|
|
|
|
|
|
|
||
Articles |
James Ewing Laboratory of Developmental Hematopoiesis, Memorial Sloan-Kettering Cancer Center, New York 10021; the Department of Immunology, Berlex Biosciences, Richmond, California 94804; the || Department of Medicine II, University of Tübingen, Tübingen 72076, Germany; and the ¶ Division of Hematology-Oncology, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania 19104
 |
Abstract |
|---|
 Top Abstract Materials and MethodsResultsDiscussionReferences |
|---|
Key Words: megakaryocyte • endothelium • chemotaxis • stromal cell–derived factor 1 • E-selectin
The development of megakaryocytes (MKs)1 can be divided into two distinct stages or compartments. The initial stage of MK development involves sequential proliferation of CD34+ hematopoietic stem cells into rapidly proliferating megakaryoblasts. The second phase of MK development is comprised of a population of morphologically identifiable MKs that undergo nuclear endoreplication and ultimately fragment into functional platelets.
Thrombopoietin (TPO) has been shown not only to regulate MK proliferation, maturation, and endoreplication (1, 2), but also to induce platelet formation in vitro (3–5). However, whether the process of platelet formation in vivo is also mediated by TPO, alone or in combination with other factors, is not known (6). Recently it has been shown that NF-E2 (a hematopoietic specific transcription factor of erythroid cells and MKs) knockout mice, despite physiological levels of TPO and an increased level of MKs, have profound reduction in platelet counts (7, 8), suggesting that final stages of platelet formation and release may be regulated by as yet undefined cellular signaling pathways. Furthermore, bone marrow stromal cells have been shown to inhibit platelet release, suggesting that MKs may have to exit the bone marrow microenvironment to release platelets (9).
Several studies have shown that mature polyploid MKs reside in close proximity to bone marrow endothelial cells (BMEC; references 10 and 11). Adherence of MKs to the bone marrow stromal cell extracellular matrix induces extension of pseudopodia and final fragmentation into platelet-like particles (12–16). MKs also express certain adhesion molecules that may enable them to interact with resting as well as cytokine-activated endothelial cells (17, 18). Electron microscopic analysis of bone marrow biopsy samples has shown that MKs residing at the subluminal surface of the BMEC are capable of transmigrating through the BMEC junctions (14, 19). Several lines of evidence suggest that MKs generated in the bone marrow also have the capacity to travel to the lungs, where MKs fragment into platelets (20–22). Collectively, these data suggest that one mechanism whereby mature MKs release platelets is transmigration through BMEC—releasing platelets in the sinusoidal space or by stochastic fragmentation in the lung capillaries.
To date, very few reports have explored the effect of chemokines on MK trafficking. Chemokines such as platelet factor 4 (PF4) and neutrophil-activating peptide 2 (NAP-2) have been shown to have an inhibitory effect on MK development (23–25). However, there has been no report of a chemokine that may induce transmigration of MKs. In search for factors that may mediate transendothelial migration of MKs, we have found that mature polyploid MKs express the HIV coreceptor CXCR4. Stromal cell–derived factor 1 (SDF-1), the chemokine ligand for CXCR4, has been shown to induce transmigration of lymphocytes (26, 27) and CD34+ cells (28, 29). In addition, mice lacking SDF-1 have profound defects in hematopoiesis (30). In this report, we demonstrate that SDF-1, the ligand for CXCR4, increases the affinity of mature MKs for BMEC and induces transmigration of MKs through BMEC monolayers, resulting in augmentation of platelet release. These data suggest that expression of CXCR4 by mature MKs may be one of the major signaling pathways that may induce transmigration of MKs, thus initiating the process of platelet formation.
MK Preparation.
Transmigration Assay.
For migration inhibition studies, MKs were preincubated with 40 µg/ml of mAb to CXCR4 (12G5) for 30 min or with 2.5 µg/ml pertussis toxin in Xvivo-20 for 90 min, washed once with HBSS, then placed in bare transwells as described above. For chemotactic versus chemokinetic analysis of SDF-1 function, MKs were suspended in medium containing the same concentration of SDF-1 as the lower chamber, and placed in bare transwells as described above. To examine the nature of the interaction between BMEC and MKs, transmigration through IL-1β–activated BMEC was inhibited by preincubation with 40 µg/ml of anti–E-selectin (CD62E), anti–vascular cell adhesion molecule 1 (VCAM-1) or intercellular adhesion molecule 1 (ICAM-1) mAb, alone or in combination, in the upper and lower chambers for 15 min. BMEC monolayers were stimulated with 10 U/ml of IL-1β for 12–16 h. After removal of the IL-1β, the stimulated BMEC monolayers were incubated with MKs.
Flow Cytometry.
Cell Counts and Cytospin Preparation.
SDF-1 Binding to MKs.
Cell Cycle Analysis of MKs.
