1.4NA oil objective. To avoid cross-talk between different fluorochromes, images were acquired in a sequential fashion.Western blot analysis Cells were harvested at indicated times, washed with PBS, and lysed in Nonidet P-40 (NP-40) buffer. Protein concentration was determined with Bio-Rad protein assay (Biorad Laboratories, Hercules, CA). Equal amounts were boiled for 5 minutes in sodium dodecyl sulfate (SDS) sample buffer (40 mM Tris-HCl, pH 7.4, 5% glycerol, 5% mercaptoethanol, 2% SDS, 0.05% bromophenol blue) and subjected to SDS-polyacrylamide gel electrophoresis. The proteins were electrotransferred to nitrocellulose membranes (Hybond; Biorad) and blocked overnight at 4°C in PBS, 0.1% Tween, containing 5% nonfat dried milk. Blots were incubated with an anti-caspase-3 polyclonal Ab (CPP-32; Pharmingen), an anti-caspase-9 polyclonal Ab (Cell Signaling Technology), an antigelsolin mAb (Transduction Laboratories), and an anti-PARP (Oncogene) mAb and detected with appropriate secondary antibodies conjugated with horseradish peroxidase (Amersham Bioscience Europe, Orsay, France). Filters were developed with an enhanced chemiluminescence system (ECL kit; Amersham).Retroviral plasmid constructs The MFG-GFP plasmid was constructed by inserting a green fluorescent protein (GFP) insert (pIRES-EGFP; Clontech, Palo Alto, CA) into Nco1-BamH1 restriction sites of MFG backbone plasmid 27 under the control of the retroviral long terminal repeats (LTR) promoter. To construct the MFG-Bcl-2/internal ribosomal entry site (IRES)/GFP (MFG-BIG), 2 polymerase chain reaction (PCR)-amplified fragments containing, respectively, the 0.9 kilobase (kb) of the human Bcl-2 cDNA flanked by Nco1-EcoR1 restriction sites and the encephalomyocarditis virus (EMCV) IRES flanked by EcoR1-Nco1 restriction sites, were coinserted into the Nco1 site of the MFG-GFP plasmid. A Nco1 site generating a consensus Kozak sequence which replaced the original Bcl-2 ATG in order to strengthen the traduction.Generation and culture of producer lines The TELCeB6 packaging cell line (kindly provided by F. L. Cosset, Ecole Normale Supérieure, Villeurbanne, France) was cotransfected with the MFG and p respiratory syncytial virus glycoprotein G (pRSV) neoplasmids (10/1) using DOTAP (1,2-dioleoyl trimethylammonium propane), according to the manufacturer's guidelines (Gibco BRL, Invitrogen, Groningen, the Netherlands). After G418 selection (Gibco BRL; 1 mg/mL) and cloning, supernatants were collected and tested for their ability to transduce NIH3T3 cells. The best producer clone for each construct was identified by Southern blot. The titers obtained were 2.5 × 106 plaque-forming units (pfu)/mL for the MFG-BIG retrovirus and 5 × 106 pfu/mL for MFG-GFP retrovirus.MK infection CD34+ cells were cultured in the presence of PEG-rHuMGDF. At day 6, 3 × 105 cells were cocultured for 2 days at 37°C in 6-well plates containing subconfluent monolayers of either MFG-BIG or MFG-GFP virus-producing cells. Coculture medium consisted of -minimum essential medium (MEM) supplemented with 10%
FCS (Gibco BRL) in the presence of PEG-rHuMGDF. Nonadherent cells were
sorted 48 hours later after incubation with the R-PE anti-CD41a mAb.
Caspase inhibition blocks proplatelet formation Studies using animal models have suggested that caspase inhibition induces thrombocytopenia. We therefore hypothesized that a platelet release defect might be due to an inhibition of proplatelet formation. Indeed, in the absence of formation of such extensions, a decrease in platelet release is observed. We therefore cultured human CD34+ cells in a liquid serum-free culture system in the presence of TPO until day 7. At day 8, a purified MK population (CD34+/CD41a+) was sorted by flow cytometry and cultured in 96-well tissue culture plates to quantify proplatelet-displaying MKs.29 Cells were then cultured in the presence of TPO, with or without caspaselike inhibitors (Z-VAD.fmk, z-LEDH.fmk, z-DEVD.fmk) or an apoptosis inducer (STS). Control MKs had a round morphology until day 8, when some MKs began to change their form (Figure 1A) and harbored long extensions, corresponding to proplatelet formation from day 9 to day 14 (Figure 1B,C,E). Similar results were obtained for adult blood-derived MKs from day 10 to day 14 and for cord blood-derived MKs from day 12 to day 16, although the percentage of MKs exhibiting proplatelet formation was lower than that observed with peripheral blood and bone marrow MK cultures. The use of the pan-caspase inhibitor Z-VAD.fmk, over a dose range of 50 to 100 µM added at day 8 in the culture medium, led to an almost complete inhibition of proplatelet formation (Figure 1D-F). In contrast, when MKs had started to display proplatelets, the addition of Z-VAD.fmk had almost no effect on further proplatelet production. No differences were observed between control MKs and MKs cultured in the presence of Z-VAD.fmk in terms of viability, assessed by 7 amino-actinomycin D (7-AAD). The use of STS led to cell death and no proplatelet formation occurred. To further investigate the contribution of the 2 main caspases involved in the intrinsic pathway, we used the caspase-9 inhibitor z-LEDH.fmk and the caspase-3 inhibitor z-DEVD.fmk. Both these inhibitors reduced proplatelet formation, but to a slightly lesser extent than z-VAD.fmk (Figure 1E,F). In striking contrast, calpain inhibitor I had no significant effects on proplatelet formation (Figure 1F).
