Myeloperoxidase (MPO), a pivotal lineage marker for acute myeloid leukemia (AML), has been also shown to have a prognostic value: a high percentage of MPO-positive blasts correlates to favorable prognosis. To understand the relationship between the expression of MPO in leukemia cells and the response to chemotherapeutic agents, we established MPO-expressing K562 leukemia cell lines and then treated them with cytosine arabinocide (AraC). Cells expressing wild-type MPO, but not mutant MPO that could not mature, died earlier of apoptosis than control K562 cells. Reactive oxygen species (ROS) were generated more in leukemia cells expressing MPO, and the generation was abrogated by MPO inhibitors or antioxidants. Tyrosine nitration of cellular protein also increased more in MPO-expressing K562 cells than control cells after treatment with AraC. In clinical samples, CD34-positive AML cells from high-MPO cases showed a tendency to be sensitive to AraC in the colony-formation assay, and the generation of ROS and the nitration of protein were observed only when the percentage of MPO-expressing cells was high. These data suggest that MPO enhances the chemosensitivity of AML through the generation of ROS and the nitration of proteins.
It is widely accepted that the expression of myeloperoxidase (MPO), a microbiocidal protein, is a golden marker for the diagnosis of acute myeloid leukemia (AML) utilized by the French–American–British and WHO classifications1, 2 to determine the hematopoietic lineage of immature blasts as myeloid. Apart from its role in the diagnosis of AML, MPO has also been shown to have a prognostic value by several groups including ours.3, 4, 5 These reports demonstrated that the percentage of MPO (or Sudan black B)-positive blasts assessed by cytochemical methods was related to the prognosis of AML patients; those with a higher percentage of MPO-positive blasts had better survival rates. Our previous report5 showed significant differences in complete remission rate, disease-free survival and overall survival using multivariate analysis. However, so far there is no clear explanation as to how the expression of MPO relates to the prognosis of AML.
Chemotherapeutic agents create various reactions in leukemia cells when administered. One of the effects triggered by chemotherapy is the production of reactive oxygen species (ROS).6, 7 ROS are known to modulate the regulators of a wide variety of cellular biological processes including calcium signaling, protein phosphorylation, gene expression, cell growth and differentiation, and chemotaxis.8, 9 They also induce cellular damage associated with lipid peroxidation and alteration of proteins and nucleic acids.10 Mainly on the basis of in vitro studies, it is believed that ROS produced by chemotherapeutic agents play a role in the induction of apoptosis in target cells, which could directly relate to the efficacy of chemotherapy.7 MPO catalyzes the production of hypochlorous acid using hydrogen peroxide (H2O2) as a substrate.11 Since hypochlorous acids are highly toxic for cells, it is presumed that higher amounts of hypochlorous acids produced by MPO would result in higher toxicity for cells. For example, in the HL60 leukemia cell line, the amount of MPO in cells was directly related to cytotoxicity elicited by chemotherapeutic agents.12 MPO in HL60 cells was also demonstrated to be involved in the induction of apoptosis by H2O2.13
The clinical and experimental importance of MPO in the cytotoxicity of chemotherapeutic agents prompted us to directly evaluate the influence of MPO on the efficacy of cytosine arabinocide (AraC), an important antileukemia drug for AML, on leukemia cells. We generated MPO-expressing K562 leukemia cell lines that were originally negative for MPO expression to test for changes in sensitivity to AraC. In this report, we show that the activity of MPO directly enhanced the cytotoxicity of AraC by producing increased amounts of ROS and nitrated tyrosine residues in cellular proteins. In accordance with the observation on leukemia cell line, in samples from AML patients, AraC inhibited colony formation of AML cells more efficiently when MPO expression was high. The production of ROS and nitrated tyrosine was also partly related to the percentage of MPO-positive blasts in clinical samples. These observations suggest important roles for MPO in the cytotoxicity of chemotherapeutic agents during the treatment of AML.
