Circulating CD34+, CD133+, and Vascular Endothelial Growth Factor Receptor 2–Positive Endothelial Progenitor Cells in Myelofibrosis With Mye
http://www.100md.com
《临床肿瘤学》
the Laboratory of Biotechnology
the Transplant Research Area
the Unit of Internal Medicine III
the Unit of Clinical Immunology, Immunohematology, and Transfusion Service
the Department of Pediatrics
the Laboratory of Clinical Epidemiology, Istituto di Ricovero e Cura a Carattere Scientifico Policlinico S. Matteo, Pavia, Italy
the Section of Hematology/Oncology
Department of Pathology, University of Illinois Cancer Center
University of Illinois College of Medicine
the Myeloproliferative Disorders Research Consortium, Chicago, IL
ABSTRACT
PURPOSE: Endothelial progenitor cells (EPCs) are present in circulation and contribute to vasculogenesis in adults. We measured the number of circulating EPCs in patients with myelofibrosis with myeloid metaplasia (MMM), and we examined the relationship between the number of EPCs and severity of the MMM disease process.
PATIENTS AND METHODS: The number of EPCs was measured by assaying the CD34+CD133+ vascular endothelial growth factor receptor 2 (VEGFR2) –positive cell phenotype in 110 MMM patients, 16 patients with other Philadelphia-negative chronic myeloproliferative disorders (Ph-negative CMPDs), and 14 healthy participants. In four MMM patients, the capacity of selected CD34+ cells to form endothelial colonies (CFU-End) in vitro was tested.
RESULTS: CD34+, CD133+, and VEGFR2-positive EPCs were detectable in unselected peripheral-blood cells of 50.9% MMM patients, 37.5% control patients, and 21% healthy participants. Patients with MMM had a median of 0.26% EPCs, significantly higher than that in healthy controls (median, 0%) and in patients with other Ph-negative CMPDs (median, 0.1%). In 14.5% of MMM patients, the numbers of EPCs were greater than the highest value found in patients with other Ph-negative CMPDs. CD34+ selected cells produced colony-forming unit–endothelial (CFU-End), which were vascular endothelial (VE) -cadherin positive, CD31+, von Willebrand factor positive, and CD45–. In MMM patients, the larger the number of EPCs, the smaller the number of circulating immature myeloid cells and circulating CD45+CD34+ hematopoietic progenitor cells. Increased numbers of EPCs were associated with younger age and a diagnosis of prefibrotic MMM.
CONCLUSION: Circulating EPCs are elevated in MMM patients in the early stage of the disease. Heightened mobilization of EPCs may represent an important mechanism for development of neoangiogenesis in MMM.
INTRODUCTION
Myelofibrosis with myeloid metaplasia (MMM) is a clonal Philadelphia-negative chronic myeloproliferative disorder (Ph-negative CMPD), characterized by a variable degree of bone marrow fibrosis, extramedullary hematopoiesis, and presence of immature myeloid cells in the peripheral blood.1,2 A high degree of trafficking of hematopoietic progenitor cells (HPCs) is a characteristic of the disease that discriminates MMM from other Ph-negative CMPDs (ie, polycythemia vera [PV] and essential thrombocythemia [ET]), and is predictive of disease severity and risk of blast transformation.3-6
There is strong evidence that a population of endothelial progenitor cells (EPCs) can be isolated from circulating mononuclear cells of normal patients,7 that different pathologic conditions can affect their number,7-9 and that they contribute to vasculogenesis in adult organisms.10 Immunophenotype identification of EPCs relies on coexpression of specific cell-surface proteins; in particular, endothelial markers, including CD34 and vascular endothelial growth factor receptor 2 (VEGFR2), and the human stem-cell marker CD133.8,9,11
Bone marrow releases EPCs as a result of mechanisms that are in part similar to those that regulate the release of HPCs.12 We therefore expected a high rate of EPC release from the bone marrow of MMM patients and we hypothesized that circulating EPCs may sustain the prominent increase of neoangiogenesis in the bone marrow13 and spleen.14 Hence, we assayed the number of circulating EPCs by cell phenotype from the peripheral blood of a large, well-characterized population of MMM patients. The aim of the study was to assess the ability of the number of EPCs to distinguish MMM from other Ph-negative CMPDs, and to examine the relationship among number of EPCs and type (primary v secondary), phase, and severity of the MMM disease process.
PATIENTS AND METHODS
Patients and Study Protocol
All consecutive patients with a diagnosis of MMM referred to the Hospital Istituto di Ricovero e Cura a Carratere Scientifico (IRCCS) Policlinico S. Matteo (Pavia, Italy) between July 2002 and June 2004 were included in this study. Our institution is a teaching and tertiary-care hospital, and is a major referral site for patients with MMM because it coordinates the Italian Registry for Myelofibrosis, a nationwide prospective database. The study protocol was approved by the Institutional Review Board of the Hospital IRCSS Policlinico S. Matteo. All patients provided written informed consent to be included in this study. Study design, data collection, analysis, and reporting were performed independently by a study committee.
One hundred ten patients with MMM were referred by 39 Italian centers participating in the Italian Registry for Myelofibrosis. In 94 patients, the diagnosis of MMM was established according to the Italian consensus conference criteria,15 by which the diagnosis held if diffuse bone marrow fibrosis was present and Ph chromosome or breakpoint cluster region–Abelson murine leukemia (BCR-ABL) rearrangement in peripheral-blood cells was absent (necessary criteria), and if an algorithm of optional criteria was satisfied. Sixteen additional patients who initially were classified as having atypical myeloproliferative disorders were also included. They had a bone marrow biopsy with an absent or only slightly increased reticulin fibrosis at diagnosis, and intense hyperplasia of megakaryocytes with abnormal or bizarre megakaryocytes appearing in clusters of more than four cellular elements. In these patients, the diagnosis of MMM was based on the absence of the Ph chromosome, and on the algorithm for optional criteria set by the Italian consensus conference.15 These patients frequently had a slight increase in platelet count; however, in no patient was the platelet count greater than 600 x109/L. These patients exhibited the corresponding prefibrotic stage of the disease accepted in the classification of myeloproliferative disorders proposed by the WHO.16
Ninety patients (81.8%) were classified as having primary MMM and 20 were classified as having secondary MMM (15 had post-PV MMM and five had post-ET MMM). Twenty-five patients (22.7%) were studied at the time of diagnosis, whereas 85 were enrolled during the course of their disease. Seventy-three patients (66.3%) were studied before the start of any cytoreductive treatment or when the cytoreduction had been stopped for at least 3 months. At the time of analysis, 26 patients were receiving hydroxyurea and 11 were receiving thalidomide treatment. Nine patients had been splenectomized and sampling was done more than 6 months after splenectomy.
All patients were screened according to a strict protocol. Together with the collection of demographic data, a sample of venous blood was drawn to assess a complete blood count with differential, circulating CD45+CD34+ cell count and phenotype, and circulating EPCs. The WBC count was corrected for the number of circulating erythroblasts. Circulating nucleated cells were classified as immature myeloid cells, erythroblasts, and blasts. Blasts were defined as undifferentiated cells with an immature nucleolated nucleus and basophilic cytoplasm with or without azurophilic granules. Spleen size was measured by ultrasonography. Two parameters of spleen size were recorded: the distance from the splenic tip to the costal margin, and the spleen index17 (ie, the product of the longitudinal by the transverse spleen axis, the latter defined as the maximal width of the organ). Liver enlargement was measured as the distance from the right costal margin in centimeters. Patients were assigned a prognostic score based on findings of Dupriez et al.18
Sixteen patients with other Ph-negative CMPDs were also studied for comparison (six women and 10 men; median age, 46 years; range, 22 to 70 years). They were categorized as having PV (nine patients) or ET (seven patients) according to the Polycythemia Vera Study Group (PVSG) criteria,19,20 after exclusion of MMM. Of the patients with other Ph-negative CMPDs, eight were studied at diagnosis, and eight were studied from 4 to 10 months after diagnosis but before any therapy had been started or when therapy had been stopped for at least 3 months.
