- Research Article
- Open Access
Examination of the Cytotoxic and Embryotoxic Potential and Underlying Mechanisms of Next-Generation Synthetic Trioxolane and Tetraoxane Antimalarials
© The Author(s) 2012
- Received: 4 April 2012
- Accepted: 22 May 2012
- Published: 29 May 2012
Semisynthetic artemisinin-based therapies are the first-line treatment for P. falciparum malaria, but next-generation synthetic drug candidates are urgently required to improve availability and respond to the emergence of artemisinin-resistant parasites. Artemisinins are embryotoxic in animal models and induce apoptosis in sensitive mammalian cells. Understanding the cytotoxic propensities of antimalarial drug candidates is crucial to their successful development and utilization. Here, we demonstrate that, similarly to the model artemisinin artesunate (ARS), a synthetic tetraoxane drug candidate (RKA182) and a trioxolane equivalent (FBEG100) induce embryotoxicity and depletion of primitive erythroblasts in a rodent model. We also show that RKA182, FBEG100 and ARS are cytotoxic toward a panel of established and primary human cell lines, with caspase-dependent apoptosis and caspase-independent necrosis underlying the induction of cell death. Although the toxic effects of RKA182 and FBEG100 proceed more rapidly and are relatively less cell-selective than that of ARS, all three compounds are shown to be dependent upon heme, iron and oxidative stress for their ability to induce cell death. However, in contrast to previously studied artemisinins, the toxicity of RKA182 and FBEG100 is shown to be independent of general chemical decomposition. Although tetraoxanes and trioxolanes have shown promise as next-generation antimalarials, the data described here indicate that adverse effects associated with artemisinins, including embryotoxicity, cannot be ruled out with these novel compounds, and a full understanding of their toxicological actions will be central to the continuing design and development of safe and effective drug candidates which could prove important in the fight against malaria.
Semisynthetic artemisinin-based therapies are the recommended first-line treatment for P. falciparum malaria, due to their high efficacy against blood-borne stages of multidrug-resistant forms of the parasite (1). Although relatively well tolerated in patients, artemisinins are reported to induce neurotoxicity (2) and embryotoxicity (3) in a number of animal species, with the latter risk prompting the contraindication of artemisinin-based therapies in the first trimester of pregnancy unless suitable alternatives are unavailable (4). At the cellular level, artemisinin embryotoxicity appears to involve the selective depletion of primitive erythroblasts during defined early periods of gestation (3).
At present there is a lack of consensus on the pharmacological mechanism of action of the artemisinins. It is clear, however, that the endoperoxide bridge within the 1,2,4-trioxane unit is essential for antimalarial activity of these compounds, as exemplified by the lack of antiparasitic activity associated with artemisinin counterparts in which the endoperoxide moiety is replaced with an ether linkage (5). It has been hypothesized that iron-catalyzed reductive cleavage of the endoperoxide bridge results in the generation of toxic carbon-centered radicals, which alkylate and disrupt macromolecules that are vital for parasite homeostasis (6). It also has been proposed that artemisinins are redox-active molecules and that the intrinsic activity of the endoperoxide moiety serves to exacerbate levels of oxidative stress within the parasite by interfering with the function of important redox-sensitive enzymes (7). Inhibition of the parasite sarco/endoplasmic reticulum calcium ATPase (SERCA/PfATP6) has been proposed as an alternative mechanism of antimalarial activity (8), although this has been contested (9).
In addition to its critical role in driving the antiparasitic activity of the artemisinins, the endoperoxide bridge appears to represent a toxicophore in sensitive mammalian cells (10). We have demonstrated that the selective activation of the endoperoxide bridge is the chemical basis for the differential cytotoxicity of the synthetic analogue 10β-(p-bromophenoxy)dihydroartemisinin toward sensitive HL-60 promyelocytic leukemia cells and insensitive peripheral blood mononuclear cells (PBMCs) (11). Bioactivation of the endoperoxide bridge is sensitive to pharmacological manipulation of cellular heme levels, and triggers the generation of reactive oxygen species (ROS), a process that is dependent upon the integrity of the mitochondrial electron transport chain (12). These biochemical events induce the onset of apoptotic cell death, characterized by mitochondrial membrane depolarization, activation of caspases 3 and 7, and DNA fragmentation (11,13).