Antibodies and Reagents.
Electron Microscopy.
Statistical Analysis.
In the first few days of MK expansion, the number of CD41a+CXCR4+ cells comprised a very small percentage of the total CD41a+ population. However, by days 12–16 of expansion, the majority of CD41a+ cells were CXCR4+ (n = 5; Fig. 1 A). Moreover, two-color flow cytometry with CXCR4-PE and CD42b-FITC revealed that all mature CD42b+ cells expressed CXCR4 (n = 5; Fig. 1 B). These data suggest that CXCR4 is expressed on MKs, and that its expression is enhanced with maturation of MKs. The majority of the CD41a and CD42b–negative population was comprised of immature CD41a+ MKs and CD15+ myeloid progenitor cells.
Materials and Methods
Top
Abstract
Materials and Methods
Results
Discussion
References
Preparation of BMEC Monolayer.
Primary BMEC were isolated by a standard method as described previously (31). For transmigration assays, 5-µm pore transwells covered with confluent monolayers of BMEC were prepared as described previously (32). Confluent monolayers of BMEC were cultured on bare transwells in medium 199 (M199; GIBCO BRL, Gaithersburg, MD) with 20% fetal bovine serum (FBS; Hyclone, Logan, UT), or with 20% FBS, 5 ng/ml basic fetal growth factor (bFGF), and 50 ng/ml heparin. Before transmigration assay, the integrity of the monolayer was confirmed by placing 100 µl medium 199/ 20% FBS containing [14C]albumin (American Radiolabeled Chemicals, St. Louis, MO) on the cells for 6 h and measuring the amount of radioactivity accumulating in the lower chamber (32).
Mononuclear cells derived from human cord blood (CB) or G-CSF–mobilized peripheral blood leukopheresis were separated from whole blood obtained from consenting donors, by Ficoll-Paque (d = 1.077 g/ml; Pharmacia Biotech AB, Uppsala, Sweden). CD34+ progenitor cells were purified from mononuclear cell preparations using magnetic cell sorting Dyna-beads (Dynal A.S., Oslo, Norway). CD34+ cells derived from CB were used for ex vivo expansion of MKs for the transmigration studies. MKs were ex vivo expanded with TPO (100 ng/ml) and Kit ligand (KL, 10 ng/ml) from CD34+ cells in Xvivo-20 (BioWhittaker, Walkersville, MD) medium for 14–16 d. The number of CD41a+ and CD42b+ cells on day 12, generated from ex vivo expansion of CB, was 57 ± 10 and 36 ± 4% of total expanding cells, respectively.
Ex vivo-expanded MKs were washed once with HBSS solution and resuspended with fresh Xvivo-20 to a density of 106 cells/ml. Aliquots of the MK cell suspension (100 µl) were applied on 5-µm pore transwells, covered with or without confluent BMEC monolayer, in a 24-well plate (Boyden chamber; Costar Corp., Cambridge, MA [32]). Immediately, 600 µl of serum-free chemotactic medium, containing Xvivo-20 with or without various concentrations of chemokines, or MS5-conditioned medium prepared in Xvivo-20, was then placed in the lower chamber. After 3 h of incubation in bare transwells or 24 h in BMEC-covered transwells at 37°C in a CO2 incubator, MKs floating in the transwell, attached to BMEC, or transmigrated to the lower chamber were manually counted with a hemacytometer, and the phenotype was analyzed by two-color flow cytometry.
A total of 104–105 cells were incubated for 30 min at 4°C with the FITC- or PE-conjugated mAb CD41a-FITC, CD42b-FITC, or CXCR4-PE (clones HIP8, HIP1, and 12G5; PharMingen, San Diego, CA). Isotype-matched antibodies served as controls (IgG1 and IgG2a, FITC- or PE-conjugated; PharMingen). BMEC detached by collagenase or MKs grown in suspension were harvested by centrifugation and washed twice with PBS (pH 7.4). The cells were analyzed using an Elite flow cytometer (Coulter Corp., Hialeah, FL). For coexpression analysis, an FL-1/FL-2 contour plot was used to calculate the percentage of positive cells; a proportion of 1% false positive events was accepted in the negative control sample. The mean fluorescence intensity was calculated from the fluorescence histogram and expressed in arbitrary units.
Cell numbers and concentrations were assessed using a hemacytometer or automated cell counter (Coulter Corp.). Viability of cells, always >95%, was confirmed manually with trypan blue, or by flow cytometry with propidium iodide staining. Standard cytospin preparations were stained with Wright-Giemsa. A differential count of at least 100 cells was performed for each cytospin preparation.
