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 Previous Article | Table of Contents | Next Article 
Blood, Vol. 95 No. 5 (March 1), 2000:
pp. 1633-1641
HEMATOPOIESIS
Existence of a differentiation blockage at the stage of a
megakaryocyte precursor in the thrombocytopenia and absent radii (TAR)
syndrome
Rémi Letestu,
Natacha Vitrat,
Aline Massé,
Jean-Pierre Le Couedic,
Vladimir Lazar,
Philippe Rameau,
Françoise Wendling,
Jacqueline Vuillier,
Patrick Boutard,
Emmanuel Plouvier,
Mireille Plasse,
Rémi Favier,
William Vainchenker, and
Najet Debili
From INSERM U 362, Laboratoire associé no. 5 du comité
de Paris de la Ligue Nationale, Institut Gustave Roussy, Villejuif,
France; Plateau technique, Institut Gustave Roussy, Villejuif, France;
Service d'hématologie, CHU Jean Minjoz, Besançon, France;
Service d'hématologie, CHU Côte de Nacre, Caen, France;
Unité d'hématologie infantile, CHU St Jacques,
Besançon, France; Service de pédiatrie, centre hospitalier
d'Albertville, Albertville, France; and Service d'hématologie
biologique, Hôpital Armand Trousseau, Paris, France.
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Abstract |
The thrombocytopenia and absent radii (TAR) syndrome is a rare
disease associating bilateral radial agenesis and congenital thrombocytopenia. Here, we investigated in vitro megakaryocyte (MK)
differentiation and expression of c-mpl in 6 patients. Using blood or
marrow CD34+ cells, the colony-forming unit (CFU)-MK
number was markedly reduced. CD34+ cells were also
cultured in liquid medium in the presence of a combination of 3 cytokines (stem cell factor, interleukin-3, and interleukin-6) or
megakaryocyte growth and development factor (PEG-rHuMGDF) with or
without SCF. In the presence of PEG-rHuMGDF, the majority of mature
megakaryocytes (CD41 high, CD42 high) underwent apoptosis. This
phenomenon was also observed in cultures stimulated by three cytokines.
However, this last combination of cytokines allowed a more complete
terminal MK differentiation. Surprisingly, a homogeneous population of
CD34-CD41+CD42- cells accumulated
during the cultures. This population was unable to differentiate along
the myeloid pathways. This result suggests that a fraction of MK cells
is unable to differentiate in the TAR syndrome. We subsequently
investigated whether this could be related to an abnormality in c-mpl.
No mutation or rearrangement in the c-mpl gene was found by
Southern blots or by sequencing of the c-mpl coding region and its
promoter in any of the patients. Using Western blot analysis, a
decreased level of Mpl was found in patient platelets. A decreased
level of c-mpl messenger RNA in TAR platelets was also detected with a
lower c-mpl-P to c-mpl-K ratio in comparison to adult platelets.
Altogether, these results demonstrate that the thrombocytopenia of the
TAR syndrome is associated with a dysmegakaryocytopoiesis characterized
by cells blocked at an early stage of differentiation.
(Blood. 2000;95:1633-1641)
© 2000 by The American Society of Hematology.
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Introduction |
The thrombocytopenia and absent radii (TAR) syndrome is
a rare disease occurring with an approximate prevalence of 1 case in
500 000 to 1 million births. The TAR syndrome is characterized by the
association of skeletal malformations and hematologic
abnormalities.1-3 Among the skeletal defects, bilateral
absence of the radii and the presence of thumbs are the most typical
features. Thrombocytopenia is present in all cases; it is extremely
profound at birth and during the first 4 months of life, with platelet
counts below 10 × 109/L.1-3
Thrombocytopenia has 2 main characteristics: (1) according to most
authors, it is due to the absence of megakaryocytes (MK) in the marrow,
with a profound defect in the growth of colony-forming units (CFU)-MK
in vitro4,5; according to others, it may be due to a defect
in megakaryocytopoiesis with the presence of small MKs in the marrow as
well as in CFU-MK-derived colonies6; and (2) the
thrombocytopenia improves during the first 2 years of life, and
platelet counts may reach nearly normal values in
adulthood.1-3
The pathophysiology of TAR syndrome is poorly understood. One of the
possible candidate genes was a HOX gene. The HOX gene family is known to play a key role during embryogenesis and cell differentiation, including hematopoietic lineages. Moreover, targeted disruption of the HOXA11 and HOXD11 genes resulted in
radio-ulnar aplasia in mice.7 Furthermore, HOXA10
is the principal HOX gene expressed during MK differentiation,
and its overexpression in murine hematopoietic cells increases
megakaryocytic differentiation.8,9 However, HOXA10, HOX
A11, and HOXD11 genes had a normal nucleotide sequence, and
HOXA10 expression could be detected in patient cells by reverse
transcriptase-polymerase chain reaction (RT-PCR), thus ruling out a
direct involvement of these HOX genes in the TAR syndrome.10 Recently, Ballmaier et al4 have
shown that the thrombocytopenia in the TAR syndrome may be related to a
defect in the thrombopoietin (TPO) signaling pathway. For this reason, we reinvestigated in vitro megakaryocytopoiesis from 6 patients with
TAR syndrome. We found a profound defect in MK progenitors associated
with a blockage in MK differentiation with accumulation of cells
expressing CD41 without CD42. MK differentiation was poorly stimulated
by TPO and remained extremely abnormal, even when TPO was replaced by a
combination of cytokines. This abnormal MK differentiation was
associated with a decrease in c-mpl transcripts and Mpl protein. An
increased c-mpl-K to c-mpl-P ratio was also found.
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Materials and methods |
Patients
Nine patients with a TAR syndrome, including 1 fetus, were studied.
The clinical features and the investigations performed with each
samples are detailed in Table 1. Informed
consent was obtained in all cases (patients, parents, and controls) in
accordance with the institutional guidelines of the Committee on Human
Investigation.
