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The role of the liver in the production of thrombopoietin compared with erythropoietin

Jelkmann, Wolfgang

European Journal of Gastroenterology & Hepatology: July 2001 - Volume 13 - Issue 7 - p 791-801
Review in Depth

The liver plays an important role in the production of haemopoietic hormones. It acts as the primary site of synthesis of erythropoietin (EPO) in the fetal stage, and it is the predominant thrombopoietin (TPO)-producing organ for life. In contrast to that of EPO and other liver proteins, the hepatic synthesis of TPO is influenced little by external signals. Hepatocytes express the TPO gene in a constitutive way, i.e. irrespective of the level of platelets in blood. Megakaryocytes and platelets remove the hormone from blood by means of their high-affinity TPO receptors. Normally, the plasma level of TPO is relatively low ( 10−−12 mol/l). However, in thrombocytopenic states due to marrow failure or bleeding, the concentration of circulating TPO may increase greatly. The simple feedback regulation by TPO and its target cells is efficient in maintaining constant platelet numbers in healthy people. Persisting thrombocytopenia develops only in severe liver or marrow failure. On the other hand, an increase in circulating TPO and interleukin 6 (IL-6) may cause reactive thrombocytosis in inflammatory diseases, including cancer. The indications for recombinant human thrombopoietin (rHuTPO) therapy and its impact on transfusion medicine are still under investigation.

Institute of Physiology, Medical University of Lubeck, Lubeck, Germany

Correspondence to Wolfgang Jelkmann, Institute of Physiology, Medical University of Lubeck, Ratzeburger Allee 160, 23538 Lubeck, Germany Tel: +49 451 500 4150; fax: +49 451 500 4151; e-mail:

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The liver is the principal site of synthesis of most of the plasma proteins, with the exception of immunoglobulins. The liver also removes proteins from the circulation by means of its specific cell-surface receptors. Acute or chronic liver diseases are therefore often associated with abnormalities in protein metabolism. Clinical interest has focused on the fatal haematological consequences of liver disease, which include thrombocytopenia, coagulation defects, and anaemia [1].

There are important ontogenetic changes with respect to the role of the liver in human haematopoiesis. Early in the fetal stage, the liver is the main site of production of blood cells. Erythropoiesis and megakaryopoiesis reach their maximum level at the thirteenth to sixteenth week of gestation in parenchymatous hepatic tissue [2]. Interestingly, the fetal liver is also the primary site of production of the relevant haemopoietic hormones. These are the glycoproteins thrombopoietin (TPO) and erythropoietin (EPO), and the somatomedins. TPO is the primary regulator of megakaryopoiesis and platelet formation. Its synthesis continues in the liver for life. In contrast, after birth the kidneys take over from the liver as the main site of the synthesis of EPO, which is a primary regulator of erythropoiesis. Somatomedin C (insulin-like growth factor I) may also promote erythropoiesis [3,4], but the physiological significance of this still needs to be elucidated. Other functions of the liver related to erythropoiesis include the production of transferrin and the storage of iron, vitamin B12 and folate (Table 1).

Table 1

Table 1

After decades of laborious search [5], convincing evidence has been provided recently for the existence of a specific thrombopoiesis-stimulating factor (TPO) originating from the liver and other organs [6,7]. This article will focus on the biochemical and pathophysiological relevance of this novel hepatic hormone. Furthermore, initial reports on the clinical use of recombinant human thrombopoietin (rHuTPO) for treatment of thrombocytopenia in tumour patients undergoing chemotherapy will be critically reviewed. In the face of structural and biological similarities between TPO and EPO, some aspects of the role of the liver in EPO synthesis will also be summarized. For more comprehensive information on EPO and the liver, the reader is referred to previous reviews [8–10].

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Molecular biology of thrombopoietin

Earlier clinical observations and laboratory studies had provided evidence suggesting that megakaryopoiesis is a regulated process and under the control of a humoral factor [5]. The normal number of platelets in human blood is 2.3 (range 1.5–4.0) × 1011 platelets/litre [11]. Intraindividually, the number of platelets is maintained relatively constantly. The rate of production of platelets increases significantly for 1–2 weeks following the induction of thrombocytopenia, e.g. after haemorrhage. The proliferation and differentiation of megakaryocytic progenitors is stimulated by a variety of growth factors and cytokines, including stem cell factor, granulocyte monocyte colony-stimulating factor (GM-CSF), EPO, and the interleukins (IL) 1, 3, 4, 6, 7 and 11 [12]. Prior to the identification of the specific thrombopoiesis-stimulating hormone TPO, its receptor was discovered as a member of the haematopoietic growth factor receptor superfamily class I [13]. The TPO receptor was originally termed MPL, because it is the human homologue of the murine myeloproliferative leukaemia oncogene-encoded receptor, v-mpl oncogene [14]. TPO binding to its receptor induces tyrosine phosphorylation of several cellular proteins [15], but the details of the TPO signalling cascade are only beginning to be understood. In vitro studies have shown that TPO promotes the viability of multipotent CD34+CD38− haemopoietic progenitor cells [16]. TPO potently stimulates the proliferation of megakaryocyte colony-forming units (CFU-Meg) and the maturation and sequestration of megakaryocytes. In the absence of TPO, immature megakaryocytes undergo apoptosis [17]. TPO also augments agonist-induced platelet aggregation and secretion [18].

In 1994, several groups reported the isolation of TPO cDNA of human and a few other species [19–24]. The primary translation product of the six exons of the human TPO gene is composed of 353 amino acids, including a secretory leader sequence of 21 amino acids (Table 2). Circulating TPO presents with truncated forms [25]. Alternative splicing of TPO mRNA may be one reason for the heterogeneity of circulating TPO [26,27]. In addition, there are a number of potential proteolytic cleavage sites in the human TPO molecule, suggesting that some of the various forms may result from proteolysis through the action of thrombin [28]. At one of the two basic amino acid sequences (Arg–Arg at positions 153 and 154) the molecule can be separated into two domains [29]. The NH2-terminal part (amino acids 1–153) exhibits 23% sequence identity with EPO. It is this domain that binds to the TPO receptor [23]. In contrast, the CO2-terminal domain shows no homology with any known protein. This region contains multiple N- and O-linked carbohydrate side chains and appears to prolong the in vivo survival of the hormone. Its deletion does not abolish the in vitro activity of TPO [30]. Accordingly, for treatment of severe thrombocytopenias, not only glycosylated full-length recombinant human thrombopoietin (rHuTPO) but also a truncated recombinant protein of the 163 NH2-terminal amino acids has been produced. The latter is termed PEG-conjugated recombinant human megakaryocyte growth and development factor (PEG-rHuMGDF), because polyethylene glycol (PEG) is added to stabilize the molecule in vivo[31].

Table 2

Table 2

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Control of hepatic thrombopoietin and erythropoietin synthesis

Initial studies of the localization of the site of synthesis of TPO were carried out in experimental animals. TPO mRNA was detected in liver extracts from fetal animals, and in liver and kidney extracts from adult animals [19,22]. Thus, TPO derives from the same organs that produce EPO. Other tissues and cells that are reportedly capable of expressing TPO mRNA are in vivo smooth muscle, spleen and bone marrow, and in vitro endothelial cells and fibroblasts [6,7]. Because TPO mRNA is also abundant in human amygdala and hippocampus, and the NH2-terminus of TPO shares a significant homology with neurotrophins, it has been proposed that TPO could have specific functions in the brain [32].