Control samples were obtained from 15 healthy individuals (13 men and two women; median age, 49 years; range, 27 to 60 years), including four first-time donors. Each control participant was free of wounds, ulcers, recent surgery, or inflammatory or malignant disease that might influence the number of EPCs.7,21
Flow Cytometry Analysis of Circulating HPCs
To measure the numbers of HPCs in peripheral blood, 50 μL of EDTA-anticoagulated blood was stained with fluorescein isothiocyanate (FITC) –conjugated, phycoerythrin (PE) -conjugated, peridinin chlorophyll protein (PerCp) –conjugated, and laser dye styryl (LDS751) –conjugated monoclonal antibodies. The CD45-FITC/CD34-PE/LDS751 and CD38-FITC/CD34-PE/CD45-PerCp antibodies were used. Antibodies were obtained from Becton Dickinson (San Jose, CA), and the analyses were performed by a fluorescence activated cell sorter (FACS) calibur flow cytometer (Becton Dickinson). For each sample, 20,000 events were acquired and the percentage of positive cells was calculated based on the appropriate isotype control. The analysis was completed using the cell-gating guidelines recommended by the International Society of Hematotherapy and Graft Engineering.22
Flow Cytometry Analysis of Circulating Endothelial Progenitor Cells
Circulating EPCs were measured by phenotypic analysis in unselected peripheral-blood cells in all patients and normal controls. Blood samples were processed within 2 hours after they were drawn. Fifty microliters of EDTA-anticoagulated blood was incubated for 30 minutes at 4°C with 20 μL of FITC-conjugated anti-CD34, 10 μL of PE-conjugated anti-CD133, and 6 μL of 1:10 diluted biotin-conjugated anti-VEGFR2 (Sigma Chemical Co, St Louis, MO). Biotin conjugated anti-VEGFR2 was revealed using 5 μL of 1:5 diluted PerCp-streptavidin (Becton Dickinson, Pharmingen, San Diego, CA). Appropriate isotype controls were used for each staining procedure. One milliliter of lysis solution (Dako, Glostrup, Denmark) was added for 5 minutes at room temperature. The samples were then centrifuged and pellets were resuspended in 300 μL of phosphate buffer with 0.5% fetal calf serum (HyClone, Logan, UT). Two x 105 cells were acquired by a FACS calibur flow cytometer, and analyzed by CellQuest software (BD Biosciences, San Diego, CA). Because the number of dead/apoptotic cells was negligible, the analysis was performed excluding cellular debris in a side scatter/forward scatter (SSC/FSC) dot plot. CD34+ cells were electronically gated and the percentage of cells coexpressing CD133 and VEGFR2 was evaluated. For each sample a minimum of 100,000 events was acquired. Detection of EPCs by flow cytometry was defined by the presence of at least 0.03% nucleated cells coexpressing the three antigens over the background fluorescence. Results were expressed as percentage of CD34+ cells that coexpressed CD133 and VEGFR2. On the basis of the peripheral-blood nucleated cell count, we also calculated the absolute number of CD34+CD133+VEGFR2-positive cells.
In a subgroup of 11 MMM patients and in five patients with other Ph-negative CMPDs, cells expressing the phenotype of EPCs were also assessed in samples enriched in CD34+ cells by means of immunomagnetic selection. Fifteen milliliters of heparinized peripheral blood underwent density gradient centrifugation (1,077 g/mL; Biocoll; Seromed, Berlin, Germany) and CD34+ cell purification using a MiniMacs CD34+ separation kit (Miltenyi Biotech; Bergish Gladback, Germany) in accordance with the manufacturer's instructions. The selected cells were stained with FITC–anti-CD34, PE–anti-CD133 (Miltenyi Biotech), and biotin-conjugated anti-VEGFR2 monoclonal antibodies revealed with PerCp-streptavidin. Selected CD34+ cells were also stained with the appropriate isotype controls, FITC–anti-CD34, PE-immunoglobulin G1, and biotin-conjugated immunoglobulin G1 monoclonal antibodies revealed with PerCp streptavidin. For each sample a minimum of 100,000 events was acquired.
In a subgroup of four healthy volunteer donors, the cell phenotype of EPCs was also assessed from buffy coats obtained during the preparation of transfusion products. Fifty milliliters of buffy coats was obtained by donation of one blood unit (450 mL) in a plastic bag (Maco-Pharma, Tourcoing, France) in preservative-free anticoagulant (citrate, phosphate, and dextrose) after centrifugation at 230 x g for 20 minutes at 20°C in a refrigerated centrifuge (Cryofuge 8500; Heraeus Instruments, Hanau, Germany). The buffy coats were then diluted in 100 to 200 mL of saline and centrifuged onto a Ficoll gradient (1,077 g/mL; Seromed) at 400 x g for 30 minutes at room temperature. Peripheral-blood mononuclear cells were washed twice with saline; immunomagnetic selection of CD34+ cells was then performed in accordance with the same procedure used for the CD34+ cell selection of MMM patients.
Functional Characterization of CD34+ Progenitor Cells
In four MMM patients, the endothelial differentiation potential of circulating CD34+ cells was assessed. CD34+ cells were purified using a MiniMacs CD34+ separation kit (Miltenyi Biotech) as described above. In all four patients, cell purity was greater than 90%. One to 2 x 105 cells (according to the number of CD34+ cells recovered after the selection procedure) were plated in liquid culture in a 24-well plate coated with fibronectin in EBM-2 medium (Cambrex Bio Science, Walkersville, MD) supplemented with 10% fetal calf serum, VEGF, and basic fibroblast growth factor (bFGF; Cambrex Bio Science). Medium was changed every 3 days. After 10 to 14 days of incubation at 37°C and 5% CO2, the colonies were counted using an inverted microscope. Colony-forming unit–endothelial (CFU-End) colonies were identified according to previously published morphologic criteria,23 and their endothelial phenotype confirmed by in situ fluorescence staining with antibodies to VE-cadherin (Bender Med System, San Bruno, CA), CD31 (1F11; Immunotech, Marseille, France), von Willebrand factor (Dako), and CD45 (2D1; BD Bioscience), as previously described.9 Alexa Fluor 488 or Alexa Fluor 594 conjugated goat antirabbit or goat antimouse antibodies (Molecular Probes, Eugene, OR) were used as secondary antibodies. Stained colonies were analyzed using an inverted fluorescence microscope (Leitz GmbH, Hoberochen, Germany).
Statistical Methods
Skewed variables were logarithmically transformed before they were entered for parametric analysis. Comparisons between groups were performed using the Mann-Whitney U test. Associations between patient characteristics (covariates) were assessed for pairs of numerical variables by Spearman's correlation, and for categoric and continuous variables by Wilcoxon-Mann-Whitney statistics. Multivariate logistic models were obtained by performing a backward elimination with a P value cutoff of .05, and then allowing any variable previously deleted to enter the final model if its P value was less than .05. Results were considered statistically significant when P values were less than .05. All computations were performed with Statistica software (Statsoft, Tulsa, OK).
RESULTS
Patient Characteristics
The hematologic and clinical characteristics of the population of 110 MMM patients studied are summarized in Table 1. Risk stratification according to the Dupriez-based prognostic scoring system at the time of blood sampling showed a preponderance of low- and intermediate-risk classes (90.9%). The median absolute number of circulating CD45+CD34+ HPCs in the overall population of MMM patients was 46.1 x 106/L (range, 0.58 to 1,300 x106/L). In 16 patients, the count was higher than 300 x 106/L, a value we have previously shown to be highly predictive of occurrence of blast transformation.5 When CD45+ cells were double stained with anti-CD34 and anti-CD38, the median percentage of CD45+CD34+ cells that also expressed CD38 was 27.5%, but the values ranged from 4% to 76%. Disease duration, spleen size, and the number of circulating immature myeloid cells and blasts had a significant univariate direct correlation with the CD45+CD34+ cell count (all P < .005).
Patients in whom prefibrotic MMM was diagnosed, at the time of blood sampling had a disease duration longer than patients with MMM diagnosed in the fibrotic stage (median, 79.5 v 20 months; P = .05). However, they were younger in age, and had milder splenomegaly, milder anemia, lower WBC count, higher platelet count, lower number of circulating CD34+ cells, and lower value of Dupriez and severity score (all P < .05).