Ex vivo–expanded MK cells derived from CB (6.3 x 106 cells per 200 ml) were incubated in PBS with 125I-labeled SDF-1 in the presence and absence of various concentrations of unlabeled SDF-1 at room temperature for 30 min. The incubation was terminated by removing aliquots from the cell suspension and separating cells from buffer by centrifugation through a silicone/paraffin oil mixture. Nonspecific binding was determined in the presence of 1 mM unlabeled ligand. The binding data were curve-fitted with the computer program IGOR (WaveMetrics, Inc., Lake Oswego, OR) to determine binding affinity (Kd), and Scatchard analysis was used for number of sites per cell.
Ploidy of nonmigrated and migrated CD41a+ MKs was analyzed by propidium iodide. Migrated and nonmigrated MKs were stained with CD41a-FITC and fixed overnight in 70% ethanol at 4°C. After staining with 50 µg/ml propidium iodide containing 100 U/ml RNase A in PBS (pH 7.4) Ca2+Mg2+, the MKs were analyzed by flow cytometry focusing on the FITC-positive gate.
The chemokines IL-8, monocyte chemotactic protein (MCP)-1, macrophage-inflammatory protein (MIP)-1
and β, thymus and activation–regulated chemokine (TARC), MIP-3 (liver and activation–regulated chemokine [LARC]), MIP-3β (EBV-induced molecule 1 [EBI-1] ligand chemokine [ELC]), SDF-1, and Eotaxin were a gift from Berlex BioSciences, or were purchased from PeproTech (Rocky Hill, NJ). Radiolabeled SDF-1, 2,200 Ci/mmol, was from NEN Research Products (Boston, MA). The mAb to CXCR4 (12G5) was as described (33). Pertussis toxin, heparin, propidium iodide, and RNase A were from Sigma Chemical Co. (St. Louis, MO). mAbs to E-selectin, VCAM-1, and ICAM-1 were from R&D Systems. The MS5 stromal cell line, a rich source of SDF-1, was obtained by incubating MS5 with serum-free medium Xvivo-20 for 48 h.
Electron microscopy was performed on the migrated MKs and platelets formed in the lower chamber of the transwell plates. Immediately and 24 h after migration of the MKs in response to SDF-1, the MKs and generated platelets were fixed immediately in 2% paraformaldehyde, 0.5% glutaraldehyde. After dehydration, the samples were embedded for electron microscopy. 60–70-nm (Silver grade) sections were cut using a diamond knife (Diatome, Fort Washington, PA). Sections were stained with 0.1% lead citrate and examined using an electron microscope (model 100CXII; JEOL, Ltd., Tokyo, Japan) at an accelerating voltage of 80 kV.
Data are expressed as mean ± SEM of four to five independent experiments. To detect differences between migrating and nonmigrating cells, the t test for paired samples was applied. A P value <0.05 was considered statistically significant.
Results
Top
Abstract
Materials and Methods
Results
Discussion
References
CXCR4 Is Expressed on Mature MKs.
Human MKs derived from CB were ex vivo expanded with TPO (100 ng/ ml) and KL (10 ng/ml). The time course of expression of CXCR4 on ex vivo–expanded MKs was measured by two-color flow cytometry using PE-conjugated mAb to CXCR4 (CXCR4-PE, 12G5) and FITC-conjugated mAb to various specific MK markers, including GPIIb/IIIa (CD41a-FITC) and GPIb (CD42b-FITC). CD41a is expressed on early megakaryoblasts, and its expression persists with maturation of MKs. CD42b is another specific MK marker which is mainly expressed on the mature polyploid MKs.
|
|
(CCR1 ligand), MIP-1β, RANTES (regulated upon activation, normal T cell expressed and secreted, CCR5 ligand), MCP-1 (CCR2 ligand), Eotaxin (CCR3 ligand), TARC (CCR4 ligand), MIP-3 (LARC, CCR6 ligand), MIP-3β (ELC, CCR7 ligand), failed to induce migration of MKs (Fig. 2 A). We and others have shown that MKs express vascular endothelial growth factor (VEGF)R1 (fms-like tyrosine kinase 1 [FLT-1]; reference 34) and c-kit receptor (35). Both VEGF and KL have been shown to induce chemotaxis and chemokinesis of certain hematopoietic cells (36). However, KL failed to induce migration of CD41a+CXCR4+ MKs (Fig. 2 A). Furthermore, VEGF failed to induce migration of MKs (data not shown). In addition, replacement of TPO with all other known megakaryopoietins, including IL-6, IL-11, bFGF, and PF4, failed to induce migration of MKs. These data suggest that among the numerous known chemokines and megakaryopoietins, only SDF-1 has a substantial effect on the migration of MKs. Preincubation of MK cells with the blocking CXCR4 mAb (12G5) was able to block SDF-1–induced chemotaxis (n = 4, P <0.05; Fig. 2 B). 12G5 mAb was also able to block chemotaxis induced by SDF-1 containing conditioned medium from the stromal cell line MS5 (data not shown). Addition of the G-coupled protein inhibitor pertussis toxin (2.5 µg/ml) completely abrogated the migration of MKs (n = 4, P <0.05; Fig. 2 B). Neutralization of the SDF-1 gradient by adding SDF-1 (200 ng/ml) to both upper and lower chambers resulted in inhibition of migration of MKs, indicating that SDF-1's predominant action on MKs is chemotactic rather than chemokinetic (n = 5). Decrease in the concentration of SDF-1 to 50 ng/ml in the upper chamber while keeping the SDF-1 concentration constant in the lower chamber (200 ng/ml) resulted in recovery of 18 ± 5% of SDF-1 chemotactic activity (n = 3). These results suggest that the physiological activity of SDF-1 may depend on the generation of a gradient secretion of the chemokine within the bone marrow microenvironment.