Elegant studies by Qian et al. have shown clearly that the liver is the major site of TPO production [33]. When these authors replaced normal livers in wild-type mice by livers from TPO knock-out mice, platelet numbers in the recipients decreased to < 50% of normal. An in situ hybridization study on human tissues has shown that the TPO mRNA-expressing cells are hepatocytes in the liver and proximal tubular cells in the kidney [34]. The latter finding is of interest since EPO synthesis has been localized to interstitial peritubular fibroblasts in rodent kidneys [35–38]. Recent studies from our laboratory utilizing reverse transcription/competitive polymerase chain reaction (RT-PCR) have shown that the liver accounts for 95% of the total body TPO mRNA [39] and for 80% of the total body EPO mRNA [40] in human fetuses. The TPO mRNA content relative to the total cellular RNA is almost equal in liver, kidney, spleen and bone marrow. It is the high organ weight that makes the liver the dominant site of TPO production [39]. The liver remains the principal site of synthesis of TPO after birth [41]. Thrombocytopenia associated with liver cirrhosis is associated with low plasma TPO concentrations and reduced hepatic TPO mRNA levels both in children [42] and in adults [43,44].

In contrast, there is a switch of the main production site of EPO from the fetal liver to the kidneys after birth. Kochling et al. recently identified DNA sequences upstream of the EPO gene that are required for kidney-specific induction [45]. Both renal and hepatic EPO gene expression are induced by hypoxia. Studies in experimental animals have revealed that the hepatic EPO mRNA amounts to about 15% of the total body EPO mRNA in anaemic mice [46] and 20–50% in anaemic or hypoxaemic rats [47,48]. Faced with the considerable contribution of the liver to total body EPO mRNA in adult rodents, we have raised the question as to why the liver is incapable of substituting for the kidneys more effectively as an EPO-producing site in humans suffering from anaemia due to chronic renal failure [10]. There are three possible explanations. First, there may be species differences in that the adult human liver expresses relatively little EPO mRNA. Second, hepatic EPO mRNA may be insufficiently translated. Along these lines, attempts have failed to extract bioactive EPO from livers of hypoxic animals [49]. Third, chronically diseased kidneys may suppress the hepatic production of EPO by an unknown humoral mechanism. Evidence for the latter concept has been provided in animal models of chronic renal failure [50,51].

In situ hybridization and cell separation studies have shown that EPO mRNA is expressed predominantly by hepatocytes in the liver of rodents [36,52]. In addition, EPO mRNA has been localized in transgenic animals to Ito cells, which lie in a perisinusoidal position within the space of Disse [53]. Hepatic EPO production increases under conditions of lowered oxygen supply. The human hepatoma cell lines Hep3B and HepG2 are a widely used in vitro model for the study of oxygen-dependent EPO gene expression [54,55]. In these cells, a hypoxia-inducible factor (HIF-1) binds to the hypoxia-responsive enhancer in the 3′-flanking sequence of the EPO gene. HIF-1 is a dimeric protein composed of two different subunits, the 120-kDa HIF-1α and the 91–94-kDa HIF-1β [56]. HIF-1 is a ubiquitous transcription factor. It also controls the expression of other genes that encode proteins that are protective against hypoxia, such as vascular endothelial growth factor and distinct glycolytic enzymes [57,58]. HIF-1 is thought to play an important role in the oxygen-dependent zonation of the key enzymes of carbohydrate metabolism in the liver [59]. Apart from the effect of hypoxia, several agents may modulate EPO production in human hepatoma cultures similar to their effects in vivo. The EPO synthesis-increasing factors include IL-6 [60], thyroid hormones [61], and vitamin A [62], while inhibition is exerted by the pro-inflammatory cytokines IL-1 and tumour necrosis factor alpha (TNF-α) [60,63] and by reactive O2 species [64].

Unlike that of EPO, the synthesis of TPO in human hepatoma cultures appears to be little influenced by external signals, as demonstrated in cell cultures treated with various cytokines and growth factors [65,66]. Evidence for unregulated, i.e. constitutive, TPO gene expression in vivo has been provided by RT-PCR and Northern blot analyses of TPO mRNA levels in hepatic and renal tissue extracts from normal versus thrombocytopenic or thrombocytotic mice [67–70], and by in situ hybridization on human tissues [34]. TPO mRNA expression is induced in bone marrow of thrombocytopenic people [34], but the contribution of this site to circulating TPO is still unproven. In view of the constitutive expression of the TPO gene, other mechanisms must be involved in the regulation of the rate of thrombopoiesis and the number of platelets in blood. Fielder et al. have shown that the injection of platelets into TPO receptor knock-out mice produces a decrease in the plasma TPO concentration [68]. There is good evidence to assume that platelets internalize and degrade circulating TPO [68,70] by means of their high-affinity TPO receptors [71]. Megakaryocytes also appear to take up the hormone [72–74]. Figure 1 demonstrates the simple feedback regulation between the plasma TPO level on the one side and the mass of megakaryocytes and platelets on the other. As a result, the plasma TPO concentration increases with the development of thrombocytopenia, such as following chemotherapy for cancer (Fig. 2).

Fig. 1

Fig. 1

Fig. 2

Fig. 2

Whether the liver is also involved in the metabolism of TPO is currently unclear. Previous studies on the role of the liver in the catabolism of EPO have provided evidence that the rate of hepatic catabolism of this glycoprotein hormone is minor at best. Dinkelaar et al. have shown that the plasma disappearance half-time of endogenous EPO is little elevated in rats in which liver failure is induced by the administration of the hepatotoxic agent D-galactosamine-HCl [75].

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Concentration of circulating thrombopoietin in health and liver disease

The present immunoassays for circulating TPO have not been standardized. Thus, while intrastudy differences in TPO concentrations may provide valuable scientific and diagnostic information, the comparison of data between different studies and laboratories is usually not valid. Median or mean values reported for the concentration of plasma or serum TPO have ranged from 20 to 240 pg/ml in normal people, independently of gender. Values in the lower range were generally measured by commercial enzyme-linked immunosorbent assays with monoclonal antibodies [76–78], whereas values in the upper range were reported by most [79–83], but not all [84,85], investigators who developed assay systems based on polyclonal antibodies from rabbits immunized with rHuTPO or rHuMGDF. Possibly, assays utilizing polyclonal antibodies are also sensitive to truncated forms of TPO in blood. Still other authors have expressed their results in molar units (normal value ∼1 fmol/ml) [86,87]. Folman et al. have reported that TPO levels in serum are on average 3.4 times higher than in plasma [88], but other investigators measured much smaller differences when serum and plasma values were compared [81,82,85,89,90]. It is recommended to use plasma rather than serum for assay of TPO, because platelets release TPO during blood clotting. Finally, it is worth noting that children have higher plasma TPO levels than adults [87].

As mentioned above, the concentration of circulating TPO is normally elevated in thrombocytopenic states not associated with hepatic disease, e.g. after chemotherapy. However, TPO levels are increased less significantly in thrombocytopenic patients with severe acute or chronic liver failure [42–44,91–95]. Evidence for impaired TPO synthesis has been provided by the demonstration of lowered TPO mRNA levels in cirrhotic liver tissue from children [42] and adults [43,44]. In the study by Wolber et al. it was shown that hepatic TPO mRNA levels, blood platelet counts and the concentration of plasma proteins of liver origin were lowest in acute liver failure, intermediate in decompensated cirrhosis, and close to normal in compensated cirrhosis [42]. Ishikawa et al. have reported that the decrease in the TPO mRNA content of the liver of rats treated with the hepatotoxic agent dimethylnitrosamine is due in part to reduced TPO mRNA expression and in part to the loss of hepatocytes [44].