Freshly drawn venous blood was collected from healthy volunteers in heparinized tubes. Peripheral blood mononuclear cells (PBMCs) and red blood cells (RBCs) were isolated using Lymphoprep (Axis-Shield, Kimbolton, UK), as described previously (20). ARS was kindly donated by Dafra Pharma International (Turnhout, Belgium). The anti-caspase 3 antibody (#9662) was obtained from Cell Signaling Technology (Danvers, MA, USA). Unless noted, all other reagents were from Sigma-Aldrich (Poole, UK).
Chemical Synthesis of RKA182 and FBEG100
The protocols described here were undertaken in accordance with criteria outlined in a license granted under the Animals (Scientific Procedures) Act 1986 and approved by the Institution local animal ethics committee. Eleven-wk-old Crl:CD (Sprague Dawley) male and primiparous female rats were supplied by Charles River (Kent, UK). Rats were maintained under standard conditions (room temperature 21.5 ± 1.5°C, humidity 55 ± 5%, artificial lighting from 6 AM to 6 PM, feed and tap water provided ad libitum). Paired female and male rats were left in cohabitation from 4 PM to 9 AM the next morning. Mating is assumed to have occurred at the midpoint of the dark cycle, hence noon of the next day is defined as gestation d 0.5. On gestation d 9.5, pregnant female rats were euthanized via increasing concentrations of CO2 and embryos were explanted. These were distributed randomly to experimental groups and exposed to the indicated concentrations of ARS, RKA182 or FBEG100 for 48 h. The vehicle was methanol, and the concentration of the solvent in the media was controlled to 0.05%, which is known to be nonembryotoxic in this model (22). Embryos were cultured essentially as described by New (23), in sterile 50 mL glass bottles containing four embryos per bottle in 100% serum (5 mL) collected from male Sprague Dawley rats. At the start of culture, the bottles were flushed for 1 min with a gas mixture of 5/5/90% O2/CO2/N2 and placed on a nonrocking roller mixer (60 rpm) at 37°C. The next morning the bottles were flushed again with a gas mixture of 20/5/75% O2/CO2/N2, and in the afternoon with 40/5/55% O2/CO2/N2. The total culture time was 48 h. At the end of the culture period the embryos were transferred into Tyrode salt solution and examined under a stereomicroscope. Yolk sac diameter, crown-rump length and head length were measured and total morphological scores were assigned using the method of Brown and Fabro (24).
Cull Culture and Treatments
HL-60 cells, RBCs and PBMCs were cultured in RPMI-1640 media supplemented with 25 mmol/L 4-(2-hydroxyethyl)-1-piperazine ethanesulfonic acid (HEPES), 300 mg/L L-glutamine, 10% fetal bovine serum (Biowest, Nuaillé, France), 100 U/mL penicillin and 100 µg/mL streptomycin. HEK293T, HeLa and HepG2 cells were cultured in Dulbecco’s modified Eagle medium (DMEM) supplemented with 584 mg/L l-glutamine, 10 % fetal bovine serum (FBS), 100 U/mL penicillin and 100 µg/mL streptomycin. All cells were cultured at 37°C in a 5% CO2 atmosphere. For drug treatments, ARS, RKA182 and FBEG100 were dissolved in methanol, with the concentration of the solvent in the media controlled to 0.5%. For biochemical manipulation experiments, cells were pretreated for 1 h with succinyl acetone (SA; 1 mmol/L), δ-aminolevulinic acid (ALA; 1 mmol/L), protoporphyrin IX (PPIX; 1 µmol/L), desferroxamine (DFO; 10 µmol/L), holotransferrin (HTF; 10 µmol/L), tiron (1 mmol/L), N-acetylcysteine (NAC; 10 mmol/L) or Z.VAD.FMK (100 µmol/L), or for 16 h with buthionine sulphoximine (BSO; 30 µmol/L), before exposure to the indicated concentrations of RKA182, FBEG100 or ARS for a further 24 h.