In a second set of experiments, we overexpressed Bcl-2 in MKs to
investigate whether Bcl-2, by its ability to inhibit MMP and
mitochondria-mediated caspase activation, could block platelet formation. CD34+ cells were cocultured with a TELCeB6
producer cell clone containing either the MFG-GFP (control) or MFG-BIG
vector. The latter cell line produces a bicistronic retrovirus
containing a bicistronic vector coding for Bcl-2 and GFP cDNAs
separated by an IRES. After 2 days of coculture, cells were sorted by
flow cytometry into CD41a+/GFP+ and
CD41a+/GFP
Caspase activation during MK maturation To demonstrate that caspases were effectively activated during proplatelet formation, we performed Western blots with Abs directed against caspase-9 on samples from marrow CD34+-derived MKs at different days of culture (Figure 3). At day 8 of culture, the anti-caspase-9 Ab recognized a 47-kd band, corresponding to unprocessed procaspase-9, and a 35-kd band, corresponding to the cleaved form (Figure 3A). The small cleavage fragment (10 kd) was not detected, suggesting that only a minor fraction of procaspase 9 was cleaved. In contrast, at day 12 (lane 2), presence of the 10-kd form was markedly increased, as it was also after STS treatment. At day 12 the proteolytic maturation of procaspase-9 was totally inhibited by Z-VAD.fmk. Procaspase-3 (32 kd) was found to be cleaved to yield the active form (17 kd) (Figure 3B), while in contrast, no processed caspase-3 was detected at earlier time points (day 6 or 8). As before, the proteolytic maturation of caspase-3 was totally inhibited by Z-VAD.fmk. At day 6 of the culture the caspase-3 substrate, gelsolin, was essentially found in its native form (93 kd) (Figure 4A), while at days 11 and 13 it was partially cleaved into a 46-kd form when the MKs were shedding platelets. This process was reversed by z-VAD.fmk and was increased following STS treatment. Similar results were obtained for PARP, another executioner caspase substrate (Figure 4B). PARP was essentially found in its native form (116 kd) at day 6 of culture. It was partially cleaved into an 86-kd apoptotic-related cleavage fragment when platelet-shedding MKs were present. Together, these experiments demonstrate a functional activation of caspase during MK maturation, before the terminal phase of its life span.
Early caspase activation occurs before proplatelet formation and is sequestered in the cytoplasm of maturing MKs To precisely determine the relationship between caspase activation and proplatelet formation, we investigated the precise kinetics of caspase activation at the cellular level. A confocal analysis revealed the presence of activated caspase-3 (caspase-3a) in a fraction of the day-8 round MKs, which continued to be detected until day 13 (Figure 5A,B). This labeling was granular with a variable number of patches, from 4 or 5 to 100. This labeling did not colocalize with the anti-VWF (Figure 5B) or the anti-CD63 staining (Figure 5D), suggesting that caspase-3a was not localized in-granules or in lysosomes. In contrast, a diffuse
caspase-3a staining was detected in senescent MKs (Figure 5C) or in MKs
induced to undergo apoptosis by STS. In MKs associated with a diffuse caspase-3 activation, VWF staining was weak in contrast to the MKs with
a localized activation of caspase-3 (compare Figures 5B and 5C). We
next questioned whether the procaspase-3 was present in the whole cell
or compartmentalized in the cytoplasm, using an anti-procaspase-3 Ab.