Materials and methods
Vectors, cDNA constructs and mutagenesis
Full-length cDNA for human MPO (kindly provided by Dr Nagata, Institute of Medical Science, University of Tokyo)14 was cloned into pCI-neo, a mammalian expression vector (Promega, Madison, WI, USA). The R569W mutation of the MPO gene (arginine at the 569th amino-acid position was changed into tryptophan) was generated by PCR-based methods that replaced the C nucleotide at the 1868 bp position by T. Mutagenesis was confirmed by using the BigDye Terminator v3.1 cycle sequencing kit (Applied Biosystems, Foster City, CA, USA) and ABI PRISM3100 Genetic analyzer (Applied Biosystems). All PCR experiments were performed using the GeneAmp PCR System9700 and GeneAmp High Fidelity Enzyme Mix (Applied Biosystems).
Cell culture and electroporation
The human leukemia cell line, K562, obtained from the American Type Culture Collection (ATCC, Manassas, VA, USA; CCL-243) was maintained in Iscove's modified Dulbecco's medium (Invitrogen, Carlsbad, CA, USA) supplemented with 10% fetal bovine serum (SAFC biosciences, Lenexa, KA, USA) and antibiotics at 37 °C under 5% CO2. In some experiments, cytosine arabinoside (Sigma, St Louis, MO, USA), H2O2 (Wako Pure Chemical Industries, Osaka, Japan), N-acetylcysteine (Sigma) or 4-aminobenzoic hydrazide (Sigma) were added alone or in various combinations into the culture medium. Peroxynitrite tetramethylammonium (Alexis Biochemicals, San Diego, CA, USA) was used as a source of reactive nitrogen species (RNS). Cell growth was assessed using the Premix WST-1 Cell Proliferation Assay System (Takara Biochem., Tokyo, Japan). pCI-neo carrying normal or mutated cDNA for MPO was transfected into log-phase K562 cells by electroporation. In brief, cells suspended at the concentration of 1 × 107 cells per ml in Nucleofector solution (Amaxa biosystems, Gaithersburg, MD, USA) were mixed with 1 μg of plasmid DNA and then electroporation was performed with Nucleofector (program T-16; Amaxa biosystems). Stable lines that were transfected with various plasmids were selected as a single clone in the presence of 800 μg ml−1 of G418 (Sigma).
Flow cytometry analysis
For the detection of Annexin V, cells were stained with an Annexin V Fluos staining kit (Roche, Mannheim, Germany). To measure the mitochondrial membrane potential, cells were incubated with the J-aggregate-forming cationic dye, JC-1 (Molecular Probes, Karlsruhe, Germany), at a concentration of 10 μg ml−1 for 10 min at 37 °C. ROS in cells were measured by flow cytometry using 2-[6-(4′-amino)phenoxy-3H-xanthen-3-on9-yl]benzoic acid (APF; Daiichi pure chemicals, Tokyo, Japan) fluorescence and 2-[6-(4′-hydroxy) phenoxy-3H-xanthen-3-on9-yl] benzoic acid (HPF; Daiichi pure chemicals) fluorescence. APF reacts with hydroxyl radicals, peroxinytrite and hypochlorous acid. HPF reacts with hydroxyl radicals and peroxinytrite, but not with hypochlorous acid. For the detection of nitric oxide, diaminofluorescein-2 diacetate (Daiichi pure chemicals) was used. All flow cytometric measurements were performed with a FACScan flowcytometer (Becton Dickinson, San Jose, CA, USA). Data were analyzed using CellQuest software (Becton Dickinson).
Cells spread on slide glasses were stained with standard May–Grunwald Giemusa staining and the diaminobenzidine (DAB) method for the detection of MPO activity. For analysis of MPO activity with electron microscopy (JEM-1210 electron microscope; JEOL, Tokyo, Japan), cells fixed with 1.25% glutaraldehyde were incubated with DAB.