Quantification of Cell Viability
Cell viability was measured using a CellTiter-Glo Luminescent Assay (Promega, Southampton, UK) for adenosine triphosphate (ATP) or a Cytotoxicity Detection Kit (Roche Applied Science, Burgess Hill, UK) for lactate dehydrogenase (LDH), both in accordance with the manufacturer’s instructions. The concentrations of each compound that induced a 50% loss of cellular viability (IC50) were calculated using GraFit (Erithacus Software, Horley, UK).
Live-Cell Imaging of Apoptosis/Necrosis
The ability of RKA182, FBEG100 and ARS to induce apoptosis/necrosis in adherent HepG2 cells was assessed using a live-cell fluorescent imaging assay, as previously described (25). In brief, the binding of annexin V Alexa Fluor 488 (Life Technologies Corporation, Grand Island, NY, USA) conjugate to phosphatidyl serine on the membranes of apoptotic cells was followed in real-time by imaging every 30 min after drug exposure with a BD Pathway 855 imager (Becton Dickinson, Erembodegem-Aalst, Belgium). At the same time points, the intercalation of propidium iodide (PI) with cellular DNA was quantified. The total fluorescent intensity per cell area was quantified using Image Pro (Media Cybernetics, Bethesda, MD, USA).
Liquid Chromatography Mass Spectrometry
HL-60 cells were exposed to RKA182 or FBEG100 (both 100 µmol/L) for 0–24 h. Cells and media (100 µL) were combined with ARS (10 nmol) and ice-cold acetonitrile (300 µL), vortexed and centrifuged at 18,000g for 10 min. Extracted material was filtered using MultiScreen Solvinert plates (Millipore, Watford, UK), in accordance with the manufacturer’s instructions, and analyzed by multiple reaction monitoring using an API 4000 QTRAP LC-MS/MS System (AB Sciex, Warrington, UK) interfaced to a Ultimate 3000 autosampler and pump (Dionex, Camberly, UK). The data were collected and analyzed using Analyst software version 1.5 (AB Sciex). Sample separation was achieved on an ACE C8 column (100 × 2.1 mm, 3 µm; Advanced Chromatography Technologies, Aberdeen, UK). The mobile phase consisted of acetonitrile with 10 mmol/L ammonium acetate (90:10, v/v), both supplemented with 0.1% formic acid, delivered at a flow rate of 0.2 mL/min. The mass spectrometer was operated in positive ion mode. General operating parameters and analyte-specific fragmentation transitions are detailed in Tables S1 and S2. Calibration curves for RKA182 and FBEG100 were generated by plotting the ratio of peak area for analyte versus internal standard (ARS), following extraction from media containing RKA182 or FBEG100 (1–300 µmol/L) as described above.
Data are expressed as mean ± standard deviation of the mean from at least three independent experiments. The significance of differences within the data was assessed by one-way analysis of variance (ANOVA; with the Tukey post hoc test) or unpaired t test. A P value of ≤0.05 was considered to be statistically significant.
The synthesis of deoxy-artesunate (deoxy-ARS), and procedures for determination of in vitro antiparasitic activity, flow cytometry, Western blot, DNA fragmentation and glutathione content are described in Supplementary Material.
All supplementary materials are available online at https://doi.org/www.molmed.org.
Embryotoxicity of RKA182, FBEG100 and ARS
Cytotoxicity of RKA182, FBEG100 and ARS in a Panel of Human Cells
Cytotoxicity of RKA182, FBEG100, ARS and deoxy-ARS toward a panel of human cells and antimalarial activities versus P. falciparum.