CFU-End Formation From CD34+ Circulating Cells of MMM Patients
CFU-End were assayable in four of four MMM patients who were tested in a clonogenic assay. The number of CFU-End measured at 14 days ranged from 4 to 17 x 10–5 CD34+ cells. In all cases, the morphology of CFU-End was that of spindle-shaped cells surrounding a central cluster of round cells.23 On in situ immunolabeling, colonies strongly costained for CD31 and VE-cadherin; they were positive for von Willebrand factor but did not show any significant staining for CD45, thus confirming their endothelial phenotype.
CD34+CD133+VEGFR2-Positive Cells in the Peripheral Blood of MMM Patients and Controls
In 11 MMM patients, five patients with other Ph-negative CMPDs, and four healthy controls, peripheral-blood CD34+ cells were immunomagnetically selected and analyzed by flow cytometry. In MMM patients, the number of CD34+ cells coexpressing CD133 and VEGFR2 ranged from 1.3% to 18% (median, 2.2%). These figures were constantly higher than those in healthy participants, in whom the number of CD34+ coexpressing CD133 and VEGFR2 in buffy coat–derived cells ranged from 0.4% to 0.5% (median value, 0.5%; P < .04; Fig 1). In patients with other Ph-negative CMPDs, the median of CD34+CD133+VEGFR2-positive cells was 1.4% (range, 1.0% to 2.3%), which was significantly higher than that in normal controls (P < .02) but not statistically different from that in MMM patients.
In all 110 MMM patients, peripheral-blood cells were analyzed by flow cytometry without any selection procedure. Fifty-six of 110 patients (50.9%) had a measurable population of circulating CD34+CD133+VEGFR2-positive cells (> 0.03%). In six of 16 patients with other Ph-negative CMPDs (37.5%) and in three of the 14 (21%) normal controls, this cellular phenotype could also be detected (Fig 2). Patients with MMM had a median percentage of CD34+CD133+VEGFR2-positive cells of 0.26%, which was significantly higher (P = .04) than in both healthy controls (median, 0%) and in patients with other Ph-negative CMPDs (median, 0.1%). It is notable that although for patients with other Ph-negative CMPDs the number of CD34+CD133+VEGFR2-positive cells varied across a narrow range (0% to 9.2%), in patients with MMM the values ranged from 0% to 31.2%. Moreover, in 16 MMM patients (14.5%), the proportion of cells expressing this phenotype was more than 9.2% (ie, the highest value reported in other Ph-negative CMPDs). On the basis of the peripheral-blood nucleated cell count, MMM patients had an absolute number of CD34+CD133+VEGFR2-positive cells ranging from 0 to 19.8 x 106/L of blood.
Patients receiving chemotherapy had lower median EPC levels than patients out of therapy (median, 0% v 1.05%; P = .05). There was no difference in the median EPC levels between primary MMM and post-PV or post-ET MMM.
Because we found that patients with MMM presented a great variability in the number of detectable circulating CD34+CD133+VEGFR2-positive cells, we tried to identify the clinical and biologic features of the subset of patients with a high number of EPCs. To this end, we restricted the relation analysis of circulating CD34+CD133+VEGFR2-positive cells and disease features to those MMM patients who were not receiving cytoreductive treatment (n = 73). We found that those patients with a diagnosis of prefibrotic MMM (n = 13) had a higher level of EPCs than those with a diagnosis of MMM in a fibrotic stage (n = 60; median, 2.1% v 1.05%; P = .05). Moreover, age differed between those MMM patients with assayable EPCs (n = 24) and those without (n = 49; median, 62 v 51 years; P < .05).
Patients with higher numbers of CD34+CD133+VEGFR2-positive cells had a significantly smaller number of circulating immature myeloid cells and blasts (r = –0.49; P = .005) and smaller numbers of circulating CD45+CD34+ HPCs (r = –0.72; P < .001; Fig 3). When we analyzed the population of CD45+CD34+ cells also expressing the CD38 antigen, an inverse correlation was evidenced between the number of CD34+CD133+VEGFR2-positive cells and the more immature phenotype of HPCs (ie, CD45+CD34+CD38– cells; r = 0.45; P = .020).
A multiple linear regression was performed by selection of the above-described significantly correlated variables. The analysis yielded age and a diagnosis of MMM in a prefibrotic stage as independent predictors of the number of EPCs, with an adjusted R2 of 28%.
DISCUSSION
This study demonstrated that patients with MMM had a median percentage of circulating CD34+CD133+VEGFR2-positive EPCs significantly higher than that in healthy controls, and that in 14% of the examined patients, the number of these cells was higher than in patients with other Ph-negative CMPDs. Although increased circulating EPCs have been reported in diverse conditions associated with vascular injury, including vascular trauma,21 cardiovascular risk,24 exercise-induced ischemia,25 aortic aneurysm repair,26 and myocardial infarction,9 to the best of our knowledge, this is the first study that demonstrates evidence of a high number of circulating EPCs in the peripheral blood of patients with a hematologic malignancy.
A central role for the significance of the findings reported in this work is the careful characterization of the investigated cell population. To assess the number of circulating EPCs, we counted the percentage of CD34+CD133+VEGFR2-positive cells. This cell population, isolated from cord blood, bone marrow, or fetal liver, was reported to proliferate in vitro in an anchorage-independent manner and to be induced to differentiate into adherent CD133– mature endothelial cells.11 To exclude an aberrant expression of endothelial surface markers in our neoplastic CD34+ cell population, we tested the potential for growth of CD34+ cells in endothelial colonies from four patients with MMM. In all of them, we were able to grow colonies that were stained with specific endothelial markers. Thus, we reasonably assumed that EPCs defined by the three specific surface antigens function like EPCs. However, because currently there is no uniform definition of EPCs or experimental methods to discriminate between different populations of EPCs,27 this population could represent only one of multiple development steps of EPCs.
By means of our cytofluorimetric approach, we were able to detect circulating EPCs in more than half of the MMM patients, and in more than 30% of patients with other Ph-negative CMPDs, whereas only 21% of normal controls displayed this cellular phenotype in peripheral blood without any selection procedure. In MMM patients, CD34+CD133+VEGFR2-positive cells accounted for up to 31% of the circulating CD34+ cells, thus representing up to 19 x 106 nucleated cells/L of blood.
Increased trafficking of EPCs identifies a new and distinctive biologic characteristic of MMM. However, at variance with HPCs, the increase of which is fairly constant in patients with MMM, high trafficking of EPCs is restricted to a subset of patients. In this work we tried to identify the clinical features that characterize this population of patients.
By restricting the analysis to patients without chemotherapy, a higher number of EPCs resulted that were significantly associated with a younger age, a diagnosis of prefibrotic MMM, and a reduced number of immature myeloid cells and blasts in peripheral blood. These observations support the hypothesis that increased trafficking of EPCs characterizes those patients with a mild clinical phenotype. In fact, early age at diagnosis has been reported repeatedly to be associated with a good prognosis,28-30 and previous reports indicated that those patients diagnosed with a prefibrotic form of the disease have a favorable prognosis.31 Moreover, the inverse correlation with the number of immature myeloid cells and blasts in peripheral blood with the number of CD45+CD34+ HPCs strengthens the idea that mobilization of EPCs from the bone marrow is a phenomenon that occurs during the early phase of the disease when the process of HPC mobilization is absent or just beginning. The reason for a statistically independent association of EPC mobilization from bone marrow with young age in patients with MMM is unknown. This may reflect similar age-associated variations in the number of circulating EPCs in participants without malignancy, in whom mobilization is due to cardiovascular disorders.32
The mechanisms by which the mobilization of EPCs in peripheral blood of MMM patients is restricted to an early biologic phase, and a clinically indolent phenotype of MMM remains to be determined. The lack of any information on the clonal or nonclonal nature of circulating EPCs in CMPDs, and on the differential trafficking kinetics of HPCs and EPCs, makes any consideration of their clinical and biologic significance a matter of speculation. By assuming the same bone marrow origin for EPCs and HPCs, high levels of circulating EPCs in MMM could be the result of a common aberration of bone marrow stem and progenitor cells or of bone marrow microenvironment that could directly influence their mobilization.
