| | Elevated thrombopoietin levels in patients with myelofibrosis may not be due to enhanced production of thrombopoietin by bone marrowReceived 20 February 2002; accepted 9 April 2002. Abstract Thrombopoietin (TPO) is recognized as the primary regulator of megakaryocyte and platelet production. Two alternative hypotheses for the mechanism of regulation have been proposed: (1) platelet and/or megakaryocyte mass regulate circulating TPO levels by binding to TPO through TPO receptors (c-MPL), with subsequent internalization and degradation of the protein; (2) TPO mRNA produced by bone marrow (BM) stromal cells or BM cells modulates blood TPO levels or platelet counts. In myeloproliferative disorders (MPD), including primary myelofibrosis (MF) and essential thrombocythemia (ET), elevated blood TPO levels occur despite increased platelet and megakaryocyte mass. Therefore, in these diseases, elevated blood TPO levels cannot be explained by the first mechanism. The present study, was designed to measure TPO mRNA production by BM mononuclear cells and BM stromal cells using a relative RT–PCR technique, to verify the second mechanism. We found no increase of TPO mRNA production in either BM cells or in BM stromal cells in patients with MF and ET. Furthermore, in those patients with MF who had elevated plasma TPO levels, TPO mRNA levels in bone marrow fibroblasts (BMFs) or BM cells were not elevated as compared with controls. Therefore, we concluded that in patients with MF, the elevated plasma TPO levels are not due to enhanced production of TPO mRNA either by BMF, or BM cells. The TPO receptor (c-MPL) abnormalities including reduced MPL protein levels or defective TPO induced signal transduction pathways are the likely mechanisms.
1. Introduction  Thrombopoietin (TPO), the ligand for the MPL receptor, is the primary regulator of platelet and megakaryocyte production [1], [2], [3], [4]. The mechanisms regulating the serum TPO levels and platelet counts in circulation, however, have been a subject of debate. Two possible mechanisms have been proposed: (1) platelets and/or megakaryocytes regulate serum TPO levels by binding via MPL receptors, internalizing and degrading the protein [5], [6], [7], [8], [9], [10]; (2) TPO mRNA of bone marrow (BM) stromal cells [11] or BM cells modulates circulating platelet counts and TPO levels [12]. The second hypothesis has been found that though TPO is synthesized in the liver, kidney, spleen, and lung [13], the levels of TPO mRNA in BM cells [12] or stromal cells [11], [13], and not in these tissues vary in accordance with circulating TPO concentrations or platelet numbers. The first mechanism has been favored, because platelets bind and metabolize TPO [6], [7], and because transfusion of platelets into thrombocytopenic animals reduces TPO levels towards normal [5], [7]. However, in myeloproliferative disorders (MPD), including agnogenic myeloid metaplasia (AMM) or called idiopathic myelofibrosis (MF), essential thrombocythemia (ET), and polycythemia vera (PV), different mechanisms must be proposed, because blood TPO levels are elevated [15], [16], [17], [18], despite increased total platelet and megakaryocyte mass which would be expected to have increased uptake and internalization of TPO. Reduced expression of c-MPL receptors in PV and IMF was found by Moliterno et al. [19], and in ET by Horikawa et al. [20]. No information regarding the mRNA production by BM stromal cells or BM cells has been published in this disease. Therefore, the present study was designed to measure TPO mRNA in BM stromal cells and BM mononuclear cells in patients with MPD, especially in MF/AMM. 2. Materials and methods 2.1. Patients and RNA extraction Peripheral blood and BM samples, after informed consent, were obtained from patients seen in the Hematology/Oncology Department at Brookdale Hospital. The diagnosis of myeloproliferative disorder was made according to clinical and laboratory criteria [21]. BM cells were obtained from aspiration samples and used for RNA isolation. Bone marrow fibroblasts (BMFs) cells were cultured from BM biopsy specimens as previously described [22]. Peripheral blood mononuclear cells were isolated from freshly drawn peripheral blood in EDTA, then by Ficoll-Hypaque density gradient method. Total RNA was extracted from peripheral blood cells (PB), BM cells and BMF using a commercially available kit (Ambion, Austin, TX) according to the manufacture’s instructions. 