11.6 ± 4.0a
40.6 ± 3.0
2.2 ± 0.2
14.7 ± 0.2
37.1 ± 2.5
20.1 ± 3.4
25.5 ± 5.3
58.8 ± 10.8
77.2 ± 17.2
37.7 ± 5.5
55.2 ± 5.6
87.8 ± 11.4
28.0 ± 7.3
18.9 ± 3.7
46.6 ± 0.7
56.8 ± 2.1
P. falciparum (3D7)
4.9 ± 1.2 nmol/Lb,c
1.4 ± 0.3 nmol/L
1.8 ± 0.6 nmol/Lc
Pathways of Cell Death Induced by RKA182, FBEG100 and ARS
Role of Heme and Iron in the Mechanism of RKA182 and ARS Cytotoxicity
Role of Oxidative Stress in the Mechanism of RKA182 and ARS Cytotoxicity
Previously, we have shown that artemisinins induces concentration and time-dependent elevations of ROS in mammalian cells, and that the superoxide scavenger tiron protects against the toxicity of these compounds (12). In keeping with these observations, here both ARS and RKA182 toxicity was diminished by tiron and the thiol antioxidant NAC in HL-60 cells (Figure 7C, Supplementary Figure S5). To further probe the involvement of oxidative stress in the toxicity induced by ARS and RKA182, we pretreated HL-60 cells with BSO, an inhibitor of glutathione biosynthesis (30). BSO provoked glutathione depletion (Figure 7D) and augmented the cytotoxic action of ARS and RKA182, an effect that was reversed by coincubation with NAC (Figure 7E). Taken together, these data indicate that oxidative stress is an important mechanistic component of ARS and RKA182-induced cell death (Figure 7F).
Role of Chemical Decomposition in RKA182 and FBEG100 Cytotoxicity
Synthetic trioxolanes and tetraoxanes have shown promise as next-generation antimalarial drug candidates, particularly in terms of therapeutic efficacy and lack of synthetic constraints. Indeed, we recently have identified the tetraoxane drug candidate RKA182 that possesses improved pharmacokinetic properties and outstanding antimalarial activity (16). In addition, a number of trioxolane and tetraoxane compounds currently are being supported by the Medicines for Malaria Venture (MMV) and/or are in clinical trials (15,31), yet the continuing development and optimal therapeutic utilization of these compounds must be supported by an understanding of their potential to induce cytotoxicity in mammalian cells, and an appreciation of the underpinning chemical and molecular mechanisms. Here, we have revealed that, similarly to the established artemisinin ARS, both RKA182 and its trioxolane counterpart FBEG100 can induce embryotoxicity in the rodent WEC model, as well as cytotoxicity across a panel of primary and established human cells. These findings may have important implications for the continuing development and ultimate therapeutic utilization of these promising antimalarial drug candidates.
P. falciparum infection during pregnancy poses a significant health risk to both the mother and unborn child (32), yet artemisinin-based therapies are currently contraindicated in the first trimester of pregnancy due to an increasing body of evidence indicating that artemisinins are embryotoxic in laboratory animals (3). Here, we have demonstrated that RKA182 and FBEG100 are embryotoxic in the rodent WEC model. Both compounds, together with ARS, induced embryotoxicity at >1 µmol/L, in keeping with previous reports of embryotoxicity induced by artemisinin (lower limit for toxicity 0.9 µmol/L) (33), dihydroartemisinin (1.8 µmol/L) (34) and the trioxolane OZ277 (1.3 µmol/L) (33). It is thought that the selective depletion of primitive erythroblasts, and the anemia that ensues, is the underlying cause of artemisinin embryotoxicity in rodents (3). Indeed, the sensitive period for artemisinin embryotoxicity in the rat (gestation d 10–14) correlates with the timecourse of dependence upon primitive, rather than definitive, erythroblasts for embryonic oxygen requirements (3). In keeping with this notion, we observed that overt embryotoxicity induced by ARS, RKA182 and FBEG100 was preceded by a marked paleness within the visceral yolk sac circulation, indicating that the targeting of primitive erythroblasts is a common basis for the embryotoxicity of artemisinins, tetraoxanes and trioxolanes. The ultimate risk of embryotoxicity in humans can only be established through improved translation of animal data together with robust monitoring of inadvertent exposures in pregnant patients.