Decreasing levels of circulating EPCs in patients with greater numbers of circulating HPCs may be due to a competitive relationship between EPCs and HPCs mobilization. This hypothesis was suggested by our data that document that the number of HPCs with the phenotype CD45+CD34+CD38– was inversely associated with the number of EPCs. Because low or undetectable levels of CD38 are characteristic of immature human hematopoietic cells with a high potential for self-renewal and multilineage differentiation,33 this observation is in keeping with a mechanism that leads to the increased proliferative and mobilization potential of HPCs at the expense of the mobilization of EPCs. However, the continuous stimulation of the compartment of EPCs may also lead to an eventual depletion or exhaustion of a presumed limited supply of EPCs. Finally, a progressive trapping of circulating EPCs by the spleen or other extramedullary organs could preclude their recirculation by progressively decreasing their levels in peripheral blood. This latter hypothesis is supported by a model of athymic immunodeficient nude rats in which infusion of radioactively labeled human EPCs resulted in the majority of the radioactivity being detected in the spleen and liver but not in the bones for up to 96 hours after injection.34 This is at variance with the observation that HPCs recirculate from the spleen to the peripheral blood and bone marrow.15 In support of this hypothesis, we recently reported that the percentage of CD34+CD133+VEGFR2-positive EPCs in the spleen of five MMM patients was significantly greater than that observed in the peripheral blood and greater than that in normal spleen tissue.15
In conclusion, we have demonstrated that EPCs circulate in increased numbers in the peripheral blood of MMM patients at an early phase of their disease. These results open a number of new perspectives in the pathogenesis of the disease. It remains to be determined if circulating EPCs have a role in neoangiogenesis and whether neoangiogenesis influences the disease progression.
Authors' Disclosures of Potential Conflicts of Interest
The authors indicated no potential conflicts of interest.
Acknowledgment
The following members of the Italian Registry for Myelofibrosis (RIMM) contributed patients to this study (person responsible, department, center city, listed by alphabetical order): Rosaria Agugliaro, Centro Trasfusionale, DH Ematologico, Ospedale San Biagio, Marsala. Giuliana Alimena, Ematologia, Policlinico Umberto I, Roma. Giuseppe Andreoni, Divisione di Medicina Interna, Ospedale Arnaboldi, Broni, Pavia. Mario Bazzan, Unità di Ematologia e Malattie Tromboemboliche, Ospedale Evangelico Valdese, Torino. Marilena Bertini, Divisione di Medicina, Ospedale Valdese di Torre Pellice, Torino. Enrico Bobbio Pallavicini, Servizio di Oncologia Medica ed Ematologia, Ospedale Maggiore di Crema (CR). Giorgio Broccia, Anna Di Tucci, Divisione di Ematologia-CTMO, Ospedale Businco, Cagliari. Sergio Cabibbo, Servizio di Medicina Trasfusionale, Ambulatorio Ematologico, Azienda Ospedaliera di Ragusa. Andrea Caenazzo, Clinica Medica II, Azienda Ospedaliera di Padova. Rossella Calori, Ematologia, Ospedale Maggiore Policlinico, Milano. Flavio Cardillo, Nadia Pezza, Dipartimento di Immunoematologia–DH Ematologico, Presidio Ospedaliero di Vasto (CH). Maria Ada Corbellini, Massimo Lazza, Giampiero Benetti, Divisione di Medicina 1 degrees e Servizio di Epatologia, Ospedale di Circolo Predabissi, Milano. Pietro Cirasino, Medicina Generale, Azienda Ospedaliera di Caserta. Alfonso D'Arco, UO di Medicina Interna, Ospedale Umberto I, Nocera Inferiore (SA). Giuseppina De Falco, Diego Di Maria, Divisione di Ematologia, Ospedale Ave Gratia Plena, San Felice a Castello (CE). Roberto Di Lorenzo, Reparto di Ematologia, Ospedale Civile Santo Spirito, Pescara. Dario Di Michele, Medicina Interna, Ospedale Mazzini, Teramo. Francesco Di Raimondo, Sezione di Ematologia, Ospedale V. Emanuele, Sezione Ferrarotto, Catania. Elena Elli, Enrico Pogliani, Divisione di Ematologia, Ospedale San Gerardo, Monza (MI). Adriana Gemiram, UO di Medicina Interna, Ospedale San Carlo Borromeo, Milano. Carlo Gentile, Medicina Valentini, Azienda Ospedaliera dell'Annunziata, Cosenza. Marco Giovannini, UO di Ematologia, Ospedale Umberto I, Frosinone. Paolo Gobbi, Clinica Medica I, IRCCS Policlinico S. Matteo, Pavia. Maria La Targia, UO di Medicina Interna III, Ospedale di Circolo, Busto Arsizio, Varese. Paola Lenhert, Carlo Loni, Medicina-DH Oncoematologico, Ospedale di Pietrasanta, Lucca. Marco Luoni, Oncologia, Ospedale di Legnano e Cuggiono (MI). Renata Mascolino, Servizio Talassemia, AO Vittorio Emanuele, Gela, Ragusa. Vincenzo Martinelli, Divisione di Ematologia, Policlinico Federico II, Napoli. Giuliana Nudo, Reparto di Medicina Interna Cosco, Azienda Ospedaliera di Cosenza. Daniele Perego, Medicina Generale, Ospedale di Circolo di Desio, Milano. Michele Pizzuti, Divisione di Ematologia, Azienda Ospedaliera San Carlo, Potenza. Eligio Radin, Ematologia, Ospedale Civile di Mestre. Umberto Recine, Day Hospital Oncoematologico, Ospedale Generale Santo Spirito, Roma. Maria Maddalena Ricetti, Divisione di Ematologia, Azienda Ospedaliera di Verona- Borgo Roma, Verona. Maria Savarè, Medicina I e II, Ospedale Civile Fornaroli, Magenta (Milano). Maria Rosaria Saieva, Divisione di Ematologia e CTMO, Azienda Ospedaliera San Camillo–Forlanini, Roma. Marzia Salvucci, UO di Ematologia, Ospedale Civile Santa Maria delle Croci, Ravenna. Gianfranco Silvestri, Unità di Medicina Interna, Ospedale Regionale Generale, Aosta. Emilio Usala, Emanuele Angelucci, Divisione di Ematologia-CTMO, Ospedale Businco, Cagliari.
NOTES
Supported by grant No. CS30 (2003) from Istituto Superiore di Sanità and by a grant (Ricerca Finalizzata 2002) from the Italian Ministry of Health.
Authors' disclosures of potential conflicts of interest are found at the end of this article.
