Suppression of erythroid development in vitro by Plasmodium vivax
© Panichakul et al.; licensee BioMed Central Ltd. 2012
Received: 29 February 2012
Accepted: 24 May 2012
Published: 24 May 2012
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© Panichakul et al.; licensee BioMed Central Ltd. 2012
Received: 29 February 2012
Accepted: 24 May 2012
Published: 24 May 2012
Severe anaemia due to dyserythropoiesis has been documented in patients infected with Plasmodium vivax, however the mechanism responsible for anaemia in vivax malaria is poorly understood. In order to better understand the role of P. vivax infection in anaemia the inhibition of erythropoiesis using haematopoietic stem cells was investigated.
Haematopoietic stem cells/CD34+ cells, isolated from normal human cord blood were used to generate growing erythroid cells. Exposure of CD34+ cells and growing erythroid cells to P. vivax parasites either from intact or lysed infected erythrocytes (IE) was examined for the effect on inhibition of cell development compared with untreated controls.
Both lysed and intact infected erythrocytes significantly inhibited erythroid growth. The reduction of erythroid growth did not differ significantly between exposure to intact and lysed IE and the mean growth relative to unexposed controls was 59.4 ± 5.2 for lysed IE and 57 ± 8.5% for intact IE. Interestingly, CD34+ cells/erythroid progenitor cells were susceptible to the inhibitory effect of P. vivax on cell expansion. Exposure to P. vivax also inhibited erythroid development, as determined by the reduced expression of glycophorin A (28.1%) and CD 71 (43.9%). Moreover, vivax parasites perturbed the division of erythroid cells, as measured by the Cytokinesis Block Proliferation Index, which was reduced to 1.35 ± 0.05 (P-value < 0.01) from a value of 2.08 ± 0.07 in controls. Neither TNF-a nor IFN-g was detected in the culture medium of erythroid cells treated with P. vivax, indicating that impaired erythropoiesis was independent of these cytokines.
This study shows for the first time that P. vivax parasites inhibit erythroid development leading to ineffective erythropoiesis and highlights the potential of P. vivax to cause severe anaemia.
Anaemia has frequently been associated with severe malaria and is believed to contribute to the morbidity and mortality of this disease. Most published reports on malaria-associated anaemia focus on Plasmodium falciparum with Plasmodium vivax being less well studied . However, growing evidence from several geographic regions has demonstrated that P. vivax malaria is associated with a higher frequency and more severe anaemia [2–12]. Several cases of patients infected with P. vivax that resulted in severe disease and death were found to have syndromes resembling those commonly observed with falciparum malaria [8, 13, 14]. Although the underlying causes of severe malarial anaemia are multifactorial, major causes are the destruction of parasitized erythrocytes and ineffective erythropoiesis or dyserythropoiesis. In vivax malaria patients with anaemia red cells in blood film are usually normochromic and normocytic with the absolute reticulocyte count not elevated . However, changes in haemoglobin concentrations are associated with continuing parasitaemia during infection of P. vivax in patients with anaemia . In addition, P. vivax has been observed in the bone marrow of patients who exhibited dyserythropoiesis  and one case of P. vivax malaria in Southeast Asia displayed pancytopaenia in blood and bone marrow . Investigation by light and electron microscopy of marrow aspirates from four Thai patients with P. vivax malaria and anaemic symptoms revealed morphological evidence of dyserythropoiesis and the presence of erythroblasts at various stages of degradation within the cytoplasm of macrophages . It seems likely that vivax malaria infection was associated with an activation of the pro-inflammatory response and cytokine imbalance  and experimental findings in mice are consistent with a role for TNF-a in the dyserythropoietic changes in malaria . Other mechanisms have also been suggested, including alterations in IFN-g, IL-12, IL-6, IL-1, reactive oxygen species, nitric oxide, macrophage dysfunction, or a direct effect of parasites (or parasite products) on the bone marrow [15, 20–22]. However, the mechanism responsible for anaemia in vivax malaria remains poorly understood. Here, haematopoietic stem cells (HSCs)/CD34+ from normal human cord blood were used to generate growing erythroid cells (gEC) to investigate the effect of P. vivax infection on erythropoiesis. Enhancing the understanding of the pathogenesis of anaemia caused by malaria is a prerequisite for developing effective prevention and treatment strategies.