Antibodies and reagents
Directly conjugated R-phycoerythrin (R-PE) anti-CD34 (HPCA-2, Becton
Dickinson, Mountain View, CA), R-PE-CY5 anti-CD34 (Immunotech, Lumigny,
France), R-PE anti-CD41a (anti-GPIIb/IIIa, Pharmingen, San Diego, CA),
fluorescein isothiocyanate (FITC) anti-CD42a (GPIX, Immunotech), R-PE
anti-glycophorin A (Dako, Glostrup, Denmark), and FITC anti-CD15
(Lewisx, Dako) monoclonal antibodies (MoAb)
were used for flow cytometry. FITC-, R-PE-, and R-PE-CY5-conjugated
immunoglobulin G1 (IgG1) MoAb (obtained from
Becton Dickinson and Dako) were used as isotype controls.
Unconjugated Y2/51 MoAb (anti-CD61, GPIIIa) was a generous gift from D. Mason (Oxford, UK). Alkaline phosphatase-coupled polyclonal goat
antibody against mouse immunoglobulin (Caltag Laboratories, Burlingame,
CA) was purchased from Tebu (Le Perray-en-Yvelines, France).
Different antibodies were used for Western blot analysis: a rabbit IgG
anti-human Mpl provided by T. Kato (Kirin, Tokyo, Japan), a rabbit
anti-human CD41 antibody11 obtained from D. Pidard (INSERM
U485, Institut Pasteur, Paris, France), and a rabbit anti-human glycocalicin (GPIb , CD42b) antibody.12 Lastly, a
horseradish peroxidase-conjugated donkey anti-rabbit IgG antibody was
purchased from Amersham (Life Sciences, Buckinghamshire, UK).
FITC-annexin-V (Immunotech) and 7 actinomycin D (7AAD) (Sigma, St
Louis, MO) were used for apoptosis analysis and cell death evaluation.
Phorbol 12-myristate 13-acetate (PMA) and 13-cis retinoic acid
were obtained from Sigma.
Thrombopoietin ELISA
The enzyme-linked immunosorbent assay (ELISA) for the detection of
human TPO was the Human TPO Quantikine kit (R & D Systems, Minneapolis,
MN), which was used according to the manufacturer's instructions. The
ELISA sensitivity limit was 25 ng/L of TPO.
Purification of CD34+ cells
CD34+ cells were purified either from bone marrow or
blood of TAR syndrome patients. Controls were aliquots of leukapheresis obtained from mobilized patients, blood from normal controls, and bone
marrow from patients undergoing hip surgery. Informed consent was
obtained from all these donors. CD34+ cells were purified
using a magnetic cell-sorting system (mini MACS, Miltenyi Biotec GmbH,
Bergisch Gladbach, Germany). Cells from leukapheresis samples and
normal bone marrow were passed twice through the column. Purity
evaluated by flow cytometry was about 90%. For blood or bone marrow
samples from TAR syndrome patients and blood from normal controls, the
immunomagnetic cell-sorting system was used to enrich the samples for
CD34+ cells by passing them only once on the column. Cells
eluted from the column were stained by the R-PE anti-CD34 MoAb and
sorted by means of a FACS (fluorescence-activated cell sorter)
Vantage (Becton Dickinson) equipped with a 100-µm nozzle
and an argon laser (Coherent Radiation, Palo Alto, CA) tuned to 488 nm
and operating at 500 mW.
Quantitation of clonogenic progenitors in semisolid cultures
Serum-free fibrin clot assays.
Cultures were performed in serum-free fibrin clot assays in the
presence of cytokines as previously reported.13 Target
cells were either low-density blood or marrow cells
(1 × 105 to 2.5 × 105 cells/mL)
or CD34+ cells (1 × 103 to
2 × 103 cells/mL). Three different cytokine
conditions were used: (1) megakaryocyte growth and development factor
(PEG-rHuMGDF; 10 ng/mL, a pegylated truncated form of human TPO, a
generous gift from J. L. Nichol, Amgen; Thousand Oaks, CA) alone; or
(2) with stem cell factor (SCF; 50 ng/mL, Amgen) or (3) a combination
of 3 cytokines: SCF (50 ng/mL); interleukin (IL)-6 (100 U/mL), a
generous gift from Dr S. Burstein (Oklahoma City, OK); and IL-3 (100 U/mL), kindly provided by Novartis (Basel, Switzerland). Cultures were incubated at 37°C in a fully humidified atmosphere containing 5%
carbon dioxide in air. MK colonies were enumerated after 10 to 12 days
by an indirect immuno-alkaline phosphatase-labeling technique.13
Methylcellulose assays.
Erythroid (burst-forming unit, erythroid [BFU-E]) and granulocytic
(CFU-GM [granulocyte macrophage]) progenitors were quantitated using
previously described methylcellulose assays.14
CD34+ cells were plated at a concentration varying from
0.5 × 103 to 2 × 103 cells/mL
of complete methylcellulose medium (0.8% methylcellulose in Iscove's
modified Dulbecco's medium (IMDM; Gibco Life Technologies,
Cergy-Pontoise, France), 30% fetal calf serum (FCS; Stem Cell,
Vancouver, BC), 1% deionized bovine serum albumin (Cohn
fraction V, Sigma) and 10-4 mol/L  -mercaptoethanol.
Cultures were carried out in the presence of recombinant human growth
factors: PEG-rHuMGDF (10 ng/mL), SCF (50 ng/mL), granculocyte
colony-stimulating factor (G-CSF, 20 ng/mL; Amgen), IL-6 (100 U/mL),
IL-3 (100 U/mL), and human erythropoietin (Epo, 1000 IU/L;
Janssen-Cilag, Issy les Moulineaux, France). Hematopoietic progenitors
were scored on day 12 using an inverted microscope.
In vitro liquid cultures of megakaryocytes from CD34+
cells
CD34+ cells were grown for 6 to 12 days in serum-free
medium supplemented with a combination of 2 cytokines (PEG-rHuMGDF and SCF) or 3 cytokines (SCF, IL-3, and IL-6).13
Immunolabeling and flow cytometric analysis
Cultured cells were labeled with different combinations of
fluorescent-conjugated antibodies by a 30-minute incubation at 4°C.