In humans suffering from end-stage liver failure, the concentration of TPO in blood increases after successful orthotopic liver transplantation reaching a maximum 5–6 days after surgery [42,43,93,96]. The peripheral platelet count reaches normal values 2 weeks after orthotopic liver transplantation [43,91,93]. Although some investigators have claimed that plasma TPO is not lowered in cirrhosis [92,95], one has to conclude that the respective TPO production rates are lowered, taking into account the slowed metabolism of the hormone at reduced platelet and megakaryocyte mass. The restitution of normal peripheral platelet counts following liver transplantation is not associated with reduced platelet destruction and spleen size, as shown by computed tomography volumetry [97]. If patients do not survive their liver transplantation, TPO levels reach extremely high values in association with the postoperative thrombocytopenia [98]. Non-transplanted thrombocytopenic patients with cirrhosis who undergo only portal decompression by insertion of a transjugular intrahepatic portosystemic stent shunt show no rise in the concentration of circulating TPO or platelets [91]. Thus, the thrombocytopenia associated with liver disease is due at least partly to impaired TPO production. Accordingly, plasma TPO increases in response to the thrombocytopenia caused by interferon (alpha or beta) therapy for treatment of chronic hepatitis C infection in non-cirrhotic patients [99,100], but not in patients with cirrhosis [99]. Lack of TPO due to impaired liver function is also considered a risk factor contributing to HIV-associated thrombocytopenia [101]. Lowered plasma TPO levels in patients with veno-occlusive liver disease following high-dose chemotherapy have been noted in one report [102], but not in another [103].

Compared with TPO, less interest has been paid to changes in EPO production related to acute or chronic non-malignant liver diseases. In patients with end-stage renal failure, serum EPO may increase after hepatitis B or C infection, resulting in an improvement of red cell status [104]. The mechanism of this increase still needs to be identified. Clinically relevant lack of EPO has not been reported in cirrhotic patients [105], which is probably associated with the secondary role of the liver in the synthesis of this haemopoietic hormone.

Patients with malignant liver diseases sometimes exhibit thrombocytosis or erythrocytosis. Tumour cell-associated TPO production is likely in view of the demonstration of elevated TPO concentrations in the serum of patients with hepatoblastoma and significant TPO mRNA levels in hepatoma tissue samples [106] as well as in human and rat hepatoma cell line cultures [65,107].

Erythrocytosis has been observed in 3–12% of patients with hepatocellular carcinoma [108–110]. In several other cases, the development of erythrocytosis may have been prevented by the factors causing anaemia of inflammation in tumour patients [111,112]. Kew and Fisher measured a mean serum EPO concentration of 77 U/l in 65 patients with hepatocellular carcinoma, compared with a normal value of 22 U/l [110]. By immunohistochemistry, EPO could be demonstrated in the carcinoma cells but not in the surrounding normal hepatic tissue [113].

Patients with non-hepatic solid tumours, infections or autoimmune diseases are often anaemic. This complication has been called ‘anaemia of chronic disease’ or ‘anaemia of inflammation', and been explained by increased haemolysis, bleeding, lowered EPO production, impaired iron mobilization, and reduced proliferation of erythrocytic progenitors due to the action of distinct pro-inflammatory cytokines [111,112]. In turn, patients with malignancies and inflammations often present with reactive thrombocytosis. The combination of anaemia, thrombocytosis and leucocytosis has been termed ‘haematological stress syndrome’ [114]. Bioassay [115] and immunoassay [76,78–82,87,116] measurements have shown that the concentration of circulating TPO is abnormally high in reactive thrombocytosis associated with cancer or autoimmune diseases, such as inflammatory bowel disease. TPO levels also increase greatly in patients with fulminant septicaemia [117]. Basically, the elevated TPO levels could be due to an increased lifespan of the hormone or due to stimulation of its production. Support for the latter concept is provided by our finding that the immunomodulatory peptide IL-6 enhances TPO gene expression in human hepatoma cell cultures [66]. Furthermore, an analysis of the published sequence of the 5′-flanking region of the human TPO gene [118] shows that there are several potential IL-6 responsive elements. Thus, TPO resembles acute phase proteins [66,90,117]. In rat hepatocytes in primary culture, hepatocyte growth factor/scatter factor, but not IL-6 or other cytokines, was found to increase TPO mRNA expression [119]. A recent study shows that the majority of carcinoma cell lines of various origins express TPO mRNA variants [120], thus indicating that the thrombocytosis in tumour patients could be a paraneoplastic syndrome.

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Possible role of rHuTPO versus rHuEPO for hormone replacement therapy

Several differences have to be noted when the insufficient TPO production in liver disease is compared with the insufficient EPO production in chronic renal disease. Lack of TPO and the resulting impairment of megakaryopoiesis and platelet production is only one contributing factor in the pathogenesis of the thrombocytopenia in chronic liver disease. In addition, the platelet half-life is commonly reduced in association with splenomegaly and portal hypertension. Other causes include acute severe blood loss and increased platelet consumption secondary to disseminated intravascular coagulation. The thrombocytopenia is rarely threatening enough to warrant platelet transfusion. A threshold platelet count of 10 000/μl has been proposed for prophylactic platelet transfusion if risk factors for haemorrhage, such as fever, evidence of bleeding or the need for an invasive procedure, are absent [121,122]. Note that the application of rHuTPO in emergency cases of thrombocytopenia is not useful anyway because of the 5-day latency period for significant stimulation of thrombopoiesis.

While serious episodes of bleeding are usually not to be expected until the blood platelet count falls below 5% of the normal, anaemic people suffer from symptoms of tissue hypoxia, such as shortness of breath, tachycardia and angina pectoris, if the blood haemoglobin concentration falls below just 50% of normal. In healthy people, the decrease in blood haemoglobin concentration will be prevented by an increase in EPO gene expression primarily in the kidneys. This regulation is missing in patients with chronic renal failure whose plasma EPO concentration is by one to two orders of magnitude lower than in people with normal renal function at similar blood haemoglobin concentrations. Here, the treatment with rHuEPO is a true replacement therapy. rHuEPO corrects the anaemia in virtually all predialysis and dialysis patients with chronic renal failure [123] provided the iron supply is sufficient [124]. Apart from impaired availability of iron, infectious and inflammatory diseases may reduce responsiveness to rHuEPO, because various immunomodulatory cytokines inhibit the proliferation of erythrocytic progenitors [111,125]. Measurements of C-reactive protein and baseline fibrinogen concentrations in serum may provide early recognition of the probability of response to rHuEPO [126].

During rHuEPO therapy, storage iron is mobilized for use in red cell production, which leads to a decline in serum ferritin levels. Combined with phlebotomy, this effect has been considered a tool for treatment of iron overload associated with frequent blood transfusion [127]. A histological study has proven the regression of the deposited iron in liver biopsies [128]. With respect to liver disease, it is also of interest that the treatment with rHuEPO can increase antibody titres after hepatitis B vaccination in dialysis patients [129].

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Clinical trials with recombinant thrombopoietin in patients with solid tumours

The results of preclinical studies in normal animals and in myelosuppressed thrombocytopenic mice, dogs and monkeys have been reviewed previously [6,7,130]. To sum up, the administration of recombinant TPO or MGDF produces an increase in the concentration of circulating platelets beginning after 3–5 days. Thus, the action of TPO appears to be due to a stimulation of the proliferation and differentiation of megakaryocytic progenitors rather than an immediate sequestration of platelets from preformed megakaryocytes. TPO increases the number, size and fluidity of the megakaryocytes in the bone marrow. In addition, it increases the growth of more primitive pluripotent haemopoietic progenitor cells.

rHuTPO and PEG-rHuMGDF have been administered to tumour patients before and after chemotherapy in phase-I/II trials, which are described in more detail elsewhere [131]. Vadhan-Raj et al. treated 12 sarcoma patients 3 weeks before chemotherapy with a single intravenous dose of rHuTPO (0.3–2.4 μg/kg body weight) [132]. This treatment resulted in a dose-dependent increase in the concentration of circulating platelets. The patients’ platelets were morphologically and functionally unaltered. Haematocrit and white blood cell numbers were unaffected. Note, however, that the drug did not only promote the proliferation and differentiation of cells in the megakaryocytic lineage; it also expanded the pool of multilineage progenitors in bone marrow and their mobilization into peripheral circulation [133]. Very similar observations were made when PEG-rHuMGDF was given to patients with solid tumours before [134,135] or after [136] chemotherapy. In an additional study, granulocyte colony-stimulating factor (G-CSF, 5 μg/kg body weight) was combined with PEG-rHuMGDF at doses of 0.03–5 μg/kg body weight for daily subcutaneous injection in a randomized, blinded phase-I trial in 41 patients with advanced cancer after dose-intensive chemotherapy with carboplatin and cyclophosphamide [137]. The platelet nadir and the recovery to baseline platelet count occurred earlier in the patients given PEG-rHuMGDF than in the placebo control group. Furthermore, the degree of the mobilization of peripheral blood progenitor cells (PBPC) on day 15 after chemotherapy was significantly greater in patients treated with PEG-rHuMGDF in combination with G-CSF, suggesting that this treatment might allow for more efficient collection of stem cells for autologous or allogeneic transplantation [137]. In the studies by Fanucchi et al. [136] and Basser et al. [137], thrombotic complications were seen in two patients, but the relation of these events to the treatment with PEG-rHuMGDF was not clarified.