Although the mechanism by which artemisinins deplete embryonic erythroblasts has yet to be elucidated, it is well documented that these compounds induce hallmarks of apoptotic cell death in sensitive mammalian cells in vitro. Here, we have demonstrated that RKA182, FBEG100 and ARS induce concentration, time and caspase-dependent apoptosis in sensitive cells, while also invoking a form of caspase-independent cell death that bears the hallmarks of necrosis. As such, the ultimate mechanism of cytotoxicity appears to be similar for semi-synthetic artemisinins and synthetic tetraoxanes and trioxolanes (Figure 7F). Recently, we and others have demonstrated the role of heme, iron and ROS in the pharmacological and toxicological actions of the artemisinins (6,12,28,29). Here, oxidative stress was shown to underpin the cytotoxic action of both RKA182 and ARS, indicating a convergence of mechanism between novel tetraoxanes and established artemisinins. Although not tested here, the generation of oxidative stress could perturb critical, but as yet poorly defined, cellular processes, or provoke the onset of lipid peroxidation, contributing to the induction of cell death (7,36–38). We also have demonstrated that the heme synthesis inhibitor SA and the iron chelator DFO diminish the toxic effects of ARS and RKA182, although the latter was relatively less sensitive to these interventions. In contrast, RKA182 toxicity was relatively more sensitive to elevation of cellular heme and iron to nonphysiological levels.
The above data indicate that tetraoxanes are less reliant, relative to artemisinins, on interaction with basal levels of heme and iron for the induction of cytotoxicity. However, under conditions of elevated heme and/or iron, which are notably associated with malaria infection (39), tetraoxane toxicity appears to be augmented to a greater extent than that of the artemisinins. That such subtle differences should exist in the biochemical mechanisms that underlie the toxic effects of artemisinins compared with tetraoxanes and trioxolanes is consistent with our inability to detect appreciable chemical decomposition of RKA182 or FBEG100 under conditions of substantial cytotoxicity. However, it has been shown that the peroxidic moiety is essential for the pharmacological action of tetraoxanes and trioxolanes (17–19). Therefore, it is possible that a relatively minor and/or specific bioactivation, prompted by interaction with an as yet undefined cellular target(s), may contribute to the chemical mechanism of toxicity of these novel compounds. Further work is therefore required to identify suitable chemical markers of trioxolane and tetraoxane bioactivation, to fully elucidate the latter’s role in cytotoxicity.
It is important to note that, at least based on the ex vivo and in vitro data presented here, the benefit:risk balance of these promising antimalarial drug candidates appears to be considerable, in terms of the differences between pharmacologically active and toxic concentrations of the compounds. Specifically, in terms of the ratio between their IC50 values for embryotoxicity (in the rodent WEC model) and antimalarial activity (in the P. falciparum 3D7 screen), RKA182 (904.1), FBEG100 (2619.3) and ARS (1630.0) each appear to have excellent therapeutic indices. However, in the absence of robust in vivo pharmacokinetic data for these and other novel synthetic endoperoxide-based antimalarials, adverse effects associated with artemisinins cannot yet be ruled out with next-generation tetraoxane and trioxolane drug candidates. Such knowledge is particularly important in light of the present focus on the development of synthetic drug candidates with enhanced stability and bioavailability, which will increase systemic exposure and the likelihood of a single-dose cure for malaria, but also the potential for toxicity. On the other hand, in light of proposals that the sensitivity of cancerous cells to artemisinin toxicity in vitro be exploited therapeutically (35), our finding that model tetraoxanes and trioxolanes are cytotoxic toward mammalian cells suggests that these compounds also may represent promising candidates for anticancer therapy.