Flow cytometric analyses were performed on a FacSort cytometer (Becton Dickinson).
Cell sorting of megakaryocyte subsets
MKs at different stages of differentiation were obtained after 6 days of culture and sorted on the basis of expression of CD34, CD41a,
and CD42a. Cells were incubated with a mixture of FITC anti-CD42a, R-PE
anti-CD41a, and R-PE-CY5 anti-CD34 MoAbs for 30 minutes at 4°C in
their culture medium. Cells were sorted according to their
immunophenotype into 2 populations:
CD34-CD41a+CD42a- and
CD42a-CD41a++CD42a++. Individual
CD34-CD41a+CD42a- cells were sorted
with the automatic cloning unit device of the cell sorter into 96-well
plates.13 Serum-free cultures were performed in the
presence of different combinations of the following 6 cytokines:
PEG-rHuMGDF (10 ng/mL), SCF (50 ng/mL), IL-3 (100 U/mL), IL-6 (100 U/mL), G-CSF (12 ng/mL), and Epo (1 IU/mL). Plates were
examined at days 7 and 12 after incubation at 37°C in an air
atmosphere supplemented with 5% carbon dioxide.
Western blot analysis
Platelet lysate preparation.
Peripheral blood was obtained from 4 TAR syndrome patients; platelet
controls included 2 healthy donors and 2 different cord blood samples.
Platelet-rich plasma was prepared. The platelet pellet was washed twice
with phosphate buffered-saline (PBS; Gibco, Paisley, Scotland)
containing 0.1% ethylenediaminetetraacetic acid (EDTA; Sigma).
Platelets were then lysed in a lysis buffer supplemented with protease
inhibitors (Complete, Boehringer Mannheim, Meylan, France); the soluble
material was collected after centrifugation at 12 000g for 15 minutes. Protein concentration was determined for each sample using the
Bio-Rad DC Protein colorimetric assay (Bio-Rad, Hercules, CA).
Western blotting.
Platelet lysates were diluted in an equal volume of Laemmli buffer and
fractionated by sodium dodecyl sulfate-polyacrylamide gel
electrophoresis (SDS-PAGE, 8% polyacrylamide gels); 20 µg of total
protein lysate was deposited in each lane with prestained molecular
weight markers (Bio-Rad). Proteins were then electrophoretically transferred onto a nitrocellulose membrane (Bio-Rad). Residual nonspecific protein binding sites were blocked by incubating the filter
for 2 hours at room temperature in PBS with 0.1% Tween 20, containing
5% (w/v) dried milk. For immunoblotting, either rabbit IgG anti-human
Mpl or rabbit anti-human GPIIb polyclonal or rabbit anti-glycocalicin
(GPIb) antibodies were diluted at 1:500, 1:2000, and 1:1000,
respectively, in PBS-Tween-milk buffer and were incubated with the
membrane for 1 hour at room temperature. After washes, the filter was
incubated with a donkey anti-rabbit polyclonal antibody coupled to
horseradish peroxidase diluted at 1:5000. Antibody binding was
visualized with the enhanced chemoluminescence system (ECL kit,
Amersham). After scanning, the intensity in each lane was measured by
pixel quantitation using the MacBas v2.2 software.
Quantitative titration of messenger RNA by RT-PCR
RNA and complementary DNA preparation.
Total RNA was isolated using RNA PLUS (Bioprobe Systems, Quantum,
Montreuil sous Bois, France). RNA was reverse transcribed using random
hexamers (Gibco BRL/Life Technologies) with Superscript reverse transcriptase.
Construction of double internal standard for c-mpl-P and c-mpl-K.
An internal standard was constructed that contains the specific
sequence of c-mpl-P and c-mpl-K. In a first step, we amplified 124 base
pairs (bp) ranging from the exon 10 common to c-mpl and c-mpl-K to the
exon 11 specific to c-mpl-P (sense primer: 5-GATCTCCTTGGTGACCGCTC-3; antisense primer: 5-AAGTGAGGGCCACAGGGC-3) from a human erythroleukemia cell line complementary DNA (cDNA). A 19-bp oligonucleotide
(5-CTGGTCCACCGCCAGTCT-3) that corresponds to a sequence present on the
intron 10, specific to c-mpl-K, was synthetized and subsequently
ligated to the previous product by PCR. This standard was cloned, and
its sequence was checked.
Real-time quantitative RT-PCR.
To study more precisely the ratio between Mpl-P and Mpl-K, a
quantitative approach based on the fluorescent Taqman methodology and
real-time PCR on the ABI Prism 7700 sequence detection system (Perkin-Elmer, Foster City, CA) was performed. An internal control containing the c-mpl-P and c-mpl-K was constructed. The steady state
level of GPIIb, c-mpl-P (the main c-mpl splicing variant), and c-mpl-K
messenger RNAs (mRNAs) in the specimens was compared using a relative
quantitative approach as just described. Relative quantitation was performed using the standard curve method.
Amplification of an endogenous control (18S ribosomal RNA) was
performed to standardize the amount of sample cDNA added to the reaction.
Primers and probes were designed using the Primer Express
(Perkin-Elmer) and Oligo 4 (National Biosciences, Plymouth, MN) softwares and purchased from Perkin-Elmer.
PCR amplifications were performed using the Taqman core kit (ABI) in
standard conditions according to the manufacturer's instructions. Briefly, reactions were performed in 50-µL reactions containing the
cDNA equivalent to 25 ng of total RNA, 1 × Taqman buffer A, 5 mM MgCl2, 200 µM dATP, dCTP, dGTP, and 400 µM dUTP,
1.25 U AmpliTaq Gold, and 0.5 U UNG (uracil N-glycosylase [AmpErase],
250 nM each primer and 100 nM of the probe). Each reaction
was performed in duplicate. ABI 7700 sequence detection system was set
up in the manufacturer's standard thermal cycling conditions. The SDS
software was used to analyze fluorescent signals and calculate the
cycle threshold.