The safety and activity of rHuTPO as a PBPC mobilizer in combination with G-CSF was evaluated recently in 29 breast cancer patients treated with high-dose chemotherapy followed by PBPC reinfusion. This regimen resulted in an accelerated granulocyte and platelet recovery and decreased blood cell transfusion requirements. In the majority of patients, only a single apheresis procedure was needed [138].

Thus, the isolation and in vitro expression of the human TPO gene has been followed rapidly by the clinical demonstration that both the full-length rHuTPO and the truncated pegylated rHuMGDF are capable of stimulating platelet production in tumour patients undergoing chemotherapy. Still, several medical and economic concerns will have to be overcome before TPO can be approved for use in clinical routine, thereby following in the steps of the haemopoietic growth factors EPO, G-CSF and GM-CSF. With respect to the administration of rHuEPO for prevention of chemotherapy-induced anaemia, the impact and cost-effectiveness of therapy have been well studied [139,140]. Similarly, practice guidelines for the use of G-CSF and GM-CSF for protection against post-chemotherapy fibril neutropenia have been published [141,142]. In contrast, the significance of recombinant thrombopoiesis-stimulating growth factors has not been defined so far. First, severe thrombocytopenia is not commonly seen with standard chemotherapy regimens [143]. Clinically significant bleeding does not usually occur, even if the platelet concentration falls to 20 000/μl blood [144]. Second, rHuTPO as a new supportive treatment modality will have to compete with the established standard, i.e. platelet transfusions. Note here that despite major advances in safety, allogeneic platelet transfusions carry a remaining risk for immunological and infectious diseases [121,122]. Third, rHuIL-11, which is another potent stimulator of the growth of megakaryocytic progenitors [145], has recently received FDA approval for the prevention of severe thrombocytopenia in high-risk patients receiving myelosuppressive chemotherapy. Fourth, one of the companies investigating PEG-rHuMGDF (Amgen Inc., Thousand Oaks, CA, USA) has discontinued further development due to evidence of neutralizing antibodies in a few patients participating in cancer clinical trials and in additional people in platelet donor clinical trials.

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The author's studies are supported by the Deutsche Forschungsgemeinschaft (DFG, SFB 367-C8). The expert secretarial work of Ms Lisa Zieske in the preparation of this manuscript is gratefully acknowledged.