Since the extracellular matrix within the bone marrow microenvironment is enriched with heparinoid molecules, we explored the possibility that heparin-bound SDF-1 within the bone marrow may be less functionally active. Introduction of heparin significantly inhibited migration of MKs (n = 4, P <0.05; Fig. 2 B), suggesting that binding of SDF-1 to heparin reduces SDF-1 activity, creating a functional gradient within the bone marrow microenvironment.
The ED50 for migration of CD41a+ MKs in response to SDF-1 is 
125 pmol/liter (n = 3; Fig. 2 C). To determine the relative amount of CXCR4 receptors expressed on the surface of these cells, we performed SDF-1 ligand binding studies on TPO/KL ex vivo–expanded immunomagnetic purified MK cells. 125I-labeled SDF-1 displacement binding studies revealed a dissociation constant (Kd) for SDF-1 binding to MKs of 17 ± 5 nM (data not shown). By Scatchard analysis of the SDF-1 binding data, 2.14 x 105 receptors per cell were estimated. Collectively, the binding and chemotaxis data suggest that SDF-1 is a potent chemokine for mature MKs.
SDF-1–induced Transmigration of MKs through BMEC.
To explore the possibility that interaction between the BMEC and mature MKs in response to SDF-1 may be critical for MK migration, ex vivo–expanded MKs were placed in transwell plates coated with confluent monolayers of BMEC. Subsequently, SDF-1 at 200 ng/ml was added to the lower chamber. In the absence of SDF-1, there was no migration or increased adhesion of MKs to BMEC. However, in the presence of SDF-1, 14 ± 3% of MKs migrated through BMEC (n = 5, P <0.05; Fig. 3 A). Furthermore, in the presence of SDF-1, there was a significant increase in the adhesion of MKs to BMEC monolayers (n = 5, P <0.05; Fig. 3 A).
|
SDF-1 Induce Transendothelial Migration of Polyploid Mature MKs.
Migration of MKs through BMEC was associated with profound morphological changes in MKs, including unilateral pseudopodia formation (Fig. 4 A). Light microscopy of the transmigrated Wright/Giemsa-stained MKs showed a predominance of intact polyploid MKs, suggesting that polyploid MKs preferentially express CXCR4 and migrate in response to SDF-1. Ploidy analysis of transmigrating MKs in response to SDF-1 (200 ng/ml) through the 5-µm pore showed preferential migration of polyploid (>2, 4, 8, and 16N; n = 4, P <0.05) MKs (Fig. 4).
|
|
Electron microscopy of the cells in the lower chamber of the transwells recovered 3 h after transmigration in response to SDF-1 showed the presence of large polyploid MKs (Fig. 6 A), demonstrating that SDF-1 induces the transendothelial migration of whole intact MKs. In addition, electron microscopy of cells in the lower chamber 12 h after transendothelial migration demonstrated the presence of MKs in the process of fragmentation into platelets (Fig. 6 B). These data demonstrate that SDF-1 induces rapid migration (3–5 h) of intact polyploid MKs through BMEC, followed by fragmentation of MKs into platelets within 12–24 h after migration.
|
|
Discussion |
|---|
|
TopAbstractMaterials and MethodsResultsDiscussionReferences |
|---|
and β, and IL-8. However, among the known factors that may potentially influence MK function and induce chemotaxis, including TPO, VEGF, KL, IL-6, IL-11, and myriad other CXC or CC chemokines, only SDF-1 induced transmigration of MKs. The majority of the transmigrated MKs were large polyploid mature CD41a+CD42b+ MKs, suggesting that expression of CXCR4 by mature MKs may initiate cellular signaling pathways that in turn initiate the process of migration and platelet formation. Interaction of MKs with BMEC is critical, since adhesion molecules expressed on the BMEC monolayers seem to play a crucial role in regulating the migration of MKs through endothelial cell junctions. Many studies have shown that BMEC in vivo assume the phenotype of activated endothelium and constitutively express low levels of VCAM-1 and E-selectin (37). In addition, mature MKs are capable of producing physiological levels of stimulatory cytokines, including IL-1β (38). In this regard, in order to mimic the in vivo phenotype of BMEC, BMEC monolayers were activated with IL-1β. Activation of BMEC results in the upregulation of adhesion molecules, including E-selectin, VCAM-1, and ICAM-1. Activation of BMEC with IL-1β not only increased the number of BMEC-bound MKs, it also significantly increased the number of migrating MKs. In addition, migration of MKs through activated BMEC was partially blocked by neutralizing mAb to E-selectin, but not to VCAM-1 or ICAM-1. This finding demonstrates that migration of MKs is not a random process, and involves sequential interaction of mature MKs with specific adhesion molecules on endothelial cells. These series of events may facilitate egress of MKs and initiate platelet release.