DNA sequencing
Genomic DNA was isolated from peripheral blood cells (leukocytes and
mononuclear cells), bone marrow cells, or patient Epstein-Barr virus
cell lines using the proteinase K/SDS treatment and phenol/chloroform extraction.15 Each of the individual 12 exons
of the c-mpl gene was amplified and sequenced using a PCR
technique previously described16 with the same primers
(Table 2). The gene promoter, a 903-bp fragment that includes GATA and its binding sites, was also sequenced. PCR was performed with the primers described in Table 2.
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Results |
Selective defect in megakaryocyte progenitors of TAR syndrome
patients
In a first set of experiments, CFU-MK was grown from low-density
marrow or blood cells from 3 TAR syndrome patients. MK colonies were
almost undetectable in serum-free fibrin clot assays after 12 days of
culture. Therefore, further experiments were performed using purified
CD34+ cells to obtain a much higher number of colonies.
Because most experiments in TAR syndrome patients were performed with
blood samples, we evaluated the number of CFU-MK present in the
peripheral blood CD34+ cells from 2 healthy donors obtained
in steady state conditions. Three conditions of cytokine stimulation
were compared: (1) PEG-rHuMGDF alone, (2) SCF plus PEG-rHuMGDF, and (3)
combination of 3 cytokines (SCF, IL-3, and IL-6) (Figure
1A). The best growth of
normal progenitors was obtained with the combination of PEG-rHuMGDF and
SCF, leading to 255 and 271 CFU-MK per 2000 CD34+ control
cells.
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| Fig 1.
Colony formation in the TAR syndrome.
(A) CFU-MK colony formation from peripheral blood CD34+
cells of TAR syndrome patients. CD34+ cells from blood of 4 TAR patients and 2 normal controls were purified by an immunomagnetic
procedure followed by cell sorting and were cultured in the serum-free
fibrin clot assay; 2000 CD34+ cells were seeded in the
presence of PEG-rHuMGDF alone, SCF plus PEG-rHuMGDF, or a combination
of 3 cytokines (IL-3, SCF, and IL-6) in triplicate. Colonies were
scored at day 12 after staining with an anti-CD61 MoAb. CFU-MK was
considered as aggregates of more than 2 CD61+ cells. A
marked reduction in CFU-MK colony formation was observed in TAR
syndrome patients compared to controls. In addition, colonies were
composed of aggregates of a maximum of 5 cells. (B) Comparison of
CFU-MK colony formation from blood and marrow CD34+ cells
in 2 other TAR syndrome patients. The same technique was used as in
Figure 1A. A marked parallel decrease in CFU-MK colony formation was
observed with both blood and marrow CD34+ cells.
Combination of 3 cytokines was more efficient than PEG-rHuMGDF alone or
SCF plus PEG-rHuMGDF in the induction of CFU-MK growth. (C) BFU-E and
CFU-GM colony formation from peripheral blood CD34+ cells
of TAR syndrome patients. Peripheral blood CD34+ cells were
purified as described above and were plated in methylcellulose in the
presence of a combination of PEG-rHuMGDF, SCF, G-CSF, IL-6, IL-3, and
Epo for 14 to 16 days. Colonies were scored under an inverted
microscope. No alteration in CFU-GM and BFU-E colony formation was
found in 4 TAR syndrome patients. All results are expressed per 2000 CD34+ cells.
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In the case of TAR syndrome patients, a profound decrease in colony
number, from 10-fold to 25-fold, was observed in all culture conditions
depending on the cytokines used (Figure 1A). In contrast to what was
observed with normal cells, the best cytokine combination (SCF, IL-3,
and IL-6) did not include PEG-rHuMGDF, and the combination of SCF plus
PEG-rHuMGDF was slightly superior to PEG-rHuMGDF alone. In addition,
colonies were composed of only a few cells (from 3 to 5 cells).
To confirm that this profound quantitative defect in MK progenitors was
not related to abnormal migration of MK progenitors in blood, CFU-MK
growth of marrow and blood CD34+ cells from 2 patients was
compared (Figure 1B). Similar numbers of MK progenitors and responses
to cytokines were found in blood and marrow CD34+ cells.
To investigate if this quantitative defect was restricted to the MK
lineage, other hematopoietic progenitors were assayed in
methylcellulose in the presence of 5 cytokines (SCF, IL-3, IL-6, Epo,
and G-CSF). Growth of BFU-E- and CFU-GM-derived colonies from
peripheral blood CD34+ cells was similar in TAR syndrome
and controls (Figure 1C).
We next investigated if this abnormal MK colony formation was due to a
pure quantitative defect in the number of MK progenitors or to a
maturation defect that impairs MK colony formation.
Presence of an in vitro MK maturation defect in the TAR syndrome
To precisely study MK maturation, cultures from CD34+
cells were performed in liquid medium in the presence of PEG-rHuMGDF, or SCF plus PEG-rHuMGDF, or the combination of 3 cytokines (SCF, IL-3,
and IL-6). In the case of normal CD34+ cells, these
experimental conditions induced terminal MK differentiation, including
platelet-shedding MKs, on day 9-10 of culture.17
With PEG-rHuMGDF alone, no cell amplification was observed at day 8 with TAR syndrome samples. When SCF was added to PEG-rHuMGDF, only a
2.5-fold increase in cell number was noted, whereas the combination of
3 cytokines (SCF, IL-3, and IL-6) elicited a 5-fold increase in cell
number. No terminal MK maturation was noted except in the presence of 3 cytokines, a condition in which rare proplatelet-bearing MKs (less than
1%) were observed. These experiments demonstrate impaired in vitro MK
maturation in the TAR syndrome and an abnormal response to cytokines,
particularly to TPO.