As expected, the proenzyme form of the procaspase-3 was present in the
whole cell, including proplatelets, with a diffuse staining (Figure
5E). We then quantified the number of MKs with a granular labeling with
an anti-caspase-3a Ab. Their number was low at day 7 (6%, n = 2)
and peaked between day 10 and day 12 (30%, n = 10), prior to
maximum platelet shedding. Caspase activation under a granular form was
increased in MKs shedding platelets (about 50%) in comparison with the
other round MKs. This suggests that localized activation of the
caspase-3 precedes proplatelet formation and persists during this
process. Similar experiments were performed with an antiactivated
caspase-9 antibody. Although labeling was much weaker, some MKs
exhibited the characteristic patchy appearance (data not
shown).
Early caspase activation is functional but does not lead to cell death To demonstrate a functional activation of caspase-3, we used a fluorescent substrate of caspase-3, PhiPhiLux. Cultured cells were stained with an anti-VWF Ab and PhiPhiLux. A fraction of mature MKs (about 10%) at day 10, including platelet-shedding MKs (Figure 6A-C), were labeled by the PhiPhiLux. None of the mature MKs exhibited clear signs of apoptosis, based on the chromatin condensation. Moreover, we performed double staining with an Ab specific to activated caspase-3, and DNA strand breaks revealed by the TUNEL reaction. Senescent MKs or STS-induced apoptotic MKs with diffuse caspase-3a exhibited DNA fragmentation (Figure 7A-C), while in contrast, the MKs with patchy and discrete activated caspase-3 retained their DNA integrity (Figure 7D-F).
Signs of mitochondrial membrane permeabilization (MMP) are present in MKs with a localized activation of caspase To confirm that caspase activation involves a critical Bcl-2-controlled step, we explored the intrinsic (mitochondrial) pathway of caspase activation. Cyt-c release was measured using double immunostaining with antibodies specific for Cyt-c (TRITC) and caspase-3a (Figure 8). In MKs having no detectable caspase-3 activation, staining with the anti-Cyt-c was strong and punctuate (Figure 8A). Cyt-c and Bcl-2 have the same pattern of labeling, suggesting that Cyt-c is mainly localized in mitochondria (data not shown). In contrast, MKs that exhibited a patchy activation of caspase-3 had a much weaker and diffuse staining, suggesting that Cyt-c is released from mitochondria and gains access into the cytosol (Figure 8B). A similar observation was made in MKs with a diffuse caspase-3 activation and nuclear signs of apoptosis, although Cyt-c expression was at the threshold of detection.
Taken together, these experiments clearly demonstrate an activation of caspase preceding proplatelet formation. However, active forms of caspases were specifically localized in granules; this compartmentalization was lost in apoptotic MKs. Sequestration of caspase activation may lead to regional cell death (sparing proplatelets), while extended activation would lead to death of the entire cell. Signs of apoptosis are also compartmentalized on the plasma membrane surface of MKs shedding platelets Exposure of phosphatidylserine residues on the plasma membrane surface (determined with annexin V) was not detectable in maturing MKs, except in MKs exhibiting features of apoptosis (Figure 9). In proplatelet-displaying MKs, surface phosphatidylserine was detected on the cell membrane except along the proplatelets. Proplatelets disrupted from MKs were also negative for surface phosphatidylserine.
Platelet release depends on the formation of MK cytoplasmic extensions called proplatelets. In the absence of such extensions, only a few, if any, platelets are released.16 The appearance of proplatelets delineates 2 main regions in mature MKs, with distinct fates: proplatelets, which will engender functional platelets in the blood flow,5,10,11 and the central area of the MK surrounding the nucleus, which will give rise to senescent MKs. This denuded MK remains in the bone marrow, undergoes an apoptotic process, and is phagocytosed by macrophages.18,19 Previous studies have suggested that at the terminal phase of the MK life span, platelet production and MK apoptosis are closely related events.19 Moreover, nitric oxide-induced apoptosis facilitates platelet production during the final stages of megakaryocytopoiesis.31 Here, we provide evidence that activation of the apoptotic cell machinery is directly involved in differentiation and platelet shedding: its activation is necessary not only for cell death but also for the formation of proplatelets. The first line of evidence that platelet formation has some similarities to apoptosis is that proplatelet formation was associated with a cleavage of caspase-9 and caspase-3 in MKs derived from cord blood, mobilized blood, or bone marrow CD34+ cells. Procaspase-3 cleavage gave rise to the enzymatically active caspase-3 fragment because gelsolin and PARP were cleaved when MKs were shedding platelets. The second line of evidence is that the pan-caspase inhibitor Z-VAD.fmk could inhibit proplatelet formation. This process was temporally controlled. Indeed, once proplatelet formation had been initiated, Z-VAD.fmk had few, if any, effects. However, Z-VAD.fmk is also capable of inhibiting calpain, which has recently been reported to promote apoptosislike events during platelet activation.32-34 However, proplatelet formation is not a calpain-dependent process, because a calpain inhibitor had no inhibitory effect. Furthermore, overexpression of Bcl-2 in CD41+ cells largely blocked proplatelet formation. This result also supports the contention that the thrombocytopenia observed in Bcl-2 transgenic mice is related to an inhibition of platelet production.20 A surprising finding was related to the granular distribution of