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 Of special interest

•• Of outstanding interest

1. Chisholm M. Haematological disorders in liver disease. In: Liver and Biliary Disease. Wright R, Millward-Sadler GH, Alberti KGMM, Karran S (editors). London: W.B. Saunders Company; 1995. pp. 189–214.
2. Ohkita T, Enzan H. Morphological studies on the hepatic hematopoiesis of human fetuses. In: Genetics, Structure and Function of Blood Cells. Hollan SR, Gardos G, Sarkadi B (editors). Budapest: Akademiai Kiado; 1980. pp. 53–64.
3. Kurtz A, Jelkmann W, Bauer C. A new candidate for the regulation of erythropoiesis: insulin-like growth factor I. FEBS Lett 1982; 149: 105–108.
4. Kurtz A, Zapf J, Eckardt KU, Clemons G, Froesch ER, Bauer C. Insulin-like growth factor I stimulates erythropoiesis in hypophysectomized rats. Proc Natl Acad Sci USA 1988; 85: 7825–7829.
5. •McDonald TP. Thrombopoietin: its biology, purification, and characterization. Exp Hematol 1988; 16: 201–205. An interesting survey of early attempts to identify the specific thrombopoiesis-regulating hormone.
6. Eaton DL, de Sauvage FJ. Thrombopoietin: the primary regulator of megakaryocytopoiesis and thrombopoiesis. Exp Hematol 1997; 25: 1–7.
7. Kaushansky K. Thrombopoietin. N Engl J Med 1998; 339: 746–754.
8. Fried W. The liver as a source of extrarenal erythropoietin production. Blood 1972; 40: 671–677.
9. Fisher JW. Extrarenal erythropoietin production. J Lab Clin Med 1979; 93: 695–699.
10. Jelkmann W. Erythropoietin: structure, control of production, and function. Physiol Rev 1992; 72: 449–489.
11. Trowbridge EA, Martin JF, Slater DN, Kishk YT, Warren CW, Harley PJ, Woodcock B. The origin of platelet count and volume. Clin Phys Physiol Meas 1984; 5: 145–170.
12. Avraham H. Regulation of megakaryocytopoiesis. Stem Cells (Dayt) 1993; 11: 499–510.
13. ••Vigon I, Mornon JP, Cocault L, Mitjavila MT, Tambourin P, Gisselbrecht S, Souyri M. Molecular cloning and characterization of MPL, the human homolog of the v-mpl oncogene: identification of a member of the hematopoietic growth factor receptor superfamily. Proc Natl Acad Sci USA 1992; 89: 5640–5644. Initial report on MPL, a novel member of the haemopoietic growth factor receptor superfamily that was later shown to mediate thrombopoietic activity and identified as thrombopoietin receptor.
14. Souyri M. Mpl: from an acute myeloproliferative virus to the isolation of the long sought thrombopoietin. Semin Hematol 1998; 35: 222–231.
15. Drachman JG, Griffin JD, Kaushansky K. The c-Mpl ligand (thrombopoietin) stimulates tyrosine phosphorylation of Jak2, Shc, and c-Mpl. J Biol Chem 1995; 270: 4979–4982.
16. Borge OJ, Ramsfjell V, Cui L, Jacobsen SE. Ability of early acting cytokines to directly promote survival and suppress apoptosis of human primitive CD34+CD38– bone marrow cells with multilineage potential at the single-cell level: key role of thrombopoietin. Blood 1997; 90: 2282–2292.
17. Osada M, Komeno T, Todokoro K, Takizawa M, Kojima H, Suzukawa K. et al. Immature megakaryocytes undergo apoptosis in the absence of thrombopoietin. Exp Hematol 1999; 27: 131–138.
18. Kojima H, Hamazaki Y, Nagata Y, Todokoro K, Nagasawa T, Abe T. Modulation of platelet activation in vitro by thrombopoietin. Thromb Haemost 1995; 74: 1541–1545.
19. •De Sauvage FJ, Hass PE, Spencer SD, Malloy BE, Gurney AL, Spencer SA. et al. Stimulation of megakaryocytopoiesis and thrombopoiesis by the c-Mpl ligand. Nature 1994; 369: 533–538.
20. •Wendling F, Maraskovsky E, Debili N, Florindo C, Teepe M, Titeux M. et al. cMpl ligand is a humoral regulator of megakaryocytopoiesis. Nature 1994; 369: 571–574.
21. •Kaushansky K, Lok S, Holly RD, Broudy VC, Lin N, Bailey MC. et al. Promotion of megakaryocyte progenitor expansion and differentiation by the c-Mpl ligand thrombopoietin. Nature 1994; 369: 568–571.
22. •Lok S, Kaushansky K, Holly RD, Kuijper JL, Lofton-Day CE, Oort PJ. et al. Cloning and expression of murine thrombopoietin cDNA and stimulation of platelet production in vivo. Nature 1994; 369: 565–568.
23. •Bartley TD, Bogenberger J, Hunt P, Li YS, Lu HS, Martin F. et al. Identification and cloning of a megakaryocyte growth and development factor that is a ligand for the cytokine receptor Mpl. Cell 1994; 77: 1117–1124.
24. •Sohma Y, Akahori H, Seki N, Hori T, Ogami K, Kato T. et al. Molecular cloning and chromosomal localization of the human thrombopoietin gene. FEBS Lett 1994; 353: 57–61. Simultaneous reports on the isolation and characterization of the MPL ligand, TPO, as well as the cloning and in vitro expression of its gene.
25. Matsumoto A, Tahara T, Morita H, Usuki K, Ohashi H, Kokubo-Watarai A. et al. Characterization of native human thrombopoietin in the blood of normal individuals and of patients with haematologic disorders. Thromb Haemost 1999; 82: 24–29.
26. Gurney AL, Kuang WJ, Xie MH, Malloy BE, Eaton DL, de Sauvage FJ. Genomic structure, chromosomal localization, and conserved alternative splice forms of thrombopoietin. Blood 1995; 85: 981–988.
27. Ghilardi N, Wiestner A, Skoda RC. Thrombopoietin production is inhibited by a translational mechanism. Blood 1998; 92: 4023–4030.
28. Kato T, Oda A, Inagaki Y, Ohashi H, Matsumoto A, Ozaki K. et al. Thrombin cleaves recombinant human thrombopoietin: one of the proteolytic events that generates truncated forms of thrombopoietin. Proc Natl Acad Sci USA 1997; 94: 4669–4674.
29. Wada T, Nagata Y, Nagahisa H, Okutomi K, Ha SH, Ohnuki T. et al. Characterization of the truncated thrombopoietin variants. Biochem Biophys Res Commun 1995; 213: 1091–1098.
30. Hokom MM, Lacey D, Kinstler OB, Choi E, Kaufman S, Faust J. et al. Pegylated megakaryocyte growth and development factor abrogates the lethal thrombocytopenia associated with carboplatin and irradiation in mice. Blood 1995; 86: 4486–4492.
31. Foster D, Hunt P. The biological significance of truncated and full-length forms of mpl ligand. In: Thrombopoiesis and Thrombopoietins: Molecular, Cellular, Preclinical, and Clinical Biology. Kuter DJ, Hunt P, Sheridan W, Zucker-Franklin D (editors). Totowa, NJ: Humana Press; 1997. pp. 209–214.
32. Li B, Pan H, Winkelmann JC, Dai W. Thrombopoietin and its alternatively spliced form are expressed in human amygdala and hippocampus [letter]. Blood 1996; 87: 5382–5384.
33. •Qian S, Fu F, Li W, Chen Q, de Sauvage FJ. Primary role of the liver in thrombopoietin production shown by tissue-specific knockout. Blood 1998; 92: 2189–2191. Evaluation of the hepatic contribution to platelet homeostasis in liver-specific TPO gene knock-out mice.
34. ••Sungaran R, Markovic B, Chong BH. Localization and regulation of thrombopoietin mRNA expression in human kidney, liver, bone marrow, and spleen using in situ hybridization. Blood 1997; 89: 101–107. Identification of the cells expressing TPO mRNA in the liver and other organs of humans with normal and lowered platelet counts by means of in situ hybridization.
35. Semenza GL, Koury ST, Nejfelt MK, Gearhart JD, Antonarakis SE. Cell-type-specific and hypoxia-inducible expression of the human erythropoietin gene in transgenic mice. Proc Natl Acad Sci USA 1991; 88: 8725–8729.
36. Schuster SJ, Koury ST, Bohrer M, Salceda S, Caro J. Cellular sites of extrarenal and renal erythropoietin production in anaemic rats. Br J Haematol 1992; 81: 153–159.
37. Bachmann S, Le Hir M, Eckardt KU. Co-localization of erythropoietin mRNA and ecto-5′-nucleotidase immunoreactivity in peritubular cells of rat renal cortex indicates that fibroblasts produce erythropoietin. J Histochem Cytochem 1993; 41: 335–341.
38. Maxwell PH, Ferguson DJ, Nicholls LG, Iredale JP, Pugh CW, Johnson MH, Ratcliffe PJ. Sites of erythropoietin production. Kidney Int 1997; 51: 393–401.