To better investigate if this poor response to TPO was due to a
decreased sensitivity, we studied the effect of increasing concentrations of PEG-rHuMGDF in the absence or presence of SCF. CD34+ cells (105 cells/mL) were grown in
96-well plates in the presence of 4 different PEG-rHuMGDF
concentrations (1, 10, 50, and 100 ng/mL supplemented or not with SCF
25 ng/mL), each condition being performed in triplicate. For a control,
normal CD34+ cells were cultured in presence of 10 ng/mL
PEG-rHuMGDF and 25 ng/mL SCF. After 10 days of culture, each well was
harvested separately and cells were labeled by FITC anti-CD42a and R-PE
anti-CD41a MoAbs. Dead cells were excluded by 7AAD. Total cell numbers
and CD41a+ cells were directly quantitated by flow
cytometry through a time-dependent acquisition and analyzed after
removing dead cells by gating 7AAD-positive events. Figure
2 illustrates the results obtained in 1 patient. The dose-response assay did not differ from normal controls,
with a plateau obtained at 1 ng/mL of PEG-rHuMGDF for CD41+
cells.18 SCF had no synergistic effects with PEG-rHuMGDF on the growth of MK. In contrast, it markedly increased the number of
non-MK cells (CD34+ and myeloid cells). Interestingly,
patient cells grown in the presence of 3 cytokines exhibited a higher
proliferation rate, but the content in CD41a+ cells was
very similar to that of cultures stimulated by PEG-rHuMGDF.
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| Fig 2.
Effects of PEG-rHuMGDF in liquid culture medium on the
growth of MKs from CD34+ cells of TAR syndrome patients.
PEG-rHuMGDF was tested at different concentrations, alone or in
combination with SCF. CD34+ cells (2000 cells) from bone
marrow of a TAR syndrome patient were grown in a 96-well plate (100 µL) in triplicate. At day 10, the numbers of viable cells and viable
CD41+ cells were quantitated by flow cytometry using
staining with an R-PE anti-CD41a MoAb and 7AAD.
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In these experiments, we observed a dissociation between the expression
of CD41 and CD42 (data not shown). To further characterize the cellular
abnormality associated with the TAR syndrome, we studied the expression
of CD41a and CD42a by flow cytometry in MK liquid cultures from 6 different patients. CD34+ cells from mobilized
leukapheresis samples, normal bone marrow, or normal peripheral blood
grown in similar conditions were used as a control. Analyses were
performed on different days of culture. At day 8, a very low proportion
of cells expressed CD42a in comparison to the controls. More
surprisingly, cultures were extremely depleted in cells coexpressing
CD41a and CD42a, corresponding to maturing MKs (Figure
3). At day 15, unlike the situation
observed in normal cultures, the proportion of
CD41a+CD42a+ cells decreased in the TAR
syndrome cultures, and a well-defined subset of
CD41a+CD42a- cells appeared (Figure 3). This MK
subpopulation seemed to be blocked at an early stage of maturation. In
control cultures, most cells coexpressed CD41a and CD42a because a
shift along the X axis (CD42a, FITC) was observed for
CD41a+ cells, including most of those expressing a low
level of CD41a (Figure 3). Only rare cells with low expression of CD41a
appeared to be negative for CD42a. In contrast, cells with an
intermediate and low level of CD41a did not express CD42a in the TAR
syndrome cultures (Figure 3). Similar results were observed in the 6 patients studied, either in the presence of PEG-rHuMGDF or 3 cytokines. However, some differences were observed depending on the cytokines used. When PEG-rHuMGDF alone was used, most of the cells expressed the
CD41a antigen, but no terminal maturation was observed. In addition,
marked apoptosis detected by binding of annexin V to CD41a+
cells was present (Figure 4). Three
cytokines appeared to be the best cytokine mix to support TAR syndrome
MK cultures.
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| Fig 3.
Double staining by anti-CD41a and CD42a MoAbs of
CD34+ cells cultured for 8 and 15 days in liquid culture
medium.
Blood CD34+ cells from a TAR syndrome patient were grown in
liquid medium in the presence of SCF plus PEG-rHuMGDF or a combination
of 3 cytokines (IL-3, SCF, and IL-6). At days 8 and 15, cells were
labeled by an R-PE anti-CD41a MoAb and FITC anti-CD42a MoAb and
analyzed by flow cytometry. This figure illustrates the results
obtained in 1 patient. Similar results were obtained in the 4 other
patients studied.
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| Fig 4.
Quantitative analysis of apoptosis in
CD41a+ cells from TAR syndrome patients.
Blood CD34+ cells were grown in liquid culture in the
presence of PEG-rHuMGDF (A), SCF plus PEG-rHuMGDF (B), or a combination
of 3 cytokines (SCF, IL-3, and IL-6) (C). At day 9, cells were stained
by annexin V-FITC, R-PE anti-CD41a MoAb, and 7AAD. Dead cells were
excluded on the basis of 7-AAD staining (FL3). Viable cells were
studied for CD41a and annexin V staining. The horizontal axis shows
cells stained by annexin V and the vertical axis cells stained by the
anti-CD41a MoAb. This figure illustrates a typical experiment from 1 patient. Similar results were obtained in 3 other patients.
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The rare maturing MKs, ie, CD41a+CD42a+ cells,
were also sorted and plated in liquid culture to investigate terminal
maturation. Cultures were examined under an inverted microscope, and
platelet-shedding MKs were scored daily for 5 days. In 3 different
patients, very few platelet-shedding MKs were observed in culture with
PEG-rHuMGDF alone, whereas the best results were obtained with the
combination of 3 cytokines (SCF, IL-3, and IL-6) (data not shown).
All of these results suggest that MK progenitors mature abnormally,
with a partial block in maturation at a cellular stage characterized by
the presence of CD41 and absence of CD42. Rare cells escaped to this block.
Characterization of the CD41a+CD42a- cell
subset
Cells were first characterized by their immunophenotype using
3-color labeling. CD41a+CD42a- cells did not
express CD34 or other lineage markers, such as glycophorin A, CD15, and
CD14 (data not shown).