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Moore MA, Hattori K, Heissig B, et al: Mobilization of endothelial and hematopoietic stem and progenitor cells by adenovirus-mediated elevation of serum levels of SDF-1, VEGF, and angiopoietin-1. Ann N Y Acad Sci 938:36-45, 2001
Mesa RA, Hanson CA, Rajkumar SV, et al: Evaluation and clinical correlations of bone marrow angiogenesis in myelofibrosis with myeloid metaplasia. Blood 96:3374-3380, 2000
Barosi G, Rosti V, Ramajoli I, et al: Spleen neoangiogenesis in patients with myelofibrosis with myeloid metaplasia. Br J Haematol 124:618-625, 2004
Barosi G, Ambrosetti A, Finelli C, et al: The Italian Consensus Conference on diagnostic criteria for myelofibrosis with myeloid metaplasia. Br J Haematol 104:730-737, 1999
Thiele J, Pierre R, Imbert M, et al: Chronic idiopathic myelofibrosis, in Jaffe ES, Harris N, Stan N, et al (eds): World Health Organization of Tumors of Hematopoietic and Lymphoid Tissues. Washington, DC, IARC Press, 2001, pp 35-38
Goulis J, Chau TN, Jordan S, et al: Thrombopoietin concentrations are low in patients with cirrhosis and thrombocytopenia and are restored after orthotopic liver transplantation. Gut 44:754-758, 1999
Dupriez B, Morel P, Demory JL, et al: Prognostic factors in agnogenic myeloid metaplasia: A report on 195 cases with a new scoring system. Blood 88:1013-1018, 1996
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Murphy S, Iland H, Rosenthal D, et al: Essential thrombocythemia: An interim report from the Polycythemia Vera Study Group. Semin Hematol 23:177-182, 1986
Gill M, Dias S, Hattori K, et al: Vascular trauma induces rapid but transient mobilization of VEGFR2+AC133+ endothelial precursor cells. Circ Res 88:167-174, 2001
Keeney M, Chin-Yee I, Weir K, et al: Single platform flow cytometric absolute CD34+ cell counts based on the ISHAGE guidelines: International Society of Hematotherapy and Graft Engineering. Cytometry 34:61-70, 1998
Asahara T, Mutohara T, Sullivan A, et al: Isolation of putative progenitor endothelial cells for angiogenesis. Science 275:964-967, 1997
Hill JM, Zalos G, Halcox JPJ, et al: Circulating endothelial progenitor cells, vascular function, and cardiovascular risk. N Engl J Med 348:593-600, 2003
Adams V, Lenk K, Linke A, et al: Increase of circulating endothelial progenitor cells in patients with coronary artery disease after exercise-induced ischemia. Arterioscler Thromb Vasc Biol 24:684-690, 2004
Eizawa T, Ikeda U, Murakami K, et al: Increase in circulating endothelial progenitor cells after aortic aneurysm repair. Heart Vessels 19:107-110, 2004
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Barosi G, Berzuini C, Liberato LN, et al: A prognostic classification of myelofibrosis with myeloid metaplasia. Br J Haematol 70:397-401, 1988
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Thiele J, Kvasnicka HM, Boeltken B, et al: Initial (prefibrotic) stages of idiopathic (primary) myelofibrosis (IMF): A clinicopathological study. Leukemia 13:1741-1748, 1999
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the Transplant Research Area
the Unit of Internal Medicine III
the Unit of Clinical Immunology, Immunohematology, and Transfusion Service
the Department of Pediatrics
the Laboratory of Clinical Epidemiology, Istituto di Ricovero e Cura a Carattere Scientifico Policlinico S. Matteo, Pavia, Italy
the Section of Hematology/Oncology
Department of Pathology, University of Illinois Cancer Center
University of Illinois College of Medicine
the Myeloproliferative Disorders Research Consortium, Chicago, IL
ABSTRACT
PURPOSE: Endothelial progenitor cells (EPCs) are present in circulation and contribute to vasculogenesis in adults. We measured the number of circulating EPCs in patients with myelofibrosis with myeloid metaplasia (MMM), and we examined the relationship between the number of EPCs and severity of the MMM disease process.
PATIENTS AND METHODS: The number of EPCs was measured by assaying the CD34+CD133+ vascular endothelial growth factor receptor 2 (VEGFR2) –positive cell phenotype in 110 MMM patients, 16 patients with other Philadelphia-negative chronic myeloproliferative disorders (Ph-negative CMPDs), and 14 healthy participants. In four MMM patients, the capacity of selected CD34+ cells to form endothelial colonies (CFU-End) in vitro was tested.
RESULTS: CD34+, CD133+, and VEGFR2-positive EPCs were detectable in unselected peripheral-blood cells of 50.9% MMM patients, 37.5% control patients, and 21% healthy participants. Patients with MMM had a median of 0.26% EPCs, significantly higher than that in healthy controls (median, 0%) and in patients with other Ph-negative CMPDs (median, 0.1%). In 14.5% of MMM patients, the numbers of EPCs were greater than the highest value found in patients with other Ph-negative CMPDs. CD34+ selected cells produced colony-forming unit–endothelial (CFU-End), which were vascular endothelial (VE) -cadherin positive, CD31+, von Willebrand factor positive, and CD45–. In MMM patients, the larger the number of EPCs, the smaller the number of circulating immature myeloid cells and circulating CD45+CD34+ hematopoietic progenitor cells. Increased numbers of EPCs were associated with younger age and a diagnosis of prefibrotic MMM.
CONCLUSION: Circulating EPCs are elevated in MMM patients in the early stage of the disease. Heightened mobilization of EPCs may represent an important mechanism for development of neoangiogenesis in MMM.
INTRODUCTION
Myelofibrosis with myeloid metaplasia (MMM) is a clonal Philadelphia-negative chronic myeloproliferative disorder (Ph-negative CMPD), characterized by a variable degree of bone marrow fibrosis, extramedullary hematopoiesis, and presence of immature myeloid cells in the peripheral blood.1,2 A high degree of trafficking of hematopoietic progenitor cells (HPCs) is a characteristic of the disease that discriminates MMM from other Ph-negative CMPDs (ie, polycythemia vera [PV] and essential thrombocythemia [ET]), and is predictive of disease severity and risk of blast transformation.3-6
There is strong evidence that a population of endothelial progenitor cells (EPCs) can be isolated from circulating mononuclear cells of normal patients,7 that different pathologic conditions can affect their number,7-9 and that they contribute to vasculogenesis in adult organisms.10 Immunophenotype identification of EPCs relies on coexpression of specific cell-surface proteins; in particular, endothelial markers, including CD34 and vascular endothelial growth factor receptor 2 (VEGFR2), and the human stem-cell marker CD133.8,9,11
Bone marrow releases EPCs as a result of mechanisms that are in part similar to those that regulate the release of HPCs.12 We therefore expected a high rate of EPC release from the bone marrow of MMM patients and we hypothesized that circulating EPCs may sustain the prominent increase of neoangiogenesis in the bone marrow13 and spleen.14 Hence, we assayed the number of circulating EPCs by cell phenotype from the peripheral blood of a large, well-characterized population of MMM patients. The aim of the study was to assess the ability of the number of EPCs to distinguish MMM from other Ph-negative CMPDs, and to examine the relationship among number of EPCs and type (primary v secondary), phase, and severity of the MMM disease process.
PATIENTS AND METHODS
Patients and Study Protocol
All consecutive patients with a diagnosis of MMM referred to the Hospital Istituto di Ricovero e Cura a Carratere Scientifico (IRCCS) Policlinico S. Matteo (Pavia, Italy) between July 2002 and June 2004 were included in this study. Our institution is a teaching and tertiary-care hospital, and is a major referral site for patients with MMM because it coordinates the Italian Registry for Myelofibrosis, a nationwide prospective database. The study protocol was approved by the Institutional Review Board of the Hospital IRCSS Policlinico S. Matteo. All patients provided written informed consent to be included in this study. Study design, data collection, analysis, and reporting were performed independently by a study committee.
One hundred ten patients with MMM were referred by 39 Italian centers participating in the Italian Registry for Myelofibrosis. In 94 patients, the diagnosis of MMM was established according to the Italian consensus conference criteria,15 by which the diagnosis held if diffuse bone marrow fibrosis was present and Ph chromosome or breakpoint cluster region–Abelson murine leukemia (BCR-ABL) rearrangement in peripheral-blood cells was absent (necessary criteria), and if an algorithm of optional criteria was satisfied. Sixteen additional patients who initially were classified as having atypical myeloproliferative disorders were also included. They had a bone marrow biopsy with an absent or only slightly increased reticulin fibrosis at diagnosis, and intense hyperplasia of megakaryocytes with abnormal or bizarre megakaryocytes appearing in clusters of more than four cellular elements. In these patients, the diagnosis of MMM was based on the absence of the Ph chromosome, and on the algorithm for optional criteria set by the Italian consensus conference.15 These patients frequently had a slight increase in platelet count; however, in no patient was the platelet count greater than 600 x109/L. These patients exhibited the corresponding prefibrotic stage of the disease accepted in the classification of myeloproliferative disorders proposed by the WHO.16
Ninety patients (81.8%) were classified as having primary MMM and 20 were classified as having secondary MMM (15 had post-PV MMM and five had post-ET MMM). Twenty-five patients (22.7%) were studied at the time of diagnosis, whereas 85 were enrolled during the course of their disease. Seventy-three patients (66.3%) were studied before the start of any cytoreductive treatment or when the cytoreduction had been stopped for at least 3 months. At the time of analysis, 26 patients were receiving hydroxyurea and 11 were receiving thalidomide treatment. Nine patients had been splenectomized and sampling was done more than 6 months after splenectomy.