Plasmodium vivax was obtained from patients attending the malaria clinic in Mae Sot, Tak Province, Thailand. Patient blood with 0.1-0.3% parasitaemia, as determined by examining thick and thin blood smears, was collected. The ethical and methodological aspects of this study for parasite collection (MU-IRB 2010/344.1612) have been approved by the Mahidol University Institutional Review Board, Mahidol University, Bangkok, Thailand. Infected erythrocytes (IE) were separated from patient blood using a 60% Percoll solution as previously described . Briefly, whole blood with vivax parasites was collected and then filtrated using a Plasmodipur filter (Euro-Diagnostic B.V., Netherlands) to remove white blood cells. To obtain asexual parasites, packed, infected RBCs from 20 ml of patient blood was diluted 1:2 with RPMI1640 (Invitrogen®, CA, USA), layered on 60% Percoll and centrifuged at 1,200 g for 20 mins at 20°C. The purity of IE after isolation was 95% and the pure fraction of isolated IE contained 80% schizontes and 20% of other stages. The isolated IEs were used either intact or as lysed cells prepared by freezing and thawing.
Umbilical cord blood from normal full-term deliveries in Ramathibodi Hospital, Bangkok, Thailand was collected into cord blood bags containing anticoagulant solution (CPDA-1 solution) (Kawasumi Laboratories, Thailand). Cord blood collection (ID 04-45-16) was approved by the Ethical Committee of Research on Human Beings of the Ramathibodi Hospital, Faculty of Medicine, Mahidol University. Haematopoietic stem cells/CD34+ cells were isolated from cord blood mononuclear cells (MNC) using a CD 34 isolation kit with magnetic microbead selection with Mini-MACS columns (Miltenyi Biotech, Geramany) as described by Panichakul et al. The purity of CD34+ cells after isolation was 97% as judged by flow cytometry analysis.
HSCs/CD34+ cells at a density of 1 × 105 cell/well in 24-well tissue culture plates (Corning Incorporated Costar®, NY, USA) were cultured in 0.5 ml of complete medium containing StemlineII medium (Sigma-Aldrich Corporation, Missouri, USA) supplemented with cytokines . IE at the indicated concentration were added to cell cultures on days 1, 5, 8 and 11 and cultured for three additional days. Intact and lysed IE from the same patients were utilized in this study. Recombinant human tumour necrosis factor-alpha (TNF-a), human interferon gamma (IFN-g) (Prospec-Tany TechnoGene Ltd., Rehovot, Israel), and uninfected erythrocytes (UE) from normal donor blood were included in this study. All cultures were incubated at 37°C in 5% CO2 and viable cells were determined by trypan blue dye exclusion.
Cell surface markers were detected using immunofluorescence with mouse antibodies to human CD34 (Miltenyi Biotech), CD71, CD45 (eBioscience, Inc. CA, USA) and glycophorin A (Serotec Inc., NC, USA) to confirm cell types of HSC and derived erythroid cells. Cells (5–10 × 104 cells in 100 μl of medium) were stained with 5 μl of each antibody for 30 min at 4°C. After washing twice with phosphate-buffer saline, stained cells were fixed with 1% paraformaldehyde. Cell death was also determined by staining with 20 μg/ml of propidium iodide (BenderMedSystems®, GmSH). All cell markers were analysed by flow cytometry using an Epics XL-MCL analyser (Beckman Coulter, Inc. CA, USA).