Blocking mAb to VCAM-1 and ICAM-1 did not significantly block the migration of MKs. Furthermore, a combination of neutralizing mAbs to VCAM-1, ICAM-1, and E-selectin did not block the migration of MKs beyond that originally blocked by E-selectin mAb. E-selectin expressed by activated endothelial cells seems to play a role in the tethering of leukocytes. In addition, engagement of E-selectin with its ligand has been shown to result in inside-out signaling (39, 40). Therefore, it is possible that engagement of E-selectin results in activation of signaling pathways that mobilize MKs for migration.
Based on the data presented in this paper, 75% of day 12 to day 14–expanded MKs express CXCR4. IL-1β stimulation of BMEC monolayers induces migration of up to 45 ± 5% of the MKs. However, since close to 25% of CXCR4+ MKs do not migrate, it is possible that either another factor is needed for migration or not all CXCR4+ MKs express the appropriate adhesion molecules. Therefore, based on these data we can infer that the mere functional expression of CXCR4 on MKs is not sufficient for migration, but that attachment through the appropriate adhesion molecules or stimulation by an as yet unrecognized chemokine is critical for transendothelial migration.
Although transmigration of MKs through bare transwells or BMEC was rapid (3–5 h), platelet generation took place maximally 24–48 h after migration. However, compared with bare transwell plates, migration of MKs through BMEC resulted in higher numbers of functional platelets, suggesting that collective interaction of MKs with BMEC and SDF-1 is critical for optimal platelet production.
However, both flow cytometric and electron microscopic analyses of transmigrated MKs demonstrated that even after 48 h, close to 20% of MKs remained intact, without any indication of generating platelets. These data suggest that CXCR4 may be expressed on polyploid MKs at different stages of maturation. However, the more mature MKs will give rise to platelets immediately after migration, whereas the less mature may migrate and give rise to platelets after the process of platelet compartmentalization has been completed. Therefore, it is conceivable that these relatively immature MKs, not having enough time within the sinusoidal space to release platelets, may travel to the lungs and stochastically release platelets in the lung capillaries.
SDF-1 induces a significant increase in the adhesion of mature MKs to resting or activated endothelial cells, suggesting that SDF-1 may either modulate the adhesion molecule repertoire in the MKs, or indirectly modulate endothelial cell adhesion molecule expression. Recently it has been shown that endothelial cells also express CXCR4 (41). However, to date we have been unable to demonstrate that SDF-1 has any effect on the adhesion molecule repertoire of BMEC. Therefore, it is likely that SDF-1 may directly regulate adhesion molecule repertoire on MKs.
CXCR4 is a coreceptor for T-tropic HIV strains, and has been shown to mediate HIV entry into T cells in the presence of CD4 (42, 43). It has also been demonstrated that the mAb (12G5) directed to CXCR4 can inhibit HIV entry into cells (33, 44, 45). Furthermore, some subsets of patients with HIV infection have been shown to have profound thrombocytopenia on initial clinical presentation. Soluble gp120 from T-tropic HIV has also been shown recently to induce apoptosis in human neurons or to be chemotactic for T cells (J. Hesselgesser, unpublished data). Based on the data presented here, it is possible that thrombocytopenia in patients with HIV may be due to direct entry of HIV into MKs via CXCR4. It also conceivable that direct interaction of soluble gp120 with CXCR4 on MK cells could induce apoptosis. This may result in dysregulation of chemotaxis of MKs and disruption of platelet production.
Based on our results, in order for SDF-1 to induce migration of mature MKs in vivo, SDF-1 concentration has to be much higher within the sinusoidal space than in the bone marrow microenvironment. In fact, most studies have shown that SDF-1 is produced by bone marrow stromal cells, including bone marrow fibroblasts such as the MS5 murine fibroblastic cell line (26). Given that heparin significantly blocked SDF-1 activity, it is conceivable that SDF-1 produced within the bone marrow microenvironment, particularly in the vicinity of the BMEC, is functionally inactive. Therefore, assuming that SDF-1 within the sinusoidal space may exist in an unbound form, this could produce a functional gradient resulting in selective migration of MKs from the bone marrow microenvironment to the sinusoidal space.