Western blot analysis
After disruption in lysis buffer (50 mM Tris-HCl (pH 8.0), 150 mM NaCl, 1% Nonidet P40, 1 mM EDTA, 10 μg ml−1 of aprotinin, 10 μg ml−1 of leupeptin and 1 mM phenylmethylsulfonyl fluoride), samples were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis, then transferred to nitrocellulose membranes (Millipore, Billerica, MA, USA). Target proteins were visualized using a rabbit polyclonal antibody against MPO (DakoCytomation, Carpinteria, CA, USA), a rabbit polyclonal antibody to nitrotyrosine (Chemicon, Temecula, CA, USA), mouse monoclonal antibody to β-actin (Abcam, Cambridge, UK) or to heat-shock protein 90α/β (Santa Cruz Biotechnology, Santa Cruz, CA, USA) with peroxidase-labeled secondary antibodies (Amersham Bioscience, Buckinghamshire, UK) and an enhanced chemiluminescence system (ECL Advance Western Blotting Detection Kit; GE Healthcare Bio-Sciences, Buckinghamshire, UK).
Patients' samples for ROS and nitrotyrosine detection
Peripheral blood or bone marrow samples were obtained from 14 AML patients prior to treatment with informed consent. CD34-positive (+) cells were selected using an immunomagnetic column (Miltenyi Biotech, Auburn, CA, USA). The purity of CD34+ cells was assessed by flow cytometry, demonstrating that more than 95% of cells was CD34-positive after selection. In six cases, CD34-positive AML cells (1 × 105 per well in a 24-well culture plate) with or without 20 nM AraC were cultured in semisolid media (MethoCalt GF H4434; StemCell Technologies, Vancouver, BC, Canada). The number of colonies containing 30 or more cells was counted 7–14 days after plating. In other eight cases, cells were cultured in Iscove's modified Dulbecco's medium with 10% fetal bovine serum incubated with 10 μM AraC or saline as a control for 6 h with or without H2O2, and then analyzed for the detection of ROS. Using four out of eight samples treated with 10 μM AraC up to 6 h, the nitration of tyrosine residues was assessed by western blot analysis with anti-nitrotyrosine antibody as mentioned above. Quantification of bands of western blot was performed using FluoChem IS-8800 and AlphaEase FC Software (Alpha Innotech Corp., san Leandro, CA, USA). The intensity of bands was shown as an average value (AVG). The pixel value and area of each band were counted; then AVG was calculated as follows: AVG=[Σ(each pixel value−background)]/area, which was suggested by the system manual. Expression of MPO in CD34+ cells was examined by flow cytometry.
Results are presented as the mean±s.d. of three independent experiments. Differences between experimental groups were compared using one-way analysis of variance followed by the Scheffe's multiple comparison procedure. Statistical significance was considered at a P-value of 0.05.
Establishment of cell lines expressing wild-type or mutant MPO
K562 cell lines expressing wild-type and mutant (R569W) MPO were established as single clones. R569W mutation of the MPO protein,15 originally found in an MPO-deficient person, resulted in a defective maturation process. MPO protein with the R569W mutation attains apopro-myeloperoxidase status but cannot mature further; it remains in the non-functional stage. Western blot analysis demonstrated the presence of immature MPO protein (apopro-myeloperoxidase, 89 kDa) in both wild-type and mutant MPO-expressing cell lines (MPO-21 and R569W-2, respectively; Figure 1). On the other hand, as expected, the α-subunit of mature MPO protein at 64 kDa and the β-subunit at 14 kDa were detected only in MPO-21 cells since these subunits are generated at the late maturation process of MPO. No apparent difference in the morphological features of MPO-21 and R569W-2 were detected by May-Grunwald Giemusa staining (Figure 2a). Cytochemical analysis using light microscopy detected MPO activity in MPO-21 but not R569W-2 cells (Figure 2a). Other two MPO-expressing lines (MPO-6 and MPO-18) also had the same-size MPO protein as MPO-21 and showed the MPO activity (Supplementary data, Figure 1). Electron microscopy revealed that enzymatically active MPO protein was localized to the cytoplasm (Figure 2b).