All patients were screened according to a strict protocol. Together with the collection of demographic data, a sample of venous blood was drawn to assess a complete blood count with differential, circulating CD45+CD34+ cell count and phenotype, and circulating EPCs. The WBC count was corrected for the number of circulating erythroblasts. Circulating nucleated cells were classified as immature myeloid cells, erythroblasts, and blasts. Blasts were defined as undifferentiated cells with an immature nucleolated nucleus and basophilic cytoplasm with or without azurophilic granules. Spleen size was measured by ultrasonography. Two parameters of spleen size were recorded: the distance from the splenic tip to the costal margin, and the spleen index17 (ie, the product of the longitudinal by the transverse spleen axis, the latter defined as the maximal width of the organ). Liver enlargement was measured as the distance from the right costal margin in centimeters. Patients were assigned a prognostic score based on findings of Dupriez et al.18
Sixteen patients with other Ph-negative CMPDs were also studied for comparison (six women and 10 men; median age, 46 years; range, 22 to 70 years). They were categorized as having PV (nine patients) or ET (seven patients) according to the Polycythemia Vera Study Group (PVSG) criteria,19,20 after exclusion of MMM. Of the patients with other Ph-negative CMPDs, eight were studied at diagnosis, and eight were studied from 4 to 10 months after diagnosis but before any therapy had been started or when therapy had been stopped for at least 3 months.
Control samples were obtained from 15 healthy individuals (13 men and two women; median age, 49 years; range, 27 to 60 years), including four first-time donors. Each control participant was free of wounds, ulcers, recent surgery, or inflammatory or malignant disease that might influence the number of EPCs.7,21
Flow Cytometry Analysis of Circulating HPCs
To measure the numbers of HPCs in peripheral blood, 50 μL of EDTA-anticoagulated blood was stained with fluorescein isothiocyanate (FITC) –conjugated, phycoerythrin (PE) -conjugated, peridinin chlorophyll protein (PerCp) –conjugated, and laser dye styryl (LDS751) –conjugated monoclonal antibodies. The CD45-FITC/CD34-PE/LDS751 and CD38-FITC/CD34-PE/CD45-PerCp antibodies were used. Antibodies were obtained from Becton Dickinson (San Jose, CA), and the analyses were performed by a fluorescence activated cell sorter (FACS) calibur flow cytometer (Becton Dickinson). For each sample, 20,000 events were acquired and the percentage of positive cells was calculated based on the appropriate isotype control. The analysis was completed using the cell-gating guidelines recommended by the International Society of Hematotherapy and Graft Engineering.22
Flow Cytometry Analysis of Circulating Endothelial Progenitor Cells
Circulating EPCs were measured by phenotypic analysis in unselected peripheral-blood cells in all patients and normal controls. Blood samples were processed within 2 hours after they were drawn. Fifty microliters of EDTA-anticoagulated blood was incubated for 30 minutes at 4°C with 20 μL of FITC-conjugated anti-CD34, 10 μL of PE-conjugated anti-CD133, and 6 μL of 1:10 diluted biotin-conjugated anti-VEGFR2 (Sigma Chemical Co, St Louis, MO). Biotin conjugated anti-VEGFR2 was revealed using 5 μL of 1:5 diluted PerCp-streptavidin (Becton Dickinson, Pharmingen, San Diego, CA). Appropriate isotype controls were used for each staining procedure. One milliliter of lysis solution (Dako, Glostrup, Denmark) was added for 5 minutes at room temperature. The samples were then centrifuged and pellets were resuspended in 300 μL of phosphate buffer with 0.5% fetal calf serum (HyClone, Logan, UT). Two x 105 cells were acquired by a FACS calibur flow cytometer, and analyzed by CellQuest software (BD Biosciences, San Diego, CA). Because the number of dead/apoptotic cells was negligible, the analysis was performed excluding cellular debris in a side scatter/forward scatter (SSC/FSC) dot plot. CD34+ cells were electronically gated and the percentage of cells coexpressing CD133 and VEGFR2 was evaluated. For each sample a minimum of 100,000 events was acquired. Detection of EPCs by flow cytometry was defined by the presence of at least 0.03% nucleated cells coexpressing the three antigens over the background fluorescence. Results were expressed as percentage of CD34+ cells that coexpressed CD133 and VEGFR2. On the basis of the peripheral-blood nucleated cell count, we also calculated the absolute number of CD34+CD133+VEGFR2-positive cells.
In a subgroup of 11 MMM patients and in five patients with other Ph-negative CMPDs, cells expressing the phenotype of EPCs were also assessed in samples enriched in CD34+ cells by means of immunomagnetic selection. Fifteen milliliters of heparinized peripheral blood underwent density gradient centrifugation (1,077 g/mL; Biocoll; Seromed, Berlin, Germany) and CD34+ cell purification using a MiniMacs CD34+ separation kit (Miltenyi Biotech; Bergish Gladback, Germany) in accordance with the manufacturer's instructions. The selected cells were stained with FITC–anti-CD34, PE–anti-CD133 (Miltenyi Biotech), and biotin-conjugated anti-VEGFR2 monoclonal antibodies revealed with PerCp-streptavidin. Selected CD34+ cells were also stained with the appropriate isotype controls, FITC–anti-CD34, PE-immunoglobulin G1, and biotin-conjugated immunoglobulin G1 monoclonal antibodies revealed with PerCp streptavidin. For each sample a minimum of 100,000 events was acquired.
In a subgroup of four healthy volunteer donors, the cell phenotype of EPCs was also assessed from buffy coats obtained during the preparation of transfusion products. Fifty milliliters of buffy coats was obtained by donation of one blood unit (450 mL) in a plastic bag (Maco-Pharma, Tourcoing, France) in preservative-free anticoagulant (citrate, phosphate, and dextrose) after centrifugation at 230 x g for 20 minutes at 20°C in a refrigerated centrifuge (Cryofuge 8500; Heraeus Instruments, Hanau, Germany). The buffy coats were then diluted in 100 to 200 mL of saline and centrifuged onto a Ficoll gradient (1,077 g/mL; Seromed) at 400 x g for 30 minutes at room temperature. Peripheral-blood mononuclear cells were washed twice with saline; immunomagnetic selection of CD34+ cells was then performed in accordance with the same procedure used for the CD34+ cell selection of MMM patients.
Functional Characterization of CD34+ Progenitor Cells
In four MMM patients, the endothelial differentiation potential of circulating CD34+ cells was assessed. CD34+ cells were purified using a MiniMacs CD34+ separation kit (Miltenyi Biotech) as described above. In all four patients, cell purity was greater than 90%. One to 2 x 105 cells (according to the number of CD34+ cells recovered after the selection procedure) were plated in liquid culture in a 24-well plate coated with fibronectin in EBM-2 medium (Cambrex Bio Science, Walkersville, MD) supplemented with 10% fetal calf serum, VEGF, and basic fibroblast growth factor (bFGF; Cambrex Bio Science). Medium was changed every 3 days. After 10 to 14 days of incubation at 37°C and 5% CO2, the colonies were counted using an inverted microscope. Colony-forming unit–endothelial (CFU-End) colonies were identified according to previously published morphologic criteria,23 and their endothelial phenotype confirmed by in situ fluorescence staining with antibodies to VE-cadherin (Bender Med System, San Bruno, CA), CD31 (1F11; Immunotech, Marseille, France), von Willebrand factor (Dako), and CD45 (2D1; BD Bioscience), as previously described.9 Alexa Fluor 488 or Alexa Fluor 594 conjugated goat antirabbit or goat antimouse antibodies (Molecular Probes, Eugene, OR) were used as secondary antibodies. Stained colonies were analyzed using an inverted fluorescence microscope (Leitz GmbH, Hoberochen, Germany).
Statistical Methods
Skewed variables were logarithmically transformed before they were entered for parametric analysis. Comparisons between groups were performed using the Mann-Whitney U test. Associations between patient characteristics (covariates) were assessed for pairs of numerical variables by Spearman's correlation, and for categoric and continuous variables by Wilcoxon-Mann-Whitney statistics. Multivariate logistic models were obtained by performing a backward elimination with a P value cutoff of .05, and then allowing any variable previously deleted to enter the final model if its P value was less than .05. Results were considered statistically significant when P values were less than .05. All computations were performed with Statistica software (Statsoft, Tulsa, OK).