In summary, we have revealed that model tetraoxane and trioxolane antimalarials are embryotoxic in the rodent WEC model and induce rapid cytotoxicity in mammalian cells. The subtle differences in mechanisms and timecourses of toxicity for these compounds and established artemisinins reported here serve to further emphasize the need to better understand pharmacological and toxicological mechanisms of action of next-generation antimalarial drug candidates to enable a more informed assessment of the ultimate risk of adverse effects in patients. In the continuing search for more potent endoperoxide-based antimalarials, it is vital that we explore and exploit differences in mechanisms of action within the parasite and mammalian cells to ensure the successful development of safe and efficacious new drugs.
The authors declare that they have no competing interests as defined by Molecular Medicine, or other interests that might be perceived to influence the results and discussion reported in this paper.
The authors thank Richard Amewu and Fatima Bousejra-El Garah (Department of Chemistry, The University of Liverpool, Liverpool, United Kingdom) for the synthesis and provision of RKA182 and FBEG100; Nuna Araújo (Departamento de Química e Farmácia, Universidade do Algarve, Portugal) for helpful suggestions on the synthesis of deoxy-ARS; Claire Taylor and Victoria Daniels (Liverpool School of Tropical Medicine, Liverpool, United Kingdom) for technical assistance with the WEC experiments; Anahi Santoyo Castelazo and James Maggs (Centre for Drug Safety Science, The University of Liverpool. Liverpool, United Kingdom) for assistance with mass spectrometry.
This work was supported by the European Union Seventh Framework Programme (ARTEMIP, 200805) and the UK Medical Research Council (MRC), as part of the Centre for Drug Safety Science (G0700654). Work at Leiden University was facilitated by the award of a UK Royal Society International Travel Grant (TG102742) and the British Toxicology Society’s Norman Aldridge Travelling Fellowship to IM Copple. J Firman’s PhD is supported by the MRC Integrative Toxicology Training Partnership.
Open Access This article is licensed under a Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International License, which permits any non-commercial use, sharing, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, and provide a link to the Creative Commons license. You do not have permission under this license to share adapted material derived from this article or parts of it.
The images or other third party material in this article are included in the article’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.
To view a copy of this license, visit (https://doi.org/creativecommons.org/licenses/by-nc-nd/4.0/)
- WHO. (2010) Guidelines for the treatment of malaria [Internet]. 2nd ed. Geneva: WHO. [cited 2011 Jun 1]. Available from: https://doi.org/www.who.int/malaria/publications/atoz/9789241547925/en/index.html.Google Scholar
- Toovey S. (2006) Safety of artemisinin antimalarials. Clin. Infect. Dis 42:1214–5.View ArticleGoogle Scholar
- Clark RL. (2009) Embryotoxicity of the artemisinin antimalarials and potential consequences for use in women in the first trimester. Reprod. Toxicol 28:285–96.View ArticleGoogle Scholar
- WHO. (2007) Assessment of the safety of artemisinin compounds in pregnancy: report of two joint informal consultations convened in 2006 by: the Special Programme for Research and Training in Tropical Diseases 1055 (TDR) sponsored by UNICEF/UNDP/World Bank/WHO and the Global Malaria Programme of the World Health Organization [Internet]. Geneva: WHO. [cited 2011 Jun 1]. https://doi.org/www.who.int/malaria/publications/atoz/9789241596114/en/index.html.Google Scholar