MPO activity enhanced the cytotoxic effect of AraC by inducing apoptosis
Proliferation of wild-type K562 (WT), control K562 transfected with an empty vector (MOCK), MPO-21 and R569W-2 cells was similar as assessed by WST-1 assay, keeping maximum absorbance that showed a log phase in growth after day 2 of culture (Figure 3a). However, when cells were treated with AraC at 10 μM, MPO-21 showed an earlier decline than others (Figure 3b). MPO-6 also showed similar pattern as MPO-21 (Supplementary data, Figure 2). Since it is known that AraC induces apoptosis in leukemia cells,16 we next analyzed whether the introduction of MPO in K562 cells accelerated this process or not. As shown in Figure 4a, on day 4 of AraC treatment, a larger proportion of MPO-21 cells (38%) were found to have Annexin V (and propidium iodide-negative) on their surface than wild-type K562 cells (7%). On the other hand, no change was observed between wild-type K562 and R569W-2. Data from three independent experiments showed statistical differences in the expression of Annexin V between MPO-21 and wild-type K562 or R569W-2 (P<0.05, Figure 4b). An earlier marker for apoptosis, the change of mitochondrial membrane potential detected using JC-1, was also significantly increased in MPO-21 cells than other two lines on day 2 (P<0.05, Figure 4c).
MPO enhanced the generation of ROS
Since MPO catalyzes the formation of hypochlorous acid, an ROS, we examined whether the generation of ROS was enhanced by the expression of MPO using fluorescent markers for ROS. After treatment with AraC, the amount of ROS detected by APF but not by HPF was increased in MPO-21 cells when compared to wild-type K562 cells or R569W-2, suggesting the production of hypochlorous acid among ROS (Figures 5a and b). To clarify differences in ROS production, H2O2 was added into the culture medium to enhance MPO-dependent ROS production. ROS production was increased with H2O2 alone (Supplementary data, Figures 3a and b); however, combining H2O2 (40 μM) with AraC significantly enhanced the generation of ROS in MPO-21 cells but not in wild-type K562 cells or R569W-2 (Figures 5c and d). ROS were also generated in other MPO-expressing cell lines, MPO-6 and MPO-18 (Supplementary data, Figures 3c–f). The increase of ROS was completely abolished by 4-aminobenzoic hydrazide (100 μM), an inhibitor of MPO (Figure 5e) or by N-acetylcysteine (1 mM), a thiol antioxidant (Figure 5f). These results suggested that the activity of MPO was directly related to the production of ROS when cells were treated with AraC. We did not observe any change in the fluorescent intensity of diaminofluorescein-2 diacetate, a probe for nitric oxide, even after treatment with H2O2 and AraC (data not shown).
To further analyze the effect of H2O2 on cell growth, we treated wild-type K562 and MPO-21 cells with H2O2 for a short period in the presence or absence of AraC. As shown in Figure 6a, after treatment with 40 μM H2O2 for 45 min, the value of WST-1 assay decreased in both wild-type K562 and MPO-21 cells transiently and recovered on day 2. However, in the presence of AraC, the same treatment with H2O2 suppressed cell growth more significantly in MPO-21 than in K562 cells (Figure 6b). In addition, only wild-type K562 cells recovered from the suppression. In this system, the combination of AraC and H2O2 was not enough to suppress the growth of leukemia cells; MPO was also necessary.
Generation of nitrotyrosine was enhanced by MPO
Since MPO was shown to catalyze the generation of not only ROS but also nitrotyrosine in the presence of nitrogen dioxide, we next examined whether the introduction of MPO in K562 cells also changed the amount of nitrotyrosine. Western blot analysis using an anti-nitrotyrosine antibody detected strong nitration of proteins in the positive control lysate of wild-type K562 cells incubated with RNS (Figure 7). Without AraC or RNS, wild-type K562, R569W-2 and MPO-21 cells showed similar patterns and intensities in the expression of nitrotyrosine, which were all much weaker than those of the positive control. After treatment with AraC, the intensity of bands only increased in MPO-21 cells.
Colony formation of AML cells in semisolid media
In six AML cases, colony formation of CD34+ AML cells was tested in the presence or absence of AraC (Table 1). CD34+ cells were selected from the bone marrow or peripheral blood to avoid the influence of MPO present in mature myeloid cells. The number of colonies generated was increased among three cases with low MPO (3, 6 and 10%) than three with high MPO positivity (90, 96 and 100%). AraC (20 nM in culture) suppressed colony formation in three cases with high MPO compared to low-MPO cases: the number of colonies decreased to 0–10% of control in the presence of AraC, whereas 33–89% of control in low-MPO cases.