On average, on day 12 of ex vivo expansion, 57 ± 5% of cells derived from expansion of CB CD34+ cells in the presence of TPO and KL were MKs expressing CD41a+ cells. We have tried to purify MKs from ex vivo–expanded cells by flow cytometric cell sorting or elutriation. However, given the fragile nature of polyploid MKs, any manipulation of ex vivo–expanded MKs results in significant loss of mature polyploid MKs. In addition, we have found that in contrast to CD34+, T, or dendritic cells, minor physical manipulation of MKs by multiple centrifugation results in random disintegration into platelet-like particles, nonspecific calcium fluxes, and loss of response to SDF-1. Therefore, in the experiments described in this paper, we have used day 12 to 14 ex vivo–expanded MKs with minimal physical manipulation. This step has remarkably enhanced the yield of intact viable MKs that can effectively be used in migration studies.
Our data suggest that MKs may release platelets during transmigration through BMEC or after exiting through BMEC. SDF-1 induces transendothelial migration of MKs, suggesting that MKs have the capacity to release platelets during transmigration or after exiting the bone marrow. Therefore, expression of CXCR4 by mature MKs may be the critical cellular signal for transmigration of MKs and platelet formation. The cellular signaling pathways activated as the result of transendothelial MK migration that may mediate platelet formation are the subject of ongoing experiments.
|
Acknowledgments |
|---|
Submitted: 4 March 1998
Revised: 5 May 1998
1 Abbreviations used in this paper: BMEC, bone marrow endothelial cell(s); CB, cord blood; ELC, EBV-induced molecule 1 (EBI-1) ligand chemokine; FBS, fetal bovine serum; bFGF, basic fibroblast growth factor; ICAM-1, intracellular adhesion molecule 1; KL, Kit ligand; LARC, liver and activation–regulated chemokine; MCP, monocyte chemotactic protein; MIP, macrophage-inflammatory protein; MK, megakaryocyte; NAP-2, neutrophil-activating peptide 2; PF4, platelet factor 4; RANTES, regulated upon activation, normal T cell expressed and secreted; SDF-1, stromal cell–derived factor 1; TARC, thymus and activation–regulated chemokine; TPO, thrombopoietin; VCAM-1, vascular cell adhesion molecule 1; VEGF, vascular endothelial growth factor.
|
References |
|---|
|
TopAbstractMaterials and MethodsResultsDiscussionReferences |
|---|
1 Kaushansky K, Lok S, Holly RD, Broudy VC, Lin N, Bailey MC, Forstrom JW, Buddle MM, Oort PJ, Hagen FS et al.. Promotion of megakaryocyte progenitor expansion and differentiation by the c-Mpl ligand thrombopoietin, Nature, 1994, 369, 568–571.[Medline]
2 Wendling F, Maraskovsky E, Debili N, Florindo C, Teepe M, Titeux M, Methia N, Breton-Gorius J, Cosman D & Vainchenker W. cMpl ligand is a humoral regulator of megakaryocytopoiesis, Nature, 1994, 369, 571–574.[Medline]
3 Zucker-Franklin D & Kaushansky K. Effect of thrombopoietin on the development of megakaryocytes and platelets: an ultrastructural analysis, Blood, 1996, 88, 1632–1638.
4 Kaushansky K, Broudy VC, Lin N, Jorgensen MJ, McCarty J, Fox N, Zucker-Franklin D & Lofton-Day C. Thrombopoietin, the Mp1 ligand, is essential for full megakaryocyte development, Proc Natl Acad Sci USA, 1995, 92, 3234–3238.
5 de Sauvage FJ, Carver-Moore K, Luoh SM, Ryan A, Dowd M, Eaton DL & Moore MW. Physiological regulation of early and late stages of megakaryocytopoiesis by thrombopoietin, J Exp Med, 1996, 183, 651–656.
6 Bunting S, Widmer R, Lipari T, Rangell L, Steinmetz H, Carver-Moore K, Moore MW, Keller GA & de Sauvage FJ. Normal platelets and megakaryocytes are produced in vivo in the absence of thrombopoietin, Blood, 1997, 90, 3423–3429.
7 Shivdasani RA, Rosenblatt MF, Zucker-Franklin D, Jackson CW, Hunt P, Saris CJ & Orkin SH. Transcription factor NF-E2 is required for platelet formation independent of the actions of thrombopoietin/MGDF in megakaryocyte development, Cell, 1995, 81, 695–704.[Medline]
8 Shivdasani RA, Fielder P, Keller GA, Orkin SH & de Sauvage FJ. Regulation of the serum concentration of thrombopoietin in thrombocytopenic NF-E2 knockout mice, Blood, 1997, 90, 1821–1827.