39. Wolber E-M, Dame C, Fahnenstich H, Hofmann D, Bartmann P, Jelkmann W, Fandrey J. Expression of the thrombopoietin gene in human fetal and neonatal tissues. Blood 1999; 94: 97–105.
40. Dame C, Fahnenstich H, Freitag P, Hofmann D, Abdul NT, Bartmann P, Fandrey J. Erythropoietin mRNA expression in human fetal and neonatal tissue. Blood 1998; 92: 3218–3225.
41. Nomura S, Ogami K, Kawamura K, Tsukamoto I, Kudo Y, Kanakura Y. et al. Cellular localization of thrombopoietin mRNA in the liver by in situ hybridization. Exp Hematol 1997; 25: 565–572.
42. ••Wolber E-M, Ganschow R, Burdelski M, Jelkmann W. Hepatic thrombopoietin mRNA levels in acute and chronic liver failure of childhood. Hepatology 1999; 29: 1739–1742.
43. ••Martin TG, Somberg KA, Meng YG, Cohen RL, Heid CA, de Sauvage FJ, Shuman MA. Thrombopoietin levels in patients with cirrhosis before and after orthotopic liver transplantation. Ann Intern Med 1997; 127: 285–288.
44. ••Ishikawa T, Ichida T, Matsuda Y, Sugitani S, Sugiyama M, Kato T. et al. Reduced expression of thrombopoietin is involved in thrombocytopenia in human and rat liver cirrhosis. J Gastroenterol Hepatol 1998; 13: 907–913. Measurements of TPO mRNA levels in normal versus cirrhotic human livers indicating that impaired synthesis of the hormone contributes to thrombocytopenia in liver failure.
45. Kochling J, Curtin PT, Madan A. Regulation of human erythropoietin gene induction by upstream flanking sequences in transgenic mice. Br J Haematol 1998; 103: 960–968.
46. Bondurant MC, Koury MJ. Anemia induces accumulation of erythropoietin mRNA in the kidney and liver. Mol Cell Biol 1986; 6: 2731–2733.
47. Tan CC, Eckardt KU, Ratcliffe PJ. Organ distribution of erythropoietin messenger RNA in normal and uremic rats. Kidney Int 1991; 40: 69–76.
48. Fandrey J, Bunn HF. In vivo and in vitro regulation of erythropoietin mRNA: measurement by competitive polymerase chain reaction. Blood 1993; 81: 617–623.
49. Jelkmann W, Bauer C. Demonstration of high levels of erythropoietin in rat kidneys following hypoxic hypoxia. Pflugers Arch 1981; 392: 34–39.
50. Eckardt KU, Ratcliffe PJ, Tan CC, Bauer C, Kurtz A. Age-dependent expression of the erythropoietin gene in rat liver and kidneys. J Clin Invest 1992; 89: 753–760.
51. Zhang F, Laneuville P, Gagnon RF, Morin B, Brox AG. Effect of chronic renal failure on the expression of erythropoietin message in a murine model. Exp Hematol 1996; 24: 1469–1474.
52. Koury ST, Bondurant MC, Koury MJ, Semenza GL. Localization of cells producing erythropoietin in murine liver by in situ hybridization. Blood 1991; 77: 2497–2503.
53. •Maxwell PH, Ferguson DJ, Osmond MK, Pugh CW, Heryet A, Doe BG. et al. Expression of a homologously recombined erythopoietin-SV40 T antigen fusion gene in mouse liver: evidence for erythropoietin production by Ito cells. Blood 1994; 84: 1823–1830. Identification of hepatocytes and Ito cells as EPO-producing tissue in liver.
54. ••Goldberg MA, Glass GA, Cunningham JM, Bunn HF. The regulated expression of erythropoietin by two human hepatoma cell lines. Proc Natl Acad Sci USA 1987; 84: 7972–7976. Establishment of the human hepatoma cell lines Hep3B and HepG2 as suitable models for in vitro study of the molecular mechanisms of oxygen-dependent EPO synthesis.
55. Herkens C, Wolff M, Fandrey J, Schuler F, Jelkmann W. Immunocytochemical demonstration of erythropoietin in hypoxic human hepatoma cultures. Histochemistry 1993; 100: 303–309.
56. ••Wang GL, Jiang BH, Rue EA, Semenza GL. Hypoxia-inducible factor 1 is a basic-helix-loop-helix-PAS heterodimer regulated by cellular O2 tension. Proc Natl Acad Sci USA 1995; 92: 5510–5514. Characterization of hypoxia-inducible factor 1, a transacting dimeric protein, now known to control the rates of the expression of a number of genes encoding proteins involved in oxygen and glucose homeostasis, including EPO, vascular endothelial growth factor and glucose transporter 1.
57. Wenger RH, Gassmann M. HIF-1 and the molecular response to hypoxia in mammals. In: Environmental Stress and Gene Regulation. Storey KB (editor). Oxford: BIOS Scientific Publishers Ltd; 1999. pp. 25–45.
58. Ebert BL, Bunn HF. Regulation of the erythropoietin gene. Blood 1999; 94: 1864–1877.
59. Jungermann K, Kietzmann T. Role of oxygen in the zonation of carbohydrate metabolism and gene expression in liver. Kidney Int 1997; 51: 402–412.
60. Faquin WC, Schneider TJ, Goldberg MA. Effect of inflammatory cytokines on hypoxia-induced erythropoietin production. Blood 1992; 79: 1987–1994.
61. Fandrey J, Pagel H, Frede S, Wolff M, Jelkmann W. Thyroid hormones enhance hypoxia-induced erythropoietin production in vitro. Exp Hematol 1994; 22: 272–277.
62. Jelkmann W, Pagel H, Hellwig T, Fandrey J. Effects of antioxidant vitamins on renal and hepatic erythropoietin production. Kidney Int 1997; 51: 497–501.
63. Jelkmann W, Pagel H, Wolff M, Fandrey J. Monokines inhibiting erythropoietin production in human hepatoma cultures and in isolated perfused rat kidneys. Life Sci 1992; 50: 301–308.
64. Fandrey J, Frede S, Jelkmann W. Role of hydrogen peroxide in hypoxia-induced erythropoietin production. Biochem J 1994; 303: 507–510.
65. •Hino M, Nishizawa Y, Tagawa S, Yamane T, Morii H, Tatsumi N. Constitutive expression of the thrombopoietin gene in a human hepatoma cell line. Biochem Biophys Res Commun 1995; 217: 475–481. Establishment of human hepatoma cells (HepG2) as an in vitro model for study of TPO gene expression.
66. •Wolber E-M, Jelkmann W. Interleukin-6 increases thrombopoietin production in human hepatoma cells HepG2 and Hep3B. J Interferon Cytokine Res 2000; 20: 499–506. First evidence for in vitro modulation of TPO mRNA expression by a cytokine.
67. ••McCarty JM, Sprugel KH, Fox NE, Sabath DE, Kaushansky K. Murine thrombopoietin mRNA levels are modulated by platelet count. Blood 1995; 86: 3668–3675. Evidence for TPO mRNA in bone marrow cells and for its up-regulation in thrombocytopenic mice.
68. •Fielder PJ, Gurney AL, Stefanich E, Marian M, Moore MW, Carver-Moore K, de Sauvage FJ. Regulation of thrombopoietin levels by c-mpl-mediated binding to platelets. Blood 1996; 87: 2154–2161.
69. •Cohen-Solal K, Villeval JL, Titeux M, Lok S, Vainchenker W, Wendling F. Constitutive expression of Mpl ligand transcripts during thrombocytopenia or thrombocytosis. Blood 1996; 88: 2578–2584.
70. •Stoffel R, Wiestner A, Skoda RC. Thrombopoietin in thrombocytopenic mice: evidence against regulation at the mRNA level and for a direct regulatory role of platelets. Blood 1996; 87: 567–573.
71. •Broudy VC, Lin NL, Sabath DF, Papayannopoulou T, Kaushansky K. Human platelets display high-affinity receptors for thrombopoietin. Blood 1997; 89: 1896–1904.
72. •Emmons RV, Reid DM, Cohen RL, Meng G, Young NS, Dunbar CE, Shulman NR. Human thrombopoietin levels are high when thrombocytopenia is due to megakaryocyte deficiency and low when due to increased platelet destruction. Blood 1996; 87: 4068–4071.
73. •Nagata Y, Shozaki Y, Nagahisa H, Nagasawa T, Abe T, Todokoro K. Serum thrombopoietin level is not regulated by transcription but by the total counts of both megakaryocytes and platelets during thrombocytopenia and thrombocytosis. Thromb Haemost 1997; 77: 808–814. Studies indicating that TPO gene expression proceeds in an unregulated constitutive way, with the level of circulating TPO being dependent on the mass of platelets and megakaryocytes, which internalize the hormone.
74. Sato T, Fuse A, Niimi H, Fielder PJ, Avraham H. Binding and regulation of thrombopoietin to human megakaryocytes. Br J Haematol 1998; 100: 704–711.
75. Dinkelaar RB, Engels EY, Hart AA, Schoemaker LP, Bosch E, Chamuleau RA. Metabolic studies on erythropoietin (EP): II. The role of liver and kidney in the metabolism of EP. Exp Hematol 1981; 9: 796–803.