Production of ROS and nitrotyrosine in AML cells treated with AraC
We next examined whether the expression of MPO in CD34+ AML cells related to the production of ROS when treated with AraC. As shown in Figures 8a and b, in one out of eight samples tested, ROS production was increased by AraC in the presence of H2O2. The MPO positivity in this case was 98% by flow cytometry. In other seven cases, regardless of the percentage of MPO-positive cells (0, 0.4, 94, 1, 5.5, 96 and 25% among CD34+ cells), no ROS were detected. Figures 8c and d are the representative histograms of negative samples in which ROS were not detected in leukemia cells even after treatment with AraC, H2O2 or the combination of both.
Nitration of tyrosine residues was tested with western blot analysis among four cases of AML: two with high MPO (case 8, 72% of MPO positivity, and case 10, 81%) and two with low MPO (case 7, 0.4%, and case 9, 14%). The intensity of bands in each lane was measured as described in Materials and Methods, then shown as an average intensity (AVG) in Figure 9 (raw data of this procedure is in Supplementary data, Figure 4 and Table 1), which increased along with the incubation time with AraC. The increment of AVG after 6 h of treatment was larger among cases with high MPO (136 and 118% in cases 8 and 10, respectively) than among those with low MPO (110 and 113% in cases 7 and 9, respectively).
In this study, we demonstrated that MPO-expressing K562 leukemia cells showed an increased sensitivity to AraC when compared to wild-type or non-functional MPO-expressing K562 cells. After treatment with AraC, these cells generated a higher amount of ROS and nitrated tyrosine residues, resulting in an earlier induction of apoptosis. These reactions were abrogated by inhibitors of MPO or ROS. The results above strongly suggested the relationship between the expression of MPO and the production of ROS or tyrosine nitration in leukemia cells when treated with AraC. Since ROS and protein nitration were already shown to be toxic for target cells, it is likely that the active MPO protein itself worked with AraC to increase its cytotoxicity. Accordingly, using fresh AML cells, the inhibition of colony formation by AraC tended stronger in cases with high MPO than in those with low MPO expression. It is interesting that the number of colony in high-MPO cases was less than that in low-MPO cases inspite of the fact that the forced expression of MPO in K562 did not influence their proliferation. It seemed that MPO itself does not change growth of cells, but the characteristics of AML cells that express MPO might relate to one of the many factors that control their growth, at least, in some cases. The generation of ROS and the nitration of tyrosine residues, though not so apparent as in colony-formation experiments, were observed only when CD34+ blasts expressed MPO at high levels. It is conceivable that similar reactions were triggered by AraC in high-MPO AML cells as in MPO-expressing K562 cells. MPO did not enhance the fluorescence of diaminofluorescein-2 diacetate, which reacts with NO, in MPO-expressing K562 cells after AraC treatment; however, the nitration of tyrosine residues in these cells was observed by western blot analysis. It seemed that the ROS generated by MPO were involved in the nitration of tyrosine residues as reported previously.17, 18, 19
Clinical observation has repeatedly shown a significant impact of the percentage of MPO-positive blasts on the prognosis of AML patients.3, 4, 5 From data in this study, we postulate that MPO itself could enhance the cytotoxicity of chemotherapeutic agents through the generation of ROS or the nitration of cellular proteins, and that it could contribute, at least in part, to favorable responses to chemotherapy. It is very interesting that AML cases with favorable karyotypes such as t(15;17), t(8;21) and inv(16) usually have a high percentage of MPO-positive blasts.20, 21 Recently, a polymorphism in the promoter region of the MPO gene was shown to relate to survival of breast cancer patients after chemotherapy:22 patients having lower transcriptional activity of MPO (G to A conversion at the −463 nucleotide of the MPO gene) showed significantly worse prognosis. The authors of this report concluded, in concordance with our current observation, that the oxidative stress would modify prognosis after chemotherapy for breast cancer.