The presence of inflammatory cytokines TNF-a and IFN-g in culture medium of gEC was determined using the Bio-Plex Pro™ Magnetic Cytokine assay (Bio-Rad Laboratories, Inc. CA, USA). Briefly, 50 μl of beads coated with different anti-cytokine antibodies were added to a pre-wet filter plate and after washing 50 μl of culture medium or cytokine standards were added to duplicate wells and incubated for 30 min at room temperature in the dark. The plate was washed three times with washing buffer and 50 μl of streptavidin-PE was added and the reaction was incubated for 10 min. Following three washes, 125 μl of assay buffer was added and the plate was analysed immediately using an array reader (Bio-Plex Manager 5.0 Software, Bio-Rad Laboratories).
Five-day old cells were exposed to lysed IE for three days followed by incubation with 3 μg/ml cytochalasin B (Sigma-Aldrich Corporation) for 24 h as previously described . Cells (5–10 x 104 in 100 μl of medium) were spun onto a slide using cytospin (Cytospin 3, Thermo Shandon, UK) at 800 rpm for 10 min then fixed with 95% ethanol for 10 min. Thereafter, cells were stained for 10 min with Giemsa, then washed and dried. Stained cells were examined under a light microscope (Olympus BX50, Japan) and the proportion of mono-, bi-, tri- and tetranucleated cells was evaluated in samples of 2,000 cells. The Cytokinesis Block Proliferation Index (CBPI) was calculated using the formula CBPI = X(1 N) + Y(2 N) + Z(3 N) / X + Y + Z; where X, Y and Z are the number of cells with one, two and three nuclei (N), respectively .
Data were analysed using the SPSS program (version 11.0). The unpaired Mann–Whitney - Wilcoxon test was used to compare means between independent groups as appropriate and for statistical evaluation of CBPI. Results are reported as statistically significant if the P-value was less than 0.01.
Inhibition of erythroid expansion by lysate of infected erythrocytes to CD34 + cells /erythroid progenitor cells
Day-old of cell cultures
% of control (mean ± S.D.)
67 ± 9.5*
43 ± 5.5*
100 ± 4.5
98 ± 5.5
54 ± 12*
72 ± 6.5*
58 ± 4.5*
99 ± 4.0
98 ± 4.8
69 ± 8*
88 ± 4.5
65 ± 7.5*
99 ± 2.0
99 ± 5.5
74 ± 7*
98 ± 8.5
87 ± 7.0*
100 ± 4.5
100 ± 5.0
78 ± 6*
Numerous reports have shown that P. vivax can be associated with severe anaemia [2–12]. Haematologic profiles of pancytopaenia in blood and bone marrow  and dyserythropoiesis in bone marrow  have been reported in vivax malaria patients. However, the cause of reduction in blood cell production in the bone marrow of patients with vivax malaria is not completely understood. Anaemia in malaria is caused by excessive removal of non-parasitized erythrocytes, the immune destruction of parasite-infected red cells, as well as by the impaired compensation due to bone marrow dysfunction [1, 15, 17, 26–28]. Plasmodium vivax requires reticulocytes for expansion of the blood stages  and parasitaemia is generally low, therefore it is unlikely to be the primary cause of anaemia . This suggests that in addition to the simple destruction of infected red cells another mechanism is involved in anaemia in vivax malaria.
Recently, the production of erythrocytes from the in vitro cultures of haematopoietic stem cells was achieved, as previously described  and this model is now being applied to dissect the complexity of anaemia in malaria. The results presented in this study have revealed for the first time that P. vivax can directly inhibited erythropoiesis, as shown by the reduction of erythroid growth in the presence of either lysed or intact IE. Erythroid progenitor cells were susceptible to the inhibitory effect of P. vivax on cell expansion and this result is consistent with the previous report that young stages of erythroid cells were more susceptible to P. vivax infection . The suppression of erythropoiesis in malarial anaemia is not unique to P. vivax and has also been observed in infections from other Plasmodium species. In the complicated Plasmodium falciparum infection, erythroid suppression is indicated by a decrease in the number of erythroid precursors as well as colony-forming units-erythroid (CFU-E) and burst-forming units-erythroid (BFU-E) in the bone marrow cultures . Plasmodium chabaudi can directly suppress the proliferation, differentiation and maturation of erythroid progenitor cells and causes inadequate reticulocytosis in mice . However, deficient erythropoietin production does not appear to be the cause of inadequate erythropoiesis in malaria .