9 Nagahisa H, Nagata Y, Ohnuki T, Osada M, Nagasawa T, Abe T & Todokoro K. Bone marrow stromal cells produce thrombopoietin and stimulate megakaryocyte growth and maturation but suppress proplatelet formation, Blood, 1996, 87, 1309–1316.
10 Tavassoli M & Aoki M. Localization of megakaryocytes in the bone marrow, Blood Cells, 1989, 15, 3–14.[Medline]
11 Lichtman MA, Chamberlain JK, Simon W & Santillo PA. Parasinusoidal location of megakaryocytes in marrow: a determinant of platelet release, Am J Hematol, 1978, 4, 303–312.[Medline]
12 Levine RF, Eldor A, HyAm E, Gamliel H, Fuks Z & Vlodavsky I. Megakaryocyte interaction with subendothelial extracellular matrix is associated with adhesion, platelet-like shape change, and thromboxane A2 production, Blood, 1985, 66, 570–576.
13 Radley JM & Scurfield G. The mechanism of platelet release, Blood, 1980, 56, 996–999.
14 Zucker-Franklin D, Stahl C & Hyde P. Megakaryocyte ultrastructure. Its relationship to normal and pathologic thrombocytopoiesis, Ann NY Acad Sci, 1987, 509, 25–33.[Medline]
15 Leven RM. Megakaryocyte motility and platelet formation, Scanning Microsc, 1987, 1, 1701–1709.[Medline]
16 Caine, Y.G., I. Vlodavsky, M. Hersh, A. Polliack, D. Gurfel, R. Or, R.F. Levine, and A. Eldor. 1986. Adhesion, spreading and fragmentation of human megakaryocytes exposed to subendothelial extracellular matrix: a scanning electron microscopy study. Scan. Electron Microsc. Pt. 3:1087–1094.
17 Avraham H, Cowley S, Chi SY, Jiang S & Groopman JE. Characterization of adhesive interactions between human endothelial cells and megakaryocytes, J Clin Invest, 1993, 91, 2378–2384.[Medline]
18 Hagiwara T, Nagasawa T, Nagahisa H, Takizawa M, Osada M & Abe T. Expression of adhesion molecules on cytoplasmic processes of human megakaryocytes, Exp Hematol, 1996, 24, 690–695.[Medline]
19 Tavassoli M & Aoki M. Migration of entire megakaryocytes through the marrow–blood barrier, Br J Haematol, 1981, 48, 25–29.[Medline]
20 Martin JF, Slater DN & Trowbridge EA. Abnormal intrapulmonary platelet production: a possible cause of vascular and lung disease, Lancet, 1983, 1, 793–796.[Medline]
21 Levine RF, Eldor A, Shoff PK, Kirwin S, Tenza D & Cramer EM. Circulating megakaryocytes: delivery of large numbers of intact, mature megakaryocytes to the lungs, Eur J Haematol, 1993, 51, 233–246.[Medline]
22 Davis RE, Stenberg PE, Levin J & Beckstead JH. Localization of megakaryocytes in normal mice and following administration of platelet antiserum, 5-fluorouracil, or radiostrontium: evidence for the site of platelet production, Exp Hematol, 1997, 25, 638–648.[Medline]
23 Gewirtz AM, Calabretta B, Rucinski B, Niewiarowski S & Xu WY. Inhibition of human megakaryocytopoiesis in vitro by platelet factor 4 (PF4) and a synthetic COOH-terminal PF4 peptide, J Clin Invest, 1989, 83, 1477–1486.[Medline]
24 Han ZC, Sensebe L, Abgrall JF & Briere J. Platelet factor 4 inhibits human megakaryocytopoiesis in vitro, Blood, 1990, 75, 1234–1239.
25 Gewirtz AM, Keefer M, Bien R & Mangan KF. Cellular regulation of human megakaryocytopoiesis in vitro, Prog Clin Biol Res, 1986, 215, 129–135.[Medline]
26 Bleul CC, Fuhlbrigge RC, Casasnovas JM, Aiuti A & Springer TA. A highly efficacious lymphocyte chemoattractant, stromal cell–derived factor 1 (SDF-1), J Exp Med, 1996, 184, 1101–1109.
27 D'Apuzzo M, Rolink A, Loetscher M, Hoxie JA, Clark-Lewis I, Melchers F, Baggiolini M & Moser B. The chemokine SDF-1, stromal cell-derived factor 1, attracts early stage B cell precursors via the chemokine receptor CXCR4, Eur J Immunol, 1997, 27, 1788–1793.[Medline]
28 Aiuti A, Webb IJ, Bleul C, Springer T & Gutierrez-Ramos JC. The chemokine SDF-1 is a chemoattractant for human CD34+ hematopoietic progenitor cells and provides a new mechanism to explain the mobilization of CD34+ progenitors to peripheral blood, J Exp Med, 1997, 185, 111–120.