Leukemia stem cells that consist of a small fraction of the overall leukemia cell population have been reported to maintain leukemia.23 It is highly possible that the chemosensitivity of leukemia stem cells is an important and vital factor for obtaining a good response to chemotherapy leading to a favorable prognosis. We previously reported that expression of the MPO gene in CD133-positive leukemia cells related to the prognosis of AML.24 As the CD133-positive fraction of AML cells contained leukemia stem cells,25 the results of the present study could be interpreted as events occurring in the growth fraction of AML cells.
Myeloperoxidase cannot be the sole marker of a good response to chemotherapy. For example, defenses against oxidative stress would also affect the response to ROS generated by anticancer drugs. In this regard, the results in Figures 8 and 9, the ROS and nitrotyrosine generation in clinical samples needed to be re-evaluated. It therefore is necessary to fully understand the biology of the immature (stem cell) fraction of leukemia, including the expression of MPO and defense mechanism against ROS and its relationship with other factors such as the karyotype of leukemia cells and other genetic abnormalities.
Bennett JM, Catovsky D, Daniel MT, Flandrin G, Galton DA, Gralnick HR et al. Proposed revised criteria for the classification of acute myeloid leukemia. A report of the French–American–British Cooperative Group. Ann Intern Med 1985; 103: 620–625.
World Health Organization. Classification of Tumors. In: Jaffe ES, Harris NL, Stein H and Vardiman JW (eds). Pathology and Genetics of Tumors of Haematopoietic and Lymphoid Tissues. IARC Press: Lyon, 2001, pp. 79–80.
Hoyle CF, Gray RG, Wheatley K, Swirsky D, de Bastos M, Sherrington P et al. Prognostic importance of Sudan Black positivity: a study of bone marrow slides from 1386 patients with de novo acute myeloid leukaemia. Br J Haematol 1991; 79: 398–407.
Matsuo T, Cox C, Bennett JM . Prognostic significance of myeloperoxidase positivity of blast cells in acute myeloblastic leukemia without maturation (FAB: M1): An ECOG study. Hematol Pathol 1989; 3: 153–158.
Matsuo T, Kuriyama K, Miyazaki Y, Yoshida S, Tomonaga M, Emi N et al. The percentage of myeloperoxidase-positive blast cells is a strong independent prognostic factor in acute myeloid leukemia, even in the patients with normal karyotype. Leukemia 2003; 17: 1538–1543.
Modica-Napolitano JS, Singh KK . Mitochondria as targets for detection and treatment of cancer. Expert Rev Mol Med 2002; 4: 1–19.
Weijl NI, Cleton FJ, Osanto S . Free radicals and antioxidants in chemotherapy-induced toxicity. Cancer Treat Rev 1997; 23: 209–240.
Thannickal VJ, Fanburg BL . Reactive oxygen species in cell signaling. Am J Physiol Lung Cell Mol Physiol 2000; 279: 1005–1028.
Fruehauf JP, Meyskens Jr FL . Reactive oxygen species: a breath of life or death? Clin Cancer Res 2007; 13: 789–794.
Mignotte B, Vayssiere JL . Mitochondria and apoptosis. Eur J Biochem 1998; 252: 1–15.
Winterbourn CC, Vissers MC, Kettle AJ . Myeloperoxidase. Curr Opin Hematol 2000; 7: 53–58.
Myzak MC, Carr AC . Myeloperoxidase-dependent caspase-3 activation and apoptosis in HL-60 cells: protection by the antioxidants ascorbate and (dihydro) lipoic acid. Redox Rep 7: 47–53.
Wagner BA, Buettner GR, Oberley LW, Darby CJ, Burns CP . Myeloperoxidase is involved in H2O2-induced apoptosis of HL-60 human leukemia cells. J Biol Chem 2000; 275: 22461–22469.
Morishita K, Kubota N, Asano S, Kaziro Y, Nagata S . Molecular cloning and characterization of cDNA for human myeloperoxidase. J Biol Chem 1987; 262: 3844–3851.