To further investigate their biologic properties, these cells were
sorted and plated either in semisolid or in liquid medium in the
presence of different cytokine combinations. Clonogenic and limiting
dilution assays failed to detect any progenitors among
CD41a+CD42a- TAR syndrome cells (data not
shown). In liquid cultures, cells were immunophenotyped to investigate
their differentiation in response to different growth factor
combinations (including SCF, IL-3, IL-6, Epo, G-CSF, and PEG-rHuMGDF).
Even after prolonged cultures, no phenotypic changes were observed in
sorted cells, except that very few cells (less than 0.1%) acquired
CD42 before undergoing apoptosis (Figure
5). Cells did not acquire markers of other
myeloid cell lineages (glycophorin A, CD15, CD14). Addition of phorbol
ester (PMA 20 and 40 nM) to CD41+CD42- cells
did not induce any differentiation (data not shown). In contrast, PMA
markedly increased the expression of CD41a and CD42a when added to
purified CD41+CD42- cells obtained after a
7-day stimulation by PEG-rHuMGDF and SCF of normal CD34+
cells (data not shown).
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| Fig 5.
Differentiation of
CD41a+CD42a cells in the presence of 6 cytokines.
CD34+ cells from a TAR syndrome patient and a control were
grown in the presence of SCF plus PEG-rHuMGDF for 8 days.
CD41a+CD42a- cells were sorted and cultured for
10 additional days in the presence of 6 cytokines (SCF, IL-3, IL-6,
G-CSF, PEG-rHuMGDF, and Epo). At the end of the culture, cells were
restained with an R-PE anti-CD41a MoAb and an anti-CD42a MoAb. The TAR
syndrome patient cells remained CD41a+ and did
not acquire CD42a or new markers of differentiation. The majority of
normal CD41a+CD42a- cells lost the CD41 antigen
and acquired other markers of differentiation, in particular,
glycophorin A (data not shown), and the remaining differentiated along
the MK pathway with the appearance of CD42.
|
|
No genomic alteration in c-mpl gene of TAR syndrome patients
Because c-mpl, the TPO receptor, plays a crucial role in MK
differentiation, we searched for an abnormality of c-mpl in TAR syndrome. We first investigated the c-mpl gene.
A Southern blot was performed that excluded a large deletion in the
32-kilobase long genomic region that was analyzed (data not shown).
Then, a subtle genomic alteration was sought by sequencing the coding
region of the gene, the intron-exon junctions and, finally, the c-mpl
promoter. Neither mutation nor recurrent polymorphisms could be
detected in 8 patients tested, leading to the conclusion that the
defect in MK differentiation was not the result of an alteration in the
c-mpl gene. However, this result does not exclude a defect in
c-mpl expression.
Impaired expression of Mpl in TAR syndrome platelets
Mpl expression was studied by Western blot analysis in platelets
from 4 different patients (ages ranging from 7 to 55 years, platelet
counts from 50 × 109/L to
80 × 109/L). Platelets from adult and cord blood
were used as controls. Mpl was revealed first by probing with an IgG
polyclonal antibody (Figure 6); filters
were stripped and reprobed with an anti-GPIIb antibody for
normalization. Mpl and GPIIb bands were scanned and submitted to pixel
quantitation. Figure 7 shows that
normalized Mpl expression in TAR syndrome platelets was significantly
lower than in adult control platelets. Two patients (P7 and P8) had extremely low levels of Mpl in their platelets. The 2 other patients also had a decreased level of Mpl in comparison to adult platelets, as
shown in Figure 7, at a 10-minute exposure time. It is noteworthy that
cord blood platelets expressed much lower amounts of Mpl than platelets
from adults. On average, the levels of Mpl in neonatal and TAR syndrome
platelets were quite similar. Using our technique of Western blot, we
could not differentiate the Mpl-P and Mpl-K isoforms.
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| Fig 6.
Western blot analysis of Mpl, GPIIb, and GPIb
expression in platelets from normal adult controls, newborns, and TAR
syndrome patients.
Platelet lysates (20 µg) were separated by SDS-PAGE and probed with
polyclonal antibodies against Mpl, GPIIb, and GPIb.
|
|
View larger version (34K):
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| Fig 7.
Comparison of Mpl expression in platelets from normal
adult controls, newborns, and TAR syndrome patients.
Western blots were scanned, and the intensity of each lane was measured
by pixel quantitation using the MacBas v2.2 software. Mpl levels were
normalized to GPIIb expression and compared. Results are expressed in
arbitrary units resulting from the ratio of the Mpl to GPIIb band. Four
patients with a TAR syndrome, 4 adult controls, and 4 newborns were
studied.
|
|
GPIb was normally expressed in TAR syndrome, further demonstrating that
platelet production arises from CD41+CD42+
cells and not from the CD41+CD42- cell subset.
Expression level of c-mpl mRNA in platelets and MKs from TAR
syndrome patients
To investigate the mechanisms of the low level of Mpl in platelets
from TAR patients, we studied the expression of c-mpl by means of a
real-time quantitative RT-PCR assay. We studied 2 isoforms of
c-mpl c-mpl-P and c-mpl-Kwhich arise by alternative splicing with a
premature termination of transcription in intron 10, and normalized
their expression to that of GPIIb and 18S mRNA. GPIIb was expressed at
the same level in TAR, adult, and cord blood platelets in comparison to
18S. Thus, the GPIIb transcript was used for normalization. The ratio
between c-mpl and GPIIb is not the real ratio because in these
experiments we have not used an internal standard for each marker, but
it can be used to compare the difference in c-mpl content in each
platelet samples. As illustrated in Figure
8A, c-mpl-P in TAR platelets was markedly
decreased in comparison to adult platelets (a 3-fold decrease,
n = 6). c-mpl-P was also expressed at a lower level in cord blood
platelets than in adult platelets, and only a 50% decrease was
observed in TAR platelets in comparison to neonate platelets. The
c-mpl-P to c-mpl-K ratio was studied by using a single standard that
contains both forms and thus allows to precisely determine the real
ratio between c-mpl-P and c-mpl-K in each cell sample. c-mpl-P was the
predominant form of c-mpl in platelets, as previously
demonstrated.19,20 The c-mpl-P to c-mpl-K ratio was reduced
in TAR syndrome platelets in comparison to adult platelets (Figure 8B)
or neonate platelets but with some variation among patients (2 patients
having a 5-fold decreased level). The c-mpl-P to c-mpl-K ratio was also
studied in cultured MKs from adult cord blood and TAR MKs.