2.2. Optimization of relative quantitative-RT–PCR Multiplex PCR was used for the relative quantification of TPO mRNA levels. Quantum RNA 18S internal standards (Ambion, Austin, TX) were used with gene specific primers in these experiments. The Quantum RNA module allows rRNA to be used as an internal control in multiplex RT–PCR experiments by using blocked 18S rRNA primers (competimer) to adjust 18S rRNA amplification [23], [24], [25], [26]. The 18S primers were mixed in different ratios with 18S competimers (Ambion, Austin, TX) according to manufacturer’s instructions. A concentration of 5:5, primer:competimer was used to measure the relative expression levels of TPO as compared to the amounts of 18S products. For semi-quantitative RT–PCR, various cycle numbers and cDNA concentrations were tested to ensure linear range of amplification. Reverse transcription was performed with two-fold dilution of total RNA (16–500 ng) followed by PCR amplifications at different cycles (10–45) in presence of 4μCi α dATP (Amersham). Products were visualized by electrophoresis on 5% polyacrylamide gel and exposed to X-ray film as described below. Signal was measured (BioRad Molecular Imager) and plotted with cycle number, and a straight line was obtained for samples in linear range of amplification [23], [24], [25], [26], [27]. A constant increase in amplified products was observed with diluted total RNA (16–250ng) at 20–40 cycles indicating a linear increase in final product. On the basis of these results TPO and 18S rRNA, amplifications were carried out for 35 cycles with 0.2μg of total RNA. 2.3. RT–PCR analysis Specific RNA transcripts were amplified using 0.2μg of total RNA. RNA was reverse transcribed with AMV reverse transcriptase from Avian myeloblastosis virus for first strand cDNA synthesis and random hexamers (Promega, Madison, WI) at 48°C for 45min. Tfl DNA polymerase (Promega, Madison, WI) was used for second strand cDNA synthesis and DNA amplifications with TPO specific primers according to manufacturer’s instructions. cDNA was amplified in 50μl with 0.2mM of each dNTPs, 1mM of MgSO4, 50pmol of each primer, 5U of Tfl DNA polymerase and trace amount of α dATP (Amersham). Samples were denatured at 94°C for 2min, followed by 35 cycles of amplification (94°C for 30s, 60°C for 1min, 68°C for 2min) and one cycle of final extension at 68°C for 7min, in a thermocycler (Perkin-Elmer Cetus Corporation). Primers used for PCR were as follows: TPO sense primer (5′-GTAGGGGTGGGCGTTGGAGCAG-3′ and anti-sense primer 5′-AAGTGGCAGCAGGGATTCAGAGC-3′ [28]. Quantum RNA 18S internal standards (Ambion, Austin, TX) were used with gene specific primers. 18S primers were mixed in different ratios with 18S Competimers™ (Ambion, Austin, TX) according to manufacturer’s instructions. A concentration of 5:5, primer:competimer was used in further experiments. For relative quantitative RT–PCR, 35 cycles and 0.2μg RNA were used as described which were tested to have linear range of amplification. The 6μl aliquots of final PCR product were size fractionated on 5% polyacrylamide gel at 140V for 100min. The gel was transferred on a 3M filter paper, covered with plastic and exposed to X-ray film (Kodak, Arlington Heights, IL) for 1–3h at −70°C with intensifying screens. 2.4. Quantitation and statistical analysis Quantification of the amplified PCR products and the northern blot signals were performed by densitometric scanning using the soft ware (BioRad, GS-710 calibrated imaging) as per the manufacturer’s instructions. The quantity of TPO mRNA calculated was then normalized with 18S rRNA and analyzed with one-way analysis of variance (ANOVA) using statistical package PRIZM. 