Nauseef WM, Brigham S, Cogley M . Hereditary myeloperoxidase deficiency due to a missense mutation of arginine 569 to tryptophan. J Biol Chem 1994; 269: 1212–1216.
Bullock O, Ray S, Krajewski S, Ibrado AM, Huang Y, Bhalla K . Intracellular metabolism of Ara-C and resulting DNA fragmentation and apoptosis of human AML HL-60 cells possessing disparate levels of Bcl-2 protein. Leukemia 1996; 10: 1731–1740.
Eiserich JP, Hristova M, Cross CE, Jones AD, Freeman BA, Halliwell B et al. Formation of nitric oxide-derived inflammatory oxidants by myeloperoxidase in neutrophils. Nature 1998; 391: 393–397.
Eiserich JP, Baldus S, Brennan ML, Ma W, Zhang C, Tousson A et al. Myeloperoxidase, a leukocyte-derived vascular NO oxidase. Science 2002; 296: 2391–2394.
Brennan ML, Wu W, Fu X, Shen Z, Song W, Frost H et al. A tale of two controversies defining both the role of peroxidases in nitrotyrosine formation in vivo using eosinophil peroxidase and myeloperoxidase-deficient mice, and the nature of peroxidase-generated reactive nitrogen species. J Biol Chem 2002; 227: 17415–17427.
Kuriyama K, Tomonaga M, Kobayashi T, Takeuchi J, Ohshima T, Furusawa S et al. Morphological diagnoses of the Japan adult leukemia study group acute myeloid leukemia protocols: central review. Int J Hematol 2001; 73: 93–99.
Miyazaki Y, Kuriyama K, Miyawaki S, Ohtake S, Sakamaki H, Matsuo T et al. Cytogenetic heterogeneity of acute myeloid leukaemia (AML) with trilineage dysplasia: Japan Adult Leukaemia Study Group-AML 92 study. Br J Haematol 2003; 120: 56–62.
Ambrosone CB, Ahn J, Singh KK, Rezaishiraz H, Furberg H, Sweeney C et al. Polymorphisms in genes related to oxidative stress (MPO, MnSOD, CAT) and survival after treatment for breast cancer. Cancer Res 2005; 65: 1105–1111.
Bonnet D, Dick JE . Human acute myeloid leukemia is organized as a hierarchy that originates from a primitive hematopoietic cell. Nat Med 1997; 3: 730–737.
Taguchi J, Miyazaki Y, Tsutsumi C, Sawayama Y, Ando K, Tsushima H et al. Expression of the myeloperoxidase gene in AC133 positive leukemia cells relates to the prognosis of acute myeloid leukemia. Leuk Res 2006; 30: 1105–1112.
Vercauteren SM, Sutherland HJ . CD133 (AC133) expression on AML cells and progenitors. Cytotherapy 2001; 3: 449–459.
This work was supported in part by grant from the Ministry of Health, Labour and Welfare of Japan. We deeply appreciate Dr T Matsuo for his thoughtful suggestions.
About this article
Cite this article
Sawayama, Y., Miyazaki, Y., Ando, K. et al. Expression of myeloperoxidase enhances the chemosensitivity of leukemia cells through the generation of reactive oxygen species and the nitration of protein. Leukemia 22, 956–964 (2008) doi:10.1038/leu.2008.8
- reactive oxygen species
- acute myeloid leukemia
BCG-induced formation of neutrophil extracellular traps play an important role in bladder cancer treatment
Clinical Immunology (2019)
Distinct gene alterations with a high percentage of myeloperoxidase-positive leukemic blasts in de novo acute myeloid leukemia
Leukemia Research (2018)
Induction of oxidative stress and sensitization of cancer cells to paclitaxel by gold nanoparticles with different charge densities and hydrophobicities
Journal of Materials Chemistry B (2018)
The critical role of myeloperoxidase inStreptococcus pneumoniaeclearance and tissue damage during mouse acute otitis media
Innate Immunity (2017)
Tetrandrine antagonizes acute megakaryoblastic leukaemia growth by forcing autophagy-mediated differentiation
British Journal of Pharmacology (2017)