RESULTS
Patient Characteristics
The hematologic and clinical characteristics of the population of 110 MMM patients studied are summarized in Table 1. Risk stratification according to the Dupriez-based prognostic scoring system at the time of blood sampling showed a preponderance of low- and intermediate-risk classes (90.9%). The median absolute number of circulating CD45+CD34+ HPCs in the overall population of MMM patients was 46.1 x 106/L (range, 0.58 to 1,300 x106/L). In 16 patients, the count was higher than 300 x 106/L, a value we have previously shown to be highly predictive of occurrence of blast transformation.5 When CD45+ cells were double stained with anti-CD34 and anti-CD38, the median percentage of CD45+CD34+ cells that also expressed CD38 was 27.5%, but the values ranged from 4% to 76%. Disease duration, spleen size, and the number of circulating immature myeloid cells and blasts had a significant univariate direct correlation with the CD45+CD34+ cell count (all P < .005).
Patients in whom prefibrotic MMM was diagnosed, at the time of blood sampling had a disease duration longer than patients with MMM diagnosed in the fibrotic stage (median, 79.5 v 20 months; P = .05). However, they were younger in age, and had milder splenomegaly, milder anemia, lower WBC count, higher platelet count, lower number of circulating CD34+ cells, and lower value of Dupriez and severity score (all P < .05).
CFU-End Formation From CD34+ Circulating Cells of MMM Patients
CFU-End were assayable in four of four MMM patients who were tested in a clonogenic assay. The number of CFU-End measured at 14 days ranged from 4 to 17 x 10–5 CD34+ cells. In all cases, the morphology of CFU-End was that of spindle-shaped cells surrounding a central cluster of round cells.23 On in situ immunolabeling, colonies strongly costained for CD31 and VE-cadherin; they were positive for von Willebrand factor but did not show any significant staining for CD45, thus confirming their endothelial phenotype.
CD34+CD133+VEGFR2-Positive Cells in the Peripheral Blood of MMM Patients and Controls
In 11 MMM patients, five patients with other Ph-negative CMPDs, and four healthy controls, peripheral-blood CD34+ cells were immunomagnetically selected and analyzed by flow cytometry. In MMM patients, the number of CD34+ cells coexpressing CD133 and VEGFR2 ranged from 1.3% to 18% (median, 2.2%). These figures were constantly higher than those in healthy participants, in whom the number of CD34+ coexpressing CD133 and VEGFR2 in buffy coat–derived cells ranged from 0.4% to 0.5% (median value, 0.5%; P < .04; Fig 1). In patients with other Ph-negative CMPDs, the median of CD34+CD133+VEGFR2-positive cells was 1.4% (range, 1.0% to 2.3%), which was significantly higher than that in normal controls (P < .02) but not statistically different from that in MMM patients.
In all 110 MMM patients, peripheral-blood cells were analyzed by flow cytometry without any selection procedure. Fifty-six of 110 patients (50.9%) had a measurable population of circulating CD34+CD133+VEGFR2-positive cells (> 0.03%). In six of 16 patients with other Ph-negative CMPDs (37.5%) and in three of the 14 (21%) normal controls, this cellular phenotype could also be detected (Fig 2). Patients with MMM had a median percentage of CD34+CD133+VEGFR2-positive cells of 0.26%, which was significantly higher (P = .04) than in both healthy controls (median, 0%) and in patients with other Ph-negative CMPDs (median, 0.1%). It is notable that although for patients with other Ph-negative CMPDs the number of CD34+CD133+VEGFR2-positive cells varied across a narrow range (0% to 9.2%), in patients with MMM the values ranged from 0% to 31.2%. Moreover, in 16 MMM patients (14.5%), the proportion of cells expressing this phenotype was more than 9.2% (ie, the highest value reported in other Ph-negative CMPDs). On the basis of the peripheral-blood nucleated cell count, MMM patients had an absolute number of CD34+CD133+VEGFR2-positive cells ranging from 0 to 19.8 x 106/L of blood.
Patients receiving chemotherapy had lower median EPC levels than patients out of therapy (median, 0% v 1.05%; P = .05). There was no difference in the median EPC levels between primary MMM and post-PV or post-ET MMM.
Because we found that patients with MMM presented a great variability in the number of detectable circulating CD34+CD133+VEGFR2-positive cells, we tried to identify the clinical and biologic features of the subset of patients with a high number of EPCs. To this end, we restricted the relation analysis of circulating CD34+CD133+VEGFR2-positive cells and disease features to those MMM patients who were not receiving cytoreductive treatment (n = 73). We found that those patients with a diagnosis of prefibrotic MMM (n = 13) had a higher level of EPCs than those with a diagnosis of MMM in a fibrotic stage (n = 60; median, 2.1% v 1.05%; P = .05). Moreover, age differed between those MMM patients with assayable EPCs (n = 24) and those without (n = 49; median, 62 v 51 years; P < .05).
Patients with higher numbers of CD34+CD133+VEGFR2-positive cells had a significantly smaller number of circulating immature myeloid cells and blasts (r = –0.49; P = .005) and smaller numbers of circulating CD45+CD34+ HPCs (r = –0.72; P < .001; Fig 3). When we analyzed the population of CD45+CD34+ cells also expressing the CD38 antigen, an inverse correlation was evidenced between the number of CD34+CD133+VEGFR2-positive cells and the more immature phenotype of HPCs (ie, CD45+CD34+CD38– cells; r = 0.45; P = .020).
A multiple linear regression was performed by selection of the above-described significantly correlated variables. The analysis yielded age and a diagnosis of MMM in a prefibrotic stage as independent predictors of the number of EPCs, with an adjusted R2 of 28%.
DISCUSSION
This study demonstrated that patients with MMM had a median percentage of circulating CD34+CD133+VEGFR2-positive EPCs significantly higher than that in healthy controls, and that in 14% of the examined patients, the number of these cells was higher than in patients with other Ph-negative CMPDs. Although increased circulating EPCs have been reported in diverse conditions associated with vascular injury, including vascular trauma,21 cardiovascular risk,24 exercise-induced ischemia,25 aortic aneurysm repair,26 and myocardial infarction,9 to the best of our knowledge, this is the first study that demonstrates evidence of a high number of circulating EPCs in the peripheral blood of patients with a hematologic malignancy.
A central role for the significance of the findings reported in this work is the careful characterization of the investigated cell population. To assess the number of circulating EPCs, we counted the percentage of CD34+CD133+VEGFR2-positive cells. This cell population, isolated from cord blood, bone marrow, or fetal liver, was reported to proliferate in vitro in an anchorage-independent manner and to be induced to differentiate into adherent CD133– mature endothelial cells.11 To exclude an aberrant expression of endothelial surface markers in our neoplastic CD34+ cell population, we tested the potential for growth of CD34+ cells in endothelial colonies from four patients with MMM. In all of them, we were able to grow colonies that were stained with specific endothelial markers. Thus, we reasonably assumed that EPCs defined by the three specific surface antigens function like EPCs. However, because currently there is no uniform definition of EPCs or experimental methods to discriminate between different populations of EPCs,27 this population could represent only one of multiple development steps of EPCs.
By means of our cytofluorimetric approach, we were able to detect circulating EPCs in more than half of the MMM patients, and in more than 30% of patients with other Ph-negative CMPDs, whereas only 21% of normal controls displayed this cellular phenotype in peripheral blood without any selection procedure. In MMM patients, CD34+CD133+VEGFR2-positive cells accounted for up to 31% of the circulating CD34+ cells, thus representing up to 19 x 106 nucleated cells/L of blood.
Increased trafficking of EPCs identifies a new and distinctive biologic characteristic of MMM. However, at variance with HPCs, the increase of which is fairly constant in patients with MMM, high trafficking of EPCs is restricted to a subset of patients. In this work we tried to identify the clinical features that characterize this population of patients.