76. Hou M, Carneskog J, Mellqvist UH, Stockelberg D, Hedberg M, Wadenvik H, Kutti J. Impact of endogenous thrombopoietin levels on the differential diagnosis of essential thrombocythaemia and reactive thrombocytosis. Eur J Haematol 1998; 61: 119–122.
77. Dettke M, Hlousek M, Kurz M, Leitner G, Rosskopf K, Stiegler G. et al. Increase in endogenous thrombopoietin in healthy donors after automated plateletpheresis. Transfusion 1998; 38: 449–453.
78. •Heits F, Stahl M, Ludwig D, Stange EF, Jelkmann W. Elevated serum thrombopoietin and interleukin-6 concentrations in thrombocytosis associated with inflammatory bowel disease. J Interferon Cytokine Res 1999; 19: 757–760. Attempt to relate the reactive thrombocytosis associated with inflammatory bowel disease to elevated IL-6 and TPO levels in blood.
79. Heits F, Katschinski DM, Wilmsen U, Wiedemann GJ, Jelkmann W. Serum thrombopoietin and interleukin 6 concentrations in tumour patients and response to chemotherapy-induced thrombocytopenia. Eur J Haematol 1997; 59: 53–58.
80. Cerutti A, Custodi P, Duranti M, Noris P, Balduini CL. Thrombopoietin levels in patients with primary and reactive thrombocytosis. Br J Haematol 1997; 99: 281–284.
81. Wang JC, Chen C, Lou LH, Mora M. Blood thrombopoietin, IL-6 and IL-11 levels in patients with agnogenic myeloid metaplasia. Leukemia 1997; 11: 1827–1832.
82. Uppenkamp M, Makarova E, Petrasch S, Brittinger G. Thrombopoietin serum concentration in patients with reactive and myeloproliferative thrombocytosis. Ann Hematol 1998; 77: 217–223.
83. Tacke F, Schoffski P, Trautwein C, Martin MU, Stangel W, Seifried E. et al. Endogenous serum levels of thrombopoietic cytokines in healthy whole-blood and platelet donors: implications for plateletpheresis. Br J Haematol 1999; 105: 511–513.
84. Marsh JC, Gibson FM, Prue RL, Bowen A, Dunn VT, Hornkohl AC. et al. Serum thrombopoietin levels in patients with aplastic anaemia. Br J Haematol 1996; 95: 605–610.
85. Pitcher L, Taylor K, Nichol J, Selsi D, Rodwell R, Marty J. et al. Thrombopoietin measurement in thrombocytosis: dysregulation and lack of feedback inhibition in essential thrombocythaemia. Br J Haematol 1997; 99: 929–932.
86. Tahara T, Usuki K, Sato H, Ohashi H, Morita H, Tsumura H. et al. A sensitive sandwich ELISA for measuring thrombopoietin in human serum: serum thrombopoietin levels in healthy volunteers and in patients with haemopoietic disorders. Br J Haematol 1996; 93: 783–788.
87. Ishiguro A, Ishikita T, Shimbo T, Matsubara K, Baba K, Hayashi Y. et al. Elevation of serum thrombopoietin precedes thrombocytosis in Kawasaki disease. Thromb Haemost 1998; 79: 1096–1100.
88. Folman CC, von dem Borne AE, Rensink IH, Gerritsen W, van der Schoot CE, De Haas M, Aarden L. Sensitive measurement of thrombopoietin by a monoclonal antibody based sandwich enzyme-linked immunosorbent assay. Thromb Haemost 1997; 78: 1262–1267.
89. Meng YG, Martin TG, Peterson ML, Shuman MA, Cohen RL, Lee Wong W. Circulating thrombopoietin concentrations in thrombocytopenic patients, including cancer patients following chemotherapy, with or without peripheral blood progenitor cell transplantation. Br J Haematol 1996; 95: 535–541.
90. Cerutti A, Custodi P, Duranti M, Cazzola M, Balduini CL. Circulating thrombopoietin in reactive conditions behaves like an acute phase reactant. Clin Lab Haematol 1999; 21: 271–275.
91. Peck-Radosavljevic M, Zacherl J, Meng YG, Pidlich J, Lipinski E, Langle F. et al. Is inadequate thrombopoietin production a major cause of thrombocytopenia in cirrhosis of the liver? J Hepatol 1997; 27: 127–131.
92. Shimodaira S, Ishida F, Ichikawa N, Tahara T, Kato T, Kodaira H. et al. Serum thrombopoietin (c-Mpl ligand) levels in patients with liver cirrhosis. Thromb Haemost 1996; 76: 545–548.
93. •Stiegler G, Stohlawetz P, Peck RM, Jilma B, Pidlich J, Wichlas M. et al. Direct evidence for an increase in thrombopoiesis after liver transplantation. Eur J Clin Invest 1998; 28: 755–759. Detailed study of the time course of the recovery of platelets following orthotopic liver transplantation in humans.
94. Koike Y, Yoneyama A, Shirai J, Ishida T, Shoda E, Miyazaki K. et al. Evaluation of thrombopoiesis in thrombocytopenic disorders by simultaneous measurement of reticulated platelets of whole blood and serum thrombopoietin concentrations. Thromb Haemost 1998; 79: 1106–1110.
95. Stockelberg D, Andersson P, Bjornsson E, Bjork S, Wadenvik H. Plasma thrombopoietin levels in liver cirrhosis and kidney failure. J Intern Med 1999; 246: 471–475.
96. Goulis J, Chau TN, Jordan S, Mehta AB, Watkinson A, Rolles K, Burroughs AK. Thrombopoietin concentrations are low in patients with cirrhosis and thrombocytopenia and are restored after orthotopic liver transplantation. Gut 1999; 44: 754–758.
97. Peck-Radosavljevic M, Wichlas M, Zacherl J, Stiegler G, Stohlawetz P, Fuchsjager M. et al. Thrombopoietin induces rapid resolution of thrombocytopenia after orthotopic liver transplantation through increased platelet production. Blood 2000; 95: 795–801.
98. Chang FY, Singh N, Gayowski T, Wagener MM, Mietzner SM, Stout JE, Marino IR. Thrombocytopenia in liver transplant recipients: predictors, impact on fungal infections, and role of endogenous thrombopoietin. Transplantation 2000; 69: 70–75.
99. Peck-Radosavljevic M, Wichlas M, Pidlich J, Sims P, Meng G, Zacherl J. et al. Blunted thrombopoietin response to interferon alfa-induced thrombocytopenia during treatment for hepatitis C. Hepatology 1998; 28: 1424–1429.
100. Shiota G, Okubo M, Kawasaki H, Tahara T. Interferon increases serum thrombopoietin in patients with chronic hepatitis C. Br J Haematol 1997; 97: 340–342.
101. Ciernik IF, Cone RW, Fehr J, Weber R. Impaired liver function and retroviral activity are risk factors contributing to HIV-associated thrombocytopenia. Swiss HIV Cohort Study. AIDS 1999; 13: 1913–1920.
102. Hamaguchi M, Yamada H, Morishima Y, Morishita Y, Kato Y, Sao H. et al. Serum thrombopoietin level after allogeneic bone marrow transplantation: possible correlations with platelet recovery, acute graft-versus-host disease and hepatic veno-occlusive disease. Nagoya Bone Marrow Transplantation Group. Int J Hematol 1996; 64: 241–248.
103. Oh H, Tahara T, Bouvier M, Farrand A, McDonald GB. Plasma thrombopoietin levels in marrow transplant patients with veno- occlusive disease of the liver. Bone Marrow Transplant 1998; 22: 675–679.
104. Radovic M, Jelkmann W, Djukanovic L, Ostric V. Serum erythropoietin and interleukin-6 levels in hemodialysis patients with hepatitis virus infection. J Interferon Cytokine Res 1999; 19: 369–373.
105. Ichikawa N, Kitano K, Shimodaira S, Ishida F, Ito T, Kajikawa S. et al. Changes in serum thrombopoietin levels after splenectomy. Acta Haematol 1998; 100: 137–141.
106. •Komura E, Matsumura T, Kato T, Tahara T, Tsunoda Y, Sawada T. Thrombopoietin in patients with hepatoblastoma. Stem Cells (Dayt) 1998; 16: 329–333. Demonstration of TPO mRNA and TPO receptor mRNA in tumour samples from children with hepatoblastoma.
107. •Shimada Y, Kato T, Ogami K, Horie K, Kokubo A, Kudo Y. et al. Production of thrombopoietin (TPO) by rat hepatocytes and hepatoma cell lines. Exp Hematol 1995; 23: 1388–1396. Demonstration of TPO production by primary hepatocytes and hepatoma cells in culture.
108. McFadzean AJ, Todd D, Tso SC. Erythrocytosis associated with hepatocellular carcinoma. Blood 1967; 29: 808–811.
109. Jacobson RJ, Lowenthal MN, Kew MC. Erythrocytosis in hepatocellular cancer. S Afr Med J 1978; 53: 658–660.