Surprisingly, this ratio was a 6-fold decrease in normal MKs in
comparison to platelets. Thus, this result suggests that during normal
MK differentiation expression of c-mpl-P increased at the expense of
c-mpl-K. In the TAR syndrome, only 3 patients could be studied. In 2 of
them, the ratio was in the same range as normal adult or cord blood MKs
(Figure 8C); in the third, the ratio was extremely low.
View larger version (29K):
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| Fig 8.
Expression of c-mpl-P and c-mpl-K RNA by real-time RT-PCR
in platelets and MKs from normal adult controls, newborns, and TAR
syndrome patients.
Expression of c-mpl-P, c-mpl-K, GPIIb, and 18S was studied by a
quantitative RT-PCR assay (see "Materials and Methods"). (A)
Comparison of the expression of c-mpl-P in normal adults and TAR
platelets. Expression of c-mpl-P was normalized in each sample to the
expression of GPIIb. This is expressed by a ratio that does not
correspond to the real ratio between the 2 different transcripts
because we did not use a standard. This ratio allows the comparison of
the c-mpl-P transcript expression in normal and TAR platelets. (B)
Ratio between c-mpl-P and c-mpl-K in platelets. To calculate the real
ratio between the 2 isoforms, a standard was constructed that contains
the 2 amplified sequences permitting precise evaluation of the
efficiency of each PCR. (C) Ratio between c-mpl-P and c-mpl-K in MKs.
MKs were obtained from culture of normal adult bone marrow at day 7 and
day 12, from cord blood (NN) at day 12, and TAR patients at day 12.
|
|
Plasma TPO level in 6 TAR patients
Finally, we measured plasma TPO levels in 6 TAR patients with
platelet counts ranging between 50 × 109/L and
110 × 109/L. The TPO level was slightly but
significantly elevated in these patients (mean 75 ng/L versus 40 ng/L
in adult and 50 ng/L in cord blood), as previously
reported,4 suggesting that there is no major abnormality in
the synthesis of TPO in this disease.
| |
Discussion |
The TAR syndrome is a hereditary disease associated with skeletal
malformations, including thrombocytopenia and a bilateral absence of
radii.1-3 The thrombocytopenia has very peculiar
characteristics: it is due to a defect in platelet production, and it
is extremely severe during the first years of life, but it
progressively improves, with the platelet count returning to near
normal values in adulthood. Progressive correction of the
thrombocytopenia after birth suggests that the gene(s) involved in
defective megakaryocytopoiesis is developmentally regulated.
To better understand the defects in MK differentiation in the TAR
syndrome, we studied 6 patients with this disease. All patients were
studied in infancy or adulthood at a time when thrombocytopenia was
moderate (50 × 109/L to
110 × 109/L). Nevertheless, we found that in vitro
megakaryocytopoiesis was profoundly altered. Confirming a previous
study,5 in vitro MK colony formation from either blood or
bone marrow CD34+ cells was 10-fold to 20-fold reduced in
comparison to that of normal controls, whatever cytokine was used in
the culture. In contrast, growth of BFU-E- and CFU-GM-derived
colonies was normal. Impaired MK colony formation could be related to a
true absence of MK progenitors or to a maturation defect leading to a
subsequent block in MK differentiation. To solve this problem, we
investigated the immunophenotype of MKs derived in vitro from
CD34+ cells after cytokine stimulation in liquid medium.
Very few mature MKs characterized by high levels of CD41 and CD42 were
present in the cultures, whatever cytokine was used. Surprisingly, in cultures of TAR syndrome patients, accumulation of cells with a
peculiar phenotype was detected. These cells expressed CD41 antigen in
the absence of CD42 and CD34 antigens. They began to be detectable
after day 6 of culture and, thereafter, increased in number. They had
the morphology of blasts cells and could not be induced to undergo
terminal MK differentiation by cytokines or PMA treatment. These cells
were long-lived and could be kept in culture for several weeks without
proliferation or modification of their immunophenotype. However, their
survival was dependent on the presence of cytokines in the medium.
Cells with a similar phenotype could be observed in low numbers in
liquid cultures of normal adult CD34+ cells and to a higher
degree in cultures of cord blood CD34+ cells (data not
shown). These cells from normal controls expressed much lower amounts
of CD41 than their TAR syndrome counterparts; higher levels of CD41
were associated with expression of CD42 in normal controls. These
CD41+CD42- cells could be the first
commitment step toward terminal MK differentiation because it has been demonstrated that CD41 expression precedes CD42
expression during MK differentiation.21,22 However, there is increasing evidence that CD41 antigen is not specific for the MK
lineage and can be expressed early during myeloid, erythroid, and
T-cell differentiation.23-27 In the TAR syndrome,
CD41+CD42- cells do not seem identical to those
cells because they could not be induced toward erythroid or
granulocytic differentiation in the presence of Epo or G-CSF. For this
reason, in the TAR syndrome, CD41+CD42- cells
are likely, although not totally demonstrated, MK precursors blocked in
their differentiation. It is noteworthy that these CD41+CD42- cells did not contribute to platelet
formation because they do not shed platelets in vitro and, also, that
GPIb was normally expressed in TAR syndrome platelets. In a patient
with the TAR syndrome, de Alarcon et al reported a normal number of
CFU-MK-derived colonies. Cells making up the colonies had marked
morphologic abnormalities and were of small size.6 These
cells might be the equivalent of the CD41+CD42-
cells found in our assays.