3. Results 3.1. TPO mRNA levels in bone marrow fibroblasts Five patients with AMM, six with ET, and seven with iron deficiency anemia (normal blood WBC, platelet counts and TPO levels) serving as controls were studied. TPO mRNA levels were expressed as density of signals of mRNA TPO divided by signals of 18S rRNA. As shown in Fig. 1, TPO mRNA levels (mean±S.E.) were 1.27±0.21 in MF, 0.80±0.16 in ET, and 1.27±0.16 in controls. No statistically significant difference was found among the groups. No correlation between blood TPO levels and TPO mRNA levels in BMF was found (data not shown). Of the patients with MF and ET, four with MF and two with ET had elevated plasma TPO levels (controls were  pg/ml in our laboratory [15]). The TPO mRNA levels of BMF in these four patients with MF (mean±S.E.; 1.19±0.26) also were found to have no significantly elevated levels as compared to controls. No statistical comparisons were made in these two ET patients, because of the small sizes of the samples. 3.2. TPO mRNA levels in bone marrow mononuclear cells Eight patients with MF, 12 with ET, and 17 controls (10 with iron deficiency anemia, seven with other chronic disease process, all with normal platelet counts, white blood cell counts and blood TPO levels) were studied. As shown in Fig. 2, TPO mRNA levels expressed as mean±S.E. were 1.68±0.41, 1.63±0.15, 1.18±0.12, in MF, ET and controls, respectively. No statistically significant difference was found among the groups. Furthermore, no correlation was found between blood TPO levels and BM TPO mRNA levels in these patients (r=0.27, P=0.39). No correlation was found between TPO mRNA levels and platelet counts as well (r=0.40, P=0.18). Among the patients with MF and ET, six with MF and three with ET had elevated plasma TPO levels (controls, 185pg/ml±26pg/ml). These six MF patients with elevated TPO levels also were found to have no significantly elevated TPO mRNA levels in the BM cells (mean±S.E.; 1.51±0.54). Because of the small size of the samples of these three ET patients, no statistical comparison with controls were made. 3.3. TPO mRNA levels in peripheral blood mononuclear cells Nine patients with MF, 11 with ET, seven with PV compared with 11 normal volunteer controls. As shown in Fig. 3, TPO mRNA levels (mean±S.E.) were 1.38±0.14, 1.29±0.13, 0.88±0.14 in MF, ET, PV, respectively, and with 1.06±0.11 in controls. There are no statistically significant differences found among the different groups. 3.4. Correlation between bone marrow and peripheral blood mononuclear cells TPO mRNA levels Ten patients (four, ET; six, MF) had simultaneously BM and PB collection for TPO mRNA determination that correlated with BM and PB mRNA levels. Significant correlation between mononuclear cells TPO mRNA in BM and PB was found (r=0.65, P=0.04). 3.5. Northern blot analysis Most of the samples were not be able to be detected by Northern blots analysis for TPO mRNA, probably due to too little quantity of mRNA detected (data not shown). 4. Discussion TPO mRNA levels in human BM or blood samples could not be detected by Northern Blot analysis. Relative quantitative RT–PCR analysis by analyzing the PCR product at the linear range of both the target and the reference amplicon is a reasonably accurate estimate of mRNA quantification [29], [30]. The expression of TPO mRNA has been observed in several tissues including liver, kidney, brain, skeletal muscle, intestine, spleen and BM [31], [32]. Using Northern blot analysis, the major sites are in the liver, kidney and to a lesser extent in the spleen and BM [31], [32]. The exact cell types which produce TPO, however, are not well defined. In BM, there are evidences that BM stromal cells are the main source of TPO production using in situ hybridization technique [13] and RT–PCR technique [11]. In the present study, the finding of the presence of TPO mRNA in the PB and mononuclear cells in the BM suggests that cells other than stromal cells are also the sources of TPO production. McCarty et al. [12] also demonstrated that BM cells produced TPO mRNA in mice concurred with our findings. In the thrombocytopenic murine model, TPO mRNA was not increased in liver or kidney, but was found to be increased in BM cells by McCarty et al. by RT–PCR technique [12]. However, in