Decreased responsiveness of erythroid progenitor cells to erythropoietin as well as impaired erythropoietin production mediated by inflammatory cytokines has been reported to be involved in anaemia during inflammation . Consistent with this observation, TNF-a was reported to partly inhibit proliferation of erythroid progenitor cells in bone marrow cultures . Erythroid progenitor cells produced in this model were also susceptible to inhibition by exogenous TNF-a as shown in Figure 4a. However, endogenous TNF-a and IFN-g in erythroid cultures exposed to lysates or intact P. vivax was undetectable (Figure 4b). This suggests that P. vivax can also inhibit erythropoiesis independently of TNF-a and IFN-g. Inhibition of erythroid development that is independent of TNF-a and IFN-g has also been observed by exposure with P. falciparum haemozoin [35, 36]. However, other inflammatory cytokines may be involved and high levels of IL-10 were found to correlate positively with inhibition of proliferative peripheral blood mononuclear cells in the presence of P. falciparum haemozoin . In this study, IL-10 was also detectable in supernatants from gECs in the presence of IEs and the role of this cytokine in the inhibition of erythropoiesis is currently being investigated. Interestingly, P. vivax inhibited not only growth but also the differentiation of erythroid progenitor cells as shown by the reduction of glycophorin+ and CD 71+ cells and this is similar to the inhibitory effect of P. falciparum haemozoin on erythroid cell development [35, 38]. Moreover, vivax parasites were able to perturb the cell division but did not induce the cell death of erythroid progenitor cells. Defects in the cell cycle without apoptosis has also been observed with the inhibitory effect of P. falciparum haemozoin on erythroid cell growth . It was found that falciparum haemozoin-treated erythroid cells enhanced the expression of the transcription factor p53 and cdk-inhibitor p21 in addition the retinoblastoma protein, a central regulator of G- to S-phase transition was hypophosphorylated, while GATA-1, the master transcription factor in erythropoiesis was reduced . Therefore the molecular mechanisms underlying the suppression of erythropoiesis by P. vivax or its products warrants further investigation. The findings of this study are consistent with the hypothesis that vivax parasites can suppress erythropoiesis. These results provide a better understanding of the role of chronic and persistent P. vivax infection as a cause of anaemia. Prolonged exposure to vivax parasites can suppress erythropoiesis as well as inhibit reticulocyte production, which could prevent the restoration of the erythrocyte population in chronic parasitaemic P. vivax infection. Many cases of patients with severe anaemia have been reported in vivax endemic areas in Thailand, Indonesian Papua, Korea, Pakistan, Venezuela, and Colombia [3, 5, 7, 9, 10, 14]. These patients are often infected or re-infected with the vivax parasites and parasites have the potential to inhibit erythroid development leading to ineffective erythropoiesis causing severe anaemia.
This finding suggests that suppression of erythropoiesis by P. vivax infection is potentially much more dangerous than it is commonly believed and defective erythropoiesis should be taken into consideration in the development of therapeutic strategies to treat severe malarial anaemia.
Haematopoietic stem cell
Growing erythroid cell
Tumour necrosis factor-alpha
Cytokinesis block proliferation index
This study was supported by Thailand Research Fund, Commission on Higher Education, Ministry of Education and Suan Dusit Rajabhat University, (MRG5380092), the Office of the Higher Education Commission, and Mahidol University under the National Research Universities Initiative. I would like to thank the Mahidol Vivax Research Center, Faculty of Tropical Medicine for parasite collection, Ramathibodi Hospital, Faculty of Medicine for providing cord blood specimens, the Department of Pathobiology, Faculty of Science for excellent technical assistance, and Dr Laran Jensen, Department of Biochemistry, Faculty of Science, Mahidol University, Bangkok Thailand for critical reading of this manuscript.