Therefore, one might speculate that
dysmegakaryocytopoiesis in the TAR syndrome is the
consequence of an intrinsic cellular defect related to an abnormality
in a protein that regulates late MK differentiation. The TAR syndrome
can be compared in many ways to Diamond-Blackfan anemia, in which
skeletal and hematologic abnormalities are both present. In this
syndrome, the anemia is associated with an erythroblastopenia and a
variable number of erythroid progenitors, suggesting a blockage in the
erythroid differentiation at a CFU-E stage of
differentiation.28,29 Recently, a mutation in the gene
encoding ribosomal protein S19 has been demonstrated in 30% of the
patients.30
A reduced response to TPO was also found in addition to maturation
defects: (1) unlike normal controls, a higher number of colonies were
obtained using a combination of cytokines that did not include
PEG-rHuMGDF; (2) in liquid culture, platelet-shedding MKs were observed
essentially with a combination of 3 cytokines (IL-3, IL-6, and SCF);
and (3) most maturing MKs stimulated by PEG-rHuMGDF underwent
apoptosis. These findings support the results of Ballmaier et al that
showed that platelets from TAR syndrome patients did not respond to
TPO, suggesting that this defect could be related to abnormal
signal transduction downstream of Mpl.4 A point mutation in
the c-mpl gene was not ruled out. We searched for an
abnormality of the c-mpl gene in the TAR syndrome. Our results
ruled out the presence of a point mutation in the c-mpl gene and its proximal promoter, in agreement with the recent report of Strippoli et al.31 However, we found 2 abnormalities:
(1) The level of Mpl was decreased in platelets from TAR syndrome
patients in comparison to that of adult platelets, and this reduction
was heterogeneous but quite marked in 2 patients. It is
noteworthy that the Mpl content of cord blood platelets was markedly reduced in comparison to that of adult platelets, suggesting that c-mpl is regulated during development. Levels of Mpl in neonatal and TAR syndrome platelets were quite similar. This was not due to the age of the TAR syndrome patients because 2 of them were adults.
(2) c-mpl RNA levels studied by quantitative RT-PCR assay were markedly
reduced in TAR syndrome platelets in comparison to those of adult
platelets but identical to those of neonatal platelets. In addition, we
studied the 2 main c-mpl isoforms (c-mpl-P and c-mpl-K).32,33 The c-mpl-P to c-mpl-K ratio was reduced in TAR platelets but heterogeneous among patients. However, this ratio
surprisingly varied during MK differentiation. It was much lower in
normal MKs than in platelets. The c-mpl-P to c-mpl-K ratio in TAR MKs
was only slightly diminished in comparison to normal MKs, suggesting
that the relative change in isoforms did not occur during MK
differentiation in the TAR syndrome.
Diminished expression of c-mpl has been reported in myeloproliferative
diseases, including essential thrombocythemia, polycythemia vera, and
myelofibrosis.19,34 As in the TAR syndrome, this decrease
in Mpl protein is associated with a parallel reduction in c-mpl RNA.
However, in essential thrombocythemia it was shown that the relative
expression of the different c-mpl isoforms was normal.20 In
the TAR syndrome, the decrease in c-mpl expression associated with a
marked reduction in the c-mpl-P to c-mpl-K ratio may have some biologic
consequences. Indeed, Mpl-K is an isoform that has an extracellular
sequence that is identical to that of Mpl-P but has an extensive
deletion in its intracellular domain with absence of signaling boxes 1 and 2.32,33,35 Thus, Mpl-K is theoretically unable to
transduce a signal. At present, it is not known if Mpl-K is normally
expressed on the surface of MKs and platelets. If it is expressed on
their surface, it should be able to bind TPO without transducing a
signal. Furthermore, it may behave as a dominant negative molecule
because Mpl requires homodimerization to transduce a
signal.36-38 If it is the case, the low c-mpl-P to c-mpl-K
ratio observed during the late MK differentiation in the TAR syndrome
may play a role in the defective megakaryocytopoiesis. Further
experiments to study the precise role of Mpl-K will be important to
support this hypothesis.
On the other hand, it seems unlikely that the low level of Mpl-P can
entirely explain the MK differentiation defect in the TAR syndrome, for
the following reasons: (1) the maturation defect persists, although to
a lesser extent, when cultures are stimulated by a combination of 3 cytokines, and (2) c-mpl-/- mice have decreased numbers of
MK progenitors as well as other hematopoietic
progenitors,39,40 a finding not present in the TAR
syndrome, and they have no MK maturation defect. This suggests that the
abnormal c-mpl expression in TAR syndrome may be the consequence of an
intrinsic cellular defect. However, it cannot be excluded that it
participates in the mechanism of thrombocytopenia. Future experiments
to induce c-mpl-P overexpression in TAR syndrome MK progenitors may
help to solve this question.
| |
Acknowledgments |
We thank C. Cailliot (Amgen, Neuilly, France) and J. L. Nichol (Amgen,
Thousand Oaks, CA) for the gift of PEG-rHuMGDF and stem cell factor; T. Kato (Kirin Brewery Co, Ltd, Tokyo, Japan) for the gift of polyclonal
antibodies against human Mpl; and D. Pidart (Institut Pasteur, Paris,
France) for the gift of anti-GPIIb antibody. We are grateful to
Dr F. Beaujean and F. Norol (Hôpital Henri Mondor,
Créteil, France) for providing the cytapheresis samples. We
are indebted to B. Forget for editing the English in the manuscript.
| |
Footnotes |
Submitted March 8, 1999; accepted November 3, 1999.
Supported by grants from the Institut National de la Santé et de
la Recherche Médicale. N.V. was supported by a fellowship from
Amgen and R.L. by a fellowship from the Comité de Recherche Clinique (Institut Gustave Roussy, Villejuif, France).
Reprints: Najet Debili, INSERM U 362, Laboratoire associé
no. 5 du comité de Paris de la Ligue Nationale, Institut Gustave
Roussy, Villejuif 94805, Cedex, France; e-mail: denali{at}igr.fr.
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.
| |
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J. G. Drachman
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R. A. Shivdasani
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C.-E. Dempfle, C. Burck, T. Grutzmacher, J. Wizenmann, and D. L. Heene
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Top
Abstract
Introduction