110. •Kew MC, Fisher JW. Serum erythropoietin concentrations in patients with hepatocellular carcinoma. Cancer 1986; 58: 2485–2488. Evidence that the erythrocytosis in several patients with hepatocellular carcinoma is associated with increased EPO concentrations in blood.
111. Means RT, Krantz S. Progress in understanding the pathogenesis of the anemia of chronic disease. Blood 1992; 80: 1639–1647.
112. Jelkmann W. Proinflammatory cytokines lowering erythropoietin production. J Interferon Cytokine Res 1998; 18: 555–559.
113. Sakisaka S, Watanabe M, Tateishi H, Harada M, Shakado S, Mimura Y. et al. Erythropoietin production in hepatocellular carcinoma cells associated with polycythemia: immunohistochemical evidence. Hepatology 1993; 18: 1357–1362.
114. Reizenstein P. The haematological stress syndrome. Br J Haematol 1979; 43: 329–334.
115. Estrov Z, Talpaz M, Mavligit G, Pazdur R, Harris D, Greenberg SM, Kurzrock R. Elevated plasma thrombopoietic activity in patients with metastatic cancer-related thrombocytosis. Am J Med 1995; 98: 551–558.
116. Espanol I, Hernandez A, Cortes M, Mateo J, Pujol-Moix N. Patients with thrombocytosis have normal or slightly elevated thrombopoietin levels. Haematologica 1999; 84: 312–316.
117. Bjerre A, Ovstebo R, Kierulf P, Halvorsen S, Brandtzaeg P. Fulminant meningococcal septicemia: dissociation between plasma thrombopoietin levels and platelet counts. Clin Infect Dis 2000; 30: 643–647.
118. ••Kamura T, Handa H, Hamasaki N, Kitajima S. Characterization of the human thrombopoietin gene promoter. A possible role of an Ets transcription factor, E4TF1/GABP. J Biol Chem 1997; 272: 11 361–11 368. Cloning of the 5′-flanking region of the human TPO gene and identification of relevant cis-elements in the promoter.
119. Yamashita K, Matsuoka H, Ochiai T, Matsushita R, Kubuki Y, Suzuki M, Tsubouchi H. Hepatocyte growth factor/scatter factor enhances the thrombopoietin mRNA expression in rat hepatocytes and cirrhotic rat livers. J Gastroenterol Hepatol 2000; 15: 83–90.
120. Sasaki Y, Takahashi T, Miyazaki H, Matsumoto A, Kato T, Nakamura K. et al. Production of thrombopoietin by human carcinomas and its novel isoforms. Blood 1999; 94: 1952–1960.
121. Slichter SJ. Optimizing platelet transfusions in chronically thrombocytopenic patients. Semin Hematol 1998; 35: 269–278.
122. Webb IJ, Anderson KC. Risks, costs, and alternatives to platelet transfusions. Leuk Lymphoma 1999; 34: 71–84.
123. Adamson JW, Eschbach JW. Treatment of the anemia of chronic renal failure with recombinant human erythropoietin. Annu Rev Med 1990; 41: 349–360.
124. Horl WH, Cavill I, MacDougall IC, Schaefer RM, Sunder-Plassmann G. How to diagnose and correct iron deficiency during r-huEPO therapy – a consensus report. Nephrol Dial Transplant 1996; 11: 246–250.
125. Schobersberger W, Hobisch-Hagen P, Fuchs D, Hoffmann G, Jelkmann W. Pathogenesis of anaemia in the critically ill patient. Clin Intensive Care 1998; 9: 111–117.
126. Beguin Y, Loo M, R'Zik S, Sautois B, Lejeune F, Rorive G, Fillet G. Early prediction of response to recombinant human erythropoietin in patients with the anemia of renal failure by serum transferrin receptor and fibrinogen. Blood 1993; 82: 2010–2016.
127. McCarthy JT, Johnson WJ, Nixon DE, Jenson BM, Moyer TP. Transfusional iron overload in patients undergoing dialysis: treatment with erythropoietin and phlebotomy. J Lab Clin Med 1989; 114: 193–199.
128. Onoyama K, Nakamura S, Yamamoto M, Kawadoko T, Nanishi F, Komoda T. et al. Correction of serious iron overload in a chronic hemodialysis patient by recombinant human erythropoietin and removal of red blood cells: confirmation by follow-up liver biopsy. Nephron 1990; 56: 325–328.
129. Sennesael JJ, Van-der-Niepen P, Verbeelen DL. Treatment with recombinant human erythropoietin increases antibody titers after hepatitis B vaccination in dialysis patients. Kidney Int 1991; 40: 121–128.
130. Kuter DJ. In vivo effects of Mpl ligand administration and emerging clinical applications for the Mpl ligands. Curr Opin Hematol 1997; 4: 163–170.
131. Vadhan-Raj S. Recombinant human thrombopoietin: clinical experience and in vivo biology. Semin Hematol 1998; 35: 261–268.
132. Vadhan-Raj S, Murray LJ, Bueso-Ramos C, Patel S, Reddy SP, Hoots WK. et al. Stimulation of megakaryocyte and platelet production by a single dose of recombinant human thrombopoietin in patients with cancer. Ann Intern Med 1997; 126: 673–681.
133. Murray LJ, Luens KM, Estrada MF, Bruno E, Hoffman R, Cohen RL. et al. Thrombopoietin mobilizes CD34+ cell subsets into peripheral blood and expands multilineage progenitors in bone marrow of cancer patients with normal hematopoiesis. Exp Hematol 1998; 26: 207–216.
134. Basser RL, Rasko JE, Clarke K, Cebon J, Green MD, Hussein S. et al. Thrombopoietic effects of pegylated recombinant human megakaryocyte growth and development factor (PEG-rHuMGDF) in patients with advanced cancer. Lancet 1996; 348: 1279–1281.
135. Rasko JE, Basser RL, Boyd J, Mansfield R, O'Malley CJ, Hussein S. et al. Multilineage mobilization of peripheral blood progenitor cells in humans following administration of PEG-rHuMGDF. Br J Haematol 1997; 97: 871–880.
136. Fanucchi M, Glaspy J, Crawford J, Garst J, Figlin R, Sheridan W. et al. Effects of polyethylene glycol-conjugated recombinant human megakaryocyte growth and development factor on platelet counts after chemotherapy for lung cancer. N Engl J Med 1997; 336: 404–409.
137. Basser RL, Rasko JE, Clarke K, Cebon J, Green MD, Grigg AP, et al. Randomized, blinded, placebo-controlled phase I trial of pegylated recombinant human megakaryocyte growth and development factor with filgrastim after dose-intensive chemotherapy in patients with advanced cancer. Blood 1997; 89:3118–3128. [Published erratum appears in Blood 1997; 90:2513.]
138. •Somlo G, Sniecinski I, ter-Veer A, Longmate J, Knutson G, Vuk PS. et al. Recombinant human thrombopoietin in combination with granulocyte colony-stimulating factor enhances mobilization of peripheral blood progenitor cells, increases peripheral blood platelet concentration, and accelerates hematopoietic recovery following high-dose chemotherapy. Blood 1999; 93: 2798–2806. Clinical trial indicating that the administration of recombinant human TPO combined with GC-SF may increase the yield of peripheral blood progenitor cells for retransfusion in patients treated with high-dose chemotherapy.
139. Glaspy J, Bukowski R, Steinberg D, Taylor C, Tchekmedyian S, Vadhan RS. Impact of therapy with epoetin alfa on clinical outcomes in patients with nonmyeloid malignancies during cancer chemotherapy in community oncology practice. Procrit Study Group. J Clin Oncol 1997; 15: 1218–1234.
140. Barosi G, Marchetti M, Liberato NL. Cost-effectiveness of recombinant human erythropoietin in the prevention of chemotherapy-induced anaemia. Br J Cancer 1998; 78: 781–787.
141. American Society of Clinical Oncology. Recommendations for the use of hematopoietic colony-stimulating factors: evidence-based, clinical practice guidelines. J Clin Oncol 1994; 12: 2471–2508.
142. American Society of Clinical Oncology. Update of recommendations for the use of hematopoietic colony-stimulating factors: evidence-based clinical practice guidelines. J Clin Oncol 1996; 14: 1957–1960.
143. Hofmann WK, Ottmann OG, Hoelzer D. Megakaryocytic growth factors: is there a new approach for management of thrombocytopenia in patients with malignancies? Leukemia 1999; 13: 14–18.
144. Levin J. Thrombopoietin – clinically realized? N Engl J Med 1997; 336: 434–436.
145. Maslak P, Nimer SD. The efficacy of IL-3, SCF, IL-6, and IL-11 in treating thrombocytopenia. Semin Hematol 1998; 35: 253–260.

anaemia; erythropoietin; inflammation; liver disease; thrombocytopenia; thrombocytosis; thrombopoietin

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