- Beekman AC, et al. (1997) Stereochemistry-dependent cytotoxicity of some artemisinin derivatives. J. Nat. Prod. 60:325–30.View ArticleGoogle Scholar
- O’Neill PM, Barton VE, Ward SA. (2010) The molecular mechanism of action of artemisinin—the debate continues. Molecules. 15:1705–21.View ArticleGoogle Scholar
- Haynes RK, et al. (2010) Facile oxidation of leucomethylene blue and dihydroflavins by artemisinins: relationship with flavoenzyme function and antimalarial mechanism of action. Chem Med Chem. 5:1282–99.View ArticleGoogle Scholar
- Krishna S, Pulcini S, Fatih F, Staines H. (2010) Artemisinins and the biological basis for the PfATP6/SERCA hypothesis. Trends Parasitol. 26:517–23.View ArticleGoogle Scholar
- Cardi D, et al. (2010) Purified E255L mutant SERCA1a and purified PfATP6 are sensitive to SERCA-type inhibitors but insensitive to artemisinins. J. Biol. Chem. 285:26406–16.View ArticleGoogle Scholar
- Fishwick J, McLean WG, Edwards G, Ward SA. (1995) The toxicity of artemisinin and related compounds on neuronal and glial cells in culture. Chem. Biol. Interact. 96:263–71.View ArticleGoogle Scholar
- Mercer AE, et al. (2007) Evidence for the involvement of carbon-centered radicals in the induction of apoptotic cell death by artemisinin compounds. J. Biol. Chem. 282:9372–82.View ArticleGoogle Scholar
- Mercer AE, Copple IM, Maggs JL, O’Neill PM, Park BK. (2011) The role of heme and the mitochondrion in the chemical and molecular mechanisms of mammalian cell death induced by the artemisinin antimalarials. J. Biol. Chem. 286:987–96.View ArticleGoogle Scholar
- Disbrow GL, et al. (2005) Dihydroartemisinin is cytotoxic to papillomavirus-expressing epithelial cells in vitro and in vivo. Cancer Res. 65:10854–61.View ArticleGoogle Scholar
- Jefford CW. (2007) New developments in synthetic peroxidic drugs as artemisinin mimics. Drug Discov. Today. 12:487–95.View ArticleGoogle Scholar
- Charman SA, et al. (2011) Synthetic ozonide drug candidate OZ439 offers new hope for a singledose cure of uncomplicated malaria. Proc. Natl. Acad. Sci. U. S. A. 108:4400–5.View ArticleGoogle Scholar
- O’Neill PM, et al. (2010) Identification of a 1,2,4,5-tetraoxane antimalarial drug-development candidate (RKA 182) with superior properties to the semisynthetic artemisinins. Angew. Chem. Int. Ed. Engl. 49:5693–7.View ArticleGoogle Scholar
- Fugi MA, Wittlin S, Dong Y, Vennerstrom JL. (2010) Probing the antimalarial mechanism of artemisinin and OZ277 (arterolane) with nonperoxidic isosteres and nitroxyl radicals. Antimicrob. Agents Chemother. 54:1042–6.View ArticleGoogle Scholar
- Dong Y, et al. (2005) Spiro and dispiro-1,2,4-trioxolanes as antimalarial peroxides: charting a workable structure-activity relationship using simple prototypes. J. Med. Chem. 48:4953–61.View ArticleGoogle Scholar
- Kaiser M, et al. (2007) Peroxide bond-dependent antiplasmodial specificity of artemisinin and OZ277 (RB×11160). Antimicrob. Agents Chemother. 51:2991–3.View ArticleGoogle Scholar
- Williams DP, Pirmohamed M, Naisbitt DJ, Maggs JL, Park BK. (1997) Neutrophil cytotoxicity of the chemically reactive metabolite(s) of clozapine: possible role in agranulocytosis. J. Pharmacol. Exp. Ther. 283:1375–82.PubMedGoogle Scholar