29 Kim CH & Broxmeyer HE. In vitro behaviour of hematopoietic progenitor cells under the influence of chemoattractants: stromal cell-derived factor-1, steel factor, and the bone marrow environment, Blood, 1998, 91, 100–110.
30 Nagasawa T, Hirota S, Tachibana K, Takakura N, Nishikawa S, Kitamura Y, Yoshida N, Kikutani H & Kishimoto T. Defects of B-cell lymphopoiesis and bone-marrow myelopoiesis in mice lacking the CXC chemokine PBSF/SDF-1, Nature, 1996, 382, 635–638.[Medline]
31 Rafii S, Shapiro F, Rimarachin J, Nachman RL, Ferris B, Weksler B, Moore MA & Asch AS. Isolation and characterization of human bone marrow microvascular endothelial cells: hematopoietic progenitor cell adhesion, Blood, 1994, 84, 10–19.
32 Mohle R, Moore MA, Nachman RL & Rafii S. Transendothelial migration of CD34+ and mature hematopoietic cells: an in vitro study using a human bone marrow endothelial cell line, Blood, 1997, 89, 72–80.
33 Endres MJ, Clapham PR, Marsh M, Ahuja M, Turner JD, McKnight A, Thomas JF, Stoebenau-Haggarty B, Choe S, Vance PJ et al.. CD4-independent infection by HIV-2 is mediated by fusin/CXCR4, Cell, 1996, 87, 745–756.[Medline]
34 Mohle R, Green D, Moore MA, Nachman RL & Rafii S. Constitutive production and thrombin-induced release of vascular endothelial growth factor by human megakaryocytes and platelets, Proc Natl Acad Sci USA, 1997, 94, 663–668.
35 Avraham H, Vannier E, Cowley S, Jiang SX, Chi S, Dinarello CA, Zsebo KM & Groopman JE. Effects of the stem cell factor, c-kit ligand, on human megakaryocytic cells, Blood, 1992, 79, 365–371.
36 Barleon B, Sozzani S, Zhou D, Weich HA, Mantovani A & Marme D. Migration of human monocytes in response to vascular endothelial growth factor (VEGF) is mediated via VEGF receptor flt-1, Blood, 1996, 87, 3336–3343.
37 Schweitzer KM, Drager AM, van der Valk P, Zevenbergen A, Theijsmeijer AP, van der Schoot CE & Langenhuijsen MM. Constitutive expression of E-selectin and vascular cell adhesion molecule-1 on endothelial cells of hematopoietic tissues, Am J Pathol, 1996, 148, 165–175.[Abstract]
38 Avraham H. Regulation of megakaryocytopoiesis, Stem Cells (Dayton), 1993, 11, 499–510.[Medline]
39 Steegmaier M, Levinovitz A, Isenmann S, Borges E, Lenter M, Kocher HP, Kleuser B & Vestweber D. The E-selectin-ligand ESL-1 is a variant of a receptor for fibroblast growth factor, Nature, 1995, 373, 615–620.[Medline]
40 Bevilacqua MP. Endothelial-leukocyte adhesion molecules, Annu Rev Immunol, 1993, 11, 767–804.[Medline]
41 Gupta KS, Lysko GP, Pillarisetti K, Ohlstein E & Stadel JM. Chemokine receptors in human endothelial cells, J Biol Chem, 1998, 273, 4282–4287.
42 Bleul CC, Farzan M, Choe H, Parolin C, Clark-Lewis I, Sodroski J & Springer TA. The lymphocyte chemoattractant SDF-1 is a ligand for LESTR/fusin and blocks HIV-1 entry, Nature, 1996, 382, 829–833.[Medline]
43 Bleul CC, Wu L, Hoxie JA, Springer TA & Mackay CR. The HIV coreceptors CXCR4 and CCR5 are differentially expressed and regulated on human T lymphocytes, Proc Natl Acad Sci USA, 1997, 94, 1925–1930.
44 Hesselgesser J, Halks-Miller M, Del Vecchio V, Peiper SC, Hoxie J, Kolson DL, Taub D & Horuk R. CD4-independent association between HIV-1 gp120 and CXCR4: functional chemokine receptors are expressed in human neurons, Curr Biol, 1997, 7, 112–121.[Medline]
45 Strizki JM, Turner JD, Collman RG, Hoxie J & Gonzalez-Scarano F. A monoclonal antibody (12G5) directed against CXCR-4 inhibits infection with the dual-tropic human immunodeficiency virus type 1 isolate HIV-1(89.6) but not the T-tropic isolate HIV-1(HxB), J Virol, 1997, 71, 5678–5683.[Abstract]
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Facebook 
Reddit 
Technorati 
Twitter What's this?
This article has been cited by other articles:
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| TABLE OF CONTENTS |
|