By restricting the analysis to patients without chemotherapy, a higher number of EPCs resulted that were significantly associated with a younger age, a diagnosis of prefibrotic MMM, and a reduced number of immature myeloid cells and blasts in peripheral blood. These observations support the hypothesis that increased trafficking of EPCs characterizes those patients with a mild clinical phenotype. In fact, early age at diagnosis has been reported repeatedly to be associated with a good prognosis,28-30 and previous reports indicated that those patients diagnosed with a prefibrotic form of the disease have a favorable prognosis.31 Moreover, the inverse correlation with the number of immature myeloid cells and blasts in peripheral blood with the number of CD45+CD34+ HPCs strengthens the idea that mobilization of EPCs from the bone marrow is a phenomenon that occurs during the early phase of the disease when the process of HPC mobilization is absent or just beginning. The reason for a statistically independent association of EPC mobilization from bone marrow with young age in patients with MMM is unknown. This may reflect similar age-associated variations in the number of circulating EPCs in participants without malignancy, in whom mobilization is due to cardiovascular disorders.32
The mechanisms by which the mobilization of EPCs in peripheral blood of MMM patients is restricted to an early biologic phase, and a clinically indolent phenotype of MMM remains to be determined. The lack of any information on the clonal or nonclonal nature of circulating EPCs in CMPDs, and on the differential trafficking kinetics of HPCs and EPCs, makes any consideration of their clinical and biologic significance a matter of speculation. By assuming the same bone marrow origin for EPCs and HPCs, high levels of circulating EPCs in MMM could be the result of a common aberration of bone marrow stem and progenitor cells or of bone marrow microenvironment that could directly influence their mobilization.
Decreasing levels of circulating EPCs in patients with greater numbers of circulating HPCs may be due to a competitive relationship between EPCs and HPCs mobilization. This hypothesis was suggested by our data that document that the number of HPCs with the phenotype CD45+CD34+CD38– was inversely associated with the number of EPCs. Because low or undetectable levels of CD38 are characteristic of immature human hematopoietic cells with a high potential for self-renewal and multilineage differentiation,33 this observation is in keeping with a mechanism that leads to the increased proliferative and mobilization potential of HPCs at the expense of the mobilization of EPCs. However, the continuous stimulation of the compartment of EPCs may also lead to an eventual depletion or exhaustion of a presumed limited supply of EPCs. Finally, a progressive trapping of circulating EPCs by the spleen or other extramedullary organs could preclude their recirculation by progressively decreasing their levels in peripheral blood. This latter hypothesis is supported by a model of athymic immunodeficient nude rats in which infusion of radioactively labeled human EPCs resulted in the majority of the radioactivity being detected in the spleen and liver but not in the bones for up to 96 hours after injection.34 This is at variance with the observation that HPCs recirculate from the spleen to the peripheral blood and bone marrow.15 In support of this hypothesis, we recently reported that the percentage of CD34+CD133+VEGFR2-positive EPCs in the spleen of five MMM patients was significantly greater than that observed in the peripheral blood and greater than that in normal spleen tissue.15
In conclusion, we have demonstrated that EPCs circulate in increased numbers in the peripheral blood of MMM patients at an early phase of their disease. These results open a number of new perspectives in the pathogenesis of the disease. It remains to be determined if circulating EPCs have a role in neoangiogenesis and whether neoangiogenesis influences the disease progression.
Authors' Disclosures of Potential Conflicts of Interest
The authors indicated no potential conflicts of interest.
Acknowledgment
The following members of the Italian Registry for Myelofibrosis (RIMM) contributed patients to this study (person responsible, department, center city, listed by alphabetical order): Rosaria Agugliaro, Centro Trasfusionale, DH Ematologico, Ospedale San Biagio, Marsala. Giuliana Alimena, Ematologia, Policlinico Umberto I, Roma. Giuseppe Andreoni, Divisione di Medicina Interna, Ospedale Arnaboldi, Broni, Pavia. Mario Bazzan, Unità di Ematologia e Malattie Tromboemboliche, Ospedale Evangelico Valdese, Torino. Marilena Bertini, Divisione di Medicina, Ospedale Valdese di Torre Pellice, Torino. Enrico Bobbio Pallavicini, Servizio di Oncologia Medica ed Ematologia, Ospedale Maggiore di Crema (CR). Giorgio Broccia, Anna Di Tucci, Divisione di Ematologia-CTMO, Ospedale Businco, Cagliari. Sergio Cabibbo, Servizio di Medicina Trasfusionale, Ambulatorio Ematologico, Azienda Ospedaliera di Ragusa. Andrea Caenazzo, Clinica Medica II, Azienda Ospedaliera di Padova. Rossella Calori, Ematologia, Ospedale Maggiore Policlinico, Milano. Flavio Cardillo, Nadia Pezza, Dipartimento di Immunoematologia–DH Ematologico, Presidio Ospedaliero di Vasto (CH). Maria Ada Corbellini, Massimo Lazza, Giampiero Benetti, Divisione di Medicina 1 degrees e Servizio di Epatologia, Ospedale di Circolo Predabissi, Milano. Pietro Cirasino, Medicina Generale, Azienda Ospedaliera di Caserta. Alfonso D'Arco, UO di Medicina Interna, Ospedale Umberto I, Nocera Inferiore (SA). Giuseppina De Falco, Diego Di Maria, Divisione di Ematologia, Ospedale Ave Gratia Plena, San Felice a Castello (CE). Roberto Di Lorenzo, Reparto di Ematologia, Ospedale Civile Santo Spirito, Pescara. Dario Di Michele, Medicina Interna, Ospedale Mazzini, Teramo. Francesco Di Raimondo, Sezione di Ematologia, Ospedale V. Emanuele, Sezione Ferrarotto, Catania. Elena Elli, Enrico Pogliani, Divisione di Ematologia, Ospedale San Gerardo, Monza (MI). Adriana Gemiram, UO di Medicina Interna, Ospedale San Carlo Borromeo, Milano. Carlo Gentile, Medicina Valentini, Azienda Ospedaliera dell'Annunziata, Cosenza. Marco Giovannini, UO di Ematologia, Ospedale Umberto I, Frosinone. Paolo Gobbi, Clinica Medica I, IRCCS Policlinico S. Matteo, Pavia. Maria La Targia, UO di Medicina Interna III, Ospedale di Circolo, Busto Arsizio, Varese. Paola Lenhert, Carlo Loni, Medicina-DH Oncoematologico, Ospedale di Pietrasanta, Lucca. Marco Luoni, Oncologia, Ospedale di Legnano e Cuggiono (MI). Renata Mascolino, Servizio Talassemia, AO Vittorio Emanuele, Gela, Ragusa. Vincenzo Martinelli, Divisione di Ematologia, Policlinico Federico II, Napoli. Giuliana Nudo, Reparto di Medicina Interna Cosco, Azienda Ospedaliera di Cosenza. Daniele Perego, Medicina Generale, Ospedale di Circolo di Desio, Milano. Michele Pizzuti, Divisione di Ematologia, Azienda Ospedaliera San Carlo, Potenza. Eligio Radin, Ematologia, Ospedale Civile di Mestre. Umberto Recine, Day Hospital Oncoematologico, Ospedale Generale Santo Spirito, Roma. Maria Maddalena Ricetti, Divisione di Ematologia, Azienda Ospedaliera di Verona- Borgo Roma, Verona. Maria Savarè, Medicina I e II, Ospedale Civile Fornaroli, Magenta (Milano). Maria Rosaria Saieva, Divisione di Ematologia e CTMO, Azienda Ospedaliera San Camillo–Forlanini, Roma. Marzia Salvucci, UO di Ematologia, Ospedale Civile Santa Maria delle Croci, Ravenna. Gianfranco Silvestri, Unità di Medicina Interna, Ospedale Regionale Generale, Aosta. Emilio Usala, Emanuele Angelucci, Divisione di Ematologia-CTMO, Ospedale Businco, Cagliari.
NOTES
Supported by grant No. CS30 (2003) from Istituto Superiore di Sanità and by a grant (Ricerca Finalizzata 2002) from the Italian Ministry of Health.
Authors' disclosures of potential conflicts of interest are found at the end of this article.
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