- Bousejra-El Garah F, et al. (2011) Comparison of the reactivity of antimalarial 1,2,4,5-tetraoxanes with 1,2,4-trioxolanes in the presence of ferrous iron salts, heme, and ferrous iron salts/phosphatidylcholine. J. Med. Chem. 54:6443–55.View ArticleGoogle Scholar
- Brown-Woodman PD, et al. (1995) In vitro assessment of the effect of methanol and the metabolite, formic acid, on embryonic development of the rat. Teratology. 52:233–43.View ArticleGoogle Scholar
- New DA. (1978) Whole-embryo culture and the study of mammalian embryos during organogenesis. Biol. Rev. Camb. Philos. Soc. 53:81–122.View ArticleGoogle Scholar
- Brown NA, Fabro S. (1981) Quantitation of rat embryonic development in vitro: a morphological scoring system. Teratology. 24:65–78.View ArticleGoogle Scholar
- Puigvert JC, de Bont H, van de Water B, Danen EH. (2010) High-throughput live cell imaging of apoptosis. Curr. Protoc. Cell Biol. Unit 18.10:1–13.Google Scholar
- Hou J, Wang D, Zhang R, Wang H. (2008) Experimental therapy of hepatoma with artemisinin and its derivatives: in vitro and in vivo activity, chemosensitization, and mechanisms of action. Clin. Cancer Res. 14:5519–30.View ArticleGoogle Scholar
- Gao X, et al. (2011) Dihydroartemisinin induces endoplasmic reticulum stress-mediated apoptosis in HepG2 human hepatoma cells. Tumori. 97:771–80.View ArticleGoogle Scholar
- Haynes RK, et al. (2007) The Fe2+−mediated decomposition, PfATP6 binding, and antimalarial activities of artemisone and other artemisinins: the unlikelihood of C-centered radicals as bioactive intermediates. Chem Med Chem. 2:1480–97.View ArticleGoogle Scholar
- Stocks PA, et al. (2007) Evidence for a common non-heme chelatable-iron-dependent activation mechanism for semisynthetic and synthetic endoperoxide antimalarial drugs. Angew. Chem. Int. Ed. Engl. 46:6278–83.View ArticleGoogle Scholar
- Griffith OW. (1982) Mechanism of action, metabolism, and toxicity of buthionine sulfoximine and its higher homologs, potent inhibitors of glutathione synthesis. J. Biol. Chem. 257:13704–12.PubMedGoogle Scholar
- Olliaro P, Wells TN. (2009) The global portfolio of new antimalarial medicines under development. Clin. Pharmacol. Ther. 85:584–95.View ArticleGoogle Scholar
- Desai M, et al. (2007) Epidemiology and burden of malaria in pregnancy. Lancet Infect. Dis. 7:93–104.View ArticleGoogle Scholar
- Longo M, et al. (2010) Comparative embryotoxicity of different antimalarial peroxides: in vitro study using the rat whole embryo culture model (WEC). Reprod. Toxicol. 30:583–590.View ArticleGoogle Scholar
- Longo M, et al. (2006) Effects of the antimalarial drug dihydroartemisinin (DHA) on rat embryos in vitro. Reprod. Toxicol. 21:83–93.View ArticleGoogle Scholar
- Firestone GL, Sundar SN. (2009) Anticancer activities of artemisinin and its bioactive derivatives. Expert Rev. Mol. Med. 11:e32.View ArticleGoogle Scholar
- D’Alessandro S, et al. (2011) Hypoxia modulates the effect of dihydroartemisinin on endothelial cells. Biochem. Pharmacol. 82:476–84.View ArticleGoogle Scholar
- Efferth T, Giaisi M, Merling A, Krammer PH, Li-Weber M. (2007) Artesunate induces ROSmediated apoptosis in doxorubicin-resistant T leukemia cells. PLoS One. 2:e693.View ArticleGoogle Scholar
- Haynes RK, et al. (2011) Reactions of antimalarial peroxides with each of leucomethylene blue and dihydroflavins: flavin reductase and the cofactor model exemplified. Chem Med Chem. 6:279–91.View ArticleGoogle Scholar
- Hunt NH, Stocker R. (2007) Heme moves to center stage in cerebral malaria. Nat. Med. 13:667–9.View ArticleGoogle Scholar