caspase-3a in the cytoplasm of mature MKs. Staining with Abs against
caspase-3a revealed a discrete punctuate distribution, which did not
colocalize with VWF, a marker of The mechanism of proplatelet formation may mimic the blebbing that is observed at the onset of apoptosis.37,38 However, in contrast to blebbing, which is coupled to whole-cell death, platelet shedding is associated with regional cell death. Proplatelet formation is related to profound changes in the cytoskeleton, including activation of microtubules,5,10,13,39 polymerization of actin, and phosphorylation of myosin.11,14,40,41 Several cytoskeleton molecules or actin regulators, such as gelsolin, regulators of Rho family guanosine triphosphatases (GTPases), or their effectors, have been described as substrates of caspases during apoptosis.42-48 In particular, it has been shown that Rho-kinase cleavage by caspase-3 is responsible for apoptotic membrane blebbing by inhibiting myosin phosphatase.49-51 Thus, it seems likely that caspase activation during proplatelet formation may lead to alteration in the actin cytoskeleton and myosin function, triggering a complex cytoplasm reorganization which, in contrast to blebbing, also involves cytoplasmic organelles. At present, we cannot exclude the possibility that some nonexecutioner caspases are also involved in this process. For example, it has been recently shown that caspase-14 is associated with keratinocyte terminal differentiation.25 From these experiments it can be hypothesized that a compartmentalization of caspase activation occurs during MK differentiation, which leads to proplatelet formation. This compartmentalization may affect not only the cytoplasm but also the plasma membrane. Indeed, phosphatidylserine exposure was detectable only on the cytoplasmic membrane around the nucleus, not on proplatelets (data not shown). Expression of phosphatidylserine on the external plasma membrane triggers phagocytosis,52 in agreement with the fate of the senescent MKs.18 After proplatelet formation and platelet shedding, a complete activation of the apoptotic machinery subsequently leads to apoptosis in senescent MKs. Overall, this is reminiscent of the function of caspases during terminal erythroid differentiation26 and in the enucleation of other cell types.22,23 However, there is also increasing evidence that caspase-3 may play an important role in terminal differentiation of cells that remain nucleated53 (also Sordet et al, manuscript submitted for publication). In conclusion, limited caspase activation is directly implicated in proplatelet formation. Characterization of the molecules that regulate this process will provide new insights into the mechanisms linking differentiation and apoptosis and may allow us to better understand the mechanisms of congenital or acquired thrombocytopenia.
We thank Héléne Gilgenkrantz (U129, Institut Cochin de Génétique Moléculaire, Paris, France) for providing the MFG-BIG and MFG-GFP virus-producing cells; Valérie Schiavon (U362, Villejuif, France) for cell sorting experiments; Didier Métivier (CNRS, Unité Mixte de Recherche 1599, Villejuif) for confocal analysis; Catherine Boccacio (Institut Gustave Roussy, Villejuif) for providing cytapheresis samples; and Jean Luc Pallacio and Didier Letailleur (Beauvais, France) for providing bone marrow samples. We are grateful to Pierre Golstein (Centre d'Immunologie de Marseille-Luminy, Marseille, France), for discussion and critical reading of the manuscript and to Dr Jonathan Dando for greatly improving the English.
Submitted March 5, 2002; accepted April 9, 2002.
Prepublished online as Blood First Edition Paper, June 7, 2002; DOI 10.1182/blood-2002-03-0686.
Supported by grants from INSERM, the Association pour la Recherche contre le Cancer (N.D.), la Ligue Nationale contre le Cancer (W.V. and G.K.), and the European Commission (G.K.).
E.D. and Y.Z. contributed equally to this study.
The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked "advertisement" in accordance with 18 U.S.C. section 1734.
Reprints: Najet Debili, INSERM U362, Institut Gustave Roussy, Institut Fédératif de Recherche 54, F-94805, Villejuif, France; e-mail: denali{at}igr.fr.
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Top
Abstract
Introduction
populations and were cultured in
96-well tissue culture plates in the presence of TPO. The efficacy of
Bcl-2 expression was assessed by flow cytometry and a correlation was
confirmed between Bcl-2 and GFP expression (Figure
analysis by membrane tracer and freeze-fracture studies on fresh human and cultured mouse megakaryocytes.
J Cell Biol.
1984;99:390-402