other studies by Stoffel et al. [14], using RNAase protection assay, neither BM nor spleen TPO mRNA was detectable in thrombocytopenic mice, and spleen TPO mRNA was not increased by RT–PCR. In the rat thrombocytopenic model by Yang et al. [5], liver TPO mRNA also was not increased, but BM TPO mRNA was not studied. Thus, in the presence of thrombocytopenic stress, liver, kidney TPO mRNA are not increased, and BM TPO mRNA was not detected or was not done, and was shown to be elevated in one study [12]. In human studies, by using in situ hybridization technique, Sungaran et al. [13], found TPO mRNA mainly in BM stromal cells, and it appeared to show increased intensity in staining in BM stromal cells, and to a lesser degree in other BM cells and with no significant change in intensity of staining in liver, kidney, or spleen in patients with ITP or aplastic anemia. In another study, by using RT–PCR technique, Hirayama et al. [11], found BM stromal cells were the sources of TPO production based on the finding that serum TPO levels were correlated to stromal cells TPO mRNA levels in patients with ITP and aplastic anemia [11], but in this study, TPO mRNA production by BM cells was not measured. Thus, in these two human studies, BM stromal cells, and to a lesser degree, other BM cells and not liver, kidney, or spleen cells appeared to modulate the blood TPO levels in the presence of thrombocytopenic stress. Our study deals with different diseases (MF and ET) and we found no significant difference in TPO mRNA production between patients and control subjects either in BM hematopoietic cells or in stromal cells, as shown in Fig. 1and Fig. 2. Our findings are in agreement with Hirayama’s data that in two patients with ET, TPO mRNA in BM stromal cells was not different from controls. From the present study, in patients with MF or ET, no enhanced TPO mRNA production was found in either the BMF or the BM cells. Furthermore, in those MF patients who have elevated plasma TPO levels also was found to have no enhanced TPO mRNA production in either the BMF or BM cells [11]. Therefore, we concluded that in patients with MF, the elevated plasma TPO levels are not due to enhanced production of TPO mRNA from BMF, or BM cells. Reduced expression of c-MPL by Western blot analysis and impaired response by TPO induced tyrosine phosphorylation were found in platelets from patients with PV and MF [19]. Reduced expression of c-MPL protein and mRNA was demonstrated in platelets from patients with ET [20]. These two studies suggest that in patients with MPD including MF, ET, and PV, the elevated TPO levels are likely related to reduced c-MPL receptor protein levels or defective signal transduction pathway. Mutation analysis of the c-MPL gene in patients with ET, however, disclosed no apparent mutations [33]. All these results and the findings from the present study suggest that elevated TPO levels in patients with MPD disease, including MF, may be related to c-MPL receptor abnormalities and not to TPO overproduction by BM. Induced over expression of TPO in mice by transplantation of BM cells infected with retroviral carrying TPO cDNA led to MF [34] or fatal myeloproliferative syndrome [35]. Thus, TPO may be related to the pathogenesis of MF and other MPD disorders. Further dissection of c-MPL abnormalities may lead to better understanding of this disease and may actually lead to therapeutic applications. The finding in the present study that mononuclear cells from blood also carry TPO mRNA expression suggests that PB like monocytes may be the source of TPO production, since stem cells and platelets do not have TPO mRNA by in situ hybridization technique [13]. 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Blood. 1997;90:4369. MEDLINE Division of Medical Oncology and Hematology, Brookdale University Hospital and Medical Center, Brooklyn, NY 11219, USA  Corresponding author. Present address: Maimonides Hospital Medical Center, Department of Hematology/Oncology, 953 49th Street, 5th Floor, Brooklyn, NY 11219, USA. Tel.: +1-718-283-8297; fax: 1